Multiple stressors effects on nitrate uptake vary across benthic and hyporheic compartments

doi:10.21203/rs.3.rs-3121299/v1

Download PDF

Research Article

Multiple stressors effects on nitrate uptake vary across benthic and hyporheic compartments

https://doi.org/10.21203/rs.3.rs-3121299/v1

This work is licensed under a CC BY 4.0 License

Journal Publication

published 13 Mar, 2024

Read the published version in Biogeochemistry →

You are reading this latest preprint version

Agricultural land use strongly alters nitrate (NO3) dynamics in headwater streams, but the specific mechanisms linking agricultural stressors to benthic and hyporheic NO3 uptake remain unclear. Using stream-side mesocosms and 15N-NO3 additions, we examined the individual and combined effects of fine sediment and eutrophication (i.e., increased phosphorus and light levels) on NO3 uptake in the benthic and hyporheic compartment. Eutrophication increased benthic uptake rates by 12-fold compared to the control, as phosphorus and light additions stimulated biofilm growth. Eutrophication increased hyporheic NO3 uptake by 7-fold relative to the control, this was likely due to enhanced heterotrophic uptake, which benefited from phosphorus and dissolved organic material exudated by benthic algae. The fine sediment treatment did not change benthic uptake relative to the control but increased hyporheic uptake by 14-fold. This was due to anoxic conditions, which may have stimulated hyporheic denitrification. In the combined treatment, eutrophication exerted dominance effects in the benthic compartment, while we found antagonistic stressor interactions in the hyporheic compartment.

Our findings indicate that the significant effects of agriculture on NO3 uptake observed previously in field conditions may be primarily attributed to eutrophication and only marginally to other stressors, such as fine sediment. Moreover, our compartment-specific results imply that results obtained in the benthic compartment can not be transferred to the hyporheic compartment. We advocate a compartment-specific approach when quantifying stressor effects on NO3 uptake. Such approaches will help to increase the accuracy of effect size estimates, which are essential for managing functional attributes of streams subjected to agricultural land use.

multiple stressors

agriculture

compartments

antagonism

stressor dominance

Past and present agricultural activities have exposed streams and rivers to multiple anthropogenic stressors, drastically altering their natural chemical and hydro-morphological conditions (Allan 2004). Nutrient inputs from agricultural and urban sour have altered natural stochiometric ratios (Westphal et al. 2020; Wymore et al. 2016) and caused nutrient concentrations to rise above pristine levels (Bernhardt et al. 2017). Practices such as soil tillage, drainage channelization, and riparian clear-cutting have further deteriorated the morphological conditions of streams, increasing streambed light exposure (Tank et al. 2021) and deposition of fine sediment on its surface (U.S. EPA 2000).

By altering the environmental conditions, agricultural land use has impaired stream ecosystem structure (Schürings et al. 2022) and functions, with NO3 uptake being the most strongly affected function (Brauns et al. 2022). In fact, previous studies employing ¹⁵N-NO3 tracer techniques across various biomes and land uses have consistently shown that agricultural streams have lower NO3 uptake efficiency but higher NO3 areal uptake than reference ones (Mulholland et al. 2008).

Microbial biofilms play a crucial role in many stream ecosystem functions, including NO3 uptake (Findlay and Battin 2015), and human-induced changes in light, nutrient enrichment, and fine-sediment clogging may strongly interact in their effect on stream biofilm N-NO3 uptake.

In streams, NO3 uptake occurs largely in both the benthic and hyporheic biofilms. Research on NO3 uptake has predominantly focused on the benthic compartment and our understanding of the functioning and contribution of the hyporheic compartment to total NO3 uptake is limited (but see Böhlke et al. 2004, 2009; Roche et al. 2019; Zarnetske et al. 2015). It has been suggested that some of the ¹⁵N-NO3 unaccounted for by the benthic zone could be due to overlooked hyporheic NO3 uptake (Böhlke et al. 2004; Mulholland et al. 2004a, 2008). Retention of NO3 in the hyporheic compartment can occur through assimilatory or dissimilatory processes, depending on the environmental conditions and the microbial community. Oxygenated hyporheic compartments promote aerobic processes such as assimilatory uptake by hyporheic (heterotrophic) biofilms. However, the quality and quantity of organic matter (Zeglin 2015) and the availability of other nutrients (e.g., phosphorus) can influence uptake rates. For example, carbon from benthic algal metabolites is quickly assimilated, while other components, such as lignin, cellulose, and lignocellulose, mainly from terrestrial ecosystems, are more recalcitrant. While in anoxic conditions, NO3 can be potentially removed through dissimilatory pathways, i.e., denitrification or dissimilatory nitrate reduction to ammonium (DNRA) (Arango and Tank 2008; Triska et al. 1989).

When investigating the effect of agricultural land use, it has to be recognized that agriculture is a composite stressor consisting of several individual stressors, e.g., increased nutrient concentration, light, fine sediment deposition, etc. Thus, in field conditions, it is difficult to discern the effects of each stressor on a measured variable and to understand the mechanisms through which a stressor operates. This is because stressors can interact in various ways, potentially intensifying or reducing individual responses or even producing contrasting effects that cancel each other out (Jackson et al. 2016; Morris et al. 2022).

For example, the high light and nutrient availability in agricultural streams increase gross primary production (GPP), resulting in shorter NO3 uptake lengths (Hall and Tank 2003). However, the simultaneous presence of elevated NO3 concentration counteracts this effect, lengthening NO3 uptake lengths (Hall and Tank 2003). Thereby the stressors counterbalance each other's effects and obscure the impact of agricultural land use on NO3 uptake dynamics. High nutrients and light are also known to directly affect biofilm structure(Niyogi et al. 2004; Passy 2007; Proia et al. 2012) and functioning. For example, the stochiometric imbalances due to the lower ratio of dissolved organic carbon to nitrate concentrations (DOC:NO3) in agricultural streams can reduce biofilm NO3 uptake efficiencies (Mulholland et al. 2009; Wymore et al. 2019). Other types of stressors, such as fine sediment deposition, reduce biofilm growth (Myers et al. 2007), even when light and nutrient availability are high(Atkinson et al. 2008), because fine sediment provides an unstable substrate which reduces biofilm development. In addition, fine sediment deposition can also indirectly affect biofilm functioning by changing the environmental conditions. For example, the fine sediment deposition alters the water exchange between the benthic and hyporheic compartments by quickly depleting the oxygen in the hyporheic compartment. This shift to low-oxygen environments can trigger dissimilatory processes, particularly in the presence of high NO3 concentrations, denitrification processes (Böhlke et al. 2004; Holmes et al. 1996).

So far, due to the co-occurrence of multiple stressors associated with agricultural land use and their unknown interactions, we were not able to disentangle the individual and interactive effects of those stressors on NO3 uptake. Thus, in this study, we quantified the individual and combined effects of two of the most common agricultural stressors, i.e., fine sediment and eutrophication (simultaneous increase in P and light levels) on compartmental (i.e., benthic and hyporheic) NO3 uptake rates in stream-side mesocosms.

We hypothesized that:

Eutrophication will increase NO3 uptake in the benthic compartment because light and P additions stimulate the development and uptake of benthic biofilms;
Likewise, eutrophication by P and light addition will increase hyporheic NO3 uptake as heterotrophic bacteria are stimulated due to both direct (enhanced P) and indirect (exudates from benthic algae) resource effects;
Fine sediment will reduce the benthic NO3 uptake as fine sediment provides a less stable substratum for biofilms reducing their biomass and uptake;
Also, hyporheic NO3 uptake will be lower in the treatment with fine sediment compared to the treatment with gravel. This is because the presence of fine sediment reduces the delivery of water and solutes to the hyporheic compartment, thus reducing biofilm uptake;
Combined effects of eutrophication and fine sediment on NO3 uptake will be additive in the benthic and hyporheic compartments, i.e., negative effects of fine sediment will reduce the positive effect of eutrophication on NO3 uptake.

Experimental setup

The experiment was performed in a stream-side mobile mesocosm facility (MOBICOS) (Fink et al. 2020) located at the forested, upstream part of the Holtemme River (51° 49′ 00.7″ N, 10° 43′ 29.26″ E, Germany). This river section is characterized by a closed canopy and a pristine hydromorphology, moderate NO3 concentrations, and low soluble reactive phosphorus (SRP) concentrations compared to reaches further downstream, which are influenced by urban centers and agriculture (Weitere et al. 2021). Surface water was continuously supplied to the flumes by a submersible pump (type XJ 40, ABS Sulzer, Germany). The inflowing water was filtered with a 50 µm self-cleaning filter (GEFA Process Technik GmbH, Model AP3 DA) to prevent the inflow of suspended particles and macroinvertebrates. The mesocosm setup comprised 24 flumes (72 cm length, 14 cm height, 8 cm width) (Online resource 1, Fig. 1) filled with commercially available sediment previously washed and dried at 120°C. The sediment height was 8 cm allowing for a benthic (0–2 cm depth) and a hyporheic compartment (2–8 cm depth). The height of the water column was 4 cm. Four ceramic tiles (6 x 1 x 0.5 cm) were equidistantly placed in each flume perpendicularly with the flow to facilitate water infiltration into the sediment. Discharge was set to a constant flow rate of 0.17 ± 0.006 L ${s}^{-1}$ in each flume and continuously monitored with a magnetic inductive flow meter (IFM, model: SM8000). Water passed through a twin-wall polycarbonate honeycomb with an opening diameter of 5 mm to ensure laminar flow. The light was provided using one elongated LED lamp (Solar Stinger daylight 16W) placed above each flume, and light: dark cycles were set to 10:14 h.

Treatments

We assigned the 24 flumes to four treatments: control, fine sediment, eutrophication, and fine sediment + eutrophication (Table 1). In the control treatment, we mimicked the environmental conditions found in the Holtemme River by using similar-sized gravel (2–4 mm) and comparable light intensities (i.e., 7.5 ± 1.5 µmol photons ${m}^{-2}{s}^{-1}$). The fine sediment treatment was created by using fine sand (grain size 0.1–0.4 mm). The eutrophication treatment was created by increasing the light intensity from 7.5 ± 1.5 µmol photons ${m}^{-2}{s}^{-1}$ to 51.2 ± 6.5 µmol photons ${m}^{-2}{s}^{-1}$ and increasing soluble reactive phosporous (SRP) concentrations from below the quantification limit (< 0.003 $mg {l}^{-1}$) to 0.04 mg ${l}^{-1}$ by constantly supplying a KH2PO4 solution with a peristaltic pump (Watson-Marlow 205U, Falmouth, UK). To test the effect of stressors interaction, we simultaneously manipulated substrate, SRP (from now on “P”), and light in the eutrophication + fine sediment treatment (Table 1). Each treatment was replicated with six flumes. Flumes were operated under these conditions in a constant flowing mode for two weeks.

In the hyporheic compartment, not only P but also DOC limitation can strongly influence heterotrophic microbial growth because local, autotrophic organic carbon (OC) production can be negligible (Stutter et al. 2020). We expected this to be of little relevance to our experiment due to the high bulk DOC concentrations in the stream. To make sure that DOC limitation was not the case, we tested in an auxiliary experiment with laboratory microcosms to which extent N-NO3 uptake was potentially limited by DOC availability. We used a protocol similar to Graeber et al. 2021. Briefly, we sampled water from each flume outlet once after two weeks of flume colonization and measured the uptake of DOC, dissolved N, and dissolved P fractions in a bioavailability experiment in the dark for an incubation time of eight days. Based on this, we calculated the reactive DOC, reactive dissolved N, and reactive dissolved P concentrations and calculated the C : N : P ratio of reactive, hence microbially available macronutrients. To show the potential reactive DOC or P limitation of heterotrophic NO3 uptake, we used a ternary plot approach using the ggtern package in R (Graeber et al. 2021), in which we normalized the C : N : P to the median of a large analysis of bacterial C : N : P ratios (Godwin and Cotner 2018). We also assessed potential changes in dissolved organic matter (DOM) molecular composition via fluorescence spectroscopy and parallel factor analysis using the staRdom toolbox in R (Pucher et al. 2019). A detailed protocol and results can be found in Online resource 2.

Table 1

Summary of conditions found in the flumes assigned to the different treatments. Values are mean (n = 5) and standard deviation except for sediment grain size (min-max)
	Control	Eutrophication	Fine sediment	Eutrophication+ fine sediment
Sediment grain size (mm)	2–4 (Gravel)	2–4 (Gravel)	0.1–0.4 (Fine sand)	0.1–0.4 (Fine sand)
Light intensity (µmol photons ${m}^{-2}{s}^{-1}$)	7.5 ± 1.5	51.2 ± 6.5	7.5 ± 1.5	51.2 ± 6.5
DOC (${mgl}^{-1}$)	9.4 ± 0.23	9.38 ± 0.21	9.41 ± 0.15	9.39 ± 0.23
SRP (${mgl}^{-1}$)	≤ 0.003 ± 0.00	0.04 ± 0.00	≤ 0.003 ± 0.00	0.04 ± 0.00
N-N-NO3 (${mgl}^{-1}$)	1.05 ± 0.01	1.04 ± 0.00	1.05 ± 0.01	1.05 ± 0.01

Tracer addition, sampling, and analyses

After two weeks of colonization, one flume per treatment was sampled to determine ambient isotope values (15N atom fraction, x(15N)) of benthic and hyporheic organic matter (OM). Subsequently, we added an enriched K15N-NO3 solution constantly for 24 h into the turbulent part of the flumes’ inlet section with syringe pumps (NE-1200 Twelve Channel Programmable Syringe Pump, New Era Pump Systems, Inc.). The addition was adjusted to achieve a target x(15N) of 30. After the addition, we collected sediment samples from the benthic and hyporheic compartments of each flume. To prevent sediment mixing between the benthic and hyporheic compartments, we first shoveled the upper 2 cm of sediment and stored it in a sealed container (benthic sample). We then collected a sediment sample from the hyporheic compartment and stored it in another sealed container (hyporheic sample). To separate OM from the mineral substrate, we rinsed the samples multiple times with water and passed them through a 100 µm sieve. The benthic samples exhibited visibly higher OM content compared to the hyporheic samples. Therefore, a larger volume of sediment from the hyporheic samples had to be processed to obtain sufficient OM for further analysis. The processed sediment was transported to the laboratory, where it was dried at 60°C for a minimum of 24 h and then weighed. To determine the volume of the sediment, the weight was divided by the specific density of the sediment, which was measured in the lab using a pycnometer (Blaubrand™).

In the laboratory, OM was freeze-dried (Delta 2–24 LSCplus) for at least 24 h, homogenized, and three replicates of each sample was weighed into tin caps (HEKAtech, Germany) for isotope analysis. Stable nitrogen isotopic composition was analyzed on an elemental analyzer (Flash 2000 Organic Elemental Analyzer, Thermo Fisher Scientific, Germany), directly connected via an open split system (ConFlo IV, Thermo Fisher Scientific, Germany) to an isotope ratio mass spectrometer (Delta V Advantage IRMS, Thermo Fisher Scientific, Germany).

During the analysis, only a minor enrichment was found (i.e., maximum x(15N) = 6.38). Thus, the normalization was done by analyzing the reference materials IAEA-311 (x(15N) = 2.05) (Parr and Clements 1991) and AS-3 (x(15N) = 0.36) (Kornexl et al. 1999) and applying a two-point calibration approach. The analytical precision was below ± 0.2.

Particulate organic carbon (POC) and particulate nitrogen (PN) concentrations of the OM were measured with an elemental analyzer (Elementar Analysen Systeme GmbH, Hanau).

Calculation of N uptake

N-NO3 uptake by benthic and hyporheic OM was calculated from sediment samples collected before and after the isotope tracer addition following Mulholland (2004b). Briefly, we corrected all 15N values by the background values and calculated the excess atom fraction (AFE). Then, we calculated the areal uptake rate ${U}_{Areal}$ and biomass uptake ${U}_{Biomass}$. For that, we first calculated the amount of tracer in the OM ($15{N}_{OM}$, g 15N):

$$15{N}_{OM}=OM\times \left(\frac{\%N}{100}\right)\times {AFE}_{OM}$$

where OM (g dry weight) is the organic matter stock within the samples, %N is the nitrogen elemental concentrations (%) of the OM, and ${AFE}_{OM}$ the excess 15N fraction of OM.

Then, we scaled the result by sediment volume (${15N}_{Sed}$, g 15N ${cm}^{-3}$):

$${15N}_{Sed} = \frac{{15N}_{OM}}{V}$$

Where V is the volume of sediment from which the OM was collected (${ cm}^{-3}$). The volume of sediment was calculated by dividing the dry weight (W $g$) by the sediment density (d $g{ cm}^{-3}$) as:

$$V=\frac{W}{d}$$

Where d, i.e. sand density = 2.76 $g {cm}^{-3}$, gravel density = 2.54 $g {cm}^{-3}$.

Subsequently, we calculated the N-NO3 uptake rate of the sediment (${U}_{sed}$ g 15 N ${cm}^{-3}$ ${d}^{-1}$):

$${{U}_{sed}}_{}=\frac{{15N}_{Sed}}{\left(\frac{{15N}_{Flux}}{{N}_{flux}}\right)}$$

where

$$15NFlux=\left({AFE}_{w,a}\text{*}Q\text{*}C)-({AFE}_{w,b}\text{*}Q\text{*}C\right)$$

$${N}_{flux}=Q\text{*}C$$

$15NFlux$ is the tracer mass flux, where we subtracted from the total mass flux $\left({AFE}_{w,a}\right)$ the background mass flux $\left({AFE}_{w,b}\right)$, C is the N-NO3 concentration (mg ${l}^{-1}$) and Q is flume discharge (${l s}^{-1}$).

We expressed uptake on an areal basis (${U}_{Areal}$, g N-NO3 ${m}^{-2}{d}^{-1}$), i.e. N-NO3 uptake of a 1 ${m}^{2}$ surface with a height of 1 cm by multiplying the ${U}_{sed}$ by 10,000.

We calculated biomass uptake (${U}_{Biomass}$ g N-NO3 ${{g}^{-1}}_{OM}$ ${d}^{-1}$), i.e. N-NO3 uptake of 1 gram of OM, by dividing the areal uptake by the amount of OM found in that volume as:

$${U}_{Biomass}= \frac{{U}_{Areal}}{OM}$$

We finally summed the areal uptake of the benthic and hyporheic compartments to obtain the total areal uptake (${U}_{tot}$ g N-NO3 ${m}^{-2}{d}^{-1}$).

Chlorophyll-a and microbial density

Chlorophyll-a concentrations in the sediment were measured from benthic samples after the two weeks of colonization. Sediment samples were immediately frozen at -20°C and chlorophyll-a was analyzed via high-performance liquid chromatography with a Thermo Scientific UltiMate 3000 HPLC System (Dionex, Thermo Fisher Scientific Corporation, Waltham, MA, USA). Bacterial abundance was measured by collecting a sediment sample from the benthic and hyporheic compartments of each flume in sterile Falcon tubes. To detach cells from the sediment, we used established protocols as described by Chen et al. (2021). Then cells were stained with DAPI and counted using an epifluorescence microscope at 100x magnification. A microscopic investigation was performed using a Zeiss AxioImager.Z2 epifluorescence microscope equipped with an HXP R 120W/45C UV Hg-vapor lamp, Colibri.2 LED illuminations and the following fluorescence filters: DAPI (365/10 nm excitation, 420 LP emission, FT 395 Beam Splitter), Alexa488 (472/30 excitation, 520/35 emission, 495 Beam Splitter), and Alexa594 (562/40 excitation, 624/40 emission, 593 Beam Splitter). The filter sets discriminated clearly between DAPI (365 nm) and natural autofluorescence of the diatom cells. Imaging was done with 100X oil objective numerical aperture N:A 1.4. The software for image acquisition allows for overlapping images acquired successively with different filter sets (Zen software from Carl Zeiss). Images are reported in Online resource 3.

Physical-chemical parameters monitoring

Light intensity was measured with a frequency of five minutes using a light intensity data logger MX2202 (Onset, Bourne, Massachusetts, USA) placed on the sediment of three flumes per treatment. Planar oxygen sensors (SP-PSt6-YAU, PreSens Precision Sensing GmbH, Germany) were glued to the inner wall of three flumes for each treatment to monitor oxygen concentration. Oxygen was measured every 25 seconds at two different depths: 2 cm (benthic) and 6 cm (hyporheic). Some of the cables connecting the planar sensors to the oxygen readers detached from the wall during the measurements, resulting in incomplete temporal data for certain treatments (specifically, 3 out of 9 incomplete datasets for the benthic and 2 out of 9 incomplete datasets for the hyporheic compartment). To ensure comparability of data among treatments, we selected the oxygen data from two flumes per treatment for each compartment where the oxygen data was complete and uninterrupted for a minimum of seven consecutive days (Online resource 1, Fig. 2 and Fig. 3). Water samples for N-NO3, nitrite (N-NO2), ammonium (N-NH4), reactive phosphorous (SRP) were collected and analyzed as described in Online resource 1.

Statistical analysis

To assess treatment effects on N-NO3 areal uptake, biomass uptake, OM, POC, PON, bacterial abundance, and chlorophyll-a, we applied a type I analysis of variance (ANOVA package car, function aov) and a Tukey HSD post-hoc test separately for the benthic and hyporheic compartment. Data were ln(x) transformed where necessary to approximate assumptions of normality and homogeneity of variance of residuals. Values exceeding three times the standard deviation were classified as outliers and removed from the analysis. This happened for four variables: benthic bacterial abundance (2 out of 20), hyporheic POC content (2 out of 20), hyporheic PON (2 out of 20), and benthic OM (2 out of 20). All tests were performed in R (R Core Team 2021).

Treatment effects on compartmental N-NO3 uptake

Benthic

In the benthic compartment, we found significant effects of eutrophication (Table 2), with 12- and 15-fold increases in N-NO3 areal uptake and biomass uptake in the eutrophication treatment compared to the control (Fig. 1a and 1c, Online resource 1 Table 1). Benthic areal uptake in the fine sediment treatments was on average 2.8-fold lower than in control (Fig. 1a), even though the effect of fine sediment was not significant (Table 2). The benthic compartment of the combined treatment showed no significant difference in areal and biomass uptake compared to the eutrophication treatment (Fig. 1a and Fig. 1c), and stressors did not interact significantly (Table 2). Areal and biomass uptake were significantly higher in the benthic compartment of the combined treatment than in the fine sediment and control treatment (Fig. 1a and 1c, Online resource 1 Table 1).

Hyporheic

In the hyporheic compartment, we found significant effects of fine sediment (Table 2) with a ~ 14-fold increase in areal and biomass uptake compared to the control (Fig. 1b and 1d, Online resource 1 Table 1). In contrast, eutrophication did not have a significant overall effect on areal uptake (Table 2) because it significantly interacted with fine sediment. In comparison to the control, eutrophication increased areal uptake by 7-fold. While in comparison to the fine sediment, eutrophication decreased by 2-fold areal uptake in the combined treatment (see the result of Tukey post hoc test in Fig. 1b, Online resource 1 Table 1).

Table 2

Summary of ANOVA results comparing response parameters to eutrophication, fine sediment, and their interaction in the benthic (B) and the hyporheic compartment (H). Shown are F-values and p-values in brackets. In the compartment column, a indicates that data were ln(x) transformed, b indicates that outliers were removed. When the results of the statistical tests were found to be significant at an alpha level of 0.05, an arrow was added to show the effect direction in relation to the control (i.e., ↑ increase, ↓ decrease).
Parameters	Compartment	Eutrophication	Fine sediment	Eutrophication x Fine sediment
N-NO3 areal uptake	${B}^{a}$	92.83 (< 0.001) ↑	0.50 (0.49)	2.1 (0.17)
N-NO3 areal uptake	H	0.01 (0.95)	40.33 (< 0.001) ↑	38.21 (< 0.001) ↑
N-NO3 biomass uptake	${B}^{a}$	301.03 (< 0.001) ↑	0.21 (0.65)	0.75 (0.40)
N-NO3 biomass uptake	${H}^{a}$	16.40 (< 0.001) ↑	78.40 (< 0.001) ↑	23.66 (< 0.001) ↑
Organic matter (OM)	${B}^{ab}$	0.003 (0.95)	0.06 (0.81)	4.08 (0.06)
Organic matter (OM)	${H}^{a}$	3.61 (0.07)	22.80 (< 0.001) ↓	3.45 (0.08)
Particulate organic carbon (POC)	${B}^{b}$	26.13 (< 0.001) ↑	3.24 (0.09)	0.72 (0.41)
Particulate organic carbon (POC)	${H}^{ab}$	2.67 (0.12)	4.92 (< 0.05) ↓	4.76 (< 0.05) ↓
Particulate organic nitrogen (PON)	${B}^{a}$	133.57 (< 0.001) ↑	15.14 (< 0.001) ↓	1.37 (0.26)
Particulate organic nitrogen (PON)	${H}^{ab}$	12.54 (< 0.05) ↓	4.13 (0.06)	17.08 (< 0.001) ↓
Chlorophyll-a	${B}^{a}$	103.21 (< 0.001) ↑	1.35 (0.26)	2.32 (0.15)
Bacterial abundance	${B}^{ab}$	3.7 (0.07)	0.03 (0.87)	0.47 (0.50)
Bacterial abundance	${H}^{a}$	0.35 (0.56)	0.75 (0.40)	9.35 (< 0.05) ↓

Macronutrient stoichiometry, chlorophyll-a, bacterial density, and oxygen levels

In the benthic compartment, eutrophication increased significantly chlorophyll-a concentrations while it had no significant effect on bacterial abundances (Table 2 and Table 3). In contrast, fine sediment had no significant effect on both parameters (Table 2 and Table 3). We did not find any significant stressor interaction in the benthic compartment (Table 2). In the combined treatment, the chlorophyll-a content was comparable to the eutrophication treatment and 5-fold higher than in the control and fine sediment treatments (Table 3). Fine sediment had a significant negative effect on OM content in the hyporheic compartment (Table 2), which was 1.3- fold lower than in the control and comparable to the OM content in the combined treatment (Table 3).

In the hyporheic compartment of the combined treatment, we found that stressors interacted and significantly decreased PON content compared to the control (Table 2 and Table 3).

The benthic compartment of all the treatments was oxygenated through the entire duration of the measurements (Table 3 and Online resource 1 Fig. 2). In the hyporheic compartment of the treatments with fine sediment (fine sediment and combined), we measured hypoxic and anoxic conditions (Table 3 and Online resource 1 Fig. 3). In contrast, the hyporheic compartment of the flumes with a gravel bed (control and eutrophication treatment) remained oxygenated throughout the entire experiment duration (Table 3 and Online resource 1 Fig. 3), although oxygen concentrations were lower than in the benthic compartment.

Table 3

Parameters measured in the different treatments in the benthic (B) and hyporheic compartments (H). Values are mean and standard deviations from the five flumes per treatment, except for the oxygen concentration, which was obtained by averaging the oxygen values measured every 25 seconds over seven consecutive days from two flumes per treatment. a indicates that data were ln(x) transformed, b indicates that outliers were removed. Treatments that share the same letter are not significantly different at the 0.05 level (Tukey post hoc test). No statistical test was run for oxygen concentration.
Parameters	Compartment	Control	Eutrophication	Fine sediment	Fine sediment + eutrophication
Organic matter (OM) (mg ${cm}^{-3}$)	${B}^{ab}$	154 ± 77 (a)	98 ± 40 (a)	94 ± 20 (a)	139 ± 35 (a)
Organic matter (OM) (mg ${cm}^{-3}$)	${H}^{a}$	9 ± 2 (a)	9 ± 3 (a)	6 ± 2 (ab)	3 ± 1 (b)
Particulate organic carbon (POC) (µg ${g}^{-1}$ OM)	${B}^{b}$	7.8 ± 0.3 (a)	9.5 ± 0.8 (b)	7.6 ± 0.5 (a)	8.7 ± 0.6 (ab)
Particulate organic carbon (POC) (µg ${g}^{-1}$ OM)	${H}^{ab}$	1.8 ± 0.4 (a)	1.3 ± 0.3 (ab)	1.2 ± 0.2 (b)	1.3 ± 0.2 (ab)
Particulate organic nitrogen (PON) (µg ${g}^{-1}$ OM)	${B}^{a}$	11.8 ± 1.4 (a)	24 ± 3.9 (c)	10.2 ± 0.5 (a)	18.1 ± 1.7 (b)
Particulate organic nitrogen (PON) (µg ${g}^{-1}$ OM)	${H}^{ab}$	10 ± 0.8 (a)	6.4 ± 0.6 (b)	7.1 ± 0.6 (b)	7.3 ± 1.3 (b)
Chlorophyll-a (µg ${cm}^{-3}$ )	${B}^{a}$	0.9 ± 0.3 (a)	4.6 ± 2.0 (b)	0.6 ± 0.2 (a)	5.0 ± 2.6 (b)
Bacterial abundance $\left(\text{c}\text{o}\text{u}\text{n}\text{t}\text{s}\text{*}{10}^{6} {cm}^{-3} \text{s}\text{e}\text{d}\text{i}\text{m}\text{e}\text{n}\text{t}\right)$	${B}^{ab}$	4.3 ± 3.1 (a)	3.2 ± 1.7 (a)	4.9 ± 3 (a)	2.3 ± 0.9 (a)
	${H}^{a}$	2.9 ± 1.7 (a)	6.6 ± 6.4 (a)	9.6 ± 6.3 (a)	2.4 ± 0.8 (a)
Oxygen concentration (mg ${l}^{-1}$)	B	7.0 ± 0.3	7.3 ± 0.1	5.6 ± 0.6	7.6 ± 0.6
Oxygen concentration (mg ${l}^{-1}$)	H	7.1 ± 0.3	7.3 ± 0.4	0.15 ± 0.07	0 ± 0

Auxiliary experiment on the potential DOC limitation

We measured high DOC concentrations of on average 9.4 mg ${l}^{-1}$ (Fig. 3a, Online resource 2), but low DOC reactivity – referring to DOC bioavailability for heterotrophic microorganisms - with an average of 6.9% of the DOC being taken up by microbial heterotrophs after 8 days of incubation in the dark in microcosms (Fig. 3b, Online resource 2), indicating an average concentration of microbially reactive DOC of 0.6 mg ${l}^{-1}$. We also found no removal of dissolved OM fluorophores during the incubation, instead an addition of a humic-like fluorophore (Fig. 4, Online resource 2). When combining the reactive DOC concentration with the reactive N and P (concentrations of N-NO3-N + ammonium N and SRP, and reactive dissolved organic nitrogen and P), we calculated reactive C : N : P ratios of 187 : 501 : 1 and 937 : 1326 : 1 for the control and fine-sediment treatment, respectively, and C : N : P ratios of 40 : 57 : 1 and 29 : 53 : 1 for the eutrophication and eutrophication + fine-sediment treatment, respectively (Table 3, Online resource 2). These C : N : P ratios of the microbially reactive C, N and P constituents were lower in C relative to N compared to the literature-based C : N : P average for bacteria of 64 : 14 : 1 for all treatments and lower in C relative to P for the eutrophication treatments (Fig. 5, Online resource 2).

Fine sediment inputs and eutrophication are two of the most common stressors in streams draining agricultural catchments, but their individual and combined effects on NO3 uptake are still poorly understood. Here, we manipulated P and light availability (eutrophication) together with fine sediment in a mesocosms experiment to test their individual and combined effects on NO3 uptake in the benthic and hyporheic compartments. We found that eutrophication and fine sediment individually increased NO3 uptake rates with respect to the control but through different processes in the two compartments. Moreover, stressors had interactive effects on nutrient uptake, and the interaction type differed between the benthic and hyporheic compartments.

Eutrophication increases N-NO3 uptake in both compartments – but increases the relative importance of the benthic compartment

We found a significant 12-fold increase in N-NO3 uptake in the benthic compartment of the eutrophication treatment compared to the control. Moreover, the eutrophication treatment stimulated the autotrophs resulting in a 5-fold higher benthic chlorophyll-a concentration compared to the control. Assuming that algae were the dominant mediator of NO3 uptake in the benthic compartment, this finding indicates that NO3 uptake responds more strongly to eutrophication than what is indicated by chlorophyll-a as a measure of autotrophic biomass.

Different mechanisms might explain this substantial difference. First, light and nutrient-rich conditions increase benthic biofilm biomass (Romaní et al. 2004). However, they can also induce changes in the biofilm species composition. For example, under eutrophic conditions, single-cell taxa, i.e., diatoms, can be replaced by filamentous, chain-forming green algae or cyanobacteria (Bourassa and Cattaneo 2000; McCall et al. 2017; Passy 2007). These taxa exhibit different physical characteristics, including variations in surface-to-volume ratio, nutrient uptake, and photosynthetic rates (Steinman et al. 1992). For example, larger cells have a higher nitrate uptake capacity and internal storage volume (Hillebrand et al. 2022) but a lower chlorophyll density per biovolume (Finkel et al. 2004). Thus, the higher increase in NO3 uptake, in comparison to chlorophyll-a content, could be attributed to the development of an algal community consisting of larger cells with a higher uptake and NO3 storage capacity, albeit with reduced chlorophyll-a levels. In addition, the higher phosphorous availability, together with the development of a biofilm matrix, could have stimulated heterotrophic uptake variously; for example, by internal nutrient cycling (Besemer et al. 2009; Romaní et al. 2004) and/or by providing an increased surface area for bacterial growth (Stock and Ward 1989).

We found that hyporheic N-NO3 uptake rates were 13-fold higher in the eutrophication treatment than in the control. We assume that the measured uptake is mostly attributable to assimilatory pathways, as we always measured aerobic conditions in the hyporheic compartment. The enhanced hyporheic NO3 uptake may either be associated with a release of bacteria from P limitation or an indirect relationship with the DOM exudates potentially released from the increased benthic algal production. Our auxiliary experiment showed that when considering only microbially reactive DOC, N, and P (see M. I. Graeber et al., 2021; M. I. Stutter et al., 2018 ), we found indication of severe DOC limitation of N uptake. Specifically, for all treatments, the molar C : N : P ratios of the microbially reactive C, N, and P constituents revealed lower C : N (< 1) compared to the literature-based C : N median of 4.6 for bacterial biomass (equaling 64(C) : 14(N) : 1(P) (Godwin and Cotner 2018). Furthermore, the P addition in the eutrophication treatment decreased the N : P ratio from > 500 to 53–57 and the C : P ratio from > 187 to 29–40, which are values closer to the typical bacterial N : P ratio = 14 and C : P ratios = 64 (Godwin and Cotner 2018, Fig. 5, Supplementary material 2). This suggests that heterotrophic uptake may have been DOC and P co-limited under ambient conditions (i.e., control and fine sediment treatment), and our P additions in the treatments with eutrophication may have released this limitation. Moreover, the benthic algal community could have released DOM exudates, which increased the amount of bioavailable C that is quickly assimilated in the hyporheic compartment (Zeglin 2015). However, future studies should also assess the DOC composition change due to algal exudates and whether autochthonous DOC production alleviates allochthonous DOC limitation.

We are unaware of field studies quantifying the distinct role of the benthic and hyporheic zone in whole-stream NO3 uptake. Previous field studies concluded that as long as streams did not become NO3 saturated (Bernot et al. 2006; Dodds et al. 2002; Mulholland et al. 2008), nutrient enrichment increases whole-stream NO3 uptake and attributed it to an increased uptake in the benthic compartment (Dodds et al. 2002; Earl et al. 2006; Liu et al. 2015; Von Schiller et al. 2007). Likewise, increasing light has a stimulating effect on whole-stream NO3 uptake because of increased autotrophic uptake (Hall and Tank 2003).

In this experiment, we demonstrated that eutrophication increases benthic biofilm uptake and exacerbates the role of the benthic compartment in total NO3 uptake. However, our results also expand the limited perspective of an increase in the benthic uptake as the sole response of stream N dynamics to eutrophication by demonstrating a widespread sensitivity of the hyporheic compartment to increased nutrients and light.

Contrasting effects of fine sediment in the benthic and hyporheic compartments

We observed that the areal N-NO3 uptake and chlorophyll-a content were reduced in the fine sediment treatment compared to the control. However, the effects were statistically not significant, and the magnitude of the decrease was higher for NO3 uptake than for chlorophyll-a. A possible explanation for the lower NO3 uptake in the fine sediment treatment could be attributed to the reduced stability of fine sediment (compared to gravel), which results in reduced periphyton growth and, consequently, lower NO3 uptake (Romaní et al., 2004). However, in our experimental setup, we did not witness sediment movement, and biofilm growth was comparable between treatments exposed to similar nutrient and light conditions. Thus, we suggest that the negative effects of fine sediment on NO3 uptake might not primarily be caused by reduced substrate instability but rather by a diminished mass transfer from the water column to the benthic compartment. Fine sediment exhibits lower roughness when compared to the gravel substrate and, consequently, lower turbulence near the streambed. Previous research has demonstrated that elevated near-bed turbulence enhances the downward transport of dispersal cells through the water column to the benthic compartment (Besemer et al. 2009) and increases nutrient fluxes (Anlanger et al. 2021; Risse-Buhl et al. 2017). Thus, we suggest that the low turbulence above the fine sediment substrate might be the underlying cause for the observed lower NO3 uptake in the benthic compartment of the fine sediment treatment compared to the control. Previous studies showed that heterogeneous substrates have higher algal biomass and productivity (Cardinale et al. 2002; Hoellein et al. 2009) and phosphate uptake (Marti and Sabater 1996) than homogeneous substrates. Based on our findings, it appears that this could potentially be extended to NO3 uptake, but further research is necessary to investigate this relationship more comprehensively.

In the fine sediment treatment, we measured a 14-fold higher hyporheic N-NO3 uptake than in the control, and uptake rates were 3 times higher than in the benthic compartment. Since we measured anoxic conditions in the hyporheic compartment, we assume that the increased uptake rates are likely due to a dissimilatory process, i.e., denitrification or dissimilatory NO3 reduction to ammonium (DNRA). Previous studies under field conditions concluded that denitrification has a small contribution to total NO3 removal when compared to assimilatory pathways (Mulholland et al. 2008). Nonetheless, in agricultural streams with elevated NO3 concentrations, denitrification rates tend to increase because NO3 is not limiting (Mulholland et al. 2008). However, our forested streams, which are not anthropogenically impacted, exhibit already elevated N-NO3 concentrations (e.g., 1-1.5 mg ${l}^{-1}$N-N-NO3) due to atmospheric deposition. Hence, N-NO3 is non-limiting, and denitrification can occur whenever favorable conditions are present. This might explain why a sole variation in the substrate type (i.e., from gravel to fine sand), under ambient light and P, resulted in a 14-fold increase in the uptake rates of the hyporheic compartment.

In addition, in this treatment, there was a notable difference in the relative contribution of the two compartments to the total uptake. Specifically, in comparison to the control, fine sediment reversed the roles of the benthic and hyporheic compartments in terms of contribution to total uptake, with 70% of the entire flume uptake attributed to the hyporheic compartment in the fine sediment treatment. This has at least two implications. Firstly, the hyporheic compartment might play a major role in NO3 removal in forested streams when ambient NO3 concentrations are non-limiting and fine sediment occurs, e.g., in pools. Secondly, under these conditions, rates of dissimilatory uptake in anaerobic hyporheic sediments (i.e., pools) might be comparable to rates of assimilatory uptake measured in more heterogeneous aerobic sediments (i.e., riffles). Therefore, our findings support prior research indicating substantial variation in microbial NO3 uptake rates at small scales (Peipoch et al. 2016). However, our research emphasizes the importance of extending the assessment of the spatial distribution of microhabitats beyond the benthic compartment to include the hyporheic compartment. Only a comprehensive characterization of both compartments will allow for an accurate upscaling of biogeochemical processes in streams.

Eutrophication superposes fine sediment effects in the benthic compartment and reduces hyporheic uptake

In the benthic compartment of the combined treatment, we hypothesized an additive stressor interaction where the positive effect of eutrophication would be weakened by fine sediment. The results of the experiments demonstrate that the effect of fine sediment is negligible and that, regardless of the sediment type, under eutrophication conditions, biofilm overgrows the substratum. This resulted in a 15-fold higher uptake in the benthic compartment of the combined treatment when compared to the control. In addition, uptake rates observed in the benthic compartment of the combined treatment were comparable to the ones measured in the eutrophication treatment.

We argued before that although fine sediment does not directly affect biofilm growth, the low roughness of the substrate might reduce nutrient fluxes from the water column to the benthic compartment. However, in this experiment, we found that this effect of fine sediment is completely overridden under the increased availability of nutrients and light, and uptake rates were comparable to the eutrophication treatment. As observed in other studies, biofilms can have high architectural plasticity (Besemer et al. 2009), and the same community can develop different structures to cope with the hydrodynamic conditions and nutrient availability (Risse-Buhl et al. 2017). In particular, complex biofilm can modulate their microenvironment variously, for example, by reducing shear stress (Besemer et al. 2009) or by secreting extracellular polysaccharides (EPS) to inhibit grain movement (Valentine and Mariotti 2020). We, therefore, suggest that the high light and nutrient availability might have stimulated the formation of a large and complex biofilm, which stabilized the underneath fine sediment by excreting EPS and might have formed structures to increase the uptake of nutrients from the water column. Earlier studies have already shown the importance of light and nutrient availability on biofilm structure (Proia et al. 2012). Our study builds upon this and shows eutrophication effects on benthic N-NO3 uptake are stronger than the effects of fine sediment.

In the hyporheic compartment of the combined treatment, we measured anoxic conditions, and we expected similar dissimilarity pathways to those occurring in the fine sediment treatment. However, hyporheic uptake rates were 2-fold lower than in the fine sediment treatment, indicating that eutrophication had an antagonistic effect that weakened the effect of fine sediment. Benthic biofilm growth can clog interstitial spaces, reducing the exchange of water between the stream channel and the subsurface (Aubeneau et al. 2016). This effect is exacerbated when grain sizes are small such as in our combined treatment. Hence, we attribute the reduction in hyporheic NO3 uptake to bioclogging that might have decreased the nutrient and carbon flow to the hyporheic compartment. This explanation supports previous studies which show that the benthic-hyporheic connectivity plays a crucial role in the hyporheic nitrogen uptake (Mendoza-Lera et al. 2019) by affecting solute transport and the redox conditions (Caruso et al. 2017).

Earlier studies measured a higher areal NO3 uptake rate in agricultural streams than in reference streams (Mullholland 2008, Hall et al. 2009) and attributed it to the elevated NO3 concentrations in agricultural streams. Our study expands this knowledge by showing that increased NO3 uptake in agricultural streams is not just the result of higher NO3 concentrations stimulating whole-stream uptake. Rather, the occurrence of other agricultural stressors, such as fine sediment and eutrophication, enhances the areal NO3 uptake through compartment-specific mechanisms.

Our experiment enabled us to draw two significant conclusions that extend previous knowledge on mechanisms regulating NO3 uptake in agricultural streams. Firstly, we disentangled the interactive effects of eutrophication and fine sediment deposition on NO3 uptake dynamics and demonstrated that eutrophication is more important than fine sediment in determining NO3 uptake. Based on this finding, we conjecture that the observed changes in NO3 uptake previously measured in field conditions may be primarily attributed to eutrophication and less to other stressors such as fine sediment.

Secondly, the differentiation between the benthic and hyporheic compartment allowed us to highlight the distinct contribution of both compartments to NO3 uptake. This differentiation also revealed that stressor effects are compartment-specific, which implies that the compartments are differently vulnerable to agricultural stressors. Indeed, benthic capacity was affected by eutrophication only, whereas hyporheic NO3 uptake was affected by both stressors. However, the changes induced by eutrophication on benthic NO3 uptake were more pronounced than the ones measured in the hyporheic compartment in response to any of the stressors. Furthermore, significant stressor interaction only occur in the hyporheic compartment.

The distinct vulnerability of the compartments to stressor number and intensity strongly limits the transferability of findings from one compartment to the other, especially in multiple stressor scenarios. The limited transferability is problematic because our knowledge about the effects and interactions of stressors is, so far, mostly derived from the benthic compartment. Therefore, we advocate that future studies on the effects of agricultural land use on ecosystem functions should consider both compartments in factorial, mechanisms-oriented experiments. Such approaches will help to increase the accuracy of effect size estimates, which are essential for the effective conservation and management of freshwater ecosystems subjected to agricultural land use.

Acknowledgments

The authors thank S. Bauth, A. Schlenker, F. Zander, P. Fink, C. Anlanger, J. Voigt, B. Sulejewski and A. Goeze for their immense assistance with field and laboratory work. A. Hoff and the GEWANA department for analysis of nutrients in the laboratory.

Funding

Open Access funding enabled and organized by Projekt DEAL.

This research was funded by the research program “Changing Earth – Sustaining our Future” of the German Helmholtz Association and was part of the Integration Platform ‘Freshwater resources’.

Conflict of interest

The authors have no relevant financial or non-financial interests to disclose.

Data availability

All data generated during this study are included in this published article (and in its supplementary information files).

Allan JD (2004) Landscapes and Riverscapes: The Influence of Land Use on Stream Ecosystems. Annu Rev Ecol Evol Syst 35:257–284. https://doi.org/10.1146/annurev.ecolsys.35.120202.110122
Anlanger C, Risse‐Buhl U, Schiller D, et al (2021) Hydraulic and biological controls of biofilm nitrogen uptake in gravel‐bed streams. Limnol Oceanogr 66:3887–3900. https://doi.org/10.1002/lno.11927
Arango CP, Tank JL (2008) Land use influences the spatiotemporal controls on nitrification and denitrification in headwater streams. J North Am Benthol Soc 27:90–107. https://doi.org/10.1899/07-024.1
Atkinson BL, Grace MR, Hart BT, Vanderkruk KE (2008) Sediment instability affects the rate and location of primary production and respiration in a sand-bed stream. J North Am Benthol Soc 27:581–592. https://doi.org/10.1899/07-143.1
Aubeneau AF, Hanrahan B, Bolster D, Tank J (2016) Biofilm growth in gravel bed streams controls solute residence time distributions. J Geophys Res Biogeosci 121:1840–1850. https://doi.org/10.1002/2016JG003333
Bernhardt ES, Rosi EJ, Gessner MO (2017) Synthetic chemicals as agents of global change. Front Ecol Environ 15:84–90. https://doi.org/10.1002/fee.1450
Bernot MJ, Tank JL, Royer T V, David MB (2006) Nutrient uptake in streams draining agricultural catchments of the midwestern United States. Freshw Biol 51:499–509. https://doi.org/https://doi.org/10.1111/j.1365-2427.2006.01508.x
Besemer K, Singer G, Hödl I, Battin TJ (2009) Bacterial community composition of stream biofilms in spatially variable-flow environments. Appl Environ Microbiol 75:7189–7195. https://doi.org/10.1128/AEM.01284-09
Böhlke JK, Antweiler RC, Harvey JW, et al (2009) Multi-scale measurements and modeling of denitrification in streams with varying flow and nitrate concentration in the upper Mississippi River basin, USA. Biogeochemistry 93:117–141. https://doi.org/10.1007/s10533-008-9282-8
Böhlke JK, Harvey JW, Voytek MA (2004) Reach-scale isotope tracer experiment to quantify denitrification and related processes in a nitrate-rich stream, midcontinent United States. Limnol Oceanogr 49:821–838. https://doi.org/10.4319/lo.2004.49.3.0821
Bourassa N, Cattaneo A (2000) Responses of a lake outlet community to light and nutrient manipulation: Effects on periphyton and invertebrate biomass and composition. Freshw Biol 44:629–639. https://doi.org/10.1046/j.1365-2427.2000.00610.x
BRAUNS
Cardinale BJ, Palmer MA, Swan CM, et al (2002) The influence of substrate heterogeneity on biofilm metabolism in a stream ecosystem. Ecology 83:412–422. https://doi.org/10.1890/0012-9658
Caruso A, Boano F, Ridolfi L, et al (2017) Biofilm-induced bioclogging produces sharp interfaces in hyporheic flow, redox conditions, and microbial community structure. Geophys Res Lett 44:4917–4925. https://doi.org/10.1002/2017GL073651
Chen SC, Budhraja R, Adrian L, et al (2021) Novel clades of soil biphenyl degraders revealed by integrating isotope probing, multi-omics, and single-cell analyses. ISME Journal 15:3508–3521. https://doi.org/10.1038/s41396-021-01022-9
Dodds WK, López AJ, Bowden WB, et al (2002) N uptake as a function of concentration in streams. J North Am Benthol Soc 21:206–220. https://doi.org/10.2307/1468410
Earl SR, Valett HM, Webster JR (2006) Nitrogen saturation in stream ecosystems. Ecology 87:3140–3151. https://doi.org/10.1890/0012-9658(2006)87[3140:NSISE]2.0.CO;2
Findlay RH, Battin TJ (2015) The Microbial Ecology of Benthic Environments. In: Manual of Environmental Microbiology, 4th Edition
Finkel Z V, Irwin AJ, Schofield O (2004) Resource limitation alters the ¾ size scaling of metabolic rates in phytoplankton. Mar Ecol Prog Ser 273:269–280
Fink P, Norf H, Anlanger C, et al (2020) Streamside mobile mesocosms (MOBICOS): A new modular research infrastructure for hydro‐ecological process studies across catchment‐scale gradients. Int Rev Hydrobiol 105:63–73. https://doi.org/10.1002/iroh.201902009
Godwin CM, Cotner JB (2018) What intrinsic and extrinsic factors explain the stoichiometric diversity of aquatic heterotrophic bacteria? ISME Journal 12:598–609. https://doi.org/10.1038/ismej.2017.195
Graeber D, Tenzin Y, Stutter M, et al (2021) Bioavailable DOC: reactive nutrient ratios control heterotrophic nutrient assimilation—An experimental proof of the macronutrient-access hypothesis. Biogeochemistry 155:1–20. https://doi.org/10.1007/s10533-021-00809-4
Hall RO, Tank JL (2003) Ecosystem metabolism controls nitrogen uptake in streams in Grand Teton National Park, Wyoming. Limnol Oceanogr 48:1120–1128. https://doi.org/10.4319/LO.2003.48.3.1120
Hillebrand H, Acevedo-Trejos E, Moorthi SD, et al (2022) Cell size as driver and sentinel of phytoplankton community structure and functioning. Funct Ecol 36:276–293
Hoellein TJ, Tank JL, Rosi-Marshall EJ, Entrekin SA (2009) Temporal variation in substratum-specific rates of N uptake and metabolism and their contribution at the stream-reach scale. J North Am Benthol Soc 28:305–318. https://doi.org/10.1899/08-073.1
Holmes RM, Jones JB, Fisher SG, Grimm NB (1996) Denitrification in a nitrogen-limited stream ecosystem. Biogeochemistry 33:125–146. https://doi.org/10.1007/BF02181035
Jackson MC, Loewen CJG, Vinebrooke RD, Chimimba CT (2016) Net effects of multiple stressors in freshwater ecosystems: a meta-analysis. Glob Chang Biol 22:180–189. https://doi.org/10.1111/GCB.13028
Kornexl BE, Gehre M, Höfling R, Werner RA (1999) On-line δ18O measurement of organic and inorganic substances. Rapid Communications in Mass Spectrometry 13:1685–1693. https://doi.org/10.1002/(SICI)1097-0231(19990830)13:16<1685::AID-RCM699>3.0.CO;2-9
Liu Y, Majdi N, Tackx M, et al (2015) Short-term effects of nutrient enrichment on river biofilm: N–NO3− uptake rate and response of meiofauna. Hydrobiologia 744:165–175. https://doi.org/10.1007/s10750-014-2074-3
Marti E, Sabater F (1996) High Variability in Temporal and Spatial Nutrient Retention in Mediterranean Streams. Ecology 77:854–869. https://doi.org/https://doi.org/10.2307/2265506
McCall SJ, Hale MS, Smith JT, et al (2017) Impacts of phosphorus concentration and light intensity on river periphyton biomass and community structure. Hydrobiologia 792:315–330. https://doi.org/10.1007/s10750-016-3067-1
Mendoza-Lera C, Ribot M, Foulquier A, et al (2019) Exploring the role of hydraulic conductivity on the contribution of the hyporheic zone to in-stream nitrogen uptake. Ecohydrology 12:. https://doi.org/10.1002/eco.2139
Morris OF, Loewen CJ, Woodward G, et al (2022) Local stressors mask the effects of warming in freshwater ecosystems. Ecol Lett 25:2540–2551. https://doi.org/10.1111/ELE.14108
Mulholland PJ, Valett HM, Webster JR, et al (2004a) Stream denitrification and total nitrate uptake rates measured using a field 15N tracer addition approach. Limnol Oceanogr 49:809–820. https://doi.org/10.4319/lo.2004.49.3.0809
Mulholland PJ (2004b) LINX II STREAM 15 N EXPERIMENT PROTOCOLS. 1–78
Mulholland PJ, Hall RO, Sobota DJ, et al (2009) Nitrate removal in stream ecosystems measured by 15N addition experiments: Denitrification. Limnol Oceanogr 54:666–680. https://doi.org/10.4319/LO.2009.54.3.0666
Mulholland PJ, Helton AM, Poole GC, et al (2008) Stream denitrification across biomes and its response to anthropogenic nitrate loading. Nature 2008 452:7184 452:202–205. https://doi.org/10.1038/nature06686
Myers AK, Marcarelli AM, Arp CD, et al (2007) Disruptions of stream sediment size and stability by lakes in mountain watersheds: Potential effects on periphyton biomass. J North Am Benthol Soc 26:390–400. https://doi.org/10.1899/06-086.1
Niyogi DK, Simon KS, Townsend CR (2004) Land use and stream ecosystem functioning: nutrient uptake in streams that contrast in agricultural development. Arch Hydrobiol 160:471–486. https://doi.org/10.1127/0003-9136/2004/0160-0471
Parr RM, Clements SA (1991) Intercomparison of enriched stable isotope reference materials for medical and biological studies. Vienna
Passy SI (2007) Diatom ecological guilds display distinct and predictable behavior along nutrient and disturbance gradients in running waters. Aquat Bot 86:171–178. https://doi.org/10.1016/j.aquabot.2006.09.018
Peipoch M, Gacia E, Bastias E, et al (2016) Small-scale heterogeneity of microbial N uptake in streams and its implications at the ecosystem level. Ecology 97:1329–1344. https://doi.org/10.1890/15-1210.1
Proia L, Romaní AM, Sabater S (2012) Nutrients and light effects on stream biofilms: A combined assessment with CLSM, structural and functional parameters. Hydrobiologia 695:281–291. https://doi.org/10.1007/s10750-012-1117-x
Pucher M, Wünsch U, Weigelhofer G, et al (2019) staRdom: Versatile Software for Analyzing Spectroscopic Data of Dissolved Organic Matter in R. Water (Basel) 11:2366. https://doi.org/10.3390/w11112366
Risse-Buhl U, Anlanger C, Kalla K, et al (2017) The role of hydrodynamics in shaping the composition and architecture of epilithic biofilms in fluvial ecosystems. Water Res 127:211–222. https://doi.org/10.1016/j.watres.2017.09.054
Roche KR, Shogren AJ, Aubeneau A, et al (2019) Modeling Benthic Versus Hyporheic Nutrient Uptake in Unshaded Streams With Varying Substrates. J Geophys Res Biogeosci 124:367–383. https://doi.org/10.1029/2018JG004684
Romaní AM, Giorgi A, Acuña V, Sabater S (2004) The influence of substratum type and nutrient supply on biofilm organic matter utilization in streams. Limnol Oceanogr 49:1713–1721. https://doi.org/10.4319/lo.2004.49.5.1713
Schürings C, Feld CK, Kail J, Hering D (2022) Effects of agricultural land use on river biota: a meta-analysis. Environmental Sciences Europe 2022 34:1 34:1–13. https://doi.org/10.1186/S12302-022-00706-Z
Steinman AD, Mulholland PJ, Hill WR (1992) Functional Responses Associated with Growth Form in Stream Algae. J North Am Benthol Soc 11:229–243. https://doi.org/10.2307/1467388
Stock MS, Ward AK (1989) Establishment of a Bedrock Epilithic Community in a Small Stream: Microbial (Algal and Bacterial) Metabolism and Physical Structure. Canadian Journal of Fisheries and Aquatic Sciences 46:1874–1883. https://doi.org/10.1139/f89-236
Stutter M, Graeber D, Weigelhofer G (2020) Available Dissolved Organic Carbon Alters Uptake and Recycling of Phosphorus and Nitrogen from River Sediments. Water (Basel) 12:3321. https://doi.org/10.3390/W12123321
Stutter MI, Graeber D, Evans CD, et al (2018) Balancing macronutrient stoichiometry to alleviate eutrophication. Science of The Total Environment 634:439–447. https://doi.org/10.1016/J.SCITOTENV.2018.03.298
Tank JL, Speir SL, Sethna LR, Royer T V (2021) The Case for Studying Highly Modified Agricultural Streams: Farming for Biogeochemical Insights. Limnol Oceanogr Bull 30:41–47. https://doi.org/10.1002/LOB.10436
Triska FJ, Kennedy VC, Avanzino RJ, et al (1989) Retention and Transport of Nutrients in a Third-Order Stream in Northwestern California: Hyporheic Processes. Ecology 70:1893–1905. https://doi.org/10.2307/1938120
U.S. EPA (2000) Atlas of America’s polluted waters. Washington, DC
Valentine K, Mariotti G (2020) Does eutrophication affect the ability of biofilms to stabilize muddy sediments? Estuar Coast Shelf Sci 232:106490. https://doi.org/10.1016/J.ECSS.2019.106490
Von Schiller D, Martí E, Riera JL, Sabater F (2007) Effects of nutrients and light on periphyton biomass and nitrogen uptake in Mediterranean streams with contrasting land uses. Freshw Biol 52:891–906. https://doi.org/10.1111/j.1365-2427.2007.01742.x
Weitere M, Altenburger R, Anlanger C, et al (2021) Disentangling multiple chemical and non-chemical stressors in a lotic ecosystem using a longitudinal approach. Science of the Total Environment 769:144324. https://doi.org/10.1016/j.scitotenv.2020.144324
Westphal K, Musolff A, Graeber D, Borchardt D (2020) Controls of point and diffuse sources lowered riverine nutrient concentrations asynchronously, thereby warping molar N:P ratios. Environmental Research Letters 15:104009. https://doi.org/10.1088/1748-9326/AB98B6
Wymore AS, Coble AA, Rodríguez-Cardona B, McDowell WH (2016) Nitrate uptake across biomes and the influence of elemental stoichiometry: A new look at LINX II. Global Biogeochem Cycles 30:1183–1191. https://doi.org/10.1002/2016GB005468
Wymore AS, Rodríguez-Cardona BM, Herreid A, McDowell WH (2019) LINX I and II: Lessons Learned and Emerging Questions. Front Environ Sci 7:181
Zarnetske JP, Haggerty R, Wondzell SM (2015) Coupling multiscale observations to evaluate Hyporheic nitrate removal at the reach scale. Freshwater Science 34:172–186. https://doi.org/10.1086/680011
Zeglin LH (2015) Stream microbial diversity in response to environmental changes: review and synthesis of existing research. https://doi.org/10.3389/fmicb.2015.00454

Download PDF

Journal Publication

published 13 Mar, 2024

Read the published version in Biogeochemistry →

Reviewers agreed at journal
10 Jul, 2023
Reviewers invited by journal
09 Jul, 2023
Editor invited by journal
01 Jul, 2023
Editor assigned by journal
01 Jul, 2023
First submitted to journal
28 Jun, 2023

You are reading this latest preprint version

Multiple stressors effects on nitrate uptake vary across benthic and hyporheic compartments

Status:

Journal Publication

Version 1

Abstract

Figures

Introduction

Methods

Statistical analysis

Results

Benthic

Hyporheic

Discussion

Conclusion

Declarations

References

Supplementary Files

Status:

Journal Publication

Version 1

Parameters	Compartment	Eutrophication	Fine sediment	Eutrophication x Fine sediment
N-NO3 areal uptake	\({B}^{a}\)	92.83 (< 0.001) ↑	0.50 (0.49)	2.1 (0.17)
N-NO3 areal uptake	H	0.01 (0.95)	40.33 (< 0.001) ↑	38.21 (< 0.001) ↑
N-NO3 biomass uptake	\({B}^{a}\)	301.03 (< 0.001) ↑	0.21 (0.65)	0.75 (0.40)
N-NO3 biomass uptake	\({H}^{a}\)	16.40 (< 0.001) ↑	78.40 (< 0.001) ↑	23.66 (< 0.001) ↑
Organic matter (OM)	\({B}^{ab}\)	0.003 (0.95)	0.06 (0.81)	4.08 (0.06)
Organic matter (OM)	\({H}^{a}\)	3.61 (0.07)	22.80 (< 0.001) ↓	3.45 (0.08)
Particulate organic carbon (POC)	\({B}^{b}\)	26.13 (< 0.001) ↑	3.24 (0.09)	0.72 (0.41)
Particulate organic carbon (POC)	\({H}^{ab}\)	2.67 (0.12)	4.92 (< 0.05) ↓	4.76 (< 0.05) ↓
Particulate organic nitrogen (PON)	\({B}^{a}\)	133.57 (< 0.001) ↑	15.14 (< 0.001) ↓	1.37 (0.26)
Particulate organic nitrogen (PON)	\({H}^{ab}\)	12.54 (< 0.05) ↓	4.13 (0.06)	17.08 (< 0.001) ↓
Chlorophyll-a	\({B}^{a}\)	103.21 (< 0.001) ↑	1.35 (0.26)	2.32 (0.15)
Bacterial abundance	\({B}^{ab}\)	3.7 (0.07)	0.03 (0.87)	0.47 (0.50)
Bacterial abundance	\({H}^{a}\)	0.35 (0.56)	0.75 (0.40)	9.35 (< 0.05) ↓

Parameters	Compartment	Control	Eutrophication	Fine sediment	Fine sediment + eutrophication
Organic matter (OM) (mg \({cm}^{-3}\))	\({B}^{ab}\)	154 ± 77 (a)	98 ± 40 (a)	94 ± 20 (a)	139 ± 35 (a)
Organic matter (OM) (mg \({cm}^{-3}\))	\({H}^{a}\)	9 ± 2 (a)	9 ± 3 (a)	6 ± 2 (ab)	3 ± 1 (b)
Particulate organic carbon (POC) (µg \({g}^{-1}\) OM)	\({B}^{b}\)	7.8 ± 0.3 (a)	9.5 ± 0.8 (b)	7.6 ± 0.5 (a)	8.7 ± 0.6 (ab)
Particulate organic carbon (POC) (µg \({g}^{-1}\) OM)	\({H}^{ab}\)	1.8 ± 0.4 (a)	1.3 ± 0.3 (ab)	1.2 ± 0.2 (b)	1.3 ± 0.2 (ab)
Particulate organic nitrogen (PON) (µg \({g}^{-1}\) OM)	\({B}^{a}\)	11.8 ± 1.4 (a)	24 ± 3.9 (c)	10.2 ± 0.5 (a)	18.1 ± 1.7 (b)
Particulate organic nitrogen (PON) (µg \({g}^{-1}\) OM)	\({H}^{ab}\)	10 ± 0.8 (a)	6.4 ± 0.6 (b)	7.1 ± 0.6 (b)	7.3 ± 1.3 (b)
Chlorophyll-a (µg \({cm}^{-3}\) )	\({B}^{a}\)	0.9 ± 0.3 (a)	4.6 ± 2.0 (b)	0.6 ± 0.2 (a)	5.0 ± 2.6 (b)
Bacterial abundance \(\left(\text{c}\text{o}\text{u}\text{n}\text{t}\text{s}\text{*}{10}^{6} {cm}^{-3} \text{s}\text{e}\text{d}\text{i}\text{m}\text{e}\text{n}\text{t}\right)\)	\({B}^{ab}\)	4.3 ± 3.1 (a)	3.2 ± 1.7 (a)	4.9 ± 3 (a)	2.3 ± 0.9 (a)
	\({H}^{a}\)	2.9 ± 1.7 (a)	6.6 ± 6.4 (a)	9.6 ± 6.3 (a)	2.4 ± 0.8 (a)
Oxygen concentration (mg \({l}^{-1}\))	B	7.0 ± 0.3	7.3 ± 0.1	5.6 ± 0.6	7.6 ± 0.6
Oxygen concentration (mg \({l}^{-1}\))	H	7.1 ± 0.3	7.3 ± 0.4	0.15 ± 0.07	0 ± 0