The Potential Hidden Age of Dissolved Organic Carbon Exported by Peatland Streams

2.1 Field Sampling

The DOC-rich water for incubation was collected on 17 March 2015 from Black Burn, a first-order stream draining a 3.4-km² catchment of Auchencorth Moss peatland (55°48′N, 03°15′W) in central Scotland (Dinsmore et al., 2010). This is an ombrotrophic peatland (85% of the catchment is peat), drained by a narrow open stream channel 0.7 m wide on average. Mean air temperature was 3.1 °C, total rainfall was 102 mm, and mean discharge in the Black Burn was 41 L/s for the month prior to sampling, with no unusual hydrological or climatic events for this time of year (Dinsmore et al., 2013; Pickard et al., 2017).

The aquatic C flux is dominated by DOC, with an annual yield of 19.3 ± 4.6 g C·m⁻²·year⁻¹. CO₂ evasion is the second biggest component of aquatic C flux in the study catchment (10.0 g C·m⁻²·year⁻¹; range = 2.33–24.0 g C·m⁻²·year⁻¹). Other greenhouse gas fluxes are negligible compared with DOC export and CO₂ evasion (Dinsmore et al., 2013). Peat between 0.5 and 5 m thick overlies an extensive layer of glacial-derived boulder clay, which means that the underlying bedrock (predominantly Upper Carboniferous/Lower Devonian sandstones with occasional thin bands of limestone, mudstone, and coal) is likely less important to stream CO₂ fluxes compared with deep peat layers (Billett et al., 2007). Peat soils in the riparian zone of the Black Burn have provided ¹⁴C ages of up to 1,400 years BP (83.93 pmC) at 70-cm depth (dates for deeper layers are not available but are likely older; Leith et al., 2014).

We prefiltered 60 L of stream water through a coarse nylon mesh (~225 μm) and homogenized it across two prerinsed 30-L barrels. The samples were collected at 3.7 °C (water temperature) and kept at that temperature and in the dark until the start of the incubation. Field parameters (pH, electrical conductivity [EC], and temperature) were measured at the site using a Hach PH101C pH probe and a CDC401 conductivity probe connected to an HQd portable meter; these measurements were repeated for the sample waters at the end of the incubation. Dissolved CO₂ (Hope et al., 1995) and DOC concentrations, and initial (t₀) measurements of DO¹⁴C and ¹⁴CO₂, were also collected in the field. The DO¹⁴C sample was collected in a 1-L acid-washed bottle and filtered to 0.7 μm on preashed (~500 °C for ~3 hr) Whatman glass fiber filters (GF/F) in the laboratory. The ¹⁴CO₂ sample was collected using the super-headspace method (see supporting information), collecting 3.5-ml CO₂ on a molecular sieve cartridge (Garnett, Billett, et al., 2016).

2.2 Incubations

The 60-L DOC-rich water sample was refrigerated at the University of Stirling overnight after sampling and thoroughly mixed and filtered the following day through preashed 0.7-μm GF/F filters at low pressure. This filter size isolates the DOC size fraction but allows sufficient bacteria (both heterotrophic and autotrophic) to pass through the filter for incubation (Dean, van Hal, et al., 2018). Fourteen liters of filtrate was then transferred into each of the three replicate incubation chambers. The filled chambers were refrigerated until the incubations commenced on 19 March. Three water aliquots were collected during filtration to confirm the homogeneity of the incubation water (Table S1).

Incubation chambers were 17-L white high-density polyethylene (HDPE) barrels (Ampulla UK), UN Class 1 approved for storage of pharmaceuticals and food products (Figure 3). The chambers were cleaned with dilute Decon90 and thoroughly rinsed with distilled water prior to the experiment. We conducted blank test incubations using MilliQ®-grade deionized water in the same-class HDPE barrels to account for potential leaching from the incubation vessels. After 1 month, blank water samples contained 0.006 mg C/L, equivalent to <0.05% of the sample DOC content (Table S2). After 101 days, the blank water contained 0.01 mg C/L, approximately 0.1% of the sample DOC content (blank samples were processed in the same way as the DO¹⁴C samples, but insufficient material was obtained to measure either δ¹³C or ¹⁴C content; see section 4). This shows that potential leaching from the HDPE incubation chambers used in this experiment was negligible and would not have impacted the ¹⁴C results or DOC concentrations.

When filled with sample water, chambers had a 3-L headspace that was continuously flushed at a rate of approximately 150 cm³/min (replacing the headspace in ~20 min) with outdoor ambient air scrubbed of CO₂ using an “indicating” soda lime trap (26-cm³ volume) and Mg (ClO₄)₂ moisture trap via Tygon® medical-grade E-3606 tubing and mounted CPC couplings (Colder Products Co., USA) and then sealed with plastic paint (Plasti Dip, USA; Figure 3). The soda lime trap was replaced approximately halfway through the experiment, remaining fully functional for the duration of the experiment. Unfortunately, we were not able to monitor CO₂ concentrations in the system except during sampling.

A 140-mm-diameter circular hole was drilled into the lid of each chamber and covered with a 150-mm-diameter quartz glass window that allowed all wavelengths of light to pass through while preventing gas diffusion and then sealed with silicone sealant (Figure 3). The quartz glass window allowed both primary production and photo-oxidation to take place in the incubation chambers (see below). The chambers were equally irradiated with a T5 UV-B lamp (Arcadia Reptile, UK) designed to emit the full spectrum of natural light. The irradiance value measured on the sample surface was 14.5 W/m² (3.19 W/m² between 300 and 400 nm)—this is approximately twice the typical irradiance of the site where the incubation sample was collected, but the aim of the experimental design was to allow both autotrophic and heterotrophic DOC decomposition rather than specifically focus on either decomposition process. A night/day regime was used with a 12-hr equal split (8 a.m. to 8 p.m. light).

The incubated water was recirculated continuously using an external 12-V water pump at a rate of 15 ml/s, turning over the full water volume in 16 hr via mounted CPC couplings and E-3606 tubing, in order to replicate in-stream physical processes and limit stratification and heterogeneous mixing of solutes (Figure 3). The incubation system was designed to limit the ingress of ambient CO₂ by sealing the chambers and flushing the headspace with CO₂-free outside air (flushing was not possible during CO₂ sampling, which took between 40 and 170 min). Any ingress of ambient air that did occur during CO₂ sampling was corrected for using an established methodology for field-based chamber sampling (Gaudinski et al., 2000; Walker et al., 2016). Briefly, two δ¹³C isotopic end-members, ambient lab air (Bilzon et al., 2002) and the oldest CO₂ sample collected, were used to estimate the relative contributions of each end-member to a given sample. This is then compared with our estimate of ambient air ingress into each sample based on measurements of air ingress rates into the chambers (see Supplementary Methods; Figure S2). The mixing of the incubated sample water and the regular supply of oxygen to the headspace meant that any CH₄ production was minimized. The incubations were started on 19 March 2015 and stopped on 6 August 2015. The experiment ran for 140 days to simulate longer potential residence times for DOC during aquatic transfer from the terrestrial to marine zone. While the half-life of OC within inland water systems is 2.5 ± 4.5 years (Catalán et al., 2016), average residence times in peatland streams could be much lower than this. But with drain blocking becoming an increasingly common peatland restoration technique (Evans et al., 2018), and peatlands providing a substantial proportion of water supply to reservoirs (Xu et al., 2018), residence times in the order of 140 days or longer are possible for peatland DOC within inland water systems.

2.3 Sample Collection and Analysis

Incubation aliquots for DO¹⁴C and ¹⁴CO₂ were collected from one chamber only (C2) on days 4 (t₁), 8 (t₂), 14 (t₃), 32 (t₄), 76 (t₅), and 138 (t₆; Figure 3). These were spread based on expected higher rates of DOC decomposition over the initial incubation period (Vonk et al., 2015). At the final time point, ¹⁴C samples were collected from all three chambers. Incubation water aliquots were collected from each chamber via an outlet on the water pump line; headspace ¹⁴CO₂ samples were collected via the upper mounted CPC couplings (Figure 3). DO¹⁴C samples were collected in 500-ml acid-washed clear HDPE bottles after prerinsing with sample water, filtered to 0.7 μm on preashed GF/F filters and refrigerated until analysis (less than 90 days in all cases, which should not have impacted the stable or radiogenic isotopic signatures of the DOC sample; Gulliver et al., 2010). ¹⁴CO₂ samples were collected by first purging the headspace of CO₂ by cycling it through a soda lime CO₂ trap until the concentration was <60 ppm. The headspace was then allowed to equilibrate with the water by diffusion for 1 to 2 hr (overnight in one case; see Supplementary Methods). The headspace was then cycled through a molecular sieve trap to capture the CO₂ for ¹⁴C analysis (1.5- to 4.1-ml CO₂; Table S1). Where insufficient sample was captured on the molecular sieve in one cycle, the headspace was then allowed to build up again for another 1 to 2 hr and sampled again.

Incubation aliquots for DOC concentrations (DOC and total C) and optical properties (absorbance and fluorescence) were collected in 30-ml sterile HDPE tubes (first rinsed with sample) during preincubation filtration and on days 1, 4, 7, 8, 14, 32, 34, 48, 62, 76, 97, 124, 138, and 140 (Figure 3). Dissolved organic matter (DOM—contains the DOC pool) quality parameters from absorbance and fluorescence measurements provide insights into the structure and molecular weight of the DOC pool (Fellman et al., 2010; Helms et al., 2008). Absorbance aliquots (50 mL) from the three replicate chambers were collected in sterile containers and then filtered through precombusted 0.7-μm Whatman GF/F filters under low pressure. Absorbance measurements were carried out immediately after sample collection using a dual-beam spectrophotometer (Agilent Cary 100; see Supplementary Methods). From the absorbance values, we calculated the ratio of the absorption spectral slopes between 275–295 and 350–400 nm (slope ratio, S_R), which is inversely correlated with DOM molecular weight (Fichot & Benner, 2012; Helms et al., 2008). Aliquots for the measurement of DOM fluorescence were only collected from chamber C2. DOM fluorescence was measured with a spectrofluorometer (Instant Screener®, Laser Diagnostic Instruments AS; see Supplementary Methods; Lawaetz & Stedmon, 2009; Murphy et al., 2010; Patel-Sorrentino, 2002) and used to calculate the humification index (HIX; the extent of humification in the DOM—i.e., how decomposed it is) by dividing the area under the emission spectra 435–480 nm by the peak area 300–345 nm + 435–480 nm, at excitation 254 nm (Ohno, 2002). Lower HIX values indicate more decomposition (increased humification) of the DOM pool. DOC quality and concentration aliquots collected before day 32 were not filtered prior to analysis. From day 34 onward, minor flocculation was first observed, so these samples were then filtered through 0.7-μm preashed GF/F filters, with negligible difference seen between days 32 and 34 (±1.0 mg C/L). DOC concentrations were measured at the University of Stirling on a Shimadzu Total Organic Carbon Analyzer (TOC-V series) with a lower detection limit of 0.5 μg/L and an accuracy of 1.5%.

POC concentration and ¹⁴C samples were collected on day 140 only, by first stirring to homogenize the sample thoroughly, then filtering through 0.7-μm preashed GF/F filters, and measuring the captured material. One liter of sample water was filtered to obtain the PO¹⁴C samples, and 200 ml of water was filtered for triplicate particulate organic matter concentrations; the filters were then dried at 70 °C. POC concentration samples were combusted in a Carlo-Erba EA 1108 elemental analyzer, using sulfanilamide as the standard.

¹⁴C samples were prepared for analysis at the Natural Environment Research Council Radiocarbon Facility (East Kilbride, UK). DO¹⁴C samples were processed to solids using rotary evaporation and then acid-fumigated (Dean, van der Velde, et al., 2018). DO¹⁴C and PO¹⁴C samples were combusted, and the CO₂ produced was cryogenically recovered. Molecular sieve cartridges were heated, and the desorbed sample CO₂ was cryogenically recovered (Garnett & Murray, 2013). For all ¹⁴C samples, an aliquot of the recovered CO₂ was analyzed for δ¹³C using isotope-ratio mass spectrometry (Thermo Fisher Delta V), with results reported relative to the Vienna Pee Dee Belemnite standard. A further aliquot of CO₂ was graphitized using Fe-Zn reduction and the ¹⁴C content determined by accelerator mass spectrometry (AMS) at the Scottish Universities Environmental Research Centre (Xu et al., 2004). Following convention, all ¹⁴C results were normalized using the δ¹³C values and presented as pmC and conventional radiocarbon ages (years BP, where 0 year BP is 1950 CE). The ¹⁴C background associated with these methods was quantified by analysis of ¹⁴C-dead materials (blanks); standards of known ¹⁴C content processed alongside the samples gave values within 1σ of the consensus.

3.1 Field Results

DOC and dissolved CO₂ concentrations at the field site were 18.4 ± 0.6 (Figure 4) and 2.3 ± 0.1 mg C/L, respectively, while pH was 5.9 and EC was 60.3 μS/cm. These values are usual for this time of year at the study site (Dinsmore et al., 2013), with average values in the 3 months prior to sampling being 5.1 for pH and 59.1 μS/cm for EC (Pickard et al., 2017). DO¹⁴C was 103.79 ± 0.48 pmC (i.e., modern; Figure 5), while ¹⁴CO₂ was 88.14 ± 0.38 pmC (1,014 ± 35 years BP; Figure 5). The ¹⁴C results agree with previous work at the site and similar temperate peatlands where DOC was modern, while CO₂ was consistently older (Billett et al., 2007; Billett & Garnett, 2010; Leith et al., 2014).

3.2 DOC Decomposition and ¹⁴C Dynamics

DO¹⁴C content decreased from 102.45 ± 0.47 pmC (modern) at t₁, very similar to the field sample, to 99.78 ± 0.46 pmC (17 ± 17 years BP) over 138 days of incubation (Figure 5). δ¹³C-DOC changed only slightly, from −28.7‰ to −28.2‰ during incubation (Table 1). The final DO¹⁴C concentration was consistent across all three chambers with a range of 99.16 ± 0.45 to 99.83 ± 0.44 pmC. While the reduction in DO¹⁴C content over the 138 days of incubation was relatively small (Table 1 and Figure 5), it is statistically significant (p < 0.05 for the three t₆ samples; Student's t test, R version 3.4.3) and implies a small shift toward older DOC.

DOC concentrations in chambers C2 (where the primary ¹⁴C measurements were collected) and C3 behaved similarly, remaining relatively stable for the first 8 to 14 days at a maximum of 18.6 mg C/L, before declining from day 14 to day 138, reaching a low of 13.7 mg/L (Figure 4a). DOC in chamber C1, however, increased in concentration until day 34 (19.46 mg/L) and then declined gradually to 17.8 mg/L at day 138 (Figure 4a).

S_R increased across all chambers, indicating a decline in molecular weight that suggests larger DOM molecules were continually broken down over the course of the incubation (Figure 4b). The increase in S_R was reflected in a 28.7–35.7% reduction in the absorption coefficient at 254 nm, a 31.8–40.2% reduction at 300 nm, and a 30.9–44.9% reduction at 440 nm, suggesting an overall decline in aromaticity and molecular weight (Table S1). HIX values, calculated from the DOM fluorescence measurements from chamber C2, decreased from >0.99 (measurement saturated) to 0.95, indicating a reduction in humic substances in the DOC pool (Figure 4b).

3.3 ¹⁴CO₂ dynamics

During the incubation, headspace ¹⁴CO₂ signatures fell from 89.92 ± 0.90 pmC (853 ± 100 years BP) at t₁ to 74.58 ± 3.53 (2,356 ± 767 years BP) at the final time point (Figure 5). The t₁ ¹⁴CO₂ age was slightly younger than the field ¹⁴CO₂ sample, indicating that, initially, younger DOC was decomposing to produce CO₂. The subsequent increase in ¹⁴CO₂ age was supported by replicate ¹⁴C measurements at the final time point in chambers 1 and 3 of 80.47 ± 0.84 and 83.60 ± 1.45 pmC (1,746 ± 124 and 1,439 ± 280 years BP, respectively; Table 1 and Figure 6). The pH and EC of the incubated water remained stable at 5.8–5.9 and 59.8–63.7, respectively, indicating that dissolved inorganic C was predominantly present as dissolved CO₂, which would be in isotopic equilibrium with the sampled headspace CO₂.

The δ¹³C values can also indicate CO₂ source, alongside ¹⁴C. The field CO₂ sample had a δ¹³C signature of −18.6 ‰, which was higher than the CO₂ produced directly from the decomposition of the DOC pool (−27.1 ‰; Table 1). The field δ¹³CO₂ value was matched by an old ¹⁴C age, suggesting it was sourced in part by the decomposition of deep peat layers (Billett et al., 2007; Dean, van der Velde, et al., 2018) and potentially ¹³CO₂ fractionation during evasion from the stream (Doctor et al., 2008). The lower δ¹³CO₂ value from the incubation matches that of DOC (−28.4‰ to −28.8‰; Table 1), supporting the case for the old ¹⁴CO₂ ages measured during the incubation being directly produced from the decomposition of the DOC pool.

3.4 The Production of CO₂ and POC From Decomposition of the DOC Pool

POC was formed in the incubation chambers during the experiment, and this also had an old ¹⁴C signature of 1,824 ± 37 to 2,094 ± 37 years BP (Table 1 and Figure 6). This POC formed from large-scale microbial growth, driven by heterotrophic and autotrophic production, causing flocculation of living and dead microbial biomass and OC, and the formation of microbial biofilms (Figure S2; Battin et al., 2016; Logue et al., 2016). POC was not present at the beginning of the incubation as we filtered the sample water to isolate the DOC pool.

Isotope mass balance equations can be useful for partitioning C sources (Billett et al., 2007). During the incubation, old C was predominantly partitioned into the CO₂ and POC pools (Figure 6). To satisfy isotope mass balance, the DOC age would be expected to remain constant or become younger (more ¹⁴C enriched) as old C was lost to CO₂ and POC. For example, we used equation 1 to estimate the ¹⁴C content of the DOC pool that was lost during the course of the incubation:

(1)

where Δ represent the ¹⁴C content (pmC) and M the mass of the DOC lost during the incubation (_lost) and at the start (_t0) and end (_t6) of the incubation. Because the age of DOC increased over time (i.e., ¹⁴C depleted; Figure 5), equation 1 predicts the ¹⁴C content of the CO₂ produced from the decomposition of the DOC pool to be ¹⁴C enriched (117.6 pmC). However, this value contrasts with our direct observations of the predominantly old C in the CO₂ and POC pools (Figure 6).

4.1 Carbon (Re)Cycling in the Incubation Chambers

The observed increase in DOC age in conjunction with the production of old CO₂ and POC pools during the experiment could be explained by the rapid loss of a modern component of the DOC pool early in the incubation and the reintroduction of old C into the DOC pool by autotrophic fixation. If there was rapid respiration of a labile modern component of the DOC pool (e.g., plant exudates and leachates from freshly decayed organic matter), then the resultant CO₂ could have been flushed from the headspace and thus missed in the first days of the incubation (the chambers were not closed systems). This is hinted at by the t₁ CO₂, which was generated directly from the decomposition of the DOC pool, being younger than the field CO₂, which was an integrated CO₂ pool generated from the decomposition of OC throughout the catchment peat soil profiles as well as within the stream itself (Figure 5). As this labile modern DOC pool rapidly decomposed over the early phase of the incubation and was flushed from the headspace, the old C signature would have become more pronounced in the respired CO₂, which can be seen in Figure 5. An additional mechanism, the growth of autotrophic microorganisms, would have fixed free (dissolved) CO₂ into the microbial-DOC pool through photosynthesis. This mechanism can at least partially explain the partitioning of C ages in the incubation systems. The free CO₂ in the incubation system may have been younger, initially, as described above, but rapidly aged between t₁ and t₂ (4 days; Figure 5), yielding old CO₂ for subsequent autotrophic fixation and incorporation into autotrophic biomass and POC formation. Microbial biomass and detritus less than 0.7 μm in size are also considered part of the DOC pool and are included in the DO¹⁴C and DOC concentration measurements for this experiment, so the reintroduction of old C in the DOC pool could also explain the increase in the bulk DO¹⁴C age (Fellman et al., 2010).

4.2 The Potential Hidden Age of Peatland Stream DOC

Our study supports the hypothesis that peatland stream DOC with a modern age can contain a fraction of old C that is available for decomposition during aquatic transport. Old C was identified in the CO₂ produced from the decomposition of the predominantly modern DOC pool within 20 days (Figure 5), well within the half-life of OC within inland water systems (2.5 ± 4.5 years; Catalán et al., 2016). The increase in DOC concentrations observed in chamber C1 may have been due to increased autotrophic microbial growth in this chamber compared with the others (some microbial biomass is incorporated into DOC; see section 6.4). However, shifts in DOM structure across all three incubation chambers show that the terrestrial DOC pool was continually being decomposed for the duration of the experiment, despite DOC dynamics being influenced by autotrophic and heterotrophic microbial growth, (Figure 4 and Table S1). The observed decline in HIX values (Figure 4) is likely due to the preferential decomposition of terrestrial-derived (humic) DOC, rather than the decomposition of heterotrophic and autotrophic microbial biomass that also contributes to the overall DOC pool (Fellman et al., 2010). The terrestrial-derived component of the DOC pool is likely old (Galy & Eglinton, 2011; Leith et al., 2014), and the HIX dynamics suggest that this old terrestrial OC was continually decomposed over the incubation, fueling CO₂ production as reflected in the old ¹⁴CO₂ signature we observe (Figure 5). DOM structural shifts were consistent across all the incubation chambers (Figure 4b), suggesting that although the kinetics of DOC decomposition in chamber C1 may have been different from the others, the overall decomposition of the DOC pool and ¹⁴C results are comparable across all chambers (Figure 6). However, fluxes of CO₂ produced in the different chambers were not measured to corroborate this.

There was an increase in the difference between the incubation DO¹⁴C and ¹⁴CO₂ signatures as the experiment progressed. This difference rapidly increased over the first 14 days and then plateaued (Figure 5). This is consistent with the rapid respiration of a labile modern component of the DOC pool and increased decomposition of old C once this labile modern pool was lost. Previous work also demonstrated the likely presence of a labile young C component with bulk riverine DOC loads (Galy & Eglinton, 2011; Loh et al., 2006; Raymond & Bauer, 2001). We were not able to quantify the amount of modern labile C that was rapidly lost in the initial stages of the incubation. But this fraction cannot have been the majority of the modern DOC component because the bulk DO¹⁴C signature remained close to modern for the duration of the experiment. This indicates that there was a substantial recalcitrant modern DOC component. In this experiment, we clearly demonstrate that there is also potential for an important component of labile old C to be hidden within bulk “modern” riverine DOC loads. This old C signal may have been exaggerated in our experimental system due to the rapid respiration and flushing from the system of a labile modern DOC component and the later reintroduction of C into autotrophic biomass (see section 6.4). However, that the presence of old C is observed at all, and at such clear levels in the CO₂ and POC measurements, unequivocally supports our hypothesis that labile old C can be concealed within bulk modern DOC pools.

Previous work has shown that heterotrophic respiration of temperate lake DOC produced old CO₂ (1,000 to 3,000 years BP) despite the bulk ¹⁴C signature of the incubated DOC being significantly younger (modern, with ¹⁴C contents of 101.4 to 111.0 pmC; McCallister & del Giorgio, 2012). The degradation of ancient C (>11,300 years BP) has also been demonstrated in Arctic yedoma permafrost (Drake et al., 2015; Mann et al., 2015; Spencer et al., 2015; Vonk et al., 2013). In contrast, earlier work suggests that modern C is preferentially respired from bulk riverine DOC loads, with old C remaining recalcitrant (Galy & Eglinton, 2011; Raymond & Bauer, 2001). However, concurrent in situ measurements of aquatic DO¹⁴C and ¹⁴CO₂ signatures are rare, limited to four studies, of which we are aware. Aquatic CO₂ is generally older than DOC in UK peatlands, including the study site where we collected our DOC sample for incubation (Billett et al., 2007, 2012; Leith et al., 2014). This was thought to be driven by a mixed signal comprising ¹⁴C-dead CO₂ from geological sources (~5–30%) and modern CO₂ derived from soil respiration in the upper peat layers (~70–95%). DOC was older than CO₂ in a western Canadian Arctic peatland underlain by continuous permafrost (Dean, van der Velde, et al., 2018). There, the shallow permafrost prevented contributions of ¹⁴C-dead geogenic sources, with the CO₂ and DOC sourced from both old and young shallow peat layers. The results of the incubation study presented here, where external C inputs to the ¹⁴CO₂ signal are restricted, suggest that respiration of old C within bulk DOC pool also contributes to in situ riverine ¹⁴CO₂ signals. However, this is highly system dependent, with different DOC age components available for decomposition to CO₂ during aquatic transport and storage.

4.3 Implications and Future Work

Bulk DO¹⁴C measurements are not sensitive enough to detect old C in peatland inland water systems, as demonstrated by the production of old ¹⁴CO₂ from the decomposition of a bulk modern DO¹⁴C sample in the experiment presented here—with the exception of severe cases of landscape disturbance (Figure 1). The findings of this study also suggest that bulk DO¹⁴C measurements do not represent the ¹⁴C signature of the labile OC pool exported by inland water systems more broadly (Fellman et al., 2014). In situ aquatic ¹⁴CO₂ measurements in UK peatlands were previously thought to show a mixture of modern soil and ¹⁴C-dead geological sources (Billett et al., 2007). This experiment suggests that there is also a component of old C that remains recalcitrant in oxygen-poor peat layers for many thousands of years but becomes available for decomposition after its release by variable mixing with water flowing through the peat soil layers (Dean, van der Velde, et al., 2018; van der Velde et al., 2012). This shift from recalcitrant to labile may be driven by the change from oxygen-poor to oxygen-rich conditions, differing microbial populations in the aquatic zone from the soil zone (Dean, van Hal, et al., 2018), and/or combined photodecomposition and microbial decomposition. All of these processes were operating in the experiment presented here (Figure 3). However, we did not separate them out by experimental design, seeking only to simulate natural in-stream conditions for a single site. Further studies should be carried out to distinguish the importance of these individual processes to DOC lability during transport and storage in inland water systems and across a range of ecosystems.

These findings may extend beyond aquatic respiration and may be relevant to the interpretation of ¹⁴C studies on soil respiration and porewater DO¹⁴C. Further, other aquatic C species may also be affected by components of differing C age and lability. Incubations such as the one presented here can aid this research strand, although they require multiple ¹⁴C dates and are time-consuming. For bulk POC and DOC samples, ramped pyrolysis can reveal the ¹⁴C signature of C combusted at different temperatures (i.e., indicating lability; e.g., Zhang et al., 2017), although this has only recently been developed for bulk aquatic DOC samples (Hemingway et al., 2017). Matching ramped pyrolysis DO¹⁴C signatures and integrated soil OC age distributions (Sierra et al., 2017) with bulk DO¹⁴C samples via age distribution models (e.g., Figure 2) would improve our estimates of the likely proportions of different aged C within bulk DO¹⁴C samples (Evans et al., 2014). This could be assisted by compound-specific ¹⁴C methods (e.g., Feng et al., 2013, 2017), although this can also require multiple ¹⁴C dates for a single sample depending on the target organic compounds, and it is possible that C of multiple ages can be present within the isolated pools.

Direct in situ measurements of ¹⁴CO₂ and ¹⁴CH₄ signatures are needed to improve our understanding of the contribution of old C to inland water C emissions across a range of environmental settings. Methodological advances now allow for the collection of ¹⁴CO₂ and ¹⁴CH₄ samples relatively quickly and easily in even remote field locations (Billett et al., 2006; Dean et al., 2017; Garnett, Billett, et al., 2016; Garnett, Gulliver, et al., 2016). For aquatic CO₂ and CH₄ ¹⁴C samples, age distribution analysis (e.g., Dean, van der Velde, et al., 2018) or multisource isotopic mass balance (e.g., Elder et al., 2018) can help determine relative contributions of different aged C.