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HOCl-mediated cell death and metabolic dysfunction in the yeast Saccharomyces cerevisiae
ABB Archives of Biochemistry and Biophysics 423 (2004) 170–181 www.elsevier.com/locate/yabbi
HOCl-mediated cell death and metabolic dysfunction in the yeast Saccharomyces cerevisiae David A. King,a Diane M. Hannum,b Jian-Shen Qi,b and James K. Hursta,b,* b
a Department of Chemistry, Washington State University, Pullman, WA, USA OGI School of Science and Engineering, Oregon Health and Science University, Beaverton, OR, USA
Received 30 October 2003, and in revised form 9 December 2003
Abstract The nature of oxidative damage to Saccharomyces cerevisiae caused by levels of HOCl that inhibit cell replication was explored with the intent of identifying the loci of lethal lesions. Functions of cytosolic enzymes and organelles that are highly sensitive to inactivation by HOCl, including aldolase, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), and the mitochondrion, were only marginally affected by exposure of the yeast to levels of HOCl that completely inhibited colony formation. Loss of function in membrane-localized proteins, including the hexose transporters and PMA1 Hþ -ATPase, which is the primary proton pump located within the S. cerevisiae plasma membrane, was also marginal and Kþ leak rates to the extracellular medium increased only slowly with exposure to increasing amounts of HOCl, indicating that the plasma membrane retained its intrinsic impermeability to ions and metabolites. Adenylate phosphorylation levels in fermenting yeast declined in parallel with viability; however, yeast grown on respiratory substrates maintained near-normal phosphorylation levels at HOCl doses several-fold greater than that required for killing. This overall pattern of cellular response to HOCl differs markedly from that previously reported for bacteria, which appear to be killed by inhibition of plasma membrane proteins involved in energy transduction. The absence of significant loss of function in critical oxidant-sensitive cellular components and retention of ATP-synthesizing capabilities in respiring yeast cells exposed to lethal levels of HOCl suggests that toxicity in this case may arise by programmed cell death. Ó 2003 Elsevier Inc. All rights reserved. Keywords: ATP synthesis; Fungicidal mechanisms; Glycolysis; Hypochlorous acid; Oxidative phosphorylation; Proton pumps
Myeloperoxidase (MPO)1-mediated reactions play a major role in the microbicidal action of respiring neutrophils [1,2]. Although MPO catalyzes a wide range of peroxidative reactions [3–5], including two-electron oxidations of most halides and the pseudohalide, SCN , to the corresponding hypohalites, as well as one-electron
*
Corresponding author. Fax: 1-509-335-8867. E-mail address: [email protected] (J.K. Hurst). 1 Abbreviations used: ALD, alcohol dehydrogenase; cfu, colonyforming units; DES, diethylstilbestrol; DTPA, diethylenetriaminepentaacetic acid; DTT, dithiothreitol; EC, adenylate energy charge; FDP, fructose-1,6-diphosphate; GAP, D -glyceraldehyde-3-phosphate; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; LD99 , concentration required to inhibit colony formation in 99% of the cells; LDH, lactate dehydrogenase; MPO, myeloperoxidase; PBS, phosphate-buffered saline; PMSF, phenylmethylsulfonyl fluoride; RCR, respiratory control ratio; TSA, trypticase soy agar; YPD, yeast-peptone dextrose agar. 0003-9861/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2003.12.012
oxidations of NO 2 and a variety of organic reductants, chloride ion is generally regarded as the physiological substrate. This viewpoint is supported by recent studies using bacteria [6,7] and fluorescent particulate probes [8] to trap hypochlorous acid (HOCl) generated within neutrophil phagosomes. The patterns of metabolic dysfunction observed in bacteria exposed to HOCl suggest a bactericidal mechanism wherein death occurs by interruption of the energy-transducing capabilities of the cell by inhibiting plasma membrane-localized proteins, specifically the Fo F1 -ATP synthase, proteins involved with active transport of metabolites, and the respiratory redox chain [9]. Loss of these functions renders bacteria incapable of either oxidative or substrate-level phosphorylation, and nucleotide phosphorylation levels correspondingly plummet to levels far below those necessary to maintain homeostasis, much less undertake biosynthetic repair [10,11]. In contrast, cytosolic
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enzymes that are highly susceptible to irreversible inhibition by HOCl, such as aldolase and b-galactosidase, maintained nearly full activity within bacteria that had been exposed to amounts of HOCl that were severalfold higher than is required for killing [12,13]. Thus, the observed cytosolic protection of these HOCl-responsive markers suggests that cellular inactivation by HOCl is not accompanied by extensive oxidation or chlorination of intracellular components. Furthermore, quench-flow experiments have established that these Escherichia coli are killed within 100 ms of exposure to lethal levels of HOCl [14], supporting the notion that the locus of lethal lesions is the bacterial envelope and killing involves oxidation of highly reactive nucleophilic centers located therein. Because energy transduction in all prokaryotes occurs across the ion-impermeable plasma membrane, this model is attractive in accounting for the ‘‘universal’’ bactericidal properties of HOCl [1]. However, the model is clearly not applicable to simple eukaryotes where respiration-linked ATP synthesis is mitochondrial, and therefore occurs within the relatively protected cytosol of the cell. Additionally, unlike bacteria, whose glucose transport systems are active [15], i.e., require a polarized membrane, and are inhibited by lethal levels of HOCl [13], glucose transport across plasma membranes of yeast such as Saccharomyces cerevisiae is passive [16,17], and may therefore not be particularly susceptible to inhibition by HOCl. In this case, one would expect substrate-level phosphorylation to also be impervious to HOCl in this organism. In the present study, we have explored the nature of oxidative damage caused when S. cerevisiae are exposed to lethal levels of HOCl. Our working hypothesis, based upon analogy to the bactericidal effects of HOCl, was that death would coincide with loss of function within the yeast plasma membrane and involve energy-linked phenomena such as metabolite and ion transport. Correspondingly, one focal point has been the PMA1 Hþ ATPase, a proton pump that constitutes the primary transport system of the plasma membrane in S. cerevisiae, to which all plasma-membrane secondary transport systems are chemiosmotically linked [18]. Loss of PMA1 activity is lethal [19] and would constitute the functional equivalent of loss of activity in the plasma membranelocalized Fo F1 -ATP synthase in bacteria; the latter has been shown in E. coli 25922, Pseudomonas aeruginosa 27853, and Streptococcus lactis 7962 to decline in parallel with cell viability [20]. To our surprise, the pattern of metabolic damage observed in S.cerevisiae does not support this model; although specific lethal lesions have not been identified, it is evident from the data that the microbicidal mechanisms of HOCl in this simple singlecelled eukaryote bear little relationship to those in prokaryotes.
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Materials and methods Isolation and maintenance of cell cultures Saccharomyces cerevisiae was obtained as FleishmannÕs Active Dry Yeast (Regular), which was cultured by streaking suspensions onto agar slants or plates. Cells were generally grown in a 1-L volume containing 3 g NH4 SO4 , 2 g KH2 PO4 , 0.2 g K2 HPO4 0.5 g MgSO4 7H2 O, 0.5 g NaCl, 200 lg FeCl3 6 H2 O, 0.1 g CaCl2 2H2 O, and 3 g yeast extract supplemented with an appropriate carbon source. For fermentative growth, a large amount (50 g/L) of glucose, autoclaved separately, was added to the medium; for aerobic (respiratory) growth, filter-sterilized ethanol was added to a concentration of 10 g/L (12.7 ml/L). A medium of slightly different composition was used for aerobic growth on lactate. This contained 3 g yeast extract, 1 g glucose, 1 g KH2 PO4 , 1 g NH4 Cl, 0.5 g CaCl2 2H2 O, 1 g MgSO4 7H2 O, 0.5 g NaCl, 0.3 ml of a 1% solution of FeCl3 , and 22 ml of 90% lactic acid in 1 L solution; the final pH was adjusted to 5.5 with solid KOH. Cells were first grown by inoculating a small amount of the growth medium with cells from the slants and incubating for 8 h in a shaking water bath at 30 °C, following which a portion of this suspension was used to inoculate one liter of the same medium in a 2 L Erlenmeyer flask. The cells were harvested by centrifugation, washed twice by resuspension in cold phosphate-buffered saline (0.1 M sodium phosphate, 0.15 M NaCl, pH 7.4 (PBS)), and isolated by centrifugation. They were then resuspended in cold buffer to the desired cell density and kept on ice until used. Reactions with hypochlorous acid (HOCl) and chloramines (NH2 Cl) HOCl was purified from commercial sources of NaOCl by vacuum distillation after adjusting the pH to 7–8 with phosphoric acid; NH2 Cl was prepared by reacting HOCl with a 20-fold excess of NH3 [13]. Concentrations were determined by spectrophotometric analysis using e236 (HOCl) ¼ 100 M1 cm1 [21] and e242 (NH2 Cl) ¼ 429 M1 cm1 [22]. In initial experiments, cell suspensions were warmed to 2–30 °C for 5– 10 min and then flow-mixed with an equal volume of oxidant using a Gibson-type 12-jet tangential mixer [23]. When time-dependence studies with oxidant scavengers established that S. cerevisiae killing occurred on the time scale of minutes (see Results), the simpler technique of making bolus additions of HOCl or NH2 Cl to rapidly stirring yeast suspensions was adopted. For routine studies, the cells were exposed to the oxidant for 10– 15 min and then incubated with excess (10–300 lM) sodium thiosulfate or mercaptoethanol for 5–10 min, following which portions were taken to determine
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surviving cells. Within the specified ranges, none of these variables—mixing method, reaction temperature, quencher identity and concentration, or incubation time—had any systematic effect upon the measured parameters, provided that incubation times were sufficiently long to allow complete reaction of HOCl. Viable cells were determined as colony-forming units (cfu) using either spread-plate or pour-plate techniques. Three different plating media were used over the course of these studies, consisting of the rich salts medium described above supplemented with 1–2% glucose (autoclaved separately) and 1.5% Difco technical grade agar, tripticase soy agar (TSA) plus 0.25% glucose, or a mixture of 2% yeast extract, 2% Difco Bacto-peptone, 1% glucose, and 1.5% Difco agar (YPD). Equivalent colonies were formed by glucose-grown or lactate-grown yeast with each of these plating media, although somewhat lower counts (10%) were obtained on TSA for ethanol-grown cells. Potassium ion release by HOCl-treated cells Orion Model 93-19 Kþ ion-specific and Model 90-02 double junction electrodes attached to an Orion Model 701-A digital potentiometer were used to measure potassium ion concentrations in the external medium of S. cerevisiae suspensions. The only deviation from manufacturerÕs recommended setup is that the ‘‘ionic strength adjustor’’ (5 M NaCl) used in the outer chamber of the reference electrode was diluted 50-fold, so that its Cl concentration was the same as that of the suspending PBS medium. The log½Kþ vs. mV response of this electrode assembly was linear in over the experimentally determined range (½Kþ ¼ 105 –102 M). The cells were warmed to ambient temperature and gently stirred for a few minutes to allow the electrode reading to stabilize. Appropriate amounts of HOCl were then added and the voltage changes were recorded over a period of 20 min. External concentrations of Kþ were determined by comparison to calibration curves constructed from standard [Kþ ] solutions at the time of the oxidation experiments. Adenylate phosphorylation levels Intracellular ATP, ADP, and AMP levels were analyzed by rapidly extracting the nucleotides from the cells into ice-cold 0.5 M HClO4 using a spring-loaded syringe, as previously described [20,24,25]. Nucleotide concentrations were determined by isocratic elution with 0.2 M ammonium phosphate, pH 5.6, from a 10 lm reversephase C18 HPLC column [24]. For glucose-grown cells, the suspension was supplemented by adding glucose to a final concentration of 22–270 mM for 10 min prior to extraction with HClO4 . This procedure is essential to obtain an accurate assessment of the metabolic capa-
bilities since viable glucose-grown cells suspended in media devoid of nutrients maintain very low EC values ( 60.1); similar to previous observations on bacteria [25], these EC values ‘‘step up’’ when challenged with a carbon source to levels that accurately reflect the adenine nucleotide-phosphorylating capabilities of the cells. Enzyme assays Activities of the cytosolic enzymes, aldolase (EC 4.1.2.13), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH, EC 1.2.1.12) were determined on supernatant fractions obtained by rupturing the cells by grinding with glass beads. Specifically, to determine aldolase activity, the yeast cells were pelleted, washed, and resuspended in 0.1 M glycylglycine buffer, pH 7.5. Acidwashed glass beads were added and the suspension was spun at 12,000g in a microcentrifuge at 4 °C until microscopic inspection showed that nearly all of the cells were disrupted. The supernatant and washes were collected and the aldolase activity was determined by using a standard colorimetric assay that relies upon formation of a hydrazone (kmax ¼ 240 nm) from reaction of hydrazine with 3-phosphoglyceralde, a product of the enzymatic cleavage of fructose-1,6-diphosphate (FDP) [26]. Measured initial rates of absorbance change at 240 nm were normalized according to the pelleted weights of each sample; activities of the HOCl-treated samples were expressed as percent activity of that obtained for untreated cells. The cells used for GAPDH analyses were resuspended in a medium containing 10 mM sodium pyrophosphate, 10 mM dithiothreitol (DTT), and 0.1 mM diethylenetriaminepentaacetic acid (DTPA), pH 8.5, and broken at 4 °C by spinning 30 s in a ‘‘Bead Beater’’ cell disrupter (Biospec Products, Bartlesville, OK). The beads were removed by gentle suction filtering and washed several times with buffer. The combined extract was centrifuged and activity of the supernatant was determined at 340 nm using a standard spectrophotometric assay that relies upon enzymatic reduction of NADþ by D -glyceraldehyde-3-phosphate (GAP) [27]; initial rates were expressed as percent activity of that obtained for untreated cells. Isolation of yeast plasma membranes and determination of their ATP-hydrolase activities followed, with minor modification, procedures developed by R. Serrano [28,29]. Harvested cells were washed three times in 100 mM phosphate, pH 7.4, treated with HOCl, and resuspended in a medium comprising 28 mM Tris–Cl, 5.5 mM EDTA, and 0.55 mM phenylmethylsulfonyl fluoride (PMSF) at pH 8.5. Three different methods for breaking the cells were evaluated, comprising vortexing in a test tube, shaking in a ‘‘Mini-Bead Beater’’ or vortexing in a ‘‘Bead-Beater,’’ all at 4 °C. The highest total activities and amount of protein recovered were achieved by using the ‘‘Mini-Bead Beater,’’ both of which were
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50–70% higher than the other two methods, which gave comparable total activities and protein yields. Specific activities obtained for all three methods were therefore very nearly equal. However, the low volume capacity (1.4 ml) of the ‘‘Mini-Bead Beater’’ caused the processing time of the samples to be unacceptably long. Vortexing in the Bead-Beater was chosen as the best compromise between recovery and processing time. With this method, total activity and membrane protein yields were independent of grinding time between 20 and 100 s, but beyond 100 s the activity declined progressively with time, presumably caused by inactivation of the ATPase. To determine ATPase activity, the HOCl-treated cells were vortexed for 45 s and the glass beads were removed by filtration and washed with a buffer comprising 20% glycerol and 80% of 0.2 mM EDTA, 0.2 mM DTT, and 10 mM Tris–Cl, pH 7.5. The plasma membrane was isolated by differential centrifugation and the membraneenriched pellet was washed with buffer containing 0.4 mM PMSF to decrease residual phosphate. The primary source of this phosphate appeared to be the disrupted cells themselves since parallel preparations that made use of buffers containing no inorganic phosphate gave equivalent amounts of residual phosphate at this stage of purification. Protein content was determined by the Bradford method using bovine serum albumin as a standard [30]. To determine ATPase activities, portions of the membrane suspended in the PMSF-containing wash buffer were reacted with vanadate-free ATP for 15 min at 30 °C; the inorganic phosphate released was determined as a phosphomolybdate complex [28,29] by comparing the absorbance measured at 750 nm to calibration curves obtained using standard phosphate solutions. Parallel measurements were made with 0.2 mM diethylstilbestrol (DES) added to the reaction medium. DES is a selective inhibitor of the PMA1 Hþ -ATPase; comparison of activities in its presence and absence permits determination of the relative amount of ATP hydrolytic activity attributable to this enzyme. Control measurements using analogous solutions minus ATP and minus membranes were made to correct for inorganic phosphate derived from extraneous sources and turbidity generated by the insoluble membranes, respectively. Glucose transport 14 D -[U- C]glucose
(8 mM, 12.5 mCi/mmol) in 90% ethanol was diluted 10-fold with 44 mM glucose to give a final solution that was 40 mM glucose in 9% ethanol with a specific activity of 1.25 mCi /mmol. Glucose transport rates were determined by following the general procedures described by Bisson and Fraenkel [31]. Specifically, 0.2 ml portions of the diluted radiolabeled sugar solution at 30 °C were added to 0.2 ml suspensions of 5 107 glucose-grown cells/ml that had been exposed
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to varying amounts of HOCl and warmed to 30 °C. These mixtures were briefly incubated for timed intervals, during which the suspension was vigorously vortex-mixed, and the reaction was quenched by adding 10 ml ice-cold water. The cells were then quickly filtered (<5 s) onto Whatman glass microfiber filters (GF/F, pore size 0.7 lm) and washed once with 10 ml cold water. The radioactivity accumulated on the filters was determined by placing them in Ecolite scintillation fluid and counting for 30 min on a Packard Tri-Carb 2700 Liquid Scintillation Analyzer. The counts obtained were corrected for nonspecific adsorption of the sugar onto the cells and filter by adding the radiolabeled sugar directly to the quench water prior to washing the collected cells [31]. In general, this background absorption comprised 10–20% of the total radioactivity measured for samples that had been incubated for 3 min before quenching. Isolation and O2 consumption by yeast mitochondria Mitochondria were isolated from yeast grown on lactate by a preparative sequence that involved hydrolysis of the cell wall with zymolase, homogenization of the resulting spheroplasts, and differential centrifugation. The only deviations from procedures originally described by Schatz and coworkers [32] were that: (1) 1.25 mg zymolase 20,000 per g wet weight of cells was used; (2) zymolase incubations were carried out at 21– 22 °C to minimize breakage of HOCl and NH2 Cl-treated cells; and (3) the final wash of isolated mitochondria was made with the buffer used to measure respiratory activity, i.e., 0.6 M sorbitol, 0.36 mM EDTA, 10 mM KH2 PO4 , 10 mM KCl, 10 mM Tris–Cl, and 0.3% BSA, pH 6.5 [33]. Respiration rates were measured at 25 °C in a thermostatted cell fitted with a YSI Model 5331 Clarktype polarographic electrode connected to a voltage/ time stripchart recorder. Mitochondria were added to the cell and background O2 uptake was measured for a few minutes, following which excess respiratory substrate (6 mM succinate or a-ketoglutarate) and then limiting amounts of ADP were added. The electrode response was calibrated from the pen deflection caused by introducing measured amounts of air into the O2 purged chamber, from which mitochondrial respiration rates were calculated. These were normalized to the amount of protein in each sample, which was determined by the Lowry method [34].
Results Glucose-regulated metabolism in S. cerevisiae To facilitate interpretation of the experimental data, a thumbnail summary of salient features of glucose
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metabolism is included here. Glucose exhibits broad regulatory control over pathways of intermediary metabolism in S. cerevisiae [35]. It represses expression and promotes catabolism of a wide variety of proteins, among which that are relevant to these studies are enzymes involved with mitochondrial respiration, the glyoxalate and TCA cycles, the plasma-membrane Hþ ATPase (PMA1) [36], and alcohol dehydrogenase II (ALDII), an isozyme whose function is primarily associated with oxidation of ethanol to acetaldehyde as a first step in the respiratory metabolism of ethanol (Fig. 1). Glucose also induces expression of numerous enzymes [37], including those associated with the glycolytic pathway, and alcohol dehydrogenase I (ALDI), an isozyme that is primarily involved with reduction of acetaldehyde to ethanol in fermentative metabolism (Fig. 1). Additionally, it regulates a set of genes whose products are hexose transport proteins that are the predominant glucose transporters in S. cerevisiae, the most relevant effect being induction of a low-affinity transporter that is the primary carrier in high-glucose media [16]. Growth on the nonfermentable carbon sources, lactate and ethanol, however, is driven entirely by respiration [35]; for these substrates, syntheses of enzymes associated with respiratory metabolism are derepressed. As a consequence of these regulatory patterns, aerobic growth of S. cerevisiae in batch culture on glucose can be diauxic [36,38]. The first phase, during which most of the glucose present is fermented, is characterized by rapid cell proliferation; upon exhaustion of the glucose, the metabolic changes attending
Fig. 1. Abbreviated intermediary metabolism pathway. Carbon sources used in this study are highlighted in bold type; enzymes that are highly susceptible to inactivation by HOCl are indicated in the scheme. Among others, glucose represses expression of ADHII, the enzymes of the glyoxalate and TCA cycles, the PMA1 Hþ -ATPase, and the proteins of the mitochondrial respiratory chain; among others, glucose induces expression of enzymes of the glycolytic pathway and hexose transporters.
derepression allow the cells to adapt to a slower respirative growth on the accumulated ethanol. Toxicity of HOCl and simple chloramines Saccharomyces cerevisiae exposed to bolus additions of HOCl exhibited sigmoidal dose-dependent survival curves, as measured by their ability to form colonies on nutrient-rich media; comparable toxicities were observed for cells grown on the fermentation carbon source, glucose, and on the respiratory carbon sources, ethanol or lactate. Consistent with these results, toxicities of HOCl were identical within experimental uncertainty for glucose-grown cells harvested in early log, late log, and stationary phases. Under these conditions, the primary metabolic pathway for carbohydrate utilization switches from fermentative to oxidative upon accumulation of ethanol and depletion of glucose from the growth medium [38]. Typical results for cells harvested in early to mid-log phase growth are shown in Fig. 2.
Fig. 2. Survival curves for S. cerevisiae exposed to HOCl. 3.6 107 glucose-grown cells/ml (squares) or 8.0 107 ethanol-grown cells/ml (circles) in PBS were flow-mixed through a 12-jet tangential mixer at 23 °C with an equal volume of HOCl to give the indicated final concentrations. The open symbols indicate results obtained when a stoichiometric excess of S2 O2 3 (640 lM) was present in the receiving flask; the closed symbols indicate the results obtained when S2 O2 3 addition was delayed 10 min. Individual points are averages of 2-4 replicate counts of cfu on rich-salt medium determined using pour-plate techniques.
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From the survival curves, one estimates that 20–40 lM HOCl/107 cells, or (1–2) 109 molecules HOCl/cell are required to inhibit colonial growth by 99% of the cells (LD99 ). Addition of the HOCl scavengers S2 O2 or 3 mercaptoethanol immediately after exposure to lethal concentration levels of HOCl completely protected the cells (Fig. 2). However, protection was incomplete if the subsequent addition of the scavenger was delayed and became less effective as the time interval was increased. The exposure time required for complete inactivation determined from this temporal loss of protection was 1– 2 min. Chloramine (NH2 Cl) exhibited comparable toxicities and killing rates toward S. cerevisiae grown on all three carbon sources. Inactivation of susceptible cytosolic enzymes Yeast aldolase and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) are enzymes of the glycolytic pathway [39] that are highly susceptible to inhibition by HOCl [12,40] and other oxidants [41,42]. As such, they are both convenient markers of oxidative damage by HOCl within the cytosol and potential HOCl target sites for lethal lesions. In Fig. 3, titrimetric loss of the activities of these enzymes upon exposure of glucose-
Fig. 3. HOCl inactivation of S. cerevisiae cytosolic GAPDH and aldolase. For GAPDH determinations, bolus additions of HOCl to the indicated concentrations were made at 23 °C to 1.5 107 glucosegrown cells/ml in PBS. After 10 min, residual oxidant was neutralized with 200 lM S2 O2 3 . Open circles are GAPDH activities/unit volume of suspension; open squares are normalized cfu on YPD determined using pour-plate techniques. The data are a composite of results for several cultures, with each point being an individual determination. At [HOCl] > 100 lM, both viabilities and GAPDH activities were <1% of the control values. For aldolase determinations, 3.7 107 glucosegrown cells/ml in PBS were flow-mixed at 25 °C with an equal volume of HOCl to give the indicated final concentrations; 240 lM S2 O2 3 was added 20 min after mixing. Solid circles are the aldolase activities; solid squares are cfu on rich-salt medium determined with pour-plate techniques. The aldolase data have been normalized to 1.5 107 cells/ ml to allow direct comparison to the GAPDH data.
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grown S. cerevisiae to increasing amounts of HOCl is compared to the corresponding viabilities determined for the same samples. The data indicate that 65–75% of the original activity of these enzymes is retained in the nonviable cells and that twice the amount of HOCl is required to inactivate these enzymes as is required to prevent cell division. Thus, although some damage is incurred, these enzymes do not appear to be primary target sites associated with loss of replicative capacity. Mitochondrial function The functional capabilities of mitochondria isolated from cells exposed to HOCl were assessed by measuring O2 uptake under various conditions of respiratory control [15,33]. State 3 (or uncoupled) respiration was induced by adding excess ADP or the protonophore, carbonyl cyanide m-chlorophenyl hydrazone (cccp), to mitochondrial suspensions containing the respiratory reductants, succinate or a-ketoglutarate. Under these conditions, respiratory rates are maximal and can be
Fig. 4. Functional capabilities of isolated S. cerevisiae mitochondria. 5–6 107 lactate-grown cells/ml in PBS were flow-mixed at 23 °C with equal amounts of HOCl to the indicated final concentrations and mitochondria were isolated and characterized using succinate as respiratory substrate as described in Materials and methods. Squares are cfu, determined on TSA using pour-plate methods; open circles are RCR; crossed circles are RCR measured in the presence of cccp, down triangles are the state 3 (uncoupled) respiration rates; and up triangles are the P/O ratio. All values are normalized to constant protein content of the samples and to the values obtained with control mitochondria.
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directly related to the extent of damage suffered by the respiratory chain. State 4 (or coupled) respiration is measured under conditions where the reducing substrate is in excess and the proton motive force across the mitochondrial cristae membrane is maximal, chemiosmotically restricting electron transport. The ratio of state 3 to state 4 respiration rate (respiratory control ratio (RCR)) indicates the efficiency of this chemiosmotic coupling; as such, the RCR provides information on the impermeability of the cristae membrane to the proton and other ions. The amount of O2 consumed during ADP-induced state 3 respiration (expressed as the P/O ratio, i.e., [ADP] added/[O2 ] consumed) provides a measure of the efficiency of coupling of the electron transport chain to ATP synthesis. Inhibition of the Fo F1 -ATP synthase, loss of membrane integrity, and inhibition of electron transport can all lead to reduced P/O ratios. These parameters, obtained with succinate as respiratory substrate, are compared to viabilities of lactate-grown S. cerevisiae in Fig. 4. Addition of an amount of HOCl that is sufficient to inhibit growth in 95% of the cells decreases the state 3 respiration rate to 50% of its normal value, with smaller decreases appearing in the RCR and P/O ratio. This reactivity pattern indicates that mitochondrial damage occurs primarily within the electron transport chain and that the extent of this damage may not be sufficient to significantly inhibit ATP generation by nonviable cells. Nearly identical patterns were observed when a-ketoglutarate was used as respiratory substrate and when NH2 Cl was used as oxidant. Absolute values of RCR and P/O for mitochondria isolated from untreated cells (2.5–2.7 and 1.7–1.9, respectively, with succinate as substrate and 5.1 and 2.5, respectively, with a-ketoglutarate as substrate) were nearly identical to originally reported values [33].
manner that paralleled inhibition of mitochondrial function (Fig. 4). For example, 50% reduction in phosphorylation levels of 6 107 lactate-grown or ethanolgrown cells/ml required exposure to 150 lM HOCl or 100 lM HOCl, respectively, as determined from EC vs. [HOCl] plots analogous to Fig. 3, whereas LD50 of the same suspensions was 55 lM HOCl. In contrast, glucose-grown cells harvested in mid-log phase showed EC values whose decline paralleled loss of viability. When harvested in late-log phase, the EC decline lagged behind the survival curve, and when harvested in early stationary phase, the decline in EC was very similar to that recorded for cells grown on the respiratory substrates (Fig. 5). In the experiments with glucose-grown cells, all samples were incubated with glucose following exposure to HOCl prior to extraction of the nucleotides for EC determinations. The phase-dependent changes in dose–response behavior of EC for glucose-grown cells (Fig. 5) parallel the shift in carbohydrate metabolism
Adenine nucleotide phosphorylation levels The adenylate energy charge (EC), defined as EC ¼ ([ATP] + 1/2[ADP])/([ATP] + [ADP] + [AMP]) [43], is a convenient index of the overall level of phosphorylation of the adenine nucleotide pool within the cytosol of microbial cells. Rapidly growing cells maintain EC P 0:8, whereas this value is considerably less in resting or nonviable cells [10,11,24,44]. The EC of oxidatively damaged cells will decrease if either ATP biosynthesis is impaired or consumption becomes excessive, particularly when futile cycles are established in ATPutilizing systems required to maintain homeostasis. Changes in EC within S. cerevisiae upon exposure to HOCl depended upon the carbon nutrient source and, for glucose-grown cells, upon the growth phase at the time they were isolated. EC values were normal at lethal levels of HOCl for cells grown on respiratory carbon sources, but declined at higher concentrations in a
Fig. 5. Adenylate energy charge (EC) in S. cerevisiae grown on glucose. EC values for 2.4 107 cells/ml harvested in the mid-log region of the first growth phase (open circles), 4.8 107 cells/ml harvested at the transition between growth phases (up triangles), and 3.8 107 cells/ml harvested in the second growth phase (diamonds) flow-mixed in PBS at 28 °C with an equal volume of HOCl to the indicated final concentrations. Following reaction, the cell suspensions were treated with 0.3 mM S2 O2 3 to remove any residual oxidant, incubated 20 min in 22 mM glucose, and extracted with HClO4 . Cfu (open squares) were determined on TSA using pour-plate techniques. All data have been normalized to a constant cell density of 4.3 107 cells/ml.
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from fermentative to respiratory upon depletion of glucose from the growth medium [38,39]. Insensitivity of glucose transport to lethal doses of HOCl Exposure of fermenting S. cerevisiae to 14 C-glucose resulted in rapid net accumulation of radioactivity within the cells; as previously reported [31], uptake rates were linear for 3 min and then gradually declined with increasing incubation time (data not shown). Glucose uptake was remarkably insensitive to high doses of HOCl. The initial rate of cells exposed to a LD99 dose of HOCl was two-thirds of the accumulation rate of untreated cells, as judged from the initial slopes of scintillation counts vs. incubation time, and 50% inhibition of 2.5 107 cells/ml required exposure to 470 lM HOCl in suspensions whose LD50 was 150 lM. Under the prevailing growth conditions, glucose is taken up by facilitated diffusion via an integral membrane protein that is thought to form an aqueous transmembrane channel allowing passage of the sugar across the bilayer [16,17]. As such, glucose transport is not energy-linked to membrane polarization, but is driven by enzymatic phosphorylation within the cytosol. Equilibration of intracellular glucose with the external solution takes only 2–3 min under the conditions of our experiments, as has been demonstrated by using nonmetabolizable substrate analogs and mutant cells devoid of hexokinase and glucokinase activities [31]. Consequently, the continued accumulation of radioactivity beyond 3 min incubation indicates that, in addition to glucose transport per se, glucose phosphorylation capabilities within the cell had not been severely damaged by exposure to lethal levels of HOCl.
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the solid symbols; for these data, the HOCl concentration reported on the abscissa is also 20-fold greater than indicated by the abscissa values. As we have repeatedly observed in earlier studies with bacteria [13,20,24], the amount of HOCl required for loss of function scales linearly with the density of cells in suspension. Thus, the results obtained are not dependent upon either the transient HOCl concentration generated upon bolus addition of the oxidant or the number of reactive sites presented by the yeast. Potassium ion leakage Yeast cells actively accumulate Kþ via voltage-gated channels in their plasma membranes in response to transmembrane potentials generated by the PMA1
Inactivation of the PMA1 Hþ -ATPase Comparisons of cfu with loss of ATP-hydrolytic activity in plasma membrane fractions isolated from glucose-grown cells exposed to varying concentrations of HOCl are shown in Fig. 6. These values have been corrected for ATP hydrolysis from alkaline phosphatases, apyrases, and other enzymes present in the membrane fractions by performing control experiments using diethylstilbesterol to block the PMA1 Hþ -ATPase activity. In the presence of 0.2 mM DES, the specific activity was typically 8% of the value of the untreated membrane fraction in the absence of the inhibitor, indicating that 92% of the total measured activity could be ascribed to the PMA1 Hþ -ATPase. As with the other activities that were measured, inactivation of the PMA1 lagged behind loss of colony-forming capabilities, so that three times more HOCl was required to reduce the activity of the proton pump by 50% than to completely inhibit growth. Additional studies using 20-fold greater S. cerevisiae cell densities are also reported in Fig. 6 as
Fig. 6. PMA1 Hþ -ATPase activity in isolated plasma membranes. Bolus additions of 80 mM HOCl were made at 30 °C to the final indicated concentrations to 5 107 (open symbols) or 1 109 (crosshatched symbols) glucose-grown cells/ml in 100 mM phosphate, pH 7.4. Cfu were determined by pour-plate analysis using YPD; each point represents an average of counts from five plates whose average deviation from the mean was typically within 10%. ATP-hydrolase specific activities are plotted relative to values obtained for plasma membranes from untreated control cells (0.22 0.05 lM Pi /min-mg protein). Each point is the average of at least three enzymatic determinations minus the activity found in the presence of DES; average deviations from the mean were typically less than 5%. As discussed in the text, concentrations of HOCl used for the higher cell densities (cross-hatched symbols) are 20 greater than indicated on the abscissa.
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Hþ -ATPase [18,45,46]. Oxidative injury that leads to depolarization or rupture of these membranes will be accompanied by Kþ release to the external medium. In ethanol-grown and lactate-grown cells, the rate of Kþ efflux increased in a dose-dependent manner following addition of HOCl to the medium. This slow release was nearly linear with time and underwent only very small diminution in rate when the S2 O2 3 was added 10 min later, indicating that Kþ release was not caused by ongoing oxidative damage, but was a consequence of prior irreversible damage to the cell. Initial rates of Kþ efflux measured at various levels of exposure to HOCl are plotted for the two growth media in Fig. 7. Complete release of potassium ion gave ½Kþ ¼ 0:5–0:8 mM in the medium, as determined by treating the cells with a large excess of HOCl. Saccharomyces cerevisiae grown to early-log phase on glucose exhibited a different response to added HOCl. In this case, bolus additions of up to 100 lM HOCl to 2 107 cells/ml did not increase the Kþ leak rates above a measured basal level of 1.8 0.3 lM/min (cf., Fig. 7). However, background Kþ levels in the cell suspensions increased (e.g., from 130 to 250 lM) in a dose-dependent manner following this treatment, consistent with rapid release of Kþ from a small subpopulation of the cells.
Fig. 7. Initial rates of Kþ release from respiring S. cerevisiae. Initial leak rates in PBS at 22 °C. For lactate-grown cells (at 2.1 107 cells/ ml), cfu for additions of 30, 50, 70, 80, and 100 lM HOCl were 124, 35, 16, 13, and 8%, respectively, of the control value; for ethanol-grown cells (at 3.2–3.6 107 cells/ml), cfu were 56 and 0% of controls.
Discussion Comparison of HOCl-inflicted damage in S. cerevisiae to that in bacteria One major difference in the patterns of oxidative damage caused by HOCl to bacteria and S. cerevisiae is that, in bacteria, phosphorylation levels in the adenine nucleotide pool decline in parallel with survival [9,20], whereas in S. cerevisiae grown on respiratory carbon sources phosphorylation levels in nonviable cells are nearly normal. This same behavior is observed in the late stages of growth on glucose where ethanol has become the nutrient carbon source and metabolism is respiratory (Fig. 5). Consistent with this result, mitochondria isolated from respiring cells exposed to lethal levels of HOCl retain a major fraction (60%) of their capacity for oxidative phosphorylation, with most of the damage occurring to the electron transport chain, as measured by state 3 respiration rates (Fig. 4). Other indicators of oxidative damage within the cytosol are the enzymes aldolase and GAPDH [12,40–42]. Approximately two-thirds of the normal intracellular activity of these enzymes is retained in HOCl-inactivated cells (Fig. 3). These experiments do not distinguish between oxidative inactivation of the proteins and their loss from the cytosol, i.e., by breaching of the yeast plasma membrane. However, the slow progressive increase in Kþ leak rates accompanying exposure of yeast to increasing amounts of HOCl (Fig. 7) indicates that the cellular plasma membrane remains ion-impermeable, i.e., intact, in most of the cells. Rupture of the membrane would give an immediate and abrupt increase in the extracellular Kþ concentration. Lactate dehydrogenase (LDH) and ALD are even more susceptible to inhibition by HOCl, at least in homogeneous solution [12]. Nonetheless, from the high EC values measured in HOCl-inactivated cells (Fig. 5), one can infer that the activities of all respiratory pathway enzymes, including LDH and ALDII, are sufficient to support near-normal respiratory capacity with lactate and ethanol as carbon sources (Fig. 1). In contrast to the results with respiring cells, EC values in fermenting S. cerevisiae declined in a manner that paralleled loss in cfu (Fig. 5). Because these were step-up experiments, the EC values indicate the capacity of oxidized cells to undertake substrate-level phosphorylation [25]. Obviously, loss of this capability could arise if the glycolytic pathway were inhibited by HOCl. However, the high residual activities of aldolase and GAPDH, two enzymes of the glycolytic pathway that are highly susceptible to HOCl inactivation [12], as well as the apparent limited overall damage to other vulnerable cytosolic proteins caused by lethal amounts of HOCl, render this unlikely. The alternative possibility that glucose transport across the plasma membrane was
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inhibited can also be dismissed on the basis of the insensitivity of glucose uptake by the cells to exposure to HOCl. A second major difference between these cell types is that, unlike bacteria, the plasma membrane-localized ATPase is remarkably resistant to inactivation by HOCl. Specifically, the activity of the bacterial Fo F1 -ATP synthase is lost in parallel with cfu [20], whereas significant loss of activity in the yeast PMA1 Hþ -ATPase requires addition of HOCl in amounts several-fold in excess of that required for inhibition of growth (Fig. 6). PMA1 is a plausible target for inactivation because gene-knockout experiments have established that its function is critical to cell survival [19] and the protein contains nine cysteine groups, the thiol substituents of which are vulnerable to HOC1 oxidation [12,40,47–49]. However, this and other P-type ATPases do not appear to be very sensitive to chemical modification or replacement of their cysteine residues [50]. The e-amino groups of conserved lysine residues represent possible alternative sites of attack, particularly since several reactive essential lysine groups have been identified within the ATP-binding domain of the yeast Hþ -ATPase [51,52]. Based upon results with other oxidants [53,54], N-chlorination at these sites could lead to inactivation or loss of function attending further degradation. In any event, inactivation of the yeast proton pump in these studies required exposure to a considerable excess of HOCl over that required for inhibition of growth. A third major difference is that the time scales for loss of colony-forming capabilities are dramatically different. Whereas bacteria are killed ‘‘instantaneously’’ upon exposure to lethal levels of HOCl [14], inactivation of both fermenting and respiring yeast required incubation with the toxin for several minutes (Fig. 2), indicating that the target sites in yeast are either relatively unreactive or otherwise physically protected from the external medium, e.g., as would be the case if access to the vulnerable sites required diffusion across the yeast cell membrane.
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media. In support of this viewpoint, we have previously shown that the phosphorylating capabilities of bacteria measured immediately after exposure to either bolus additions of HOCl or MPO-generated oxidant correlate directly with cfu measured 24 h later [25]. Thus, the cfu count accurately reflects the immediate HOCl-induced loss of energy-transducing capabilities in the microbial population. One consequence of this conclusion is that, although the lethal lesions have not yet been identified, death in fermenting S. cerevisiae must arise directly from HOCl inactivation of critical cellular functions. These issues are not so straightforward for respiring S. cerevisiae. Specifically, the functional capabilities of the yeast mitochondria are largely unimpaired (Fig. 4) and the adenylate phosphorylation levels in respiring cells are near-normal (Fig. 5), despite the loss in ability to undergo cell division. Thus, for yeast, it is quite possible that metabolic transformations could occur via biosynthesis during incubation of the plated cells. Moreover, there appears to be no correlation between overt loss of essential cellular function and death or localization of lethal lesions to the plasma membrane. Rather, there appears to be a global loss of function among HOCl-susceptible biomolecules located throughout the cell. In all cases, however, loss of function requires significantly greater amounts of HOCl than does inhibition of cell growth. As with other cellular oxidants [55–57], exposure to increasing amounts of HOCl causes the response of endothelial cells to vary progressively from transient inhibition of growth to apoptosis to necrosis (lysis) [58]. Programmed cell death has recently been discovered in yeast [59,60]; one of the effectors that can trigger apoptosis in these cells is oxidative stress, leading to mitochondrial damage, and/or loss of endogenous antioxidant levels. Consequently, one likely explanation for the loss of colony-forming capabilities prior to significant loss of discrete biological functions is that the oxidative damage inflicted triggers the biochemical cascades comprising S. cerevisiae apoptosis, which then ensues upon exposing the cells to nutrient agar.
Microbicidal mechanisms Biological relevance The ability to sustain colonial growth as a definition of viability in studies that attempt to correlate metabolic dysfunctions with death has been questioned primarily because there most often exists a long time interval between the measurement of specific functions, enzymatic activities or metabolite concentrations, and the measurement of colony-forming units (cfu), during which time additional damage or repair might occur that could alter the relationships. For bacteria and fermenting S. cerevisiae, at least, the inability to generate ATP and maintain adenylate phosphorylation levels ensures that biosynthesis and maintenance of cellular homeostasis will not occur, even for cells plated on nutrient-rich
Based upon survival curves obtained from bolus additions of oxidant to cell suspensions, the amount of HOCl required to inhibit cell division in S. cerevisiae is 1.5 109 HOCl/cell (Fig. 1), which is 10 times greater than that required to inhibit growth of representative laboratory strains of E. coli, S. lactis, and P. aeruginosa [20]. Recent estimates place a lower limit of myeloperoxidase-generated HOCl in the respiratory burst at 70% of the O2 consumed [61]. On average, each human neutrophil consumes as much as 4 1010 O2 molecules following respiratory stimulation [8]. Consequently, each neutrophil has, in principle, the
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capacity to kill maximally 20 S. cerevisiae using only its metabolically generated HOCl. This estimate is clearly an upper limit, however, since some of the oxidants will certainly be directed against other biological targets, including antioxidants in the serum-derived phagosomal fluid [62], and opsonins [8] and neutrophilderived protein [6,7]. Hypochlorous acid is perhaps the most toxic of the potential phagocyte-generated oxidants [62]; consequently, the most important conclusion to be drawn from these calculations may be that neutrophils cannot generate large excesses of oxidants beyond that required to inactivate phagocytosed microbes.
Acknowledgments The authors are grateful to Sirisha Dontireddy for developing protocols for isolation and functional characterization of the PMA1 Hþ -ATPase and to Professor Raymond Reeves for helpful discussions concerning yeast apoptosis. This work was supported by National Institutes of Health Grant AI15834.
[19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31] [32] [33] [34] [35]
References [1] S.J. Klebanoff, in: J.I. Gallin, I.M. Goldstein, R. Snyderman (Eds.), Inflammation: Basic Principles and Chemical Correlates, Raven Press, New York, 1998, pp. 391–444. [2] M.B. Hampton, A.J. Kettle, C.C. Winterbourn, Infect. Immun. 64 (1996) 3512–3517. [3] J.K. Hurst, in: J. Everse, K.E. Everse, M.B. Grisham (Eds.), Peroxidases in Chemistry and Biology, vol. 1, CRC Press, Boca Raton, FL, 1991, pp. 37–62. [4] H.B. Dunford, Heme Peroxidases, Wiley-VCH, New York, 1999. [5] P.G. Furtmuller, U. Burner, W. Jantschko, G. Regelsberger, C. Obdinger, Redox Rep. 5 (2000) 173–178. [6] A.L.P. Chapman, M.B. Hampton, R. Senthilmohan, C.C. Winterbourn, A.J. Kettle, J. Biol. Chem. 277 (2002) 9757–9762. [7] H. Rosen, J.R. Crowley, J.W. Heinecke, J. Biol. Chem. 277 (2002) 30463–30468. [8] Q. Jiang, D.A. Griffin, D.F. Barofsky, J.K. Hurst, Chem. Res. Toxicol. 10 (1997) 1080–1089. [9] J.K. Hurst, S.V. Lymar, Acc. Chem. Res. 32 (1999) 520–528. [10] C.J. Knowles, Symp. Soc. Gen. Microbiol. 27 (1977) 241–283. [11] A.G. Chapman, D.E. Atkinson, Adv. Microb. Physiol. 15 (1977) 253–306. [12] J.M. Albrich, C.A. McCarthy, J.K. Hurst, Proc. Natl. Acad. Sci. USA 78 (1981) 210–214. [13] J.M. Albrich, J.H. Gilbaugh III, K.B. Callahan, J.K. Hurst, J. Clin. Invest. 78 (1986) 177–184. [14] J.M. Albrich, J.K. Hurst, FEBS Lett. 144 (1982) 157–161. [15] D.G. Nicholls, S.J. Ferguson, Bioenergetics3, Academic Press, Amsterdam, 2002. [16] J. Hor ak, Biochim. Biophys. Acta 1331 (1997) 41–79. [17] M. Mueckler, C. Caruso, S.A. Baldwin, M. Panico, I. Blensch, H.R. Morris, W.J. Allard, G.E. Lienhard, H.F. Lodish, Science 229 (1985) 941–945. [18] R. Serrano, in: J.R. Broach, J. Pringle, E.W. Jones (Eds.), The Molecular and Cellular Biology of the Yeast Saccharomyces: Genome Dynamics, Protein Synthesis, and Energetics, vol. I, Cold
[36] [37] [38] [39]
[40] [41]
[42] [43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54]
Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1991, pp. 523–585. R. Serrano, M.C. Kielland-Brandt, G.R. Fink, Nature 319 (1986) 689–693. W.C. Barrette Jr., D.M. Hannum, W.D. Wheeler, J.K. Hurst, Biochemistry 28 (1989) 9172–9178. J.C. Morris, J. Phys. Chem. 70 (1966) 3798–3805. E.L. Thomas, M.B. Grisham, M.M. Jefferson, Methods Enzymol. 132 (1986) 569–585. Q.H. Gibson, L. Milnes, Biochem. J. 91 (1964) 161–171. W.C. Barrette Jr., J.M. Albrich, J.K. Hurst, Infect. Immun. 55 (1987) 2518–2525. W.C. Barrette Jr., D.M. Hannum, W.D. Wheeler, J.K. Hurst, J. Bacteriol. 170 (1988) 3655–3659. O.C. Richards, W.J. Rutter, J. Biol. Chem. 236 (1961) 3177–3181. V. Worthington (Ed.), Worthington Enzyme Manual, Worthington Biochemical Corp, Lakewood, NJ, 1993, pp. 201–206. R. Serrano, Methods Enzymol. 157 (1988) 533. R. Serrano, Biochem. Biophys. Acta 947 (1988) 1–28. M.M. Bradford, Anal. Biochem. 72 (1976) 248–254. L.F. Bisson, D.G. Fraenkel, Proc. Natl. Acad. Sci. USA 80 (1983) 1730–1734. G. Daum, P. Botini, G. Schatz, J. Biol. Chem. 257 (1982) 13028– 13033. T. Ohnishi, K. Kawaguchi, B. Hagihara, J. Biol. Chem. 241 (1966) 1797–1806. O.H. Lowry, N.J. Rosebrough, A.L. Farr, R.J. Randall, J. Biol. Chem. 193 (1951) 265–275. M. Johnston, M. Carlson, in: J.R. Broach, J.R. Pringle, E.W. Jones (Eds.), The Molecular and Cellular Biology of the Yeast Saccharomyces: Gene Expression, vol. II, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1992, pp. 193– 281. A.R. Fernandes, I. Sa-Correia, Yeast 20 (2003) 207–219. M.B. Grisham, M.M. Jefferson, D.F. Melton, E.L. Thomas, J. Biol. Chem. 259 (1984) 10404–10413. O. K€appeli, Adv. Microb. Physiol. 28 (1986) 181–209. D.G. Fraenkel, in: J.N. Strathern, E.W. Jones, J.R. Broach (Eds.), The Molecular Biology of the Yeast Saccharomyces: Metabolism and Gene Expression, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1982, pp. 1–37. J.M. Pullar, C.C. Winterbourn, M.M. Vissers, Am. J. Physiol. 277 (1999) H1505–H1512. P.A. Hyslop, D.B. Hinshaw, W.A. Halsey Jr., I.U. Schraufst€atter, R.D. Sauerheber, R.G. Spragg, J.H. Jackson, C.G. Cochrane, J. Biol. Chem. 263 (1988) 1665–1675. D.P. Buchczyk, K. Briviba, F.U. Hartl, H. Sies, Biol. Chem. 381 (2000) 121–126. D.E. Atkinson, Biochemistry 7 (1968) 4030–4034. W.J. Ball Jr., D.E. Atkinson, J. Bacteriol. 121 (1975) 975–982. A. Ambesi, M. Miranda, V.V. Petrov, C.W. Slayman, J. Exp. Biol. 203 (2000) 155–160. A. Rodriguez-Navarro, Biochim. Biophys. Acta 1469 (2000) 1–30. M.C.M. Vissers, A. Stern, F. Kuypers, J. van den Berg, C.C. Winterbourn, Free Radic. Biol. Med. 16 (1994) 703–712. E.L. Thomas, Infect. Immun. 25 (1979) 110–116. R. Hurst, Y. Bao, S. Ridley, G. Williamson, Biochem. J. 338 (1999) 723–728. V.V. Petrov, C.W. Slayman, J. Biol. Chem. 270 (1995) 28535– 28540. J.P. Pardo, C.W. Slayman, J. Biol. Chem. 263 (1988) 18664– 18668. R. Rao, C.W. Slayman, J. Biol. Chem. 268 (1993) 6708–6713. A. Valiakhmetov, D.S. Perlin, J. Biol. Chem. 278 (2003) 6330– 6336. N. Stadler, M. H€ ofer, K. Sigler, Free Radic. Res. 35 (2001) 643– 653.
D.A. King et al. / Archives of Biochemistry and Biophysics 423 (2004) 170–181 [55] A.G. Wiese, R.E. Pacifici, K.J.A. Davies, Arch. Biochem. Biophys. 318 (1995) 231–240. [56] K.-T. Lin, J.-Y. Xue, M.C. Lin, E.G. Spokas, F.F. Sun, P.Y.K. Wong, Am. J. Physiol. 274 (1998) C855–C860. [57] R.H. Burdon, V. Gill, D. Alliangana, Free Radic. Res. 24 (1996) 81–93. [58] M.C.M. Vissers, J.M. Pullar, M.B. Hampton, Biochem. J. 344 (1999) 443–449.
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[59] F. Madeo, S. Engelhardt, E. Herker, N. Lehmann, C. Maldener, A. Proksch, S. Wissig, K.-U. Fr€ ohlich, Curr. Genet. 41 (2002) 208–216. [60] M. Priault, N. Camougrand, K.W. Kinnally, F.M. Vallette, S. Manon, FEMS Yeast Res. 4 (2003) 15–27. [61] A.J. Kettle, C.C. Winterbourn, Redox Rep. 3 (1997) 3–15. [62] S.V. Lymar, J.K. Hurst, Chem. Res. Toxicol. 8 (1995) 833–840.
HOCl-mediated cell death and metabolic dysfunction in the yeast Saccharomyces cerevisiae David A. King,a Diane M. Hannum,b Jian-Shen Qi,b and James K. Hursta,b,* b
a Department of Chemistry, Washington State University, Pullman, WA, USA OGI School of Science and Engineering, Oregon Health and Science University, Beaverton, OR, USA
Received 30 October 2003, and in revised form 9 December 2003
Abstract The nature of oxidative damage to Saccharomyces cerevisiae caused by levels of HOCl that inhibit cell replication was explored with the intent of identifying the loci of lethal lesions. Functions of cytosolic enzymes and organelles that are highly sensitive to inactivation by HOCl, including aldolase, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), and the mitochondrion, were only marginally affected by exposure of the yeast to levels of HOCl that completely inhibited colony formation. Loss of function in membrane-localized proteins, including the hexose transporters and PMA1 Hþ -ATPase, which is the primary proton pump located within the S. cerevisiae plasma membrane, was also marginal and Kþ leak rates to the extracellular medium increased only slowly with exposure to increasing amounts of HOCl, indicating that the plasma membrane retained its intrinsic impermeability to ions and metabolites. Adenylate phosphorylation levels in fermenting yeast declined in parallel with viability; however, yeast grown on respiratory substrates maintained near-normal phosphorylation levels at HOCl doses several-fold greater than that required for killing. This overall pattern of cellular response to HOCl differs markedly from that previously reported for bacteria, which appear to be killed by inhibition of plasma membrane proteins involved in energy transduction. The absence of significant loss of function in critical oxidant-sensitive cellular components and retention of ATP-synthesizing capabilities in respiring yeast cells exposed to lethal levels of HOCl suggests that toxicity in this case may arise by programmed cell death. Ó 2003 Elsevier Inc. All rights reserved. Keywords: ATP synthesis; Fungicidal mechanisms; Glycolysis; Hypochlorous acid; Oxidative phosphorylation; Proton pumps
Myeloperoxidase (MPO)1-mediated reactions play a major role in the microbicidal action of respiring neutrophils [1,2]. Although MPO catalyzes a wide range of peroxidative reactions [3–5], including two-electron oxidations of most halides and the pseudohalide, SCN , to the corresponding hypohalites, as well as one-electron
*
Corresponding author. Fax: 1-509-335-8867. E-mail address: [email protected] (J.K. Hurst). 1 Abbreviations used: ALD, alcohol dehydrogenase; cfu, colonyforming units; DES, diethylstilbestrol; DTPA, diethylenetriaminepentaacetic acid; DTT, dithiothreitol; EC, adenylate energy charge; FDP, fructose-1,6-diphosphate; GAP, D -glyceraldehyde-3-phosphate; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; LD99 , concentration required to inhibit colony formation in 99% of the cells; LDH, lactate dehydrogenase; MPO, myeloperoxidase; PBS, phosphate-buffered saline; PMSF, phenylmethylsulfonyl fluoride; RCR, respiratory control ratio; TSA, trypticase soy agar; YPD, yeast-peptone dextrose agar. 0003-9861/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2003.12.012
oxidations of NO 2 and a variety of organic reductants, chloride ion is generally regarded as the physiological substrate. This viewpoint is supported by recent studies using bacteria [6,7] and fluorescent particulate probes [8] to trap hypochlorous acid (HOCl) generated within neutrophil phagosomes. The patterns of metabolic dysfunction observed in bacteria exposed to HOCl suggest a bactericidal mechanism wherein death occurs by interruption of the energy-transducing capabilities of the cell by inhibiting plasma membrane-localized proteins, specifically the Fo F1 -ATP synthase, proteins involved with active transport of metabolites, and the respiratory redox chain [9]. Loss of these functions renders bacteria incapable of either oxidative or substrate-level phosphorylation, and nucleotide phosphorylation levels correspondingly plummet to levels far below those necessary to maintain homeostasis, much less undertake biosynthetic repair [10,11]. In contrast, cytosolic
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enzymes that are highly susceptible to irreversible inhibition by HOCl, such as aldolase and b-galactosidase, maintained nearly full activity within bacteria that had been exposed to amounts of HOCl that were severalfold higher than is required for killing [12,13]. Thus, the observed cytosolic protection of these HOCl-responsive markers suggests that cellular inactivation by HOCl is not accompanied by extensive oxidation or chlorination of intracellular components. Furthermore, quench-flow experiments have established that these Escherichia coli are killed within 100 ms of exposure to lethal levels of HOCl [14], supporting the notion that the locus of lethal lesions is the bacterial envelope and killing involves oxidation of highly reactive nucleophilic centers located therein. Because energy transduction in all prokaryotes occurs across the ion-impermeable plasma membrane, this model is attractive in accounting for the ‘‘universal’’ bactericidal properties of HOCl [1]. However, the model is clearly not applicable to simple eukaryotes where respiration-linked ATP synthesis is mitochondrial, and therefore occurs within the relatively protected cytosol of the cell. Additionally, unlike bacteria, whose glucose transport systems are active [15], i.e., require a polarized membrane, and are inhibited by lethal levels of HOCl [13], glucose transport across plasma membranes of yeast such as Saccharomyces cerevisiae is passive [16,17], and may therefore not be particularly susceptible to inhibition by HOCl. In this case, one would expect substrate-level phosphorylation to also be impervious to HOCl in this organism. In the present study, we have explored the nature of oxidative damage caused when S. cerevisiae are exposed to lethal levels of HOCl. Our working hypothesis, based upon analogy to the bactericidal effects of HOCl, was that death would coincide with loss of function within the yeast plasma membrane and involve energy-linked phenomena such as metabolite and ion transport. Correspondingly, one focal point has been the PMA1 Hþ ATPase, a proton pump that constitutes the primary transport system of the plasma membrane in S. cerevisiae, to which all plasma-membrane secondary transport systems are chemiosmotically linked [18]. Loss of PMA1 activity is lethal [19] and would constitute the functional equivalent of loss of activity in the plasma membranelocalized Fo F1 -ATP synthase in bacteria; the latter has been shown in E. coli 25922, Pseudomonas aeruginosa 27853, and Streptococcus lactis 7962 to decline in parallel with cell viability [20]. To our surprise, the pattern of metabolic damage observed in S.cerevisiae does not support this model; although specific lethal lesions have not been identified, it is evident from the data that the microbicidal mechanisms of HOCl in this simple singlecelled eukaryote bear little relationship to those in prokaryotes.
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Materials and methods Isolation and maintenance of cell cultures Saccharomyces cerevisiae was obtained as FleishmannÕs Active Dry Yeast (Regular), which was cultured by streaking suspensions onto agar slants or plates. Cells were generally grown in a 1-L volume containing 3 g NH4 SO4 , 2 g KH2 PO4 , 0.2 g K2 HPO4 0.5 g MgSO4 7H2 O, 0.5 g NaCl, 200 lg FeCl3 6 H2 O, 0.1 g CaCl2 2H2 O, and 3 g yeast extract supplemented with an appropriate carbon source. For fermentative growth, a large amount (50 g/L) of glucose, autoclaved separately, was added to the medium; for aerobic (respiratory) growth, filter-sterilized ethanol was added to a concentration of 10 g/L (12.7 ml/L). A medium of slightly different composition was used for aerobic growth on lactate. This contained 3 g yeast extract, 1 g glucose, 1 g KH2 PO4 , 1 g NH4 Cl, 0.5 g CaCl2 2H2 O, 1 g MgSO4 7H2 O, 0.5 g NaCl, 0.3 ml of a 1% solution of FeCl3 , and 22 ml of 90% lactic acid in 1 L solution; the final pH was adjusted to 5.5 with solid KOH. Cells were first grown by inoculating a small amount of the growth medium with cells from the slants and incubating for 8 h in a shaking water bath at 30 °C, following which a portion of this suspension was used to inoculate one liter of the same medium in a 2 L Erlenmeyer flask. The cells were harvested by centrifugation, washed twice by resuspension in cold phosphate-buffered saline (0.1 M sodium phosphate, 0.15 M NaCl, pH 7.4 (PBS)), and isolated by centrifugation. They were then resuspended in cold buffer to the desired cell density and kept on ice until used. Reactions with hypochlorous acid (HOCl) and chloramines (NH2 Cl) HOCl was purified from commercial sources of NaOCl by vacuum distillation after adjusting the pH to 7–8 with phosphoric acid; NH2 Cl was prepared by reacting HOCl with a 20-fold excess of NH3 [13]. Concentrations were determined by spectrophotometric analysis using e236 (HOCl) ¼ 100 M1 cm1 [21] and e242 (NH2 Cl) ¼ 429 M1 cm1 [22]. In initial experiments, cell suspensions were warmed to 2–30 °C for 5– 10 min and then flow-mixed with an equal volume of oxidant using a Gibson-type 12-jet tangential mixer [23]. When time-dependence studies with oxidant scavengers established that S. cerevisiae killing occurred on the time scale of minutes (see Results), the simpler technique of making bolus additions of HOCl or NH2 Cl to rapidly stirring yeast suspensions was adopted. For routine studies, the cells were exposed to the oxidant for 10– 15 min and then incubated with excess (10–300 lM) sodium thiosulfate or mercaptoethanol for 5–10 min, following which portions were taken to determine
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surviving cells. Within the specified ranges, none of these variables—mixing method, reaction temperature, quencher identity and concentration, or incubation time—had any systematic effect upon the measured parameters, provided that incubation times were sufficiently long to allow complete reaction of HOCl. Viable cells were determined as colony-forming units (cfu) using either spread-plate or pour-plate techniques. Three different plating media were used over the course of these studies, consisting of the rich salts medium described above supplemented with 1–2% glucose (autoclaved separately) and 1.5% Difco technical grade agar, tripticase soy agar (TSA) plus 0.25% glucose, or a mixture of 2% yeast extract, 2% Difco Bacto-peptone, 1% glucose, and 1.5% Difco agar (YPD). Equivalent colonies were formed by glucose-grown or lactate-grown yeast with each of these plating media, although somewhat lower counts (10%) were obtained on TSA for ethanol-grown cells. Potassium ion release by HOCl-treated cells Orion Model 93-19 Kþ ion-specific and Model 90-02 double junction electrodes attached to an Orion Model 701-A digital potentiometer were used to measure potassium ion concentrations in the external medium of S. cerevisiae suspensions. The only deviation from manufacturerÕs recommended setup is that the ‘‘ionic strength adjustor’’ (5 M NaCl) used in the outer chamber of the reference electrode was diluted 50-fold, so that its Cl concentration was the same as that of the suspending PBS medium. The log½Kþ vs. mV response of this electrode assembly was linear in over the experimentally determined range (½Kþ ¼ 105 –102 M). The cells were warmed to ambient temperature and gently stirred for a few minutes to allow the electrode reading to stabilize. Appropriate amounts of HOCl were then added and the voltage changes were recorded over a period of 20 min. External concentrations of Kþ were determined by comparison to calibration curves constructed from standard [Kþ ] solutions at the time of the oxidation experiments. Adenylate phosphorylation levels Intracellular ATP, ADP, and AMP levels were analyzed by rapidly extracting the nucleotides from the cells into ice-cold 0.5 M HClO4 using a spring-loaded syringe, as previously described [20,24,25]. Nucleotide concentrations were determined by isocratic elution with 0.2 M ammonium phosphate, pH 5.6, from a 10 lm reversephase C18 HPLC column [24]. For glucose-grown cells, the suspension was supplemented by adding glucose to a final concentration of 22–270 mM for 10 min prior to extraction with HClO4 . This procedure is essential to obtain an accurate assessment of the metabolic capa-
bilities since viable glucose-grown cells suspended in media devoid of nutrients maintain very low EC values ( 60.1); similar to previous observations on bacteria [25], these EC values ‘‘step up’’ when challenged with a carbon source to levels that accurately reflect the adenine nucleotide-phosphorylating capabilities of the cells. Enzyme assays Activities of the cytosolic enzymes, aldolase (EC 4.1.2.13), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH, EC 1.2.1.12) were determined on supernatant fractions obtained by rupturing the cells by grinding with glass beads. Specifically, to determine aldolase activity, the yeast cells were pelleted, washed, and resuspended in 0.1 M glycylglycine buffer, pH 7.5. Acidwashed glass beads were added and the suspension was spun at 12,000g in a microcentrifuge at 4 °C until microscopic inspection showed that nearly all of the cells were disrupted. The supernatant and washes were collected and the aldolase activity was determined by using a standard colorimetric assay that relies upon formation of a hydrazone (kmax ¼ 240 nm) from reaction of hydrazine with 3-phosphoglyceralde, a product of the enzymatic cleavage of fructose-1,6-diphosphate (FDP) [26]. Measured initial rates of absorbance change at 240 nm were normalized according to the pelleted weights of each sample; activities of the HOCl-treated samples were expressed as percent activity of that obtained for untreated cells. The cells used for GAPDH analyses were resuspended in a medium containing 10 mM sodium pyrophosphate, 10 mM dithiothreitol (DTT), and 0.1 mM diethylenetriaminepentaacetic acid (DTPA), pH 8.5, and broken at 4 °C by spinning 30 s in a ‘‘Bead Beater’’ cell disrupter (Biospec Products, Bartlesville, OK). The beads were removed by gentle suction filtering and washed several times with buffer. The combined extract was centrifuged and activity of the supernatant was determined at 340 nm using a standard spectrophotometric assay that relies upon enzymatic reduction of NADþ by D -glyceraldehyde-3-phosphate (GAP) [27]; initial rates were expressed as percent activity of that obtained for untreated cells. Isolation of yeast plasma membranes and determination of their ATP-hydrolase activities followed, with minor modification, procedures developed by R. Serrano [28,29]. Harvested cells were washed three times in 100 mM phosphate, pH 7.4, treated with HOCl, and resuspended in a medium comprising 28 mM Tris–Cl, 5.5 mM EDTA, and 0.55 mM phenylmethylsulfonyl fluoride (PMSF) at pH 8.5. Three different methods for breaking the cells were evaluated, comprising vortexing in a test tube, shaking in a ‘‘Mini-Bead Beater’’ or vortexing in a ‘‘Bead-Beater,’’ all at 4 °C. The highest total activities and amount of protein recovered were achieved by using the ‘‘Mini-Bead Beater,’’ both of which were
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50–70% higher than the other two methods, which gave comparable total activities and protein yields. Specific activities obtained for all three methods were therefore very nearly equal. However, the low volume capacity (1.4 ml) of the ‘‘Mini-Bead Beater’’ caused the processing time of the samples to be unacceptably long. Vortexing in the Bead-Beater was chosen as the best compromise between recovery and processing time. With this method, total activity and membrane protein yields were independent of grinding time between 20 and 100 s, but beyond 100 s the activity declined progressively with time, presumably caused by inactivation of the ATPase. To determine ATPase activity, the HOCl-treated cells were vortexed for 45 s and the glass beads were removed by filtration and washed with a buffer comprising 20% glycerol and 80% of 0.2 mM EDTA, 0.2 mM DTT, and 10 mM Tris–Cl, pH 7.5. The plasma membrane was isolated by differential centrifugation and the membraneenriched pellet was washed with buffer containing 0.4 mM PMSF to decrease residual phosphate. The primary source of this phosphate appeared to be the disrupted cells themselves since parallel preparations that made use of buffers containing no inorganic phosphate gave equivalent amounts of residual phosphate at this stage of purification. Protein content was determined by the Bradford method using bovine serum albumin as a standard [30]. To determine ATPase activities, portions of the membrane suspended in the PMSF-containing wash buffer were reacted with vanadate-free ATP for 15 min at 30 °C; the inorganic phosphate released was determined as a phosphomolybdate complex [28,29] by comparing the absorbance measured at 750 nm to calibration curves obtained using standard phosphate solutions. Parallel measurements were made with 0.2 mM diethylstilbestrol (DES) added to the reaction medium. DES is a selective inhibitor of the PMA1 Hþ -ATPase; comparison of activities in its presence and absence permits determination of the relative amount of ATP hydrolytic activity attributable to this enzyme. Control measurements using analogous solutions minus ATP and minus membranes were made to correct for inorganic phosphate derived from extraneous sources and turbidity generated by the insoluble membranes, respectively. Glucose transport 14 D -[U- C]glucose
(8 mM, 12.5 mCi/mmol) in 90% ethanol was diluted 10-fold with 44 mM glucose to give a final solution that was 40 mM glucose in 9% ethanol with a specific activity of 1.25 mCi /mmol. Glucose transport rates were determined by following the general procedures described by Bisson and Fraenkel [31]. Specifically, 0.2 ml portions of the diluted radiolabeled sugar solution at 30 °C were added to 0.2 ml suspensions of 5 107 glucose-grown cells/ml that had been exposed
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to varying amounts of HOCl and warmed to 30 °C. These mixtures were briefly incubated for timed intervals, during which the suspension was vigorously vortex-mixed, and the reaction was quenched by adding 10 ml ice-cold water. The cells were then quickly filtered (<5 s) onto Whatman glass microfiber filters (GF/F, pore size 0.7 lm) and washed once with 10 ml cold water. The radioactivity accumulated on the filters was determined by placing them in Ecolite scintillation fluid and counting for 30 min on a Packard Tri-Carb 2700 Liquid Scintillation Analyzer. The counts obtained were corrected for nonspecific adsorption of the sugar onto the cells and filter by adding the radiolabeled sugar directly to the quench water prior to washing the collected cells [31]. In general, this background absorption comprised 10–20% of the total radioactivity measured for samples that had been incubated for 3 min before quenching. Isolation and O2 consumption by yeast mitochondria Mitochondria were isolated from yeast grown on lactate by a preparative sequence that involved hydrolysis of the cell wall with zymolase, homogenization of the resulting spheroplasts, and differential centrifugation. The only deviations from procedures originally described by Schatz and coworkers [32] were that: (1) 1.25 mg zymolase 20,000 per g wet weight of cells was used; (2) zymolase incubations were carried out at 21– 22 °C to minimize breakage of HOCl and NH2 Cl-treated cells; and (3) the final wash of isolated mitochondria was made with the buffer used to measure respiratory activity, i.e., 0.6 M sorbitol, 0.36 mM EDTA, 10 mM KH2 PO4 , 10 mM KCl, 10 mM Tris–Cl, and 0.3% BSA, pH 6.5 [33]. Respiration rates were measured at 25 °C in a thermostatted cell fitted with a YSI Model 5331 Clarktype polarographic electrode connected to a voltage/ time stripchart recorder. Mitochondria were added to the cell and background O2 uptake was measured for a few minutes, following which excess respiratory substrate (6 mM succinate or a-ketoglutarate) and then limiting amounts of ADP were added. The electrode response was calibrated from the pen deflection caused by introducing measured amounts of air into the O2 purged chamber, from which mitochondrial respiration rates were calculated. These were normalized to the amount of protein in each sample, which was determined by the Lowry method [34].
Results Glucose-regulated metabolism in S. cerevisiae To facilitate interpretation of the experimental data, a thumbnail summary of salient features of glucose
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metabolism is included here. Glucose exhibits broad regulatory control over pathways of intermediary metabolism in S. cerevisiae [35]. It represses expression and promotes catabolism of a wide variety of proteins, among which that are relevant to these studies are enzymes involved with mitochondrial respiration, the glyoxalate and TCA cycles, the plasma-membrane Hþ ATPase (PMA1) [36], and alcohol dehydrogenase II (ALDII), an isozyme whose function is primarily associated with oxidation of ethanol to acetaldehyde as a first step in the respiratory metabolism of ethanol (Fig. 1). Glucose also induces expression of numerous enzymes [37], including those associated with the glycolytic pathway, and alcohol dehydrogenase I (ALDI), an isozyme that is primarily involved with reduction of acetaldehyde to ethanol in fermentative metabolism (Fig. 1). Additionally, it regulates a set of genes whose products are hexose transport proteins that are the predominant glucose transporters in S. cerevisiae, the most relevant effect being induction of a low-affinity transporter that is the primary carrier in high-glucose media [16]. Growth on the nonfermentable carbon sources, lactate and ethanol, however, is driven entirely by respiration [35]; for these substrates, syntheses of enzymes associated with respiratory metabolism are derepressed. As a consequence of these regulatory patterns, aerobic growth of S. cerevisiae in batch culture on glucose can be diauxic [36,38]. The first phase, during which most of the glucose present is fermented, is characterized by rapid cell proliferation; upon exhaustion of the glucose, the metabolic changes attending
Fig. 1. Abbreviated intermediary metabolism pathway. Carbon sources used in this study are highlighted in bold type; enzymes that are highly susceptible to inactivation by HOCl are indicated in the scheme. Among others, glucose represses expression of ADHII, the enzymes of the glyoxalate and TCA cycles, the PMA1 Hþ -ATPase, and the proteins of the mitochondrial respiratory chain; among others, glucose induces expression of enzymes of the glycolytic pathway and hexose transporters.
derepression allow the cells to adapt to a slower respirative growth on the accumulated ethanol. Toxicity of HOCl and simple chloramines Saccharomyces cerevisiae exposed to bolus additions of HOCl exhibited sigmoidal dose-dependent survival curves, as measured by their ability to form colonies on nutrient-rich media; comparable toxicities were observed for cells grown on the fermentation carbon source, glucose, and on the respiratory carbon sources, ethanol or lactate. Consistent with these results, toxicities of HOCl were identical within experimental uncertainty for glucose-grown cells harvested in early log, late log, and stationary phases. Under these conditions, the primary metabolic pathway for carbohydrate utilization switches from fermentative to oxidative upon accumulation of ethanol and depletion of glucose from the growth medium [38]. Typical results for cells harvested in early to mid-log phase growth are shown in Fig. 2.
Fig. 2. Survival curves for S. cerevisiae exposed to HOCl. 3.6 107 glucose-grown cells/ml (squares) or 8.0 107 ethanol-grown cells/ml (circles) in PBS were flow-mixed through a 12-jet tangential mixer at 23 °C with an equal volume of HOCl to give the indicated final concentrations. The open symbols indicate results obtained when a stoichiometric excess of S2 O2 3 (640 lM) was present in the receiving flask; the closed symbols indicate the results obtained when S2 O2 3 addition was delayed 10 min. Individual points are averages of 2-4 replicate counts of cfu on rich-salt medium determined using pour-plate techniques.
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From the survival curves, one estimates that 20–40 lM HOCl/107 cells, or (1–2) 109 molecules HOCl/cell are required to inhibit colonial growth by 99% of the cells (LD99 ). Addition of the HOCl scavengers S2 O2 or 3 mercaptoethanol immediately after exposure to lethal concentration levels of HOCl completely protected the cells (Fig. 2). However, protection was incomplete if the subsequent addition of the scavenger was delayed and became less effective as the time interval was increased. The exposure time required for complete inactivation determined from this temporal loss of protection was 1– 2 min. Chloramine (NH2 Cl) exhibited comparable toxicities and killing rates toward S. cerevisiae grown on all three carbon sources. Inactivation of susceptible cytosolic enzymes Yeast aldolase and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) are enzymes of the glycolytic pathway [39] that are highly susceptible to inhibition by HOCl [12,40] and other oxidants [41,42]. As such, they are both convenient markers of oxidative damage by HOCl within the cytosol and potential HOCl target sites for lethal lesions. In Fig. 3, titrimetric loss of the activities of these enzymes upon exposure of glucose-
Fig. 3. HOCl inactivation of S. cerevisiae cytosolic GAPDH and aldolase. For GAPDH determinations, bolus additions of HOCl to the indicated concentrations were made at 23 °C to 1.5 107 glucosegrown cells/ml in PBS. After 10 min, residual oxidant was neutralized with 200 lM S2 O2 3 . Open circles are GAPDH activities/unit volume of suspension; open squares are normalized cfu on YPD determined using pour-plate techniques. The data are a composite of results for several cultures, with each point being an individual determination. At [HOCl] > 100 lM, both viabilities and GAPDH activities were <1% of the control values. For aldolase determinations, 3.7 107 glucosegrown cells/ml in PBS were flow-mixed at 25 °C with an equal volume of HOCl to give the indicated final concentrations; 240 lM S2 O2 3 was added 20 min after mixing. Solid circles are the aldolase activities; solid squares are cfu on rich-salt medium determined with pour-plate techniques. The aldolase data have been normalized to 1.5 107 cells/ ml to allow direct comparison to the GAPDH data.
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grown S. cerevisiae to increasing amounts of HOCl is compared to the corresponding viabilities determined for the same samples. The data indicate that 65–75% of the original activity of these enzymes is retained in the nonviable cells and that twice the amount of HOCl is required to inactivate these enzymes as is required to prevent cell division. Thus, although some damage is incurred, these enzymes do not appear to be primary target sites associated with loss of replicative capacity. Mitochondrial function The functional capabilities of mitochondria isolated from cells exposed to HOCl were assessed by measuring O2 uptake under various conditions of respiratory control [15,33]. State 3 (or uncoupled) respiration was induced by adding excess ADP or the protonophore, carbonyl cyanide m-chlorophenyl hydrazone (cccp), to mitochondrial suspensions containing the respiratory reductants, succinate or a-ketoglutarate. Under these conditions, respiratory rates are maximal and can be
Fig. 4. Functional capabilities of isolated S. cerevisiae mitochondria. 5–6 107 lactate-grown cells/ml in PBS were flow-mixed at 23 °C with equal amounts of HOCl to the indicated final concentrations and mitochondria were isolated and characterized using succinate as respiratory substrate as described in Materials and methods. Squares are cfu, determined on TSA using pour-plate methods; open circles are RCR; crossed circles are RCR measured in the presence of cccp, down triangles are the state 3 (uncoupled) respiration rates; and up triangles are the P/O ratio. All values are normalized to constant protein content of the samples and to the values obtained with control mitochondria.
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directly related to the extent of damage suffered by the respiratory chain. State 4 (or coupled) respiration is measured under conditions where the reducing substrate is in excess and the proton motive force across the mitochondrial cristae membrane is maximal, chemiosmotically restricting electron transport. The ratio of state 3 to state 4 respiration rate (respiratory control ratio (RCR)) indicates the efficiency of this chemiosmotic coupling; as such, the RCR provides information on the impermeability of the cristae membrane to the proton and other ions. The amount of O2 consumed during ADP-induced state 3 respiration (expressed as the P/O ratio, i.e., [ADP] added/[O2 ] consumed) provides a measure of the efficiency of coupling of the electron transport chain to ATP synthesis. Inhibition of the Fo F1 -ATP synthase, loss of membrane integrity, and inhibition of electron transport can all lead to reduced P/O ratios. These parameters, obtained with succinate as respiratory substrate, are compared to viabilities of lactate-grown S. cerevisiae in Fig. 4. Addition of an amount of HOCl that is sufficient to inhibit growth in 95% of the cells decreases the state 3 respiration rate to 50% of its normal value, with smaller decreases appearing in the RCR and P/O ratio. This reactivity pattern indicates that mitochondrial damage occurs primarily within the electron transport chain and that the extent of this damage may not be sufficient to significantly inhibit ATP generation by nonviable cells. Nearly identical patterns were observed when a-ketoglutarate was used as respiratory substrate and when NH2 Cl was used as oxidant. Absolute values of RCR and P/O for mitochondria isolated from untreated cells (2.5–2.7 and 1.7–1.9, respectively, with succinate as substrate and 5.1 and 2.5, respectively, with a-ketoglutarate as substrate) were nearly identical to originally reported values [33].
manner that paralleled inhibition of mitochondrial function (Fig. 4). For example, 50% reduction in phosphorylation levels of 6 107 lactate-grown or ethanolgrown cells/ml required exposure to 150 lM HOCl or 100 lM HOCl, respectively, as determined from EC vs. [HOCl] plots analogous to Fig. 3, whereas LD50 of the same suspensions was 55 lM HOCl. In contrast, glucose-grown cells harvested in mid-log phase showed EC values whose decline paralleled loss of viability. When harvested in late-log phase, the EC decline lagged behind the survival curve, and when harvested in early stationary phase, the decline in EC was very similar to that recorded for cells grown on the respiratory substrates (Fig. 5). In the experiments with glucose-grown cells, all samples were incubated with glucose following exposure to HOCl prior to extraction of the nucleotides for EC determinations. The phase-dependent changes in dose–response behavior of EC for glucose-grown cells (Fig. 5) parallel the shift in carbohydrate metabolism
Adenine nucleotide phosphorylation levels The adenylate energy charge (EC), defined as EC ¼ ([ATP] + 1/2[ADP])/([ATP] + [ADP] + [AMP]) [43], is a convenient index of the overall level of phosphorylation of the adenine nucleotide pool within the cytosol of microbial cells. Rapidly growing cells maintain EC P 0:8, whereas this value is considerably less in resting or nonviable cells [10,11,24,44]. The EC of oxidatively damaged cells will decrease if either ATP biosynthesis is impaired or consumption becomes excessive, particularly when futile cycles are established in ATPutilizing systems required to maintain homeostasis. Changes in EC within S. cerevisiae upon exposure to HOCl depended upon the carbon nutrient source and, for glucose-grown cells, upon the growth phase at the time they were isolated. EC values were normal at lethal levels of HOCl for cells grown on respiratory carbon sources, but declined at higher concentrations in a
Fig. 5. Adenylate energy charge (EC) in S. cerevisiae grown on glucose. EC values for 2.4 107 cells/ml harvested in the mid-log region of the first growth phase (open circles), 4.8 107 cells/ml harvested at the transition between growth phases (up triangles), and 3.8 107 cells/ml harvested in the second growth phase (diamonds) flow-mixed in PBS at 28 °C with an equal volume of HOCl to the indicated final concentrations. Following reaction, the cell suspensions were treated with 0.3 mM S2 O2 3 to remove any residual oxidant, incubated 20 min in 22 mM glucose, and extracted with HClO4 . Cfu (open squares) were determined on TSA using pour-plate techniques. All data have been normalized to a constant cell density of 4.3 107 cells/ml.
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from fermentative to respiratory upon depletion of glucose from the growth medium [38,39]. Insensitivity of glucose transport to lethal doses of HOCl Exposure of fermenting S. cerevisiae to 14 C-glucose resulted in rapid net accumulation of radioactivity within the cells; as previously reported [31], uptake rates were linear for 3 min and then gradually declined with increasing incubation time (data not shown). Glucose uptake was remarkably insensitive to high doses of HOCl. The initial rate of cells exposed to a LD99 dose of HOCl was two-thirds of the accumulation rate of untreated cells, as judged from the initial slopes of scintillation counts vs. incubation time, and 50% inhibition of 2.5 107 cells/ml required exposure to 470 lM HOCl in suspensions whose LD50 was 150 lM. Under the prevailing growth conditions, glucose is taken up by facilitated diffusion via an integral membrane protein that is thought to form an aqueous transmembrane channel allowing passage of the sugar across the bilayer [16,17]. As such, glucose transport is not energy-linked to membrane polarization, but is driven by enzymatic phosphorylation within the cytosol. Equilibration of intracellular glucose with the external solution takes only 2–3 min under the conditions of our experiments, as has been demonstrated by using nonmetabolizable substrate analogs and mutant cells devoid of hexokinase and glucokinase activities [31]. Consequently, the continued accumulation of radioactivity beyond 3 min incubation indicates that, in addition to glucose transport per se, glucose phosphorylation capabilities within the cell had not been severely damaged by exposure to lethal levels of HOCl.
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the solid symbols; for these data, the HOCl concentration reported on the abscissa is also 20-fold greater than indicated by the abscissa values. As we have repeatedly observed in earlier studies with bacteria [13,20,24], the amount of HOCl required for loss of function scales linearly with the density of cells in suspension. Thus, the results obtained are not dependent upon either the transient HOCl concentration generated upon bolus addition of the oxidant or the number of reactive sites presented by the yeast. Potassium ion leakage Yeast cells actively accumulate Kþ via voltage-gated channels in their plasma membranes in response to transmembrane potentials generated by the PMA1
Inactivation of the PMA1 Hþ -ATPase Comparisons of cfu with loss of ATP-hydrolytic activity in plasma membrane fractions isolated from glucose-grown cells exposed to varying concentrations of HOCl are shown in Fig. 6. These values have been corrected for ATP hydrolysis from alkaline phosphatases, apyrases, and other enzymes present in the membrane fractions by performing control experiments using diethylstilbesterol to block the PMA1 Hþ -ATPase activity. In the presence of 0.2 mM DES, the specific activity was typically 8% of the value of the untreated membrane fraction in the absence of the inhibitor, indicating that 92% of the total measured activity could be ascribed to the PMA1 Hþ -ATPase. As with the other activities that were measured, inactivation of the PMA1 lagged behind loss of colony-forming capabilities, so that three times more HOCl was required to reduce the activity of the proton pump by 50% than to completely inhibit growth. Additional studies using 20-fold greater S. cerevisiae cell densities are also reported in Fig. 6 as
Fig. 6. PMA1 Hþ -ATPase activity in isolated plasma membranes. Bolus additions of 80 mM HOCl were made at 30 °C to the final indicated concentrations to 5 107 (open symbols) or 1 109 (crosshatched symbols) glucose-grown cells/ml in 100 mM phosphate, pH 7.4. Cfu were determined by pour-plate analysis using YPD; each point represents an average of counts from five plates whose average deviation from the mean was typically within 10%. ATP-hydrolase specific activities are plotted relative to values obtained for plasma membranes from untreated control cells (0.22 0.05 lM Pi /min-mg protein). Each point is the average of at least three enzymatic determinations minus the activity found in the presence of DES; average deviations from the mean were typically less than 5%. As discussed in the text, concentrations of HOCl used for the higher cell densities (cross-hatched symbols) are 20 greater than indicated on the abscissa.
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Hþ -ATPase [18,45,46]. Oxidative injury that leads to depolarization or rupture of these membranes will be accompanied by Kþ release to the external medium. In ethanol-grown and lactate-grown cells, the rate of Kþ efflux increased in a dose-dependent manner following addition of HOCl to the medium. This slow release was nearly linear with time and underwent only very small diminution in rate when the S2 O2 3 was added 10 min later, indicating that Kþ release was not caused by ongoing oxidative damage, but was a consequence of prior irreversible damage to the cell. Initial rates of Kþ efflux measured at various levels of exposure to HOCl are plotted for the two growth media in Fig. 7. Complete release of potassium ion gave ½Kþ ¼ 0:5–0:8 mM in the medium, as determined by treating the cells with a large excess of HOCl. Saccharomyces cerevisiae grown to early-log phase on glucose exhibited a different response to added HOCl. In this case, bolus additions of up to 100 lM HOCl to 2 107 cells/ml did not increase the Kþ leak rates above a measured basal level of 1.8 0.3 lM/min (cf., Fig. 7). However, background Kþ levels in the cell suspensions increased (e.g., from 130 to 250 lM) in a dose-dependent manner following this treatment, consistent with rapid release of Kþ from a small subpopulation of the cells.
Fig. 7. Initial rates of Kþ release from respiring S. cerevisiae. Initial leak rates in PBS at 22 °C. For lactate-grown cells (at 2.1 107 cells/ ml), cfu for additions of 30, 50, 70, 80, and 100 lM HOCl were 124, 35, 16, 13, and 8%, respectively, of the control value; for ethanol-grown cells (at 3.2–3.6 107 cells/ml), cfu were 56 and 0% of controls.
Discussion Comparison of HOCl-inflicted damage in S. cerevisiae to that in bacteria One major difference in the patterns of oxidative damage caused by HOCl to bacteria and S. cerevisiae is that, in bacteria, phosphorylation levels in the adenine nucleotide pool decline in parallel with survival [9,20], whereas in S. cerevisiae grown on respiratory carbon sources phosphorylation levels in nonviable cells are nearly normal. This same behavior is observed in the late stages of growth on glucose where ethanol has become the nutrient carbon source and metabolism is respiratory (Fig. 5). Consistent with this result, mitochondria isolated from respiring cells exposed to lethal levels of HOCl retain a major fraction (60%) of their capacity for oxidative phosphorylation, with most of the damage occurring to the electron transport chain, as measured by state 3 respiration rates (Fig. 4). Other indicators of oxidative damage within the cytosol are the enzymes aldolase and GAPDH [12,40–42]. Approximately two-thirds of the normal intracellular activity of these enzymes is retained in HOCl-inactivated cells (Fig. 3). These experiments do not distinguish between oxidative inactivation of the proteins and their loss from the cytosol, i.e., by breaching of the yeast plasma membrane. However, the slow progressive increase in Kþ leak rates accompanying exposure of yeast to increasing amounts of HOCl (Fig. 7) indicates that the cellular plasma membrane remains ion-impermeable, i.e., intact, in most of the cells. Rupture of the membrane would give an immediate and abrupt increase in the extracellular Kþ concentration. Lactate dehydrogenase (LDH) and ALD are even more susceptible to inhibition by HOCl, at least in homogeneous solution [12]. Nonetheless, from the high EC values measured in HOCl-inactivated cells (Fig. 5), one can infer that the activities of all respiratory pathway enzymes, including LDH and ALDII, are sufficient to support near-normal respiratory capacity with lactate and ethanol as carbon sources (Fig. 1). In contrast to the results with respiring cells, EC values in fermenting S. cerevisiae declined in a manner that paralleled loss in cfu (Fig. 5). Because these were step-up experiments, the EC values indicate the capacity of oxidized cells to undertake substrate-level phosphorylation [25]. Obviously, loss of this capability could arise if the glycolytic pathway were inhibited by HOCl. However, the high residual activities of aldolase and GAPDH, two enzymes of the glycolytic pathway that are highly susceptible to HOCl inactivation [12], as well as the apparent limited overall damage to other vulnerable cytosolic proteins caused by lethal amounts of HOCl, render this unlikely. The alternative possibility that glucose transport across the plasma membrane was
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inhibited can also be dismissed on the basis of the insensitivity of glucose uptake by the cells to exposure to HOCl. A second major difference between these cell types is that, unlike bacteria, the plasma membrane-localized ATPase is remarkably resistant to inactivation by HOCl. Specifically, the activity of the bacterial Fo F1 -ATP synthase is lost in parallel with cfu [20], whereas significant loss of activity in the yeast PMA1 Hþ -ATPase requires addition of HOCl in amounts several-fold in excess of that required for inhibition of growth (Fig. 6). PMA1 is a plausible target for inactivation because gene-knockout experiments have established that its function is critical to cell survival [19] and the protein contains nine cysteine groups, the thiol substituents of which are vulnerable to HOC1 oxidation [12,40,47–49]. However, this and other P-type ATPases do not appear to be very sensitive to chemical modification or replacement of their cysteine residues [50]. The e-amino groups of conserved lysine residues represent possible alternative sites of attack, particularly since several reactive essential lysine groups have been identified within the ATP-binding domain of the yeast Hþ -ATPase [51,52]. Based upon results with other oxidants [53,54], N-chlorination at these sites could lead to inactivation or loss of function attending further degradation. In any event, inactivation of the yeast proton pump in these studies required exposure to a considerable excess of HOCl over that required for inhibition of growth. A third major difference is that the time scales for loss of colony-forming capabilities are dramatically different. Whereas bacteria are killed ‘‘instantaneously’’ upon exposure to lethal levels of HOCl [14], inactivation of both fermenting and respiring yeast required incubation with the toxin for several minutes (Fig. 2), indicating that the target sites in yeast are either relatively unreactive or otherwise physically protected from the external medium, e.g., as would be the case if access to the vulnerable sites required diffusion across the yeast cell membrane.
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media. In support of this viewpoint, we have previously shown that the phosphorylating capabilities of bacteria measured immediately after exposure to either bolus additions of HOCl or MPO-generated oxidant correlate directly with cfu measured 24 h later [25]. Thus, the cfu count accurately reflects the immediate HOCl-induced loss of energy-transducing capabilities in the microbial population. One consequence of this conclusion is that, although the lethal lesions have not yet been identified, death in fermenting S. cerevisiae must arise directly from HOCl inactivation of critical cellular functions. These issues are not so straightforward for respiring S. cerevisiae. Specifically, the functional capabilities of the yeast mitochondria are largely unimpaired (Fig. 4) and the adenylate phosphorylation levels in respiring cells are near-normal (Fig. 5), despite the loss in ability to undergo cell division. Thus, for yeast, it is quite possible that metabolic transformations could occur via biosynthesis during incubation of the plated cells. Moreover, there appears to be no correlation between overt loss of essential cellular function and death or localization of lethal lesions to the plasma membrane. Rather, there appears to be a global loss of function among HOCl-susceptible biomolecules located throughout the cell. In all cases, however, loss of function requires significantly greater amounts of HOCl than does inhibition of cell growth. As with other cellular oxidants [55–57], exposure to increasing amounts of HOCl causes the response of endothelial cells to vary progressively from transient inhibition of growth to apoptosis to necrosis (lysis) [58]. Programmed cell death has recently been discovered in yeast [59,60]; one of the effectors that can trigger apoptosis in these cells is oxidative stress, leading to mitochondrial damage, and/or loss of endogenous antioxidant levels. Consequently, one likely explanation for the loss of colony-forming capabilities prior to significant loss of discrete biological functions is that the oxidative damage inflicted triggers the biochemical cascades comprising S. cerevisiae apoptosis, which then ensues upon exposing the cells to nutrient agar.
Microbicidal mechanisms Biological relevance The ability to sustain colonial growth as a definition of viability in studies that attempt to correlate metabolic dysfunctions with death has been questioned primarily because there most often exists a long time interval between the measurement of specific functions, enzymatic activities or metabolite concentrations, and the measurement of colony-forming units (cfu), during which time additional damage or repair might occur that could alter the relationships. For bacteria and fermenting S. cerevisiae, at least, the inability to generate ATP and maintain adenylate phosphorylation levels ensures that biosynthesis and maintenance of cellular homeostasis will not occur, even for cells plated on nutrient-rich
Based upon survival curves obtained from bolus additions of oxidant to cell suspensions, the amount of HOCl required to inhibit cell division in S. cerevisiae is 1.5 109 HOCl/cell (Fig. 1), which is 10 times greater than that required to inhibit growth of representative laboratory strains of E. coli, S. lactis, and P. aeruginosa [20]. Recent estimates place a lower limit of myeloperoxidase-generated HOCl in the respiratory burst at 70% of the O2 consumed [61]. On average, each human neutrophil consumes as much as 4 1010 O2 molecules following respiratory stimulation [8]. Consequently, each neutrophil has, in principle, the
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capacity to kill maximally 20 S. cerevisiae using only its metabolically generated HOCl. This estimate is clearly an upper limit, however, since some of the oxidants will certainly be directed against other biological targets, including antioxidants in the serum-derived phagosomal fluid [62], and opsonins [8] and neutrophilderived protein [6,7]. Hypochlorous acid is perhaps the most toxic of the potential phagocyte-generated oxidants [62]; consequently, the most important conclusion to be drawn from these calculations may be that neutrophils cannot generate large excesses of oxidants beyond that required to inactivate phagocytosed microbes.
Acknowledgments The authors are grateful to Sirisha Dontireddy for developing protocols for isolation and functional characterization of the PMA1 Hþ -ATPase and to Professor Raymond Reeves for helpful discussions concerning yeast apoptosis. This work was supported by National Institutes of Health Grant AI15834.
[19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31] [32] [33] [34] [35]
References [1] S.J. Klebanoff, in: J.I. Gallin, I.M. Goldstein, R. Snyderman (Eds.), Inflammation: Basic Principles and Chemical Correlates, Raven Press, New York, 1998, pp. 391–444. [2] M.B. Hampton, A.J. Kettle, C.C. Winterbourn, Infect. Immun. 64 (1996) 3512–3517. [3] J.K. Hurst, in: J. Everse, K.E. Everse, M.B. Grisham (Eds.), Peroxidases in Chemistry and Biology, vol. 1, CRC Press, Boca Raton, FL, 1991, pp. 37–62. [4] H.B. Dunford, Heme Peroxidases, Wiley-VCH, New York, 1999. [5] P.G. Furtmuller, U. Burner, W. Jantschko, G. Regelsberger, C. Obdinger, Redox Rep. 5 (2000) 173–178. [6] A.L.P. Chapman, M.B. Hampton, R. Senthilmohan, C.C. Winterbourn, A.J. Kettle, J. Biol. Chem. 277 (2002) 9757–9762. [7] H. Rosen, J.R. Crowley, J.W. Heinecke, J. Biol. Chem. 277 (2002) 30463–30468. [8] Q. Jiang, D.A. Griffin, D.F. Barofsky, J.K. Hurst, Chem. Res. Toxicol. 10 (1997) 1080–1089. [9] J.K. Hurst, S.V. Lymar, Acc. Chem. Res. 32 (1999) 520–528. [10] C.J. Knowles, Symp. Soc. Gen. Microbiol. 27 (1977) 241–283. [11] A.G. Chapman, D.E. Atkinson, Adv. Microb. Physiol. 15 (1977) 253–306. [12] J.M. Albrich, C.A. McCarthy, J.K. Hurst, Proc. Natl. Acad. Sci. USA 78 (1981) 210–214. [13] J.M. Albrich, J.H. Gilbaugh III, K.B. Callahan, J.K. Hurst, J. Clin. Invest. 78 (1986) 177–184. [14] J.M. Albrich, J.K. Hurst, FEBS Lett. 144 (1982) 157–161. [15] D.G. Nicholls, S.J. Ferguson, Bioenergetics3, Academic Press, Amsterdam, 2002. [16] J. Hor ak, Biochim. Biophys. Acta 1331 (1997) 41–79. [17] M. Mueckler, C. Caruso, S.A. Baldwin, M. Panico, I. Blensch, H.R. Morris, W.J. Allard, G.E. Lienhard, H.F. Lodish, Science 229 (1985) 941–945. [18] R. Serrano, in: J.R. Broach, J. Pringle, E.W. Jones (Eds.), The Molecular and Cellular Biology of the Yeast Saccharomyces: Genome Dynamics, Protein Synthesis, and Energetics, vol. I, Cold
[36] [37] [38] [39]
[40] [41]
[42] [43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54]
Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1991, pp. 523–585. R. Serrano, M.C. Kielland-Brandt, G.R. Fink, Nature 319 (1986) 689–693. W.C. Barrette Jr., D.M. Hannum, W.D. Wheeler, J.K. Hurst, Biochemistry 28 (1989) 9172–9178. J.C. Morris, J. Phys. Chem. 70 (1966) 3798–3805. E.L. Thomas, M.B. Grisham, M.M. Jefferson, Methods Enzymol. 132 (1986) 569–585. Q.H. Gibson, L. Milnes, Biochem. J. 91 (1964) 161–171. W.C. Barrette Jr., J.M. Albrich, J.K. Hurst, Infect. Immun. 55 (1987) 2518–2525. W.C. Barrette Jr., D.M. Hannum, W.D. Wheeler, J.K. Hurst, J. Bacteriol. 170 (1988) 3655–3659. O.C. Richards, W.J. Rutter, J. Biol. Chem. 236 (1961) 3177–3181. V. Worthington (Ed.), Worthington Enzyme Manual, Worthington Biochemical Corp, Lakewood, NJ, 1993, pp. 201–206. R. Serrano, Methods Enzymol. 157 (1988) 533. R. Serrano, Biochem. Biophys. Acta 947 (1988) 1–28. M.M. Bradford, Anal. Biochem. 72 (1976) 248–254. L.F. Bisson, D.G. Fraenkel, Proc. Natl. Acad. Sci. USA 80 (1983) 1730–1734. G. Daum, P. Botini, G. Schatz, J. Biol. Chem. 257 (1982) 13028– 13033. T. Ohnishi, K. Kawaguchi, B. Hagihara, J. Biol. Chem. 241 (1966) 1797–1806. O.H. Lowry, N.J. Rosebrough, A.L. Farr, R.J. Randall, J. Biol. Chem. 193 (1951) 265–275. M. Johnston, M. Carlson, in: J.R. Broach, J.R. Pringle, E.W. Jones (Eds.), The Molecular and Cellular Biology of the Yeast Saccharomyces: Gene Expression, vol. II, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1992, pp. 193– 281. A.R. Fernandes, I. Sa-Correia, Yeast 20 (2003) 207–219. M.B. Grisham, M.M. Jefferson, D.F. Melton, E.L. Thomas, J. Biol. Chem. 259 (1984) 10404–10413. O. K€appeli, Adv. Microb. Physiol. 28 (1986) 181–209. D.G. Fraenkel, in: J.N. Strathern, E.W. Jones, J.R. Broach (Eds.), The Molecular Biology of the Yeast Saccharomyces: Metabolism and Gene Expression, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1982, pp. 1–37. J.M. Pullar, C.C. Winterbourn, M.M. Vissers, Am. J. Physiol. 277 (1999) H1505–H1512. P.A. Hyslop, D.B. Hinshaw, W.A. Halsey Jr., I.U. Schraufst€atter, R.D. Sauerheber, R.G. Spragg, J.H. Jackson, C.G. Cochrane, J. Biol. Chem. 263 (1988) 1665–1675. D.P. Buchczyk, K. Briviba, F.U. Hartl, H. Sies, Biol. Chem. 381 (2000) 121–126. D.E. Atkinson, Biochemistry 7 (1968) 4030–4034. W.J. Ball Jr., D.E. Atkinson, J. Bacteriol. 121 (1975) 975–982. A. Ambesi, M. Miranda, V.V. Petrov, C.W. Slayman, J. Exp. Biol. 203 (2000) 155–160. A. Rodriguez-Navarro, Biochim. Biophys. Acta 1469 (2000) 1–30. M.C.M. Vissers, A. Stern, F. Kuypers, J. van den Berg, C.C. Winterbourn, Free Radic. Biol. Med. 16 (1994) 703–712. E.L. Thomas, Infect. Immun. 25 (1979) 110–116. R. Hurst, Y. Bao, S. Ridley, G. Williamson, Biochem. J. 338 (1999) 723–728. V.V. Petrov, C.W. Slayman, J. Biol. Chem. 270 (1995) 28535– 28540. J.P. Pardo, C.W. Slayman, J. Biol. Chem. 263 (1988) 18664– 18668. R. Rao, C.W. Slayman, J. Biol. Chem. 268 (1993) 6708–6713. A. Valiakhmetov, D.S. Perlin, J. Biol. Chem. 278 (2003) 6330– 6336. N. Stadler, M. H€ ofer, K. Sigler, Free Radic. Res. 35 (2001) 643– 653.
D.A. King et al. / Archives of Biochemistry and Biophysics 423 (2004) 170–181 [55] A.G. Wiese, R.E. Pacifici, K.J.A. Davies, Arch. Biochem. Biophys. 318 (1995) 231–240. [56] K.-T. Lin, J.-Y. Xue, M.C. Lin, E.G. Spokas, F.F. Sun, P.Y.K. Wong, Am. J. Physiol. 274 (1998) C855–C860. [57] R.H. Burdon, V. Gill, D. Alliangana, Free Radic. Res. 24 (1996) 81–93. [58] M.C.M. Vissers, J.M. Pullar, M.B. Hampton, Biochem. J. 344 (1999) 443–449.
181
[59] F. Madeo, S. Engelhardt, E. Herker, N. Lehmann, C. Maldener, A. Proksch, S. Wissig, K.-U. Fr€ ohlich, Curr. Genet. 41 (2002) 208–216. [60] M. Priault, N. Camougrand, K.W. Kinnally, F.M. Vallette, S. Manon, FEMS Yeast Res. 4 (2003) 15–27. [61] A.J. Kettle, C.C. Winterbourn, Redox Rep. 3 (1997) 3–15. [62] S.V. Lymar, J.K. Hurst, Chem. Res. Toxicol. 8 (1995) 833–840.