Philippa Kaur

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Sampling site and sediment collectionPaleolimnological analysis by using sediment core samples were applied to reconstruct historical variations of Daphnia eDNA concentrations in Lake Biwa (Fig. 1b). Lake Biwa is the largest monomictic and mesotrophic lake in Japan. In this lake, during the last several decades the industrial revolution, multiple stressors of human origins impacted this ecosystem and the resident biological communities34,35,58. In our study, four 30-cm-long gravity core samples (namely LB1, LB2, LB4, and LB7; Fig. 1a–e) were collected on 17 August 2017 at the anchoring site of Hasu, a Center of Ecological Research boat from Kyoto University (Supplementary Fig. S2). A gravity corer with an inner diameter of 10.9 cm and a length of 30 cm was used to obtain the core samples. LB7 core was analyzed for chronological and reconstruction of temporal variation in Daphnia remain abundance (Fig. 1a,b). LB2 and LB1, LB4 cores were analyzed for reconstruction of temporal variation in sedimentary Daphnia DNA concentrations and resting egg production, respectively (Fig. 1b). Additionally, two 30-cm-long gravity core samples (namely IM1 and IM8; Fig. 1c,f) were collected at a pelagic site in the northern basin of Lake Biwa in August 2019 (Supplementary Fig. S2). The collected cores were sectioned at intervals of 1-cm thickness using a vertical extruder with a cutting apparatus, except for core number IM8, which was sectioned at 5-cm intervals (Fig. 1f). During the sectioning process, several millimeters of outer edge in each layer disturbed during the splitting process were carefully removed from the entire samples using a knife. After sectioning, each sliced sample were homogenized by shaking and then, all subsamples were taken from each homogenized sample. The pipes, knives, and cutting apparatus were cleaned with 0.6% sodium hypochlorite, tap water, and Milli-Q water to avoid DNA cross-contamination. Each sliced sample was transferred to lightproof bags and frozen at − 80 °C until further analysis.To examine the contamination due to core splitting, DNA extraction, and qPCR analysis, control water samples were inserted at a depth of 14.5–29.5 cm in the sediment cores, and the water samples for core IM1 were used as the negative control (Fig. 1c).Chronology of sediment coresSediment chronology was performed for the LB7 core based on the constant rate of supply (CRS) method of 210Pb dating59 and verified using the 137Cs peak traced in the period 1962 to 196360. Details of the chronological method have been reported elsewhere61. Briefly, dried samples were sealed in holders for a month to allow 222Rn and its short-lived decay product (214Pb) to equilibrate, which were determined by gamma counting using a germanium detector (GXM25P; EG & G ORTEC, Tokyo, Japan) equipped with a multi-channel analyzer (MCA7700; SEIKO EG & G, Tokyo, Japan) at the Center for Marine Environmental Studies, Ehime University. The activity of supported 210Pb was estimated by measuring the activity of 214Pb, whereas that of 210Pbexcess was determined according to the difference between the total and the supported 210Pb (210Pbexcess = 210Pbtotal − 214Pb). The age and age error of the remaining cores (LB1, LB2, and LB4) were indirectly estimated using stratigraphic correlations between the cores based on chronological controls in chlorophyll pigments and magnetic susceptibilities of the chronological LB7 core61. To compare these proxies, the marked peak or trough layers were used as reference layers (Supplementary Fig. S3).DNA extraction and purificationDNA extraction in the sediment samples was performed according to methods described in previous studies45,62. In brief, 9 g of each sediment sample was incubated at 94 °C for 50 min in a 9 mL alkaline solution comprising 6 mL of 0.33 M sodium hydroxide and a 3 mL Tris–EDTA buffer (pH 6.7). After centrifugation at 10,000×g for 60 min, 7.5 mL of the supernatant of the alkalized mixture was neutralized with 7.5 mL of 1 M Tris–HCl (pH 6.7). After adding 1.5 mL of 3 M sodium acetate (pH 5.2) and 30 mL absolute ethanol, the solution was preserved at − 20 °C for more than 1 h and then centrifuged at 10,000×g for 60 min. The pellet was transferred into a power bead tube that was installed in a fecal-soil DNA extraction kit (Power Soil DNA Isolation Kit, Qiagen, Germany). The ‘Experienced User Protocol 3 to 22’ of the Power Soil DNA Isolation Kit was followed. Finally, 200 μL of the DNA solution was obtained and stored at − 20 °C until qPCR analysis.12S rRNA gene primer-prove development for Daphnia geleata and Daphnia pulicaria
As the primer–probe for Daphnia galeata and D. pulicaria in qPCR analysis were not purchased by a company, thus we developed them for the two Daphnia species (see Supplementary Table S1). We preliminary obtained the mitochondrial 12S, 16S and COI gene of Daphnia genus from the National Center for Biotechnology Information (NCBI, http://www.ncbi.nlm.nih.gov/) and compared among them. From the preliminary results, we decided to use 12S because of the variability of sequences among Daphnia genus. Then we obtained the 12S sequences of Daphnia genus and other inhabiting plankton species in Lake Biwa, including Copepoda. We designed the primer–probe using Primer3plus (https://www.bioinformatics.nl/cgi-bin/primer3plus/primer3plus.cgi). The reference sequences for the targeted gene regions are queried for potential amplicons between 50–150 bp using NCBI primer blast. The specificities of the primers and probes were then assessed in silico with homologous sequences from other Daphnia species in Japan using NCBI targeting 154 bp of the mitochondrial 12S rRNA gene. Once suitable amplicons are found the respective primers and probes are tested against template DNA originating from the species of D. galeata and D. pulicaria to verify amplification. During the in silico screening for specificity, we performed Primer-BLAST (http://www.ncbi.nlm.nih.gov/tools/primer-blast/). We checked all species from Japan of the order Daphnia. Using the D. galeata primer-set, we did not detect any Daphnia species. However, the D. pulicaria primer can amplify D. pulex DNA, as these species are known to have very similar sequences47. In Lake Biwa, another subgenus Daphnia (D. pulex group) different from D. pulicaria was temporally found around during the 1920s, although thereafter it was never reported47. Thus, the D. pulicaria primer may temporally detect another subgenus Daphnia (D. pulex group). However, their appearance time do not overlap, therefore we used the primer for our measurement to detect D. pulicaria during the last several decades.Quantitative PCRThe DNA samples were quantified by real-time TaqMan quantitative PCR using the PikoReal Real-Time PCR system (Thermo Fisher Scientific, Waltham, MA, USA). The primer–probe sets for the two Daphnia species were used for qPCR (Supplementary Table S1). The TaqMan reaction contained 900 nM of each forward and reverse primer, 125 nM TaqMan-Probe, 5 μL qPCR master mix (TaqPath; Thermo Fisher Scientific), and 2.0 μL sedimentary DNA solution. The final volume of the PCR was 10 μL after adding distilled water (DW). The qPCR conditions were as follows: 50 °C for 2 min and 95 °C for 10 min, followed by 50 cycles of 95 °C for 15 s and 60 °C for 60 s. We used a dilution series of 10,000, 1000, 100, and 10 copies per PCR reaction (n = 4) for the standard curve using the target DNA cloned into a plasmid. The R2 values of the standard curves ranged from 0.988 to 0.996 (PCR efficiencies = 93.1–102.0%). The quantitative data of the DNA copies (copies g−1 dry sed.) were reported by mean values ± standard deviation, which were calculated from DNA copies µL−1 PCR reaction with four replicates including zero (i.e., no detection). We also performed four replicates for each sample and an NTC (n = 4). No positives were detected from the NTC and the negative control of DNA extraction, confirming that there was no cross-contamination in any of the DNA measurements.To confirm primer specificity, an in vivo test for the primer/probe set was performed using the extracted DNA (10 pg per PCR reaction, n = 4) of D. galeata and D. pulicaria. In addition, qPCR amplicons were sequenced directly from a positive PCR from each site (n = 21) after treatment with ExoSAP-IT (USB Corporation, Cleveland, OH, USA). Sequences were determined using a commercial sequencing service (Eurofins Genomics, Tokyo, Japan).Inhibitor testSpike tests were performed for the LB2 core sample to evaluate the PCR inhibition effect of several substances and minerals in the sediment samples (Fig. 1c). For the spike test, 1 µL plasmid, including the internal positive control (IPC, 207-bp, Nippon Gene Co. Ltd., Tokyo, Japan;100 copies per PCR reaction), was added to the PCR template with 1.6 µL DNA-free DW. We used the primer and probe sets for IPC as follows:

IPC1-5′: CCGAGCTTACAAGGCAGGTT

IPC1-3′: TGGCTCGTACACCAGCATACTAG

IPC1-Taq: [FAM] TAGCTTCAAGCATCTGGCTGTCGGC [TAMRA].

To measure the relative degree of PCR inhibition in the samples, the Ct shift was compared between the samples and controls with the same number of known target DNA copies. The presence of PCR inhibitors was evaluated as ΔCt = Ct sample − Ct positive control. ΔCt ≥ 3 cycles was considered evidence of inhibition63 because the presence of PCR inhibitors will delay the Ct with a given quantity of template DNA.
Daphnia abundance and resting egg production as potential sources of Daphnia DNA archived in sedimentsTo unveil the potential source of sedimentary DNA of Daphnia, we reconstructed the historical variation in Daphnia abundance by counting remains of the post abdominal claw for LB7 core. There are two dominant Daphnia species: D. galeata Sars (Hyalodaphnia) and D. pulicaria Forbes (Daphnia)47,61, which have different post-abdominal claw characteristics64 and are known to be preserved in centuries-old sediments65. The post-abdominal claw remains were counted for core LB7 from the surface to a depth layer of 21.5 cm and additionally 23.5 cm, 25.5 cm, and 29.5 cm, totaling 25 samples, though each layer was expressed as mid-depth; e.g., 0.5 cm for the 0–1 cm depth layer. The enumeration method was based on a simplified standard method65 as previously reported29.Daphnia resting eggs enveloped by thickened carapaces, referred to as ephippial cases, and these ephippia can be preserved in sediments for decades to centuries29,30,33. In Lake Biwa, Daphnia species in Lake Biwa are distinguished on the basis of the size of the ephippium, with a boundary length of approximately 860 μm between them61. We collected ephippia from the surface to a depth layer of 29.5 cm for cores LB1 and LB4 (except for several layers of the LB1 core), totaling 56 samples (Supplementary Table S4). A detailed method for collecting ephippia is described in a previous study61. The total number of collected ephippia with an almost perfect shape, namely complete formation, or with a partial body constituting more than half of the original shape, namely incomplete formation, are shown in Supplementary Table S4. In our study, at least 16 ephippia in each sample were measured by photographs taken by a digital camera, excluding those from the samples in which fewer than 16 complete ephippia were detected (Supplementary Fig. S4). Species identification was then performed based on length.To determine whether the Daphnia sedimentary DNA concentrations were regulated by DNA derived directly from Daphnia remains or ephippia included in the analytical sediment, we divided the sediment sample into two fractions to exclude the remains and ephippia (Fig. 1d). The minimum size of Daphnia remains in this lake was approximately 55 μm (Tsugeki et al., in preparation). The analytical sediments for DNA extraction were divided into particles  38 μm using 38-μm mesh sieves on three-layer samples (specifically, LB2-5; 4.5 cm, LB2-7; 6.5 cm, and LB2-17; 16.5 cm expressed in middle depth of each sample) for core sample LB2, whose layers were known to include abundant ephippia and Daphnia remains. Furthermore, to test the possibility of the vertical movement of Daphnia sedimentary DNA through pore waters, we examined the sedimentary DNA concentration in pore water and its residual sediment by qPCR analysis (Fig. 1e). All DNA extractions were evaluated for sediment with and without sieves, and pore waters and the associated residual sediment samples were evaluated according to previous studies45.Measurement of DNA concentration in sediment ephippiaTo determine the potential source of sedimentary Daphnia DNA, we quantified the DNA concentration extracted from several ephippia obtained from the 0–5 cm and 5–10 cm layers of core IM8 using qPCR analysis (Fig. 1f). We selected 34 and 23 ephippia for D. galeata and D. pulicaria, respectively. We then measured the ephippial lengths and determined whether they contained resting eggs using a microscope. Among the selected ephippia, the well-preserved 17 ephippia with almost complete formation were set aside and grouped into 6 samples together in two or three ephippia for DNA analysis (Supplementary Table S5). Grouping was performed because of the low DNA concentrations typically associated with individual ephippium61.Possible factors regulating sedimentary Daphnia DNATo explore potential factors regulating temporal variation in sedimentary DNA concentrations, we analyzed chlorophyll pigments and algal remains. Sedimentary pigments of chlorophyll a were investigated for the LB 2 core, and algal remains were investigated for the LB7 core (Fig. 1a). Details of the method used for chlorophyll-a and algal remains are described in previous study61. In short, the concentrations of chlorophyll-a and phaeopigments were calculated according to the method66 and the diatom remains were analyzed according to the simplified method67. Green algae, Micrasterias hardyi, Staurastrum dorsidentiferum, S. arctiscon, S. limneticum, S. pingue, and Pediastrum biwae, were enumerated in a Sedgewick–Rafter chamber, following the method of zooplankton enumeration.Data analysisRegression models along with the standardized major axis method were used to determine the relationship between the sedimentary DNA concentration obtained from qPCR analysis and abundance or resting egg production in the sediment layers. Since qPCR (LB2), remains (LB7), and ephippia (LB1, LB4) analyses were performed on different cores, the chronological age of each analytical sample differed slightly. Therefore, prior to performing the statistical analysis, the sedimentary DNA (LB2) and ephippia data (LB1, LB4) in each chronological age were converted to annual data by linear interpolation and averaged for the year corresponding to the period in each sample of the chronology core (LB7). This conversion was possible because the time resolution at 1-cm intervals represented several years, depending on the sediment depth29,61. We employed the Gaussian type II model because our preliminary evaluation showed higher R2 values for type II regression models with a Gaussian distribution than for those with a logarithmic distribution, in all cases. All statistical analyses were performed using R ver. 4.0.3 (R Core Team 2020) with the package “smatr” ver. 3.4-8 for type II regressions. The significant criteria of all analyses were set as α = 0.05. In addition, to explore the potential environmental factors driving temporal variation in sedimentary DNA concentrations, we performed Pearson’s correlation analysis among the sedimentary DNA concentrations, chlorophyll a concentration, and algal remains using the SPSS version 20.0 statistical package. More

Changepoint detection methodAlthough most of the reflectance time series used in the BinSeg–Normal–Mean and BinSeg–Normal–MeanVar algorithms had a normal distribution, several lagoons had distributions that were skewed or did not follow a normal distribution (Fig. S1). However, results suggested that the accuracy of the detected changepoints were not sensitive to the normality assumption or distributional characteristics.The BinSeg-Normal-Mean algorithm had the highest performance (81% of the 340 validation sites) in detecting the correct year of swine waste lagoon construction, followed by BinSeg-Normal-MeanVar (77%). The two algorithms did not detect the same year of construction for 19 waste lagoons; of these 19, the BinSeg-Normal-Mean detected the correct year for 84% of them, while the BinSeg-Normal-MeanVar detected the correct year for only 16%. Therefore, the BinSeg-Normal-MeanVar algorithm was abandoned given it did not provide additional useful information relative to the BinSeg-Normal-Mean algorithm.Despite good performance, the BinSeg-Normal-Mean algorithm consistently detected a changepoint during the period of record for all sites included in the 10% validation set (n = 340 swine waste lagoons). However, 58 of the 340 swine waste lagoons were constructed prior to 1986, before the period of record suitable for detecting an accurate changepoint. Changepoints before 1986 either (1) detected the correct construction year, or (2) incorrectly detected a changepoint due to artifact signals identified on the images taken in 1984, probably associated with the initial satellite commissioning. In the latter circumstance, if the algorithm detected a changepoint due to this signal, it meant that no land-use change was detected after 1986. Therefore, these waste lagoons were estimated as having been constructed before 1986. In some conditions, when a large number of images was available for the year 1985 and 1986, the algorithm was able to detect the changepoint occurring for the years 1985 or 1986. Further, the BinSeg-Normal-Mean algorithm detected a false year of construction for swine waste lagoons for which the mean of the segment after the changepoint (S2) had a greater average than the segment before the changepoint (S1).To increase algorithm performance, we developed a workflow to address some of the aforementioned caveats (Fig. 4). In this workflow, the BinSeg-Normal-Mean algorithm is applied to a B4 reflectance time series at location j. If the BinSeg-Normal-Mean changepoint is identified for a time in or prior to 1986 (Fig. 4a,i,b,i) we assume that the lagoon was constructed in or prior to 1986. Similarly, a lagoon is assumed to be constructed in or prior to 1986 if a BinSeg-Normal-Mean changepoint is identified after 1986 and the mean of S2 is greater than the mean of S1 (Fig. 4a,ii,b,ii). If a changepoint occurred after 1986 and the mean of S1 was greater than S2, then the changepoint was estimated as having occurred between 1987 and 2010 (Fig. 4a,iii,b,iii).Figure 4Changepoint detection algorithm for determining the year of construction of swine waste lagoons. Panel (a) summarizes the algorithm workflow, while panel (b) illustrates specific examples corresponding to each step (i–iii) in the workflow.Full size imageThe performance of the workflow was evaluated using the validation set composed of 10% of the total number of swine waste lagoons (n = 340). With the new approach, 94% of the swine waste lagoon construction years (+ /- one year) were accurately retrieved. A tolerance of + /− 1 year was chosen to account for a lack of images in some years due to issues with image quality (e.g. high cloud cover) (e.g., Fig. 5a), or because construction spanned at least a year (e.g., Fig. 5b). The changepoint detection workflow incorrectly estimated the construction years for 19 of the 340 swine waste lagoons in the validation set; the differences between the observed and predicted years of construction of these lagoons ranged from 2 to 26 years with a median of 8 years.Figure 5Examples of limitations to the changepoint detection algorithm. In some cases, an insufficient number of high-quality Landsat 5 images were available to capture the year of construction of an individual swine waste lagoon (a), resulting in errors of + /− 1 year. In other cases, the changepoint algorithms detected the start of the construction of the swine waste lagoon but the swine waste lagoon was not fully operational until later years due to prolonged construction timelines (b).Full size imageBy visually inspecting historical Google Earth images for each of the lagoon sites for which the model incorrectly estimated construction year, we identified that model errors were associated with swine waste lagoon expansion, pixel transitions to land-use classes other than swine waste lagoons, or issues with pixels being partly covered by clouds or incompletely covered by the lagoon (i.e., narrow and small waste lagoons that do not entirely cover a pixel).Estimating swine waste lagoon construction yearsUsing the newly developed algorithm (Fig. 4), construction years were estimated for each swine waste lagoon in the NC Coastal Plain (Fig. 6); the years of construction for each swine waste lagoon are included in the supplementary material. Most swine waste lagoons were built in the early 90s and prior to the moratorium of 1997. More specifically, 80% of the swine waste lagoons (n = 2,736) were built between 1987 and 1997. Sixteen percent of the swine waste lagoons were constructed in or prior to 1986. A large decrease in the construction of swine waste lagoons occurred after the moratorium of 1997, with only 3.7% of swine waste lagoons being constructed after the moratorium. These results suggest that the 1997 moratorium did not completely halt the construction of lagoons, but dramatically slowed the rate of expansion.Figure 6Spatiotemporal distribution of swine waste lagoon construction (+/- 1 year) across the HUC6 watersheds. This figure was produced using QGIS version QGIS 3.18.3 (https://www.qgis.org/).Full size imageWith regards to hydrological boundaries (Fig. 7a–h), the Cape Fear River watershed had the highest number of swine waste lagoons (i.e., 56%; Fig. 7b), followed by the Neuse River (i.e., 23%; Fig. 7d), the Lower Pee Dee River (i.e., 9%; Fig. 7c) watersheds. The Albemarle-Chowan (Fig. 7a), Onslow Bay (Fig. 7e), Pamlico (Fig. 7f), Roanoke (Fig. 7g), and Upper Pee Dee (Fig. 7h) watersheds all had less than 9% of the total lagoons within the study area.Figure 7Year of construction of the swine waste lagoons (+ /− 1 year) for the HUC6 watersheds. The y-axis scales are unequal between the plots to improve readability. The dashed red lines correspond to the establishment of the moratorium in 1997.Full size imageResults suggested that the Cape Fear River watershed was the center of the historical growth of the swine industry, where over 300 swine waste lagoons were built prior to 1987. The Cape Fear River watershed experienced a steady increase in the number of swine waste lagoons from 1987 to 1990, with an average of 46 swine waste lagoons being built annually. However, after 1991, the pace of swine waste lagoon construction increased dramatically with an average of 192 swine waste lagoons built annually between 1991 and 1997. The highest construction rate occurred in 1994, with 242 swine waste lagoons built. However, after the 1997 moratorium, the construction rate decreased dramatically; in 1997, 153 swine waste lagoons were constructed, and this number dropped to 23 in 1998. After 1998, the annual average number of swine waste lagoons constructed plunged to 5. Although the swine waste lagoon construction rate fell considerably after the 1997 moratorium, the decrease had already started in 1995. The same pattern was observed for the Neuse, Pamlico, Albemarle-Pamlico, and Onslow Bay watersheds.The spatiotemporal distribution of swine waste lagoons at the HUC12 watershed scale emphasized the historical clustering of the swine industry in the NC Coastal Plain. After the moratorium, swine waste lagoons were present within 436 HUC12 watersheds. However, before 1986, they were spread across only 197 HUC12 watersheds (Fig. 8). Before 1986, the density of waste lagoons was relatively low with an average of 3.38 swine waste lagoons per 100 km2 and a maximum of 15.13 swine waste lagoons per 100 km2 (i.e., Clayroot Swamp-Swift Creek watershed) (Fig. 8). In the 90s, swine waste lagoon construction expanded and continued to intensify in the region. After the moratorium of 1997, the average density of waste lagoons per HUC12 watersheds was 10 per 100 km2 with a maximum of 78 waste lagoons per 100 km2 identified in the Maxwell Creek-Stocking Head Creek basin. After 1997, 16 of 436 HUC12 watersheds had a swine waste lagoon density greater than 40 per 100 km2 (Fig. 8).Figure 8Cumulative swine waste lagoon density per 100 km2 reported at the HUC12 watershed scale; HUC6 watersheds shown in gray for reference. This figure was produced using QGIS version QGIS 3.18.3 (https://www.qgis.org/).Full size imageSpatiotemporal distribution of swine waste lagoons in relation to water resourcesDistance of swine waste lagoon sites to the nearest water feature (i.e., reservoir, canal/ditch, lake/pond, stream/river, estuary) were assessed using the NHD. The analysis revealed that over 150 swine waste lagoons were misclassified by the NHD and were documented in the NHD as lake/pond (n = 102) or swamp/marsh (n = 46). Further, we observed that some NHD water features were misclassified as other non-water features (e.g., forest, pasture), and most of these misclassifications were for polygons with an area less than 0.05 km2. Therefore, NHD water features with areas less than 0.05 km2 were removed from subsequent analyses. Distances between swine waste lagoons and waterways were computed from the NHD without features with areas less than 0.05 km2. The new analysis revealed that 3 swine waste lagoons remained misclassified as lake/pond (n = 1) and swamp/marsh (n = 2). Canal/Ditch, lake/pond, stream/river, and swamp/marsh were identified as the NHD features that were most commonly near swine waste lagoons (Fig. 9). Two swine waste lagoons were near a reservoir in which one was identified as a treatment-sewage pond by the NHD.Figure 9Nearest water features distance to swine waste lagoons.Full size imageThe average and median distance of all swine waste lagoons (including those built early and late in the period of record) to the nearest water features were 234 and 177 m, respectively. Further, 92% of the swine waste lagoons were less than 500 m from the nearest waterways. The Mann–Kendall results revealed a significant upward trend over time of swine waste lagoon distances to the nearest water features (alpha = 0.05, p-value = 0.01). A slight increase over time of swine waste lagoon distances to the nearest water feature is also documented in Table 1.Table 1 Temporal average and median of nearest distance (m) of swine waste lagoons to water features. NA indicated that the water feature was not the closest waterway to any of the studied swine waste lagoons for the time period.Full size table More

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