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Extensive dark biological production of reactive oxygen species in brackish and freshwater ponds Tong Zhang, Colleen Michelle Hansel, Bettina M. Voelker, and Carl H. Lamborg Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.5b03906 • Publication Date (Web): 08 Feb 2016 Downloaded from http://pubs.acs.org on February 14, 2016

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Environmental Science & Technology

Extensive dark biological production of reactive oxygen species in brackish and freshwater ponds Tong Zhanga, Colleen M. Hansela,*, Bettina M. Voelkerb, Carl H. Lamborgc

a

Department of Marine Chemistry and Geochemistry, Woods Hole Oceanographic Institution,

Woods Hole, MA 02543, USA b

Department of Chemistry and Geochemistry, Colorado School of Mines, Golden, CO 80401,

USA c

Department of Ocean Sciences, University of California-Santa Cruz, Santa Cruz, CA 95064,

USA *Corresponding Author: [email protected], phone: (508) 289-3738

1

ACS Paragon Plus Environment


1

Abstract

2

Within natural waters, photo-dependent processes are generally considered the

3

predominant source of reactive oxygen species (ROS), a suite of biogeochemically important

4

molecules. However, recent discoveries of dark particle-associated ROS production in aquatic

5

environments and extracellular ROS production by various microorganisms point to biological

6

activity as a significant source of ROS in the absence of light. Thus, the objective of this study

7

was to explore the occurrence of dark biological production of the ROS superoxide (O2-) and

8

hydrogen peroxide (H2O2) in brackish and freshwater ponds. Here we show that the ROS

9

superoxide (O2-) and hydrogen peroxide (H2O2) were present in dark waters at comparable

10

concentrations as in sunlit waters. This suggests that, at least for the short-lived superoxide

11

species, light-independent processes were an important control on ROS levels in these natural

12

waters. Indeed, we demonstrated that dark biological production of ROS extensively occurred in

13

brackish and freshwater environments, with greater dark ROS production rates generally

14

observed in the aphotic relative to the photic zone. Filtering and formaldehyde inhibition

15

confirmed the biological nature of a majority of this dark ROS production, which likely involved

16

phytoplankton, particle-associated heterotrophic bacteria, and NADH-oxidizing enzymes. We

17

conclude that biological ROS production is widespread, including regions devoid of light,

18

thereby expanding the relevance of these reactive molecules to all regions of our oxygenated

19

global habit.

20 21 22 23

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Introduction

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Reactive oxygen species (ROS), such as superoxide (O2-) and hydrogen peroxide (H2O2)

27

are intermediates formed during the sequential one-electron reduction of O2 to H2O. ROS are of

28

considerable interest due to their extremely high reactivity in mediating redox transformations in

29

the environment. For example, O2- has the ability to either reduce or oxidize biologically and

30

geochemically important metals, such as iron 1, 2, copper 2, 3, and manganese 4-6, thus impacting

31

their bioavailability and environmental behavior. H2O2, the longest-lived ROS, reacts rapidly

32

with reduced iron via the Fenton reaction to generate the highly reactive ROS, hydroxyl radical

33

(HO▪). In fact, within natural waters, the Fenton reaction is a main source of hydroxyl radical and

34

other strong oxidants, which can break down otherwise recalcitrant organic compounds 7-9. In natural aquatic environments, the dominant source of ROS has generally been assumed

35 36

to be abiotic photochemical production induced by sunlight irradiation in surface water.

37

Specifically, photo-oxidation of "chromophoric dissolved organic matter" (CDOM), the light-

38

absorbing components of natural organic matter, generates O2-, which further converts to H2O2

39

during dismutation 10-14. However, research in the past decade has discovered microorganisms

40

that are capable of extracellular ROS production, including fungi 15, 16, phytoplankton 17-20 and,

41

most recently, heterotrophic bacteria 5, 21-23 as potentially significant sources of ROS in natural

42

waters. Although a handful of field studies have demonstrated that dark production of O2- and

43

H2O2 occurred at comparable rates as photochemical production in natural waters 17, 24-28, direct

44

evidence that linked the in situ dark ROS production to a biological origin as implicated in

45

laboratory cultures or that demonstrated the biochemical pathway for this production were not

46

provided in these studies.

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47

Previous field measurements of ROS production have mostly concentrated on H2O2 in

48

marine systems14, 29-36. Few previous studies have directly examined O2- and H2O2 production

49

together 13, 25, 26, 37. In addition, no direct studies of O2- production and concentration have been

50

performed in brackish and freshwater environments, where ROS production and decay would

51

presumably be more dynamic due to the elevated abundance and variability of both ROS

52

producers (e.g., CDOM, microorganisms) and scavengers (e.g., metals).

53

Therefore, the main objectives of this study were to (1) explore the occurrence of O2- and

54

H2O2 in brackish and freshwater ponds within both the photic and aphotic zones (2) quantify the

55

rates of particle-associated light-independent production of O2- and H2O2 within these waters, (3)

56

determine the contribution of biological processes to the light-independent ROS production, and

57

(4) investigate the pathways through which biological ROS production occurred. To meet these

58

objectives, a combination of in situ field measurements and laboratory incubations were

59

conducted with water from brackish and freshwater ponds on Cape Cod (Massachusetts, U.S.A.)

60

during the summers of 2013 and 2014.

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Experimental Methods

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Sample collection. Unfiltered natural water samples were collected from two field sites

64

on Cape Cod, MA. Site 1 (41.54º N, 70.64º W) is located in the south basin of Oyster Pond and

65

was sampled in July and August of 2014. Oyster Pond is a brackish coastal kettle pond and is

66

separated from the ocean by a barrier beach. The site of sampling was in the southern basin with

67

a depth of approximately 6 m. Site 2 (41.56º N, 70.61º W) is located in the middle of a

68

freshwater pond, Weeks Pond, and was sampled in June and August of 2013. To avoid the

69

impact of H2O2 in rainwater 29, 38, all field sampling occurred after at least 24 rainless hours.

70

Triplicate or quadruplicate samples were collected in 1 L high-density polyethylene (HDPE) 4


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bottles. Additional unfiltered water was collected from the aphotic zone of Oyster Pond in 20-L

72

lightproof polyethylene carboys for the lab incubation experiments. Surface water was collected

73

by filling the sample bottles by hand, while water from deeper area of the ponds was collected

74

using a peristaltic pump and opaque flexible tubing. Bacterial, algal and filamentous

75

cyanobacterial cells were counted in surface water that was collected via peristaltic pump or

76

directly by filling the bottles by hand. A statistically significant difference was not observed

77

between these two sampling methods, indicating that pumping did not cause significant damage

78

to the microbial communities in our samples. In addition, unfiltered seawater was collected from

79

the aphotic zone (30 m below surface) in Nantucket Sound (41º 32.1’N, 70º 24.0’ W) in October,

80

2014. Seawater was collected in 20-L GoFlo bottles attached to an Amsteel line (a non-metallic

81

dyneema core rope) and dispensed into a 20-L polyethylene carboy that was covered with two

82

layers of black plastic bags. After field collection, water samples were transported on ice, in the

83

dark to the laboratory at Woods Hole Oceanographic Institution (WHOI). All sample containers

84

and equipment were acid cleaned before sampling, and “clean hands/dirty hands” techniques

85

were employed during sampling.

86 87

ROS concentrations in natural waters. O2- and H2O2 concentrations were obtained in

88

pond waters. As O2- was expected to have a short-lifetime (minutes), O2- concentrations within

89

the ponds were measured in the field. H2O2 was expected to have a significantly greater lifetime

90

(hours), and thus water samples for H2O2 concentrations were collected following the in situ O2-

91

measurements, transported back to the laboratory on ice, and analyzed within an hour. O2- concentrations. O2- concentrations in natural water samples were measured in situ by

92 93

pumping unfiltered water using a high flow rate (1 L min-1) peristaltic pump into a (clear for

5



94

surface water and opaque otherwise) sample bottle that was then immediately pumped at a

95

slower flow rate (6 mL min-1) using a high accuracy peristaltic pump directly into a flow-through

96

FeLume Mini system (Waterville Analytical, Waterville ME) positioned onboard a boat. O2-

97

detection was based on the reaction between O2- and a chemiluminescent probe, a methyl

98

cypridina luciferin analog (MCLA, Santa Cruz Biotechnology). This reagent has been widely

99

used to detect O2- in natural waters 17, 25, 26, 37, 39 as well as in microbial cultures 19, 21. MCLA

100

reagent and unfiltered natural water were independently pumped through opaque tubing, and

101

mixed in a transparent spiral flow cell. The resulting chemiluminescence signal was detected by

102

a photon-counting photomultiplier tube that was positioned immediately above the flow cell. The

103

travel time of the water samples in the opaque FeLume tubing was approximately 10 sec – in

104

total, the travel time from the water depth to the detector was less than 30 seconds. Filtered

105

natural water collected previously from the same field site and water depth and aged in the dark

106

for at least 12 h was utilized as the “blank” to establish the baseline. Procedures for baseline

107

correction are described in the Supporting Information. Following the baseline signal (~2-3 min),

108

the tubing was transferred to the opaque bottle receiving the seawater sample. Data was collected

109

for several minutes (~2-4 min) once a steady-state signal was achieved. The relative standard

110

deviation of the chemiluminescence signal during data collection was < 5%.

111

This method was calibrated following a previously established protocol 21, except that in

112

our study, the calibration matrix consisted of aged filtered natural water collected from the same

113

field site at the same water depth as the samples, which was then amended with 50 µM

114

diethylene-triaminepentaacetic acid (DTPA, Sigma) and kept in the dark for at least 12 h prior to

115

use. Briefly, a primary stock solution containing potassium dioxide (Sigma) was prepared and

116

quantified spectrophotometrically (Abs240). To prepare the calibration standards, the primary

6


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stock solution was further diluted with the calibration matrix to a final O2- concentration of 5-41

118

nM. Both primary stock solution and calibration standards were prepared immediately before the

119

analysis. The calibration standards were pumped through the FeLume Mini system using the

120

same type and length tubing used in the field samples. The corresponding chemiluminescence

121

signals were recorded and extrapolated back to the time when the primary standard was

122

quantified, using first order decay kinetics, indicating that non-metal substances (likely organics)

123

contributed to decay in the DTPA-treated waters. The half-life of O2- in the calibration matrices

124

ranged from 0.26 min to 0.49 min and the extrapolation time was 0.5-1 min. Calibration curves

125

were constructed based on the linear regression of the natural logarithm extrapolated

126

chemiluminescence signals versus O2- concentrations in the calibration standards. Calibrations

127

yielded highly linear curves (e.g., R2 > 0.91), with a sensitivity of 0.16±0.04 (average and

128

standard deviation of different water depths) and 0.10 counts per pM in Oyster Pond and

129

Nantucket Sound water, respectively.

130

To confirm that the chemiluminescence signal was indeed due to the reaction of MCLA

131

and O2-, 800 U L-1 superoxide dismutase (SOD, Sigma) was added to the natural water samples

132

at the end of each run. In all cases, the chemiluminescence signal decreased rapidly to baseline or

133

slightly below baseline levels after the addition of SOD. The detection limit of this method was

134

calculated as three times the standard deviation of blank measurements, which was 0.13 nM and

135

0.24 nM for Oyster Pond water and Nantucket Sound seawater, respectively.

136

H2O2 concentrations. H2O2 concentration was quantified based on the oxidation of

137

colorless AmplifluTM Red (AR, Sigma) to pink-colored resorufin by H2O2. This reaction is

138

catalyzed by horseradish peroxidase (HRP, Sigma) and occurs at 1:1 (AR: H2O2) stoichiometry.

139

This method detects extracellular H2O2, and has been utilized to measure H2O2 released from

7



140

biological systems 40, 41 and H2O2 present in natural waters 26. For H2O2 concentration analysis,

141

pre-mixed AR and HRP stock solution was added at a final concentration of 50 µmol L-1 AR and

142

1 kU L-1 HRP to water samples in a clear 96-well microplate. A dilution factor of 1.37 due to the

143

addition of reagents was taken into account when calculating the H2O2 concentration in water

144

samples. Light absorbance was measured at 570 nm (Abs570, maximum absorbance of resorufin)

145

and 700 nm (Abs700, to account for background absorbance) on a SpectraMax® M3 multi-mode

146

microplate reader. The difference between Abs570 and Abs700 (i.e., Abs570-700) was used for

147

calculating H2O2 concentrations in water samples based on a calibration. The calibration factor was determined by standard addition of H2O2 into 0.2-µm filtered

148 149

natural water from each site. The H2O2 stock solution was prepared daily by diluting a 30% H2O2

150

aqueous solution (Seastar Chemicals Inc.) to approximately 10 mM. The exact concentration of

151

this stock solution was determined spectrophotometrically using its absorbance at 240 nm and

152

the H2O2 molar extinction coefficient of 38.1 M-1cm-1 42. This H2O2 stock solution was then

153

diluted in 0.2-µm filtered natural water to prepare a series of standard solutions immediately

154

prior to use. The H2O2 standard solutions and the unfiltered natural water samples were analyzed

155

concurrently in the same microplate. The Abs570-700 response was linear over the concentration

156

range of H2O2 measured in the present study. The calibration factor was 4.0-4.4×10-5 and 4.0×10-

157

5

Abs570-700 per nM H2O2 in Oyster Pond and Nantucket Sound seawater, respectively. To account for autoxidation of AR, 200 kU L-1 catalase (Sigma) was added to the blanks

158 159

prior to the addition of AR and HRP. The H2O2 concentrations in natural water samples were

160

determined by applying the calibration factor to the blank-corrected Abs570-700values. The

161

detection limit of this method was calculated as three times the standard deviation of blank

162

measurements, which was 4.0 nM and 9.4 nM for Oyster Pond and Weeks Pond water,

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respectively. To test the stability of H2O2 in natural water during storage on ice, H2O2

164

concentration in Oyster Pond water was quantified immediately after the samples arrived at the

165

WHOI laboratory and then 1 h (while samples kept on ice) later, during which time the decrease

166

of H2O2 concentration was 4.9±1.1%. If initial rapid decay of H2O2 occurred during sample

167

cooling down, the in-situ H2O2 concentrations may be underestimated.

168

Dark particle-associated ROS production rates. O2- and H2O2 production rates in

169 170

natural waters were determined by conducting laboratory incubations of water samples in the

171

dark within 2 hours after field collection. O2- production rates. O2- production rates in natural water samples were quantified using

172 173

a high throughput 96-well plate assay based on the reaction between O2- and MCLA, as

174

described in detail by Godrant et al. 43. Briefly, 50 µM xanthine (Sigma), 100 µM DTPA, 12.5

175

µM MCLA, and seven types of water samples were added to a white 96-well microplate. The

176

water samples examined in this assay included unfiltered natural water, 5-µm filtered natural

177

water, 0.2-µm filtered natural water, unfiltered natural water amended with 1% (v/v)

178

paraformaldehyde (Electron Microscopy Sciences) or 0.2 mM nicotinamide adenine dinucleotide

179

(NADH, MP Biomedicals), 0.2-µm filtered natural water amended with 1% (v/v)

180

paraformaldehyde or 0.2 mM NADH. 50 kU L-1 SOD was added to all the blanks. Calibration

181

standards were prepared by adding 3.75, 8 or 20 mU L-1 xanthine oxidase (Sigma) to each

182

natural water type. After MCLA addition, chemiluminescence measurements were taken every 3.5 min for

183 184

67 min, during which time 20 data points were recorded and appeared to be constant (RSD
0.96) in both

190

brackish water and freshwater. The pH of each sample was measured at the beginning and at the

191

end of the hour-long incubation, and the pH fluctuation was within 10% of the pH of the original

192

natural water. Any effects induced by this pH shift were accounted for by conducting the

193

calibration in the same water using the same reagents. As the chemiluminscence signal was

194

relative flat during the incubation period, the pH shift was interpreted to have a negligible effect

195

on the measured signals. Procedures for converting xanthine oxidase concentration to O2-

196

production rate are described in the Supporting Information. The signals of all amended and

197

unamended samples were corrected by subtracting the signal of the blanks. All

198

chemiluminescence readings were collected on a SpectraMax® M3 multi-mode microplate reader. In each experiment, three calibrations were conducted in unfiltered, 5-µm filtered and

199 200

0.2-µm filtered natural water, and applied to the corresponding type of samples, respectively. In

201

formaldehyde and NADH amended 0.2-µm filtered water, the addition of 20 mU L-1 xanthine

202

oxidase decreased the chemiluminescence signals by 14.7±14.1% and 4.8±8.9%, respectively,

203

indicating that neither formaldehyde nor NADH reacted with O2- significantly. The O2-

204

production rates in 0.2-µm filtered samples were subtracted from those in unfiltered and 5-µm

205

filtered samples to yield particle-associated production rates. During the recording of the

206

chemiluminescence signals, water samples were incubated with MCLA in the dark at room

207

temperature. Therefore, the O2- production rates obtained from this method represent the rates of

208

dark particle-associated O2- production.

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H2O2 production rates. The absolute rates of dark particle-associated H2O2 production

209 210

were quantified based on the linear regression (R2 > 0.88) of H2O2 concentrations in water

211

samples versus reaction time of natural water, AR and HRP. The calibrations were conducted in

212

freshly filtered (0.2 µm) natural water from each site. Seven types of samples, including

213

calibration standards, unamended natural water, 5-µm filtered natural water, catalase controls,

214

killed controls, SOD controls and resorufin controls, were incubated with AR and HRP in the

215

same microplate, which was kept in the dark at ambient laboratory temperature (22-24 ºC) for 4

216

h. Catalase controls consisted of unfiltered natural water amended with 100 kU L-1 catalase every

217

hour during the incubation. Killed controls, SOD controls and resorufin controls consisted of

218

unfiltered natural water amended with 1% (v/v) paraformaldehyde, 50 kU L-1 SOD, and 0.5 µM

219

resorufin (Sigma), respectively, immediately prior to the analysis. The same amount of catalase,

220

paraformaldehyde and SOD were also added to 0.2-µm filtered waters, which were incubated

221

under the same condition as the other samples. The light absorbance of these 0.2-µm filtered

222

samples was utilized for background correction of all the controls. The H2O2 concentrations in

223

all samples were measured simultaneously at intervals during the 4-h incubation. 112.3±12.0%

224

and 104.3±8.5% of the 0.5-µM resorufin spike was recovered at the end of the incubation in

225

unfiltered freshwater and brackish water, respectively. The recovery of the standard addition (0.5

226

µM of H2O2) was 98.1±4.4% and 102.3±8.6% in SOD and formaldehyde amended 0.2-µm

227

filtered water, respectively, indicating that neither SOD nor formaldehyde reacted with H2O2

228

significantly. It should be noted that H2O2 in the air was slowly absorbed into the water samples during

229 230

the incubation, which was reflected in an increasing y-intercept of the calibration curves.

231

However, the atmospheric contamination was expected to affect all samples in adjacent

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microplate wells equally, and thus was accounted for by incubating the calibration standards with

233

the natural water samples in the same microplate for the same period of time. Photochemical H2O2 production rate estimations. An estimate of the H2O2

234 235

photoproduction rate, averaged over a 24-hour period and over a 6 m water column, was

236

generated as described in a previous study44. Briefly, we assumed that all incident sunlight (both

237

direct and downwelling diffuse) is absorbed over the 6 m water depth; we used the National

238

Center for Atmospheric Research Quick total ultraviolet (TUV) calculator to determine a 24-

239

hour average of spectral irradiance at the summer solstice at a latitude of 41.54o N. Since the

240

wavelength-dependent apparent quantum yields of H2O2 photoproduction for water from our

241

field sites are unknown, we used the log-linear function shown in Figure 6 of Yocis et al.

242

(2000)45, representing an average of a number of marine and estuarine samples. The quantum

243

yields from these different field sites ranged over about one log unit, and those of several surface

244

freshwaters measured in a different study11 fell within this range. Our estimate is therefore likely

245

to fall within a factor of three of the true production rate. O2- photoproduction can be estimated

246

as twice the H2O2 rate, based on the assumption that all photoproduced H2O2 originates from

247

dismutation of O2- and that other sinks of O2- are minor.

248 249

Results and Discussion

250

Non-zero ROS concentrations in dark water. ROS concentrations in natural water that

251

received variable levels of sunlight exposure were measured during two separate field surveys at

252

Oyster Pond in August of 2014. First, the concentrations of O2- and H2O2 were quantified at

253

different depths, including at the water surface, in the deeper photic zone (water depth 100 µmol s-1 m2) and aphotic zone (water depth > 3 m, PAR < 100 µmol s-1 m2) during

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midday. Second, water samples were collected from the surface layer and the aphotic zone

256

(water depth 3.5 m) at different time points during a day, including before sunrise, midday,

257

afternoon and after sunset. Concentrations of both O2- and H2O2 were significantly above the

258

detection limit in all dark water samples, such as samples collected from the aphotic zone and

259

surface water analyzed under little sunlight exposure (Figure 1). Depth profile measurements collected during the first field survey ranged from 2.1±0.1

260 261

nM to 7.8±0.8 nM for O2-, and from 18.8±5.4 nM to 28.6±6.9 nM for H2O2, with the maximum

262

concentrations of both ROS observed in surface water (Figure 1a). H2O2 concentrations have

263

been quantified in a variety of aquatic environments and our H2O2 measurements fell within the

264

range of previous data 26, 37, 46-48. O2- abundance in the brackish water pond exceeded most of the

265

currently available O2- concentrations, which were all measured in marine environments 17, 25, 26,

266

37

. Relatively uniform distribution of ROS was observed throughout the photic zone and the

267 268

upper aphotic zone (Figure 1a). The large drop of O2- concentration at 4.5 m may be partly

269

attributed to the lower abundance of dissolved O2 (Table S1), the precursor of O2-. However,

270

H2O2 concentration did not greatly decrease in the aphotic zone. At 3.5 m and 4.5 m, PAR

271

declined to 3.1% and 0.9% of the surface level, while the H2O2 concentration was 65.6% and

272

87.2% of that in surface water, respectively (Figure 1a). Earlier studies 29, 30, 32, 34, 36, 49 on vertical

273

profiles of H2O2 in natural waters typically showed a quasi-exponential decrease of H2O2 levels

274

with depth and minimal H2O2 concentrations in dark water. Our H2O2 results (Figure 1a)

275

demonstrate a departure from this trend, which may be attributable to the possibility of vertical

276

mixing throughout the shallow waters we examined. In any case, the present study agrees with

13



277

other recent field research conducted in marine settings that show significant levels of O2- 17 and

278

H2O2 33, 35 in the aphotic zone.

279

Both O2- and H2O2 concentrations in surface water increased as incident sunlight

280

irradiation increased from dawn to midday, and then gradually decreased until dusk (Figures 1b

281

and S2), while geochemical conditions, including the water temperature, pH, salinity and DO

282

remained similar throughout the field day (Table S2). Lowest (but non-zero) ROS

283

concentrations in surface water appeared before sunrise (Figure 1b). These results are consistent

284

with previous diel measurements in marine 17, 32, 33, estuarine 47, 48 and fresh 38, 46 photic waters.

285

Because the temporal pattern of ROS concentrations followed the variation of solar irradiation in

286

these studies, photochemical processes have been inferred as the most important source of ROS

287

in surface water.

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However, as observed in the depth profile (Figure 1a), concentrations of both ROS

289

measured in the aphotic zone did not significantly differ from those measured in the surface layer

290

(Figure 1b), even though the aphotic zone received much lower sunlight irradiation throughout

291

the sampling period (PAR < 26 µmol s-1 m2, Figure S3). More interestingly, O2- and H2O2

292

concentrations in the aphotic zone also exhibited a diel trend with a midday maximum of 3.1 nM

293

and afternoon maximum of 38.5 nM, respectively (Figure 1b). Given that we expect a relatively

294

long lifetime of H2O2, vertical mixing of photochemically generated H2O2 into deeper waters

295

might have contributed to the presence and diel cycling of H2O2 in the aphotic zone. However,

296

decomposition of O2- occurs rapidly through various light-independent pathways, including

297

abiotic redox reactions 1, 3, 50-52 with metals or organic compounds, and enzymatic degradation by

298

SOD 31. These processes render the lifetime of O2- extremely short. The non-zero O2-

299

concentrations measured in the aphotic zone and at nighttime in our study are consistent with the

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“constant background” of O2- observed in previous fieldwork 17, 25, and indicate that O2- must be

301

actively produced in the absence of light to balance rapid decomposition. This diel trend of O2-

302

levels in the aphotic water was likely due to the temporal change of some light-independent ROS

303

sources and/or sinks, such as diel variation in biological activity, which was not directly caused

304

by the change of sunlight irradiation but occurred simultaneously with it as postulated in

305

previous studies 28, 53.

306

The biological nature of dark particle-associated ROS production. In regards to the

307 308

potential light-independent source of ROS in natural waters, our study focused on particle-

309

associated processes. The rates of dark particle-associated production of O2- and H2O2 were

310

quantified in the surface water, deeper photic zone and aphotic zone of Oyster (brackish water)

311

Pond in July, 2014 and Weeks (freshwater) Pond in June and August, 2013. The boundary

312

between the photic zone and the aphotic zone was at ~3 m, determined by a PAR (100 µmol s-1

313

m2 as the threshold) sensor (Figure S4) at Oyster Pond and a Secchi disk at Weeks Pond,

314

respectively. During these three field surveys, dark particle-associated production of both O2- and

315 316

H2O2 occurred in all of the water samples (Figure 2). In brackish water, dark O2- production

317

rates ranged from 25.1±14.6 nM h-1 to 89.4±38.0 nM h-1 with a local maximum in the lower

318

photic zone (2.5 m, Figure 2a), and dark H2O2 production rates ranged from 55.4±3.9 nM h-1 to

319

95.9±4.0 nM h-1 with a local maximum in the aphotic zone (3.5 m, Figure 2b). In freshwater, the

320

June measurements exceeded the August measurements, with greater ROS production typically

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observed in the aphotic water (Figure 2c and 2d). The ROS production rates shown in Figure 2

322

are broadly similar to the findings from recent field studies. For example, Vermilyea et al.

15



323

demonstrated dark H2O2 production rates of 29-122 nM h-1 in freshwater lakes in the Denver

324

Metropolitan area 24. More recently, Marsico et al. measured dark H2O2 production rates at

325

freshwater systems with a variety of trophic states in Colorado and Massachusetts, and obtained

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a range of 3-259 nM h-1 27. Nonphotochemical O2- production in tropical 17 and non-tropical 25

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oceanic water was estimated to occur at 24 h) filtered seawater, which accounts for less than 15% of the NADH

21



461

enhancement observed in unfiltered seawater (Figure S10), indicating that substantial NADH

462

autoxidation did not occur in this water. The addition of formaldehyde into the NADH-amended

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samples decreased 43%-55% and 61%-70% of the NADH-stimulated O2- concentrations in

464

brackish water and seawater, respectively (Figure 4c and 4d). This result revealed that a large

465

fraction of the NADH-enhanced O2- in natural waters was attributed to cell-associated pathways,

466

while abiotic reactions and/or extracellular enzymatic reactions 21, 63, 64 may also contribute to

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this process. In fact, previous investigations have shown that NADH oxidases 63 and peroxidases

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64

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dislodged from the cell surface and can persist as free (not cell associated) enzymes.

involved in microbial extracellular O2- production are outer membrane enzymes that are easily

Presumably, increased dismutation of NADH-stimulated O2- production would result also

470 471

in higher H2O2 production. However, the direct impact of NADH addition on H2O2 levels could

472

not be assessed as NADH caused an interference with the Ampliflu Red assay employed here.

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Extracellular H2O2 production is often thought to represent cell leakage of photosynthetically

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reduced oxygen under high light conditions 65. In the aphotic zone waters explored here, the lack

475

of UV but minor (

Extensive Dark Biological Production of Reactive ... - ACS Publications

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