At present most knowledge on the impact of iron on
18O /16O ratios (i.e. δ18O) of dissolved oxygen
(DO) under circum-neutral conditions stems from experiments carried out
under controlled laboratory conditions. These showed that iron oxidation
leads to an increase in δ18ODO values. Here we present the
first study on effects of elevated Fe(II) concentrations on the δ18ODO in a natural, iron-rich, circum-neutral watercourse. Our
results show that iron oxidation was the major factor for rising dissolved oxygen
isotope compositions in the first 85 m of the system in the cold season
(February) and for the first 15 m during the warm season (May). Further
along the course of the stream, the δ18ODO decreased
towards values known for atmospheric equilibration around +24.6 ‰ during both seasons. Possible drivers for these changes may be reduced iron oxidation, increased atmospheric exchange and
DO production by oxygenic phototrophic algae mats. In the cold season, the
δ18ODO values stabilized around atmospheric equilibrium,
whereas in the warm season stronger influences by oxygenic photosynthesis caused values down to +21.8 ‰. In the warm season
from 145 m downstream of the spring, the δ18ODO
increased again until it reached atmospheric equilibrium. This trend can be
explained by respiratory consumption of DO combined with a relative
decrease in photosynthetic activity and increasing atmospheric influences.
Our study shows that dissolved Fe(II) can exert strong effects on the
δ18ODO of a natural circum-neutral spring system even
under constant supply of atmospheric O2. However, in the presence of
active photosynthesis, with supply of O2 to the system, direct
effects of Fe oxidation on the δ18ODO value become
masked. Nonetheless, critical Fe(II) concentrations may indirectly control
DO budgets by enhancing photosynthesis, particularly if cyanobacteria are
involved.
Introduction
Oxygen is the most abundant element (45.2 %) and iron the fourth most abundant
element (5.8 %) on Earth (Skinner, 1979). Such huge global reservoirs
render these elements critically important in global biogeochemical cycles.
In addition, their reactivity is exceptional: O2 is a powerful
oxidation agent while Fe can cover oxidation states from -4 to +7 in
extreme cases, with the most commonly known ones being 0, +2 and +3 (Lu
et al., 2016).
Iron is also an essential trace element in many biological processes,
including photosynthesis, oxygen transport and DNA biosynthesis (Kappler et
al., 2021). This closely links to the formation and dissolution of Fe
oxides. These common forms of metal oxides may enhance or reduce
availabilities of both elements in the water column and pore waters and thus
may largely regulate aqueous life.
In aqueous environments, dissolved oxygen (DO) is one of the most essential
ecosystem parameters, and, despite its moderate solubility (e.g. 9.3 mg L-1 at
20 ∘C), it assumes a central role in respiration, primary
production and Fe oxidation (Pusch, 1996). The concentration of DO coupled
to its stable isotope 18O /16O ratios (i.e. δ18O) can yield additional information about sources and sinks, including
atmospheric input, photosynthesis, respiration and mineral oxidation.
When equilibrated with the atmosphere, δ18ODO values
typically range around a value of +24.6 ‰ (Mader et
al., 2017), while photosynthesis and respiration can change these isotope
ratios (Guy et al., 1993; Kroopnick, 1975). The splitting of water molecules
during photosynthesis hardly produces isotope discrimination, and the
resulting DO should have the same isotope value as the surrounding water
(Guy et al., 1993; Eisenstadt et al., 2010). Meteoric water in temperate
climates is normally depleted in 18O, and therefore the photosynthetic
oxygen in these areas varies between -10 ‰ and -5 ‰
(Quay et al., 1995; Wang and Veizer, 2000). Respiration, on the other hand,
preferentially accumulates 16O and enriches the remaining DO in
18O. This process yields δ18ODO values between
+24.6 ‰ and +40 ‰ (Guy et al., 1993).
Additionally, oxidation of metals such as Fe also leads to increases in
δ18ODO (Lloyd, 1968; Taylor and Wheeler, 1984; Wassenaar
and Hendry, 2007; Oba and Poulsen, 2009a, b; Pati, 2016). Mostly, the
impacts of Fe oxidation on δ18ODO values have been
investigated experimentally under controlled conditions (Oba and Poulson,
2009b; Pati et al., 2016). As a new aspect, these dynamics have not been studied
in open water systems such as springs and rivers so far. New field
investigations might reconcile variations in the fractionation factors
obtained in the above-mentioned studies. Currently they are thought to
result from differences in temperature, pH and initial Fe(II) concentrations
that could be outlined under abiotic conditions.
Dissolved Fe(II) in natural systems may have primary and secondary impacts
on DO concentration and its δ18ODO values. The primary
influence originates from the O2 binding by iron oxidation (Eq. 1). This leads to decreases in the DO and causes simultaneous increases in
δ18ODO values (Wassenaar and Hendry, 2007; Smith et al.,
2011; Parker et al., 2012; Gammons et al., 2014).
4Fe2++O2+4H+→4Fe3++2H2O
Dissolved Fe(II) can also have secondary (i.e. indirect) influences on the
DO content and the δ18ODO. This happens when it acts as an
essential micronutrient to cause growth-stimulating effects on
O2-producing and respiring microorganisms. These influences of Fe(II)
on DO and δ18ODO in circum-neutral aquatic systems have so
far received little attention because of the following reasons:
Fe oxidation often masks δ18ODO values created by
respiration, photosynthetic and atmospheric oxygen.
Adequate Fe(II)-rich circum-neutral model systems are scarce on modern
Earth. This is due to the high reactivity of iron with DO.
To the best of our knowledge, no study so far has systematically
investigated the influences of elevated Fe(II) concentrations on δ18ODO values in a natural and circum-neutral iron-rich system. In
order to bridge this gap, we investigated the aqueous chemistry and δ18ODO values in the iron-rich Espan Spring in Fürth, Germany
(Fig. 1). This Fe(II)-rich artesian spring offers a complex biogeochemical
natural field site to analyse effects of different Fe(II) contents on the DO
and δ18ODO values.
Overview of the Espan Spring in Fürth, Germany. (a) and (b):
location of the spring in Bavaria and the city of Fürth. (c) Satellite
image of the spring by Google maps showing the distinct red colour. (d) to (f):
detailed photos of the system. (d) Displays the stream between sampling
points E4 and E5, (e) shows sampling point E3 (with the bank of the water
line in the upper part of the picture) and (f) displays sampling point E4.1
with algae and cyanobacteria mats.
The aims of this study were to establish an inventory of biology together
with Fe and oxygen budgets in this natural spring and stream system. We
further aimed to investigate how increased Fe(II) levels influence the
oxygen budget of the system and whether a combination of DO and δ18ODO measurements can help to assess this effect. This is also
timely because environmental impacts of Fe(II) are becoming increasingly
recognized for their negative effects on ecosystems such as with the
browning or brownification phenomenon (Kritzberg and Ekström, 2012;
Weyhenmeyer et al., 2014; Kritzberg et al., 2020). During this process,
increased iron levels can consume oxygen, cause algae blooms and reduce
water quality and thus may affect aqueous ecosystems and their services.
Here we describe a first complete spatial sampling campaign in the cold and
warm season with Fe(II), Fe(III), DO and its stable 18O /16O
isotope ratios together with field parameters (pH, T, DO, pe, electrical
conductivity). This study contributes to the knowledge of Fe oxidation in
natural systems and delivers implications of hardly explored seasonal
dynamics in Fe(II)-rich systems.
MethodsStudy site
The Espan Spring is located in the city of Fürth, Germany (49∘28′15.8′′ N, 11∘00′53.0′′ E; Fig. 1). It is an artesian spring that
originates from a confined aquifer that was tapped by a drilling project in
1935 from a depth of 448.5 m below ground. The water originates from the so-called “lower mineral water horizon”. This horizon is dominated by
artesian inflow from the lower Buntsandstein Formation. The Buntsandstein in
Fürth consists of red sandstone layers that are composed of light
reddish to yellowish-white-grey sandstones of different grain sizes. The
sandstones are intercalated with various rubble, conglomerate and clay
layers as well as thin gypsum and salt (Birzer, 1936). Three noticeable
conglomerate layers are present in the sequence of Buntsandstein layers.
Birzer (1936) distinguished the Upper Buntsandstein from the Upper Main
Buntsandstein by the Main Conglomerate which can be found at a depth of 321
to 324 m. The Middle Boulder Layer at a depth of 370 to 371 m separates the
Upper Main Buntsandstein from the Lower Main Buntsandstein and the so-called Eck'sche
Conglomerate at a depth of 433 to 440 m, which separates the Lower Main Buntsandstein from the Lower Buntsandstein (Birzer, 1936).
At a depth of 370 to 439 m, mineral water flows into the borehole from the
Upper Main Buntsandstein and Lower Main Buntsandstein and from the Eck Conglomerate. This
water, which is caught in the red sandstone and has a temperature of about
+22 ∘C, was called the lower mineral water horizon; in 1936
its yield was about 10 L s-1 at a water temperature of +23 ∘C (Kühnau 1938). The water of this lower spring horizon is under
artesian pressure and exits the spring with a head of about 13 m above ground
level (Birzer, 1936). Nowadays the Espan Spring has a constant yield of
about 5 L s-1.
After the water exits the basin in a pavilion with a temperature of
∼ 20 ∘C, it discharges into a stream of about 300 m length that is known as the Wetzendorfer Landgraben (WL). This small
stream drains into the river Pegnitz without any further tributaries (Fig. 1b, c). The water can be classified as a Na-Ca-Cl-SO4 mineral water
with initially undersaturated DO values of 2.3 mg L-1 and Fe(II) contents of
up to 6.6 mg L-1 (Table 1). Figure 1c shows an aerial image of the spring and
stream system that shows a distinct red colouring of the stream bed. The most
plausible explanation for this colouring is iron-oxide precipitates (Fig. 1d, e). The WL has a water depth between 8–10 cm with few
fluctuations.
Sampling procedures
Two field campaigns were performed in February and May 2020, during which
water was collected at 14 locations along the stream. Data from these campaigns are available in Table 1. The on-site parameters of
pH (± 0.05; instrument precision), temperature (± 0.1 ∘C), electrical conductivity, Eh and DO (all ± 2 %) were
measured with a Hach HQ40D multi-parameter instrument. Alkalinity
titrations were carried out with a Hach titrator with a bromocresol-green
indicator. Fe(II) and Fe(III) contents were measured using an iron (II–III)
cuvette test set by Hach in combination with a portable Hach
spectrophotometer (model DR 2800).
Samples for 18O /16O ratios of DO were collected in 12 mL
Exetainers (Labco Ltd., Lampeter, UK) that were prepared with 10 µL of
a saturated HgCl2 solution to prevent secondary biological activity
after sampling (Wassenaar and Koehler, 1999; Parker et al., 2005, 2010).
The Exetainers were filled with syringe-filtered water via 0.45 µm pore
size nylon filters until they were entirely full and free of air bubbles.
They were then carefully closed with screw caps with a butyl septum in order
to avoid atmospheric contamination. Test series showed that the amount of
atmospheric contamination during this filling procedure is usually
negligible (Mader et al., 2018).
Samples for water isotopes were collected in 15 mL Falcon tubes and treated
in the same manner as the ones for DO isotope measurements, except for
preservation with HgCl2. All samples were stored in a mobile
refrigerator box at 4 ∘C directly after collection and carried to
the laboratory where they were measured within 24 h.
Identification of possible mineral precipitates
In order to determine possible mineral precipitate data for the pH, pe (the
negative decadic activity of available electrons), temperature and alkalinity
(as CaCO3), as well as cations and anions, the specific sampling points
were fed into the program PHREEQC (Version 3; Parkhurst and Appelo, 2013)
for calculation of saturation indices. The database used for these
calculations was WATEQ4.
Laboratory methodsIdentification of cyanobacteria
Samples were collected in a preliminary field assessment at the anoxic
piping where the spring flows into the creek (E2), in the middle of the
creek at the first small pond after the water had contact with the atmosphere
(E3) and about 5 m downstream of this pond from an algal mat with bubbles on
the surface (E4). Samples for cyanobacterial isolation were collected in
sterile 2 mL Sarstedt tubes and sealed. Samples for microscopic analysis were
collected with a 75 % ethanol-sterilized spatula and placed in a sterile
6 cm petri dish (Sarstedt, Germany). Immediately after returning from
sampling, samples were embedded in 1.5 % agarose in de-ionized water to
preserve the structure of the bio mats during further handling and shipping.
Microscopic analysis was performed on thin sections of the embedded mats
using a confocal laser scanning microscopy (CLSM) type microscope (LSM 880, Carl Zeiss), using modified
acquisition settings from Jung et al. (2019) to discriminate between
cyanobacterial (chlorophyll a (Chl a) and phycobiliproteins (PBPs)) and green
algal (Chl a) fluorescence. Laser transmission images were also generated
using the 543 nm laser.
A spatula tip of green-coloured mat was used to inoculate 5 mL of BG11
medium (Stanier et al., 1971) in a well of a six-well plate and incubated for
3 weeks at 24 ∘C on a 16:8 day : night cycle with illumination at
15 µmol photons m-2 s-1 under an OSRAM L 30 W/840 LUMILUX Cool
White bulb. Individual cyanobacterial species were picked from the mat
cultures under a Nikon SMZ-U Zoom binocular microscope for further
subculturing on 1 % agar-solidified BG11 plates, as well as in liquid
culture. Isolates were observed under an Olympus BX53 light microscope, and
their morphologies were recorded using an Olympus DP26 camera. The number of
cells per filament and cell dimensions were measured using ImageJ 1.47v
software. DNA was extracted (Gehringer et al., 2010) from one axenic isolate
of a microscopically identified Persinema species of cyanobacteria. The 16S rDNA gene
and intergenic spacer sequence were amplified by the SSU-4-forw and ptLSU C-D-rev primer pair (Marin et al., 2005) using the Taq PCR Master Mix (Qiagen,
Germany). The PCR product was purified (NucleoSpin PCR clean-up kit,
Macherey-Nagel, Germany) and sequenced (Wilmotte et al., 1993). Sequences
were merged (HVDR Fragment Merger tool; Bell and Kramvis, 2013), and the
final 16S–ITS sequence was submitted to the National Center for Biotechnology
Information (NCBI), National Institutes of Health, USA.
Isotope measurements
Stable isotope ratios of DO (expressed as δ18ODO) were
measured on a Delta V Advantage isotope ratio mass spectrometer (IRMS;
Thermo Fisher Scientific, Bremen, Germany) coupled to an automated
equilibration unit (GasBench II). Measurements were carried out in
continuous-flow mode with a modified method by Barth et al. (2004). Here the
isolation of DO into a headspace relies on a helium extraction technique by
Kampbell et al. (1989) and Wassenaar and Koehler (1999). Different portions
of laboratory air were injected into helium-flushed Exetainers and used to
correct for linearity and instrumental drift during each
run. Water isotopes were measured with an isotope ratio infrared spectroscopy analyzer (L-1102i, Picarro Inc., Santa Clara, CA, USA) according to a method by van Geldern and Barth (2012). Here laboratory air is defined to represent atmospheric oxygen with a
ubiquitous value of +23.9 ‰ versus Vienna Standard Mean
Ocean Water (VSMOW) (Barkan and Luz, 2005). Data were normalized to this
value.
δ=Rsample/RSMOW-1(Clark and Fritz, 1997)
To obtain ratio changes in per mil (‰), the δ values were multiplied by a factor of 1000.
All samples were measured in triplicates, and isotope value standard
deviations (1σ) were less than 0.1 ‰ and 0.2 ‰ for
δ18OH2O and δ18ODO, respectively.
Field parameters, δ18ODO values, iron and uranium from sampling campaigns in February (cold season) and May (warm season) 2020 at the Espan Spring. Note that water isotopes are not listed here. They consistently ranged around a value of -9.7 ‰.
Results and discussionOn-site parameters
The on-site parameters as displayed in Table 1 show a range of pH values
between 6.1 and 8.6 in the cold season
and between 6.3 and 8.0 in the warm season. The observed changes in the pH
over the course of the spring are mostly due to the constant degassing of
CO2 from the spring. Oxygen values range from 2.3 to 11.0 mg L-1 in
the cold season and from 3.6 to 8.8 mg L-1 in the warm season.
Differences between the cold and warm season are due to the fact that cold water
can dissolve more O2 than warm water. The general increase in the
amount of DO over the course of the spring is due to a continuous
dissolution of atmospheric O2 in the spring water and due to the impact
of photosynthesis. Water temperatures ranged between 19.3 and 7.4 ∘C in the cold season and between 21.3 and 25.7 ∘C in the warm
season. The conductivity remained stable over the course of the stream and
only showed minor differences between the cold and warm season. The same
applies to the alkalinity. The behaviour of the Fe(II) and Fe(III) is
described in Sect. 3.5. Values of major ions (Cl-, SO42-,
NO3-, Na+, K+, Ca2+ and Mg2+) remained
constant over the course of the spring and show no differences between the
cold and warm season.
Precipitation calculations
Precipitating mineral phases as determined with PHREEQC showed that the
dominant phase at all measurement points was hematite (Fe2O3)
(Supplement Tables S1 and S2). Additionally, goethite (α-FeO(OH)), ferrihydrite (Fe(OH)3), siderite
(FeCO3) and K-jarosite (KFe33+(OH)6(SO4)2) as
well as CaCO3 and rhodochrosite (MnCO3) showed elevated SI values
and indicated precipitation.
Bacterial contents
CLSM showed that only the samples from
Site E4.1 have photosynthetic organisms in significant quantities during the
cold period. The photosynthetic community in this biofilm was dominated by
cyanobacteria, with very few eukaryotic algae (Fig. 2). Lyngbya was observed
along the sides of the fast-flowing stream on the smooth hard canal section
at E2; however, the loosely built Lyngbya sp. mats were only observed in the wider,
shallower sections from sampling sites E3 to E5 and predominating between
sites E3.1 and E4.1. The Lyngbya sp. filaments were not encrusted by oxidized iron
as proven by light microscopy. As these are simple cyanobacterial mats on
top of loose iron oxides, with no additional microbial layers beneath them,
the bubbles are presumably oxygen generated during photosynthesis
(Supplement Fig. S1)
CLSM images of mat sample E4.1. Overview: images of the
cross-section of the top 3 mm of the biofilm with the Chl a (C1) and Chl a plus PBP (CP1) fluorescence profile, complemented by a laser transmission
picture (LT1) and the superimposed image (O1). Detail: superimposed images
(O2, O3, O4) of Chl a (C2, C3, C4) and Chl a plus PBP (CP2, CP3, CP4) fluorescence and
laser transmission (not shown) of distinct organisms found in the bio mat.
O2: eukaryotic algae. O3: possible Klisinema- or Persinema-like sp. and a
unicellular cyanobacterium. O4: Lyngbya-like sp. and a unicellular
cyanobacterium.
Most of the cyanobacteria and all eukaryotic algae were located in the
topmost 1.2 mm of the biofilms (Fig. 2, O1). Close-up images show eukaryotic
algae (Fig. 2, O2); thin filamentous cyanobacteria, possibly Persinema sp. or Klisinema sp. (Fig. 2, O3); and
Lyngbya sp. (Fig. 2, O4). All pictures of the top layers of this sample site show an
abundance of unidentified unicellular cyanobacteria, while images from the
other sample sites show very few photosynthetic organisms at all
(Supplement Fig. S2).
In order to determine the identity of the predominant cyanobacterial species
isolated from the E4.1 enrichment cultures, a determination key was used to
compare particular features of an isolate to those already in the literature
for specific cyanobacterial species (Komárek und Anagnostidis, 2005).
Note that enrichment cultures for samples E2 and E3 did not yield enough
material for cyanobacterial determination after 5 weeks in culture.
The red-brown filamentous strain (Fig. 3c, d) exhibits single filaments,
without false branching, that are 30.9 to 38.2 µm wide (Table 2),
with a firm 9.5 to 14 µm thick sheath. The trichomes and single
cells are 21.5 to 24.2 µm wide and 1.5 to 4.1 µm long (Table 2), red-brown in colour, and constricted at the cross walls. Based on these
characteristics, the species was attributed to the cyanobacterial genus
Lyngbya.
Filament and cell dimensions of the proposed cyanobacterial species.
The blue-green filamentous strain (Fig. 3b) produces single filaments,
without false branching, that are 3.9 to 7.6 µm wide (Table 2) with a
firm 2.7 to 3.1 µm thick sheath. The trichomes and single cells are
1.2 to 4.5 µm wide and 0.3 to 0.4 µm long (Table 2) and
blue-green in colour without constriction at the cross walls. The terminal
cells in mature filaments are conical, elongated and bent to one side,
corresponding to those of the Klisinema genus recently described by Heidari et al. (2018). The thin, naked, pale green filaments (Fig. 3a, e) resembled
those of Persinema komarekii (Heidari et al., 2018) with apical cells flattened at the end. In
contrast to the observations of Heidari et al. (2018), we observed terminal
aerotopes. This species was purified in culture, and the 16S–ITS (NCBI
accession number MT708471) sequence confirmed its identity as Persinema komarekii (MF348313).
Light micrographs of the predominant isolates from sample E4.1: (a) single filament of Persinema sp., arrow indicates aerotopes. (b) Biofilm of
Klisinema sp. interspersed with Persinema sp. (arrow). (c)Lyngbya sp.
filament. (d)Lyngbya sp. sheath detail. (e) Biofilm of Persinema sp.
Dissolved oxygen (DO)
The DO concentration in the Espan System was lowest at the faucet in the
pavilion (sampling point E1a) with a saturation of 25.3 % (2.3 mg L-1)
(Fig. 4a). Over the following 100 m DO saturation increased to 88.1 % (8.7 mg L-1) in sampling point E4.1. Afterwards the saturation
continually increased to 94.6 % (11.0 mg L-1) in point E8. From an initial
depth of 435 m with the abundance of reduced species such as Fe(II) and
Mn(II), the low DO content in sampling point E1a was expected, and further along the course, more atmospheric oxygen was able to dissolve. In addition,
gas bubbles were observed in association with the Lyngbya mats. They were most
prominent at sample site E4.1 and indicate a significant contribution of
O2 from daytime photosynthesis. However, saturation with DO was not
reached during either of the sampling campaigns.
(a) Dissolved oxygen (%) and (b) Fe(II) concentrations over the
course of the Espan System in an example graph for the cold season in
February. The error for DO was 2 %, and for Fe(II) it was 0.06 mg L-1.
Errors are within the symbol size.
Fe(II) and Fe(III)
The Fe(II) content was highest at the faucet with 6.6 mg L-1, while its lowest
content was below instrument precision at sampling point E9 at 300 m
from the source (Fig. 4b). Fe(II) concentrations decreased constantly over
the stream course and were accompanied by increases in DO saturation (Fig. 4a). The decrease in Fe(II) could have been caused by three major processes:
oxidation of Fe(II) to form ferric iron minerals such as ferrihydrite,
hematite and goethite;
precipitation of Fe(II) minerals such as the iron carbonate siderite
(FeCO3) and/or an amorphous ferrous silicate phase; or
adsorption of Fe(II) on already-formed iron minerals.
All three possibilities seem plausible when taking into consideration the
saturation indices of ferric iron mineral – goethite, ferrihydrite and
hematite – precipitate at all sampling points in the system (Köhler et al.,
2020). These calculations furthermore show that siderite can precipitate in
almost all sampling points, while iron-silicate minerals are unlikely to
precipitate. Therefore, adsorption of Fe(II) onto minerals is also a
possible mechanism in the Espan System. Such adsorption of Fe(II) onto
(oxyhydr)oxides was shown to typically occur under neutral conditions and
should increase with rising pH (Zhang et al.,1992; Liger et al., 1999;
Appelo et al., 2002; Sylvester et al., 2005). Moreover, large amounts of
sulfate and chloride with average values of 2.2 and 4.5 g L-1 may have been
responsible for maintaining observed high dissolved Fe(II) contents of the
spring system at a circum-neutral pH despite rising DO concentrations. Such
elevated Cl- and SO42- contents can delay abiotic Fe(II)
oxidation (Millero, 1985).
Dissolved Fe(III) was highest (0.8 mg L-1) at sampling point E5 after 145 m
and lowest (0.05 mg L-1) at sampling point E7 after 265 m in flow distance from
the spring. The values initially increased from 0.4 mg L-1 in E1a to a maximum
of 0.8 mg L-1 in point E5 (± 0.03 mg L-1) and then decreased to their lowest
value in sampling point E7. The solubility of iron oxides in natural systems
at a circum-neutral pH and under aerobic conditions is generally very low
(Cornell and Schwertmann, 2003) with values of the solubility product
(Ksp) between 10-37 and 10-44 (Schwertmann, 1991). However,
Fe(III) could still be detected in the water, thus showing that its
dissolution was possible. The dissolution of iron oxides can occur through
several pathways including protonation, reduction and complexation that
create Fe(III) cations and Fe(II) cations as well as Fe(II) and Fe(III)
complexes (Schwertmann, 1991; Cornell and Schwertmann, 2003). Both the
protonation and the reduction would lead to the formation of
dissolved Fe(II). A steep increase in dissolved Fe(III) at 145 m downstream
of the spring (from 0.5 to 0.8 mg L-1) also indicated acceleration of
this process. One reason for this increase could be available organic
matter. However, further analyses are needed to verify this interpretation.
δ18ODO
Figure 5a and b show δ18ODO values over the course of
the spring for the cold and warm seasons, respectively. The curves are
divided into two zones for the cold season and three zones for the warm
season.
δ18ODO in the cold season (a) and the warm season
(b) over the course of the Espan Spring and stream system with the
atmospheric equilibrium value of +24.6 ‰ marked by
the horizontal dashed lines. Dashed vertical lines show borders of the different
zones of the fields labelled with 1, 2 and 3. The symbol size is larger than
the error bars.
Zone 1
In the cold season, zone 1 extended from sampling point E1a to point E4. In
this first 85 m, the δ18ODO increased from a value
of +23.7 ‰ at the faucet (E1a) to
+25.7 ‰ at E4. In the warm season, zone 1 extended
from E1a to E2 with only a 15 m distance from the spring. In this zone the
values increased from +23.4 ‰ at the faucet to a
maximum value of +24.7 ‰ at E2. In both seasons,
δ18ODO values at E1a were below the value expected for
atmospheric equilibration (+24.6 ‰). At first sight,
such 16O-enriched δ18ODO values would suggest
photosynthetic input of DO. However, the water originated from greater
depths without any exposure to light, and thus any photosynthetic influence
can be ruled out.
The occurrence of δ18ODO values below
+24.6 ‰ in groundwater has been described in the
literature (Wassenaar and Hendry, 2007; Smith et al., 2011; Parker et al.,
2014; Mader et al., 2018), and several explanations for this phenomenon
have been suggested (Wassenaar and Hendry, 2007; Smith et al., 2011; Parker
et al., 2014; Mader et al., 2018). These include
possible transfer of photosynthetic or diffusive oxygen into the shallow
aquifer (Smith et al., 2011; Parker et al., 2014; Mader et al., 2018),
radial oxygen loss of plant roots (Teal and Kanwisher, 1966; Michaud and
Richardson, 1989; Caetano and Vale, 2002; Armstrong and Armstrong, 2005b),
radiolysis of water (Wassenaar and Hendry, 2007), and
kinetic gas transfer (Benson and Krause, 1980; Knox et al., 1992; Mader
et al., 2017)
Explanations (1) and (2) are very unlikely in the Espan Spring because the
water originates from a depth of 435 m below ground through pipes that
presumably prevent any exchange with surface water or possible impacts of
plant roots. It should however be noted that water from the Espan Spring
contains up to 170 µg L-1 of uranium from easily soluble uranium
compounds that are commonly encountered in the Buntsandstein Formation
(Büttner et al., 2006; Meurer and Banning, 2019). The geogenic radiation
in the area is rather high because of the high uranium content in the
Variscian bedrocks of the area (Schwab, 1987; Büttner et al., 2006).
Because of this, radiolysis could be a possible explanation for the
unexpectedly low δ18ODO values. Kinetic gas transfer of
atmospheric oxygen during transport in the pipes or at the faucet might
be another explanation, since the sample in E1a is strongly DO undersaturated.
During non-equilibrium gas exchange the kinetically faster 16O would
cause δ18ODO below +24.6 ‰ until
equilibrium is established (Benson and Krause, 1980; Knox et al., 1992;
Mader et al., 2017).
Correlation between δ18ODO and DO over the course
of the spring for zone 1 in the cold season (a) and the warm season (b).
Correlation between δ18ODO and Fe(II) contents over the
course of the stream for zone 1 in the cold season (c) and the warm season (d).
Increases in δ18ODO values in zone 1 were accompanied by
increases in DO (Fig. 6a). In the cold season, a strong positive correlation
was evident between points E1a and E4. However, in the warm season, the same
correlation could only be observed between points E1a and E2 (Fig. 6b).
Equilibration with the atmosphere would be a reasonable explanation for this
trend until atmospheric equilibration is reached between points E2 and E3.
However, the δ18ODO values, at least in the cold season,
increased above this threshold to a value of +25.7 ‰. This shows that another process in addition to atmospheric equilibration
must have influenced the δ18ODO values in zone 1. In the
warm season, this was less evident, and the isotope atmospheric equilibrium
value was only marginally exceeded and remained within the range of the
analytical uncertainties.
Even though these processes consume DO, both respiration and iron oxidation
could be responsible for this trend when assuming that they influence the
δ18ODO values, while DO concentrations are constantly
replenished by the atmosphere. A direct negative correlation between Fe(II)
concentrations and δ18ODO values between point E1a and E4
was evident for cold-season samples and in points E1a and E2 for warm-season
samples as shown in Fig. 6c and d. This correlation between Fe(II) and
δ18ODO in the Espan System corresponds with the
experimental observations of Oba and Poulson (2009a), as well as those of
Pati et al. (2016). These studies demonstrate that Fe oxidation leads to
increases in δ18ODO values due to preferential consumption
of 16O. The increase in δ18ODO due to iron oxidation
in a natural system, which is constantly supplied with fresh oxygen,
indicates that Fe(II) oxidation must be a dominant control on δ18ODO in the first 85 m of the stream in the cold season and
in the first 15 m in the warm season. It also implies that the direct
impact of oxygen addition is subordinate in terms of DO stable isotope
changes. This is shown by iron oxidation being the dominant factor that
controls δ18ODO values, even though oxygen is constantly
supplied from the atmosphere.
Zone 2
In the cold season, zone 2 extended from sampling point E4 to point E9 with
only minor variations in δ18ODO. In this zone, the δ18ODO decreased from +25.7 ‰ in sampling
point E4 to values around atmospheric equilibrium with +24.5 ‰ in E7 and +24.8 ‰ in the
Pegnitz (Fig. 5a).
In the warm season, zone 2 extended from sampling point E2 to point E5 at
a 145 m distance from the spring. In this zone the values decreased from
+24.7 ‰ to a minimum value of
+21.8 ‰ in sampling point E5 (Fig. 5b). This decrease
in δ18ODO values can be explained by (1) a decrease in the
impact of iron oxidation on the δ18ODO values and (2) a
rising impact of atmospheric or photosynthetic oxygen. Even though a
decrease in Fe(II) values was still evident between E4 and E7 in the cold
season, as well as between E2 and E5 in the warm season, it is possible that
the decrease was not caused by Fe(II) oxidation and subsequent precipitation
as iron oxides. Alternatively, the decrease could have been caused by
adsorption of dissolved Fe(II) onto already-existing iron oxides such as
goethite, ferrihydrite and hematite (Zhang, et al., 1992; Liger et al.,
1999; Appelo et al., 2002; Sylvester et al., 2005). Because adsorbed Fe(II)
is very resistant to oxidation (Park and Dempsey, 2005), the impact of iron
oxidation on the δ18ODO values would have decreased.
No significant changes in the water chemistry were evident, and it can be
assumed that after sampling point E2 (warm season) or E4 (cold season), a
critical value was exceeded with enough Fe(II) having been adsorbed onto
iron oxides. In this case, iron oxidation – while probably still taking
place at small rates – is no longer an important factor dominating the
δ18ODO values. Downstream of points E2 and E4, oxygen
addition by the atmosphere or by photosynthesis would become more important.
Intensive growth of cyanobacterial and algal mats were observed between
point E3.1 and E5 in the cold season and between E3 and E5 in the warm
season (Fig. 1f). Because of this growth, it can be postulated that in
addition to the atmospheric O2 input, the δ18ODO
values were also influenced by photosynthetically produced oxygen. While
this effect should be less pronounced in the cold and darker season, a
stronger influence of photosynthetic oxygen on the δ18ODO
values would be expected in the warm season with higher light intensity.
Such growth of photosynthetic organisms in the Espan System is not
surprising with iron being an important micronutrient (Andrews et al.,
2003).
The fact that photosynthesizing organisms seem to preferentially grow and
impact the δ18ODO values between sampling points E3 and E5
may be due to the availability of Fe(II). In addition, the growth could also
be controlled by changes in the pH or other environmental influences, with
the site being located in a public park with the associated perturbations.
Cyanobacteria, especially aquatic strains, prefer a neutral to alkaline pH
(Brock, 1973), and the shift to higher pH values in this zone could be one of
the main factors that drive an increased supply of cyanobacterial O2. For
instance, Lyngbya spp. are diazotrophic cyanobacteria, capable of fixing nitrogen
during low availability of light, when local oxygen levels are low (Stal,
2012, p. 102). This oxygen released through oxygenic photosynthesis would
immediately react with Fe(II) and lower the partial pressure of oxygen
around the organisms in a slow-flowing stream. This could also favour
biological nitrogen fixation and limit carbon loss by reducing
photorespiration. Additionally, the reduced oxygen partial pressure induced
by Fe(II) oxidation may minimize the oxygenase activity of ribulose
1,5-bisphosphate carboxylase/oxygenase (Rubisco), thereby favouring
CO2 fixation (Stal, 2012, p. 113).
A screening of microbial ecology in several iron-rich circum-neutral springs
and experiments with the cyanobacterium Synechococcus PCC 7002 (Swanner et al., 2015)
revealed that many cyanobacteria show optimal growth between 0.4–3.1 mg L-1 of Fe(II) and that concentrations above 4.5 mg L-1 become growth-limiting. The
iron concentrations between points E3.1 and E5 in the cold season and E3 and
E5 in the warm season are thus approximately in the range of optimal
cyanobacterial growth. In order to establish a clear correlation between the
iron concentration and the decrease in δ18ODO values,
experiments would need to be carried out with the organisms found in the
Espan System. These have so far have not been assessed for their behaviour
under variable iron concentrations.
Zone 3
In the warm season, zone 3 extended from sampling point E5 to point E8. In
this zone the δ18O values rose again from
+21.8 ‰ to +24.3 ‰ (Fig. 5b).
The renewed increase in values can be explained by the influence of iron
oxidation, respiration and a decrease in photosynthetic activity. Because
Fe contents only decreased marginally, it can be assumed that decreases in
photosynthetic activities are responsible for increases in the δ18O values. This matches our observations that downstream of point E5,
only a little or no photosynthetic growth took place. Oxygen that would
dissolve in the water after point E5 would thus most likely stem from the
atmosphere. This would also explain the approach to the equilibrium value of
+24.6 ‰. Reasons for the observed decrease in
cyanobacteria are however not clear and may include changes in temperature and
light intensity and shifts in nutrient availability.
The temperature did not change significantly in this part of the watercourse
and is therefore unlikely to have caused a decrease in photosynthetic oxygen
production. In contrast, reduced light exposure could have been responsible
as downstream of point E5 trees shade the watercourse. A decrease in
nutrient availability is difficult to determine because nitrate and
phosphate were below the detection limit in the entire spring. Iron
starvation could also be a possible reason for the decrease in activity
because only ∼ 0.005 mg L-1 of Fe(II) was left in the system in the
lowest course of the stream.
Conclusions
Our study is the first systematic analysis of δ18ODO
values as a function of iron contents and oxygenic photosynthetic biofilms
in a natural iron-rich stream. We were able to confirm from field samples
that Fe oxidation leads to increases in δ18ODO values even
though oxygen was constantly replenished by atmospheric input. As soon as
photosynthetic oxygen is produced in the system, the effect of iron
oxidation on the δ18ODO values becomes negligible and can
no longer be detected. The fact that photosynthesis has a strong impact on
the δ18ODO values in specific areas of the system may be
controlled by high Fe contents. Similar iron-rich springs show
optimal growth rates of cyanobacteria in the range of 0.4–3.1 mg L-1 of
Fe(II). The presented δ18ODO values showed that
photosynthetic activity is also strongest in the Espan System within this
range of concentrations.
To what extent the changing Fe concentrations (Fe(II)–Fe(III)) influence the
growth of cyanobacteria and algae occurring in the Espan System requires
further investigation. This would ideally include isolating the organisms
from the watercourse and studying them under varying experimental levels of
Fe, pH and temperature while monitoring the δ18ODO of the
system. Further field studies with organic material from the stream bed in
combination with stable carbon isotopes would be promising to narrow down
processes for carbon and oxygen budgets in this environment.
Code availability
The code PHREEQC is available under https://www.usgs.gov/software/phreeqc-version-3 (Parkhurst and Appelo, 2009).
Data availability
All data of this study are available in this text and in the Supplement.
The supplement related to this article is available online at: https://doi.org/10.5194/bg-18-4535-2021-supplement.
Author contributions
IK, DP and JACB carried out the sample
collection and water analysis for on-site and isotope data. REM
carried out the calculation of the saturation index. MMG,
AJH and AG performed the analysis and interpretation
of cyanobacteria and algae data. IK prepared the manuscript
with contributions from all co-authors.
Competing interests
The authors declare that they have no conflict of interest.
Disclaimer
Publisher's note: Copernicus Publications remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Acknowledgements
Funding for this project was made available by the German Research
Foundation (DFG) through the Project IsoDO awarded to Johannes Barth and further DFG grants awarded to Michelle Gehringer. We also
thank Christian Hanke, Marlene Dordoni and Marie Singer for help with
sampling and analyses. Contributions by David Piatka were also carried out in the framework of the project AquaKlif in the bayklif network for investigation of regional climate change funded by the Bavarian State Ministry of Science and the Arts.
Financial support
This research has been supported by the German Research Foundation (DFG) (grant nos. BA2207/15-1, GE2558/3-1, GE2558/4-1).
Review statement
This paper was edited by Tina Treude and reviewed by two anonymous referees.
ReferencesAndrews, S. C., Robinson, A. K., and Rodríguez-Quiñones, F.:
Bacterial iron homeostasis, FEMS Microbiol. Rev., 27, 215–237,
10.1016/S0168-6445(03)00055-X, 2003.Appelo, T., Van der Weiden, M. J., Tournassat, C., and Charlet, L.: Surface
Complexation of Ferrous Iron and Carbonate on Ferrihydrite and the
Mobilization of Arsenic, Environ. Sci. Technol., 36, 3096–3103,
10.1021/es010130n, 2002.Armstrong, W. and Armstrong, J.: Stem photosynthesis not pressurised
ventilation is responsible for light-enhanced oxygen supply to submerged
roots of alder (Alnus glutinosa), Ann. Bot., 96, 591–612,
10.1093/aob/mci213, 2005.Barkan, E. and Luz, B.: High precision measurements of 17O/16O and 18O/16O ratios in H2O, Rapid Commun. Mass Sp., 19, 3737–3742, 10.1002/rcm.2250, 2005.Barth, J. A. C., Tait, A., and Bolshaw, M.: Automated analyses of O-18 / O-16
ratios in dissolved oxygen from 12-mL water samples, Limnol. Oceanogr.
Method., 2, 35–41, 10.4319/lom.2004.2.35,
2004.Bell, T. G. and Kramvis, A.: Fragment Merger: An Online Tool to Merge
Overlapping Long Sequence Fragments, Viruses, 5, 824–833,
10.3390/v5030824, 2013.Benson, B. B. and Krause D.: The concentration and isotopic fractionation
of gases dissolved in freshwater in equilibrium with the atmosphere. 1.
Oxygen, Limnol. Oceanogr., 25, 662–671,
10.4319/lo.1980.25.4.0662, 1980.
Büttner, G., Stichler, W., and Scholz M.: Hydrogeochemische
Untersuchungen in den Forschungsbohrungen Lindau 1 und Spitzeichen 1
(Fränkisches Bruchschollenland), Geol. Bavar., 109, 105–124, 2006.
Birzer, F.: Eine Tiefbohrung durch das mesozoische Deckgebirge in Fürth
in Bayern, Stuttgart, Zbl. Min. etc., Abt. B, 425–433, 1936.Brock T. D.: Lower pH limit for the existence of blue-green algae:
evolutionary and ecological implications, Science, 179, 480–483,
10.1126/science.179.4072.480 1973.Caetano, M. and Vale, C.: Retention of arsenic and phosphorus in iron-rich
concretions of Tagus salt marshes, Mar. Chem., 79, 261–271,
10.1016/S0304-4203(02)00068-3, 2002.
Clark, I. D. and Fritz, P. (Eds.): Environmental Isotopes in Hydrogeology,
CRC Press/Lewis, Boca Raton, USA, 1997.
Cornell, R. M. and Schwertmann, U. (Eds.): The Iron Oxides: Structure,
Properties, Reactions, Occurrences and Uses, Wiley-VCH Verlag, Weinheim,
Germany, 2003.Eisenstadt, D., Barkan, E., Luz, B., and Kaplan, A.: Enrichment of oxygen
heavy isotopes during photosynthesis in phytoplankton, Photosynth. Res.,
103, 97–103, 10.1007/s11120-009-9518-z, 2010.Gammons, C. H., Henne, W., Poulson, S. R., Parker, S. R., Johnston, T. B.,
Dore, J. E., and Boyd, E. S.: Stable isotopes track biogeochemical processes
under seasonal ice cover in a shallow, productive lake, Biogeochemistry,
120, 359–379, 10.1007/s10533-014-0005-z, 2014.Gehringer, M. M., Pengelly, J. J. L., Cuddy, W. S., Fieker, C., Foster, P.
I., and Neilan, B. A.: Host selection of symbiotic cyanobacteria in 21
species of the Australian cycad genus: Macrozamia (Zamiaceae), Mol. Plant.
Microbe. Interact., 23, 811–822, 10.1094/MPMI-23-6-0811,
2010.Guy, R. D., Fogel, M. L., and Berry J. A.: Photosynthetic fractionation of
the stable isotopes of oxygen and carbon, Plant Physiol., 101, 37–47,
10.1104/pp.101.1.37, 1993.Heidari, F., Zima, J., Riahi, H., and Hauer, T.: New simple trichal
cyanobacterial taxa isolated from radioactive thermal springs, Fottea.
Olomouc., 18, 137–149, 10.5507/fot.2017.024, 2018.Jung, P., Briegel-Williams, L., Schermer, M., and Büdel, B.: Strong in
combination: Polyphasic approach enhances arguments for cold-assigned
cyanobacterial endemism, Microbiologyopen, 8, e00729,
10.1002/mbo3.729, 2019.Kampbell, D. H., Wilson, J. T., and Vandegrift, S. A.: Dissolved-oxygen and
methane in water by a Gc headspace equilibration technique, Int. J. Environ.
Anal. Chem., 36, 249–257,
10.1080/03067318908026878, 1989.Kappler, A., Bryce, C., Mansor, M., Lueder, U., Byrne, J. M., and Swanner,
E. D.: An evolving view on biogeochemical cycling of iron, Nat. Rev.
Microbiol., 19, 360–374, 10.1038/s41579-020-00502-7, 2021.
Kühnau, J.: Balneologisches Gutachten über die Heilwirkungen, welche
von den im Stadtgebiet von Fürth erbohrten Mineralquellen zu erwarten
sind, Unveröff. Gutachten, 14 S., 3 Tab., Wiesbaden, 1938.Knox, M., Quay, P. D., and Wilbur, D.: Kinetic isotopic fractionation during
air-water gas transfer of O2, N2; CH4, and H2, J.
Geophys. Res.-Ocean., 97, 20335–20343,
10.1029/92JC00949, 1992.Köhler, I., Piatka, D., Barth, J. A. C., and Martinez, R. E.: Beware of
effect on isotopes of dissolved oxygen during storage of natural iron-rich
water samples: A technical note, Rapid Commun. Mass Sp., 35, e9024,
10.1002/rcm.9024, 2020.
Komárek, J. and Anagnostidis, K.: Cyanoprokaryota, 2. Oscillatoriales,
in: Süsswasserflora von Mitteleuropa, edited by: Büdel, B.,
Krienitz, L., Gärtner, G., and Schagerl, M., Elsevier/Spektrum,
Heidelberg, Germany, 760 pp., 2005.Kritzberg, E. S. and Ekström, S. M.: Increasing iron concentrations in surface waters – a factor behind brownification?, Biogeosciences, 9, 1465–1478, 10.5194/bg-9-1465-2012, 2012.Kritzberg, E. S., Maher, Hasselquist, E., Skerlep, M., Löfgren, S.,
Olsson, O., Stadmark, J., Valinia, S., Hansson, L. A., and Laudon, H.:
Browning of freshwaters: Consequences to ecosystem services, underlying
drivers, and potential mitigation measures, Ambio, 49, 375–390,
10.1007/s13280-019-01227-5, 2020.Kroopnick, P. M.: Respiration, photosynthesis, and oxygen isotope
fractionation in oceanic surface waters, Limnol. Oceanogr., 20, 988–992,
10.4319/lo.1975.20.6.0988, 1975.Liger, E., Charlet, L., and Van Cappellen, P.: Surface Catalysis of
Uranium(VI) Reduction by Iron(II), Geochim. Cosmochim. Ac., 63, 2939–2955,
10.1016/S0016-7037(99)00265-3, 1999.Llyod, R. M.: Oxygen isotope behavior in the Sulfate-Water System, J.
Geophys. Res., 73, 6099–6110,
10.1029/JB073i018p06099, 1968.Lu, J. -B. Jian, J., Huang, W., Lin, H., Li, J., and Zhou, M.: Experimental
and theoretical identification of the Fe(VII) oxidation state in
FeO4-, Phys. Chem. Chem. Phys., 18, 31125–31131,
10.1039/C6CP06753K, 2016.Mader, M., Schmidt, C., van Geldern, R., and Barth, J. A. C.: Dissolved
oxygen in water and its stable isotope effects: A review, Chem. Geol., 473,
10–21, 10.1016/j.chemgeo.2017.10.003, 2017.Mader, M., Roberts, A. M., Porst, D., Schmidt, C., Trauth, N., van Geldern,
R., and Barth, J. A. C.: River recharge versus O2 supply from the
unsaturated zone in shallow riparian groundwater: A case study from the
Selke River (Germany), Sci. Total Environ., 634, 374–381,
10.1016/j.scitotenv.2018.03.230, 2018.Marin, B., Nowack, E. C., and Melkonian, M.: A plastid in the
making: evidence for a second primary endosymbiosis, Protist,
156, 425–432, 10.1016/j.protis.2005.09.001, 2005.Meurer, M., Banning, A.: Uranmobilisierung im Helgoländer Buntsandstein
– Auswirkungen auf die Brack- und Trinkwasserqualität, Grundwasser, 24,
43–50, 10.1007/s00767-018-0408-1, 2019.
Michaud, S. C. and Richardson, C. J.: Relative radial oxygen loss in five
wetland plants, in: Constructed Wetlands for Wastewater Treatment, edited
by: Hammer, D. A., Lewis Publishers, Chelsea, USA, 501–507, 1989.Millero, F. J.: The effect of ionic interactions on the oxidation of metals
in natural waters, Geochim. Cosmochim. Ac., 49, 547–53,
10.1016/0016-7037(85)90046-8, 1985.Oba, Y. and Poulson, S. R.: Oxygen isotope fractionation of dissolved
oxygen during reduction by ferrous iron, Geochim. Cosmochim. Ac., 73, 13–24,
10.1016/j.gca.2008.10.012, 2009a.Oba, Y. and Poulson, S. R.: Oxygen isotope fractionation of dissolved oxygen
during abiological reduction by aqueous sulfide, Chem Geol., 268, 226–232,
10.1016/j.chemgeo.2009.09.002, 2009b.Park, B. and Dempsey, B. A.: Heterogeneous oxidation of Fe(II) on ferric
oxide at neutral pH and a low partial pressure of O2, Environ. Sci.
Technol., 39, 6494–6500, 10.1021/es0501058,
2005.Parker, S. R., Poulson, S. R., Gammons, C. H., and DeGrandpre, M. D.:
Biogeochemical controls on diel cycling of stable isotopes of dissolved
oxygen and dissolved inorganic carbon in the Big Hole River, Montana,
Environ. Sci. Technol., 39, 7134–7140,
10.1021/es0505595, 2005.Parker, S. R., Gammons, C. H., Poulson, S. R., DeGrandpre, M. D., Weyer, C.
L., Smith, M. G., Babcock, J. N., and Oba, Y.: Diel behavior of stable
isotopes of dissolved oxygen and dissolved inorganic carbon in rivers over a
range of trophic conditions, and in a mesocosm experiment, Chem Geol., 269,
22–32, 10.1016/j.chemgeo.2009.06.016, 2010.Parker, S. R., Gammons, C. H., Smith, M. G., and Poulson, S. R.: Behavior of
stable isotopes of dissolved oxygen, dissolved inorganic carbon and nitrate
in groundwater at a former wood treatment facility containing hydrocarbon
contamination, Appl. Geochem., 27, 1101–1110,
10.1016/j.apgeochem.2012.02.035, 2012.Parker, S. R., Darvis, M. N., Poulson, S. R., Gammons, C. H., and Stanford,
J. A.: Dissolved oxygen and dissolved inorganic carbon stable isotope
composition and concentration fluxes across several shallow floodplain
aquifers and in a diffusion experiment, Biogeochemistry, 117, 539–552,
10.1007/s10533-013-9899-0, 2014.Parkhurst, D. L. and Appelo, C. A. J.: Description of input and examples
for PHREEQC version 3, A computer program for speciation, batch-reaction,
one-dimensional transport, and inverse geochemical calculations, Volume book
6, series Techniques and Methods [code], 1–327, available at: https://www.usgs.gov/software/phreeqc-version-3 (last access: 26 July 2021), 2009.Pati, S. G., Bolotin, J., Brennwald, M. S., Kohler, H. P. E., Werner, R. A.,
and Hofstetter, T. B.: Measurement of oxygen isotope ratios
(18O /16O) of aqueous O2 in small samples by gas
chromatography/isotope ratio mass spectrometry, Rapid Commun. Mass
Sp., 30, 684–690, 10.1002/rcm.7481,
2016.Pusch, M.: The metabolism of organic matter in the hyporheic zone of a
mountain stream, and its spatial distribution, Hydrobiologia, 323, 107–118,
10.1007/BF00017588, 1996.Quay, P. D., Wilbur, D. O., Richey, J. E., Devol, A. H., Benner, R., and
Forsberg, B. R.: The 18O : 16O of dissolved oxygen in rivers and
lakes in the Amazon Basin: Determining the ratio of respiration to
photosynthesis rates in freshwater, Limnol. Oceanogr., 40, 718–729,
10.4319/lo.1995.40.4.0718, 1995.
Schwab, R. G.: Die natürliche Radioaktivität der Erdkruste, in:
Natürliche und künstliche Strahlung in der Umwelt. Eine Bilanz vor
und nach Tschernobyl, edited by: Hosemann, G., and Wirth, E., Erlanger
Forschungen Reihe B, Erlangen, Germany, 25–43, 1987.Schwertmann, U.: Solubility and dissolution of iron oxides, Plant Soil, 130,
1–25, 10.1007/BF00011851, 1991.Sylvester, P., Westerhoff, P., Boyd, O., and Sengupta, A. K.: Arsen Xnp – A new hybrid sorbent for arsenic removal from drinking water, in: ACE
′05, Proceedings of the AWWA Annual Conference and Exposition, San
Francisco, USA, 2005.Skinner, B. J.: A Second Iron Age Ahead?, Res. J. Environ. Sci., 3, 559–575,
10.1016/S0166-1116(08)71071-9, 1979.Smith, L., Watzin, M. C., and Druschel, G.: Relating sediment phosphorus
mobility to seasonal and diel redox fluctuations at the sediment-water
interface in a eutrophic freshwater lake, Limnol. Oceanogr., 56, 2251–2264,
10.4319/lo.2011.56.6.2251, 2011.Stal, L. J.: Cyanobacterial mats and stromatolites, in: The Ecology of Cyanobacteria II: their diversity in space and time, edited by: Whitton, B. A., Springer, Dordrecht, New York, London, 65–125, 10.1007/978-94-007-3855-3, 2012.Stanier, R. Y., Kunisawa R., Mandel, M., and Cohen-Bazire, G.: Purification
and properties of unicellular blue-green algae (order Chroococcales),
Bacteriol. Rev., 35, 171–205,
10.1128/mmbr.35.2.171-205.1971, 1971.Swanner, E. D., Mloszewska, A. M., Cirpka, O. A., Schoenberg, R., Konhauser,
K. O., and Kappler, A.: Modulation of oxygen production in Archaean oceans
by episodes of Fe(II) toxicity, Nat. Geosci., 8, 126–130,
10.1038/ngeo2327, 126–130, 2015.
Taylor, B. E. and Wheeler, M. C.: Sulfur- and Oxygen-Isotope Geochemistry
of Acid Mine Drainage in the Western United States, in: Environmental
Geochemistry of Sulfide Oxidation edited by: Alpers, C. N. and Blowes, D.
W., American Chemical Society Symposium Series, Washington DC, USA, 481–514,
1993.Teal, J. M. and Kanwisher, J. W.: Gas Transport in the Marsh Grass, Spartina
alterniflora, J. Exp. Bot., 17, 355–361,
10.1093/jxb/17.2.355, 1966.van Geldern, R. and Barth, J. A. C.: Optimization of instrument setup and
post-run corrections for oxygen and hydrogen stable isotope measurements of
water by isotope ratio infrared spectroscopy (IRIS), Limnol.
Oceanogr. Method., 10, 1024–1036,
10.4319/lom.2012.10.1024, 2012.
Wang, X. and Veizer, J.: Respiration-photosynthesis balance of terrestrial
aquatic ecosystems, Ottawa area, Canada, Geochim. Cosmochim. Acta., 64, 3775–3786,
10.1016/S0016-7037(00)00477-4, 2000.Wassenaar, L. I. and Koehler, G.: An on-line technique for the
determination of the δ18O and δ17O of gaseous and
dissolved oxygen, Anal. Chem., 71, 4965–4968,
10.1021/ac9903961, 1999.Wassenaar, L. I. and Hendry, M. J.: Dynamics and stable isotope composition
of gaseous and dissolved oxygen, Ground Water, 45, 447–460,
10.1111/j.1745-6584.2007.00328.x, 2007.Weyhenmeyer, G. A., Prairie, Y. T., and Tranvik, L. J.: Browning of Boreal
Freshwaters Coupled to Carbon-Iron Interactions along the Aquatic Continuum,
PLoS One, 9, e88104,
10.1371/journal.pone.0088104, 2014.Wilmotte, A., Van der Auwera, G., and De Wachter, R.: Structure of the 16S
ribosomal RNA of the thermophilic cyanobacterium Chlorogloeopsis HTF (Mastigocladus laminosus HTF') strain PCC75
18, and phylogenetic analysis, FEBS Lett., 317, 96–100,
10.1016/0014-5793(93)81499-p, 1993.Zhang, Y., Charlet, L., and Schindler P. W.: Adsorption of protons, Fe(II)
and Al(III) on lepidocrocite (γ-FeOOH), Colloids Surf., 63, 259–268,
10.1016/0166-6622(92)80247-Y, 1992.