{"id":27226,"date":"2019-01-23T00:10:01","date_gmt":"2019-01-23T00:10:01","guid":{"rendered":"http:\/\/www.ocean4future.org\/?p=27226"},"modified":"2026-04-16T13:20:32","modified_gmt":"2026-04-16T11:20:32","slug":"27226","status":"publish","type":"post","link":"https:\/\/www.ocean4future.org\/savetheocean\/archives\/27226","title":{"rendered":"Persistence of environmental DNA in marine systems – part II Rupert A. Collins, Owen S. Wangensteen, Eoin J. O\u2019Gorman, Stefano Mariani, David W. Sims & Martin J. Genne"},"content":{"rendered":"tempo di lettura: <\/span> 19<\/span> minuti<\/span><\/span>

.<\/span>
\nlivello difficile<\/span><\/a>
\n.<\/span><\/p>\n

ARGOMENTO: BIOLOGIA<\/span><\/strong>
\nPERIODO: XXI SECOLO<\/span><\/strong>
\nAREA: RICERCA<\/span><\/strong>
\nparole chiave: DNA, marine systems<\/p>\n

 <\/p>\n

Discussion<\/span><\/strong><\/span>
\nOur results show evidence for a strong spatial effect of eDNA degradation in the natural marine environment, with eDNA degrading around 1.6 times faster in the terrestrially influenced inshore environment than the ocean-influenced offshore environment. We found that eDNA also degraded around 1.1 times faster in winter than in summer, although this difference was not statistically significant. These results placed in the context of our review of eDNA decay rates in the literature (Table 4), appear to contradict the notion that eDNA degrades faster in marine environments than freshwater 18,20,29. In fact, degradation rates appear to be slower in many cases, with only marine studies or the freshwater studies at low temperature or using non-natural water sources, having a half-life of greater than 30\u2009h (Table 4). The fastest rates in freshwater systems assessed so far are of acidic stream environments (<1.2\u2009h of half-life5), while the fastest marine decay rate was 6.9\u2009h, from anchovy eDNA in Californian inshore waters at 22\u2009\u00b0C20. Most marine eDNA decay rates appear, however, to have been estimated at between 10 and 50\u2009h, and with the lowest rates corresponding to the coldest water temperatures: 63\u2009h at 4\u2009\u00b0C19 and 71\u2009h at \u22121\u2009\u00b0C15. Rates above 71\u2009h were from freshwater studies using sanitised or purified water from non-natural sources (Table 4).<\/p>\n

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table 4 – a<\/span><\/strong><\/figcaption><\/figure>\n
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table 4 – b<\/span><\/strong><\/figcaption><\/figure>\n

Compared with freshwater, marine systems are generally characterised by higher salinity and ionic content, typically higher pH, and more stable temperatures. These are factors which have been shown to promote DNA preservation, and tend to correspond to the lowest observed degradation rates5,14,23,25,30,31. Our artificial spatial gradient varied from an offshore treatment with high pH and salinity to an inshore treatment with a lower salinity and a slightly lower pH. This was designed to capture the abiotic heterogeneity that could be expected across the Western English Channel region over the period of a year, a magnitude of variation that will apply to other coastal temperate locations. We found salinity to be a better predictor of eDNA decay than pH, and with salinity varying more between locations and pH varying more over seasons (Table 3), this agrees with the finding that the spatial signal was stronger than the temporal signal, and is reflected in the correlation matrix of the combined predictor-covariate model.<\/p>\n

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table 3<\/span><\/strong><\/figcaption><\/figure>\n

The lack of a statistically significant difference over season may be due to the relatively low degree of variation in pH and temperature. Seawater pH measured in this experiment was between around 8 and 8.6, which may not have any direct impact on DNA hydrolysis, and likewise, temperature ranges in this temperate marine system (10\u201315\u2009\u00b0C) were narrower than those typically studied in terrestrial systems (e.g. 5\u201335\u2009\u00b0C12).<\/p>\n

As well as abiotic factors engaging in DNA degradation via oxidisation and hydrolysis by depurination, biotic factors are also likely to play a major role in eDNA persistence dynamics via extracellular DNases produced by heterotrophic microbes4,30. While we found support for faster degradation rates in our inshore and mixed treatments (Fig. 3), this difference did not appear to be proportional to the quantity of inshore water used in the treatment\u2014the two-third offshore treatment tended to be closer to the 100% inshore treatment than the 100% offshore treatment\u2014suggesting that biotic rather than abiotic factors are of stronger influence. Salinity itself may not be therefore entirely responsible for the difference in decay rate, rather that it is associated with particular abundances or communities of microbes. Gilbert et al. 32 reported that microbial community structure in the Western English Channel was highly dynamic seasonally. Free DNA is thought to represent an important organic phosphorus source in marine systems 29, and seasonal phosphate limitation has been identified as a key driver of eDNA turnover rates over abiotic factors such as temperature, pH and salinity 26.<\/p>\n

Therefore, the lack of seasonal difference in eDNA decay that we report may also be explained by organic phosphorous or carbon concentrations 14,26.<\/span><\/strong><\/p>\n

Taken together with the survey of rates from the literature, this implies that abiotic and biotic factors are co-implicated in eDNA degradation. Assessing the covariance and contribution among these parameters is an area that needs to be addressed, along with more sophisticated analyses of microbial communities incorporating a greater degree of spatial replication. A number of systematic biases were identified as being potentially problematic for our inferences. PCR inhibition in the samples from the inshore site could explain the faster degradation rates from that location. However, we assessed amplification efficiency of the qPCR in a serial dilution experiment, and these were near the expected 100% across treatments and season. Values well above 100% would indicate inhibition. Other studies have also indicated low instances of PCR inhibition when using kits with dedicated inhibitor removal steps such as the PowerWater kit that we used 33,34. Although not significantly different, we found that degradation rates were overall around 1.2 times faster in the crab assay than the shanny assay. This is most likely explained by differences in fragment length between the two assays (153 vs. 132\u2009bp), with longer fragments being shown to decay at a faster rate than shorter fragments 22. It was also noted that despite using a similar mass of crabs and shannys to create the eDNA, initial measured concentrations were roughly an order of magnitude lower in the crab assay (Fig. 1), perhaps indicating that the exoskeleton of the crustaceans, as well as their behaviour and breeding condition at particular times of the year may limit eDNA output 35.<\/p>\n

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Fig. 1<\/span><\/strong><\/figcaption><\/figure>\n

The treatment of qPCR non-amplifications in low-template analyses is an important source of error at the analytical stage. Due to the proportion of non-amplifications at the 192-h sample (0.56 in winter and 0.68 in summer)\u2014 i.e. outside of the experimental limit of quantification\u2014and the influence of this time point in estimating the regression slopes, our eDNA decay model was sensitive to how these missing data were treated. Excluding them, or fixing them to the limit-of-detection value resulted in the effects of season and assay becoming statistically significant. However, treating the non-amplifications this way is problematic as these missing data are not randomly distributed; the missing values will tend to be from samples of lower concentrations, and therefore the remaining positive values will then become overestimated 36. Our conservative approach was to follow Ellison et al. 37 and fix their value, although we used the lowest detectable concentration of the assay (13.7 copies\/L) rather than fixing the values at zero. Unfortunately, fixing values in this way is also problematic, creating a potential underestimate of concentration, and may interfere with the assumptions of linear regression. A better future strategy may be to avoid estimating decay rates from low copy-number time series, or to impute the missing data 36.<\/p>\n

Related to the issue of missing data is that of starting concentrations. Despite normalising each time sample as the proportion of the t\u2009=\u20090 starting concentration, we included in our model the initial value and found it to be a statistically significant predictor associated with faster degradation rates. The summer experiment and the crab assay had lower starting concentrations than the winter experiment and the shanny assay respectively (Fig. 1), but although the average crab-assay decay rate was faster than shanny, the average winter rates were faster than that of summer. Therefore, while they may not have influenced the results overall, a low starting concentration of eDNA resulted in the lower resolution of the summer crab experiment in particular, as qPCR quantification is increasingly stochastic and unreliable at low-template concentrations 38.<\/p>\n

In terms of implications for marine ecology, how do eDNA half-lives or decay rate constants relate to detectability of a given organism?<\/span><\/strong>
\nAs suggested by Sassoubre et al.20, reporting the duration of time until the detection limit is reached is misleading, as this value will depend upon the starting concentration of eDNA and the sensitivity of the assay; most studies use eDNA starting concentrations far higher than typical natural concentrations in order to generate reliable decay curves with less noise. Our negative biological controls provide an insight into natural concentrations. Sutton Harbour (our inshore treatment) is well populated with common shore crabs, and as expected, we recovered this species at approximate concentrations of 263 copies\/L (winter) and 270 copies\/L (summer). As the detection rate of the crab assay was 37% at 83 copies\/L, and the eDNA half-life inshore was around 24\u2009h, it is estimated that the chance of detection with three PCR replicates would be below the threshold after just two half-life periods (~48\u2009h). However, we did detect eDNA in at least one qPCR replicate from this control at all time points up to 192\u2009h (winter) and 48\u2009h (summer), indicating that eDNA detectability will be difficult to predict at very low concentrations. Quantitative PCR is known to be more sensitive than standard PCR combined with metabarcoding39. Thomsen et al.18 estimated similar values of natural eDNA to ours (535 copies\/L for flounder, 120 copies\/L for stickleback), and a similar detection limit (63 copies\/L). However, it must be noted that we did not consider the loss of DNA in the extraction process, which can be considerable with commercial kits that incorporate steps to remove PCR inhibitors34,40, or any loss of eDNA at the filtration stage, and therefore, real values are likely to be higher and comparisons among studies using different methodologies may be questionable.<\/p>\n

Ultimately, how eDNA persists and moves through an environment can have important repercussions for making meaningful ecological inferences, and it is important to document and understand the patterns and processes involved41,42. The combined issues of degradation, transportation and dilution of eDNA are of particular importance in the marine environment, due to the effects of tides and large water volumes 9,18,43. Fortunately, eDNA metabarcoding studies of marine systems have reported a strong local eDNA signal, either closely matching lists of expected fauna 18,44,45 or reporting an expected turnover in diversity over short spatial or temporal scales9,43,46. Most evidence therefore points to eDNA surveys offering a contemporaneous representation of a community, even over the variation encountered on a daily tide9. However, there are cases where non-resident freshwater species have been detected in marine eDNA studies47, and while this source of error can easily be discarded as clearly a riverine input, currents transporting possibly co-occurring marine species eDNA may cause a less obvious source of systematic bias. These biases may become more serious when eDNA is used in applications beyond determining occurrence, for example to monitor the spread of marine invasive species48 or correlating with animal biomass estimates49. By incorporating eDNA degradation rates in different types of water body with oceanographic modelling of tidal currents, it will be possible to build well-informed predictive probability maps of organismal distribution44,48,50. Until these are available, to our knowledge, we show for the first time that it is reasonable to assume large variation in eDNA persistence according to local factors such as salinity gradients over relatively short local scales corresponding to marine environmental stability.<\/p>\n

Methods<\/span><\/strong><\/span>
\nAssay design<\/span><\/strong>
\nStudy species were the shanny (Teleostei: Blenniidae: Lipophrys pholis) and the common shore crab (Decapoda: Portunidae: Carcinus maenas). These species were chosen because they are abundant hardy organisms amenable to transport and experimental manipulation. Reference specimens of shanny (eight individuals) and shore crab (four individuals) were obtained from the Gann estuary, Pembrokeshire, Wales (51.715, \u22125.173). Using standard molecular methods, we obtained DNA barcodes (COI; 5\u2032 mitochondrial cytochrome c oxidase I gene) for both species using the FishF1\/R1 primer set51. Additional sequence data for crabs (149 individuals) were obtained from GenBank; no GenBank COI sequences were available for shanny. Primers and hydrolysis probes were designed using Primer3 v1.1.452,53 under default settings adjusted to aim for an amplicon length between 50 and 170\u2009bp. The resulting 12 candidate primer pairs were tested in silico for general specificity against a dataset of sequences from species present in the United Kingdom. To generate a list of fishes and Malacostraca recorded from the United Kingdom, we searched the Global Biodiversity Information Facility (https:\/\/www.gbif.org) using the rgbif v0.9.9 package for R54. COI sequences for these species were then retrieved from GenBank and annotated using rentrez v1.2.155 and traits v0.3.0.931056. Each candidate primer pair was tested in an in silico PCR using MFEprimer v2.057 using liberal settings (k\u2009=\u20095). The final primers were then chosen based on a combination of amplicon length, specificity and melting temperatures, and are reported in Supplementary Table 1. The reporter dye for the shanny assay was FAM, and for the crab assay HEX; both were quenched using BHQ.<\/p>\n

Experimental setup<\/span><\/strong><\/span>
\nThe experiment was repeated twice, first in winter (water collected on 17 February, 2017) and once in late summer (water collected on 26 September, 2017). All treatments were set up in a dedicated temperature-controlled aquarium room held at temperatures consistent with natural seawater temperatures at that time of the year (10\u2009\u00b0C, winter experiment; 15\u2009\u00b0C, summer experiment). Animals were collected 2 days before the start of each experiment (also from the Gann estuary, Pembrokeshire) and placed in a separate and aerated holding tanks for each species (shanny, 50\u2009L of synthetic seawater; crabs, 25\u2009L). Approximately 300\u2009g of animal mass per species were collected (winter, 24 shannys at 343\u2009g of total weight, 18 crabs at 288\u2009g; summer, 25 shannys at 316\u2009g, 9 crabs at 304\u2009g). All animals were euthanised after the experiment was completed, and were formalin fixed and 70% alcohol preserved as voucher specimens for a reference dataset. All experiments were carried out in accordance with the University of Bristol ethical approval (UIN reference UB\/16\/012).<\/p>\n

A total of 24 aquariums at the University of Bristol Animal Services Unit were each filled with 9\u2009L of experimental water. The tanks were initially mixed but not aerated and were maintained under 12\u2009h of light\/dark LED room lighting. Five experimental water treatments were carried out as follows: 100% offshore sea surface water\u2014from herein referred to as ‘offshore’\u2014collected from Western Channel Observatory station E1 ~40\u2009km from Plymouth, Devon, UK (50.033, \u22124.367; Supplementary Fig. 2); inshore urban water\u2014from herein referred to as ‘inshore’\u2014collected from Sutton Harbour, Plymouth Sound, a site located between the estuaries of the rivers Plym and Tamar (50.370, \u22124.133; Supplementary Fig. 2); a two-thirds\/one-third mixture of offshore to inshore water; a one-third\/two-thirds mixture of offshore to inshore water; and synthetic seawater made using a proprietary aquarium salt mix. Each of the five treatments had four biological replicates (=20 tanks), plus four no-treatment controls (2\u00d7 synthetic seawater, 1\u00d7 offshore and 1\u00d7 inshore.<\/p>\n

After turning off aeration and allowing detritus to settle for an hour, 500\u2009mL of eDNA-rich surface water from both the shanny and crab stock tanks was then added to each experimental tank at the start of the experiment. At each subsequent time point, eDNA was filtered from 600\u2009mL of experimental tank water with a peristaltic pump and Sterivex 0.22-\u03bcm PES filters (Millipore part no. SVGP01050)58. Measurements were taken at six intervals from the same tanks (0, 12, 24, 48, 96 and 192\u2009h), resulting in 144 filtered water samples (24\u00d76). After being cleared of water, filters were frozen immediately at \u221220\u2009\u00b0C. DNA was subsequently extracted from the Sterivex filters using the PowerWater DNA isolation kit (MoBio\/Qiagen part no. 14900-100-NF) following manufacturers\u2019 instructions, but with 50\u2009\u03bcL of final elution volume. Extractions were carried out in a dedicated pre-PCR extraction room regularly decontaminated with 10% bleach and UV sterilisation. Environmental covariates were also measured from each tank with a Hach HQ40D multimeter, and included salinity (conductivity in mS\/cm), pH, and temperature at source. As a proxy for biological activity, we also recorded total background double-stranded DNA (dsDNA) concentration from 2\u2009L of source water with a Qubit 3 fluorometer (ThermoFisher) assay (filtered and extracted in the same way as the experimental treatments).<\/p>\n

Quantitative PCR<\/span><\/strong><\/span>
\nQuantitative PCR reactions were conducted as per the manufacturer\u2019s instructions, in multiplex, on a PCRmax Eco48 machine in 48-well plates of 5\u2009\u03bcL per reaction, with ROX normalisation. Each reaction volume comprised 2.5\u2009\u03bcL of mastermix (qPCRBIO Lo-Rox Probe mix; part no. PB20.21-05); 0.5\u2009\u03bcL of shanny\u2013crab primer-probe mix (optimised reaction concentration for shanny assay: 600\u2009nM each primer, 200\u2009nM probe; crab assay: 600\u2009nM each primer, 300\u2009nM probe); 1\u2009\u03bcL of water and 1\u2009\u03bcL of eDNA template. The cycling parameters comprised 3\u2009min at 95\u2009\u00b0C polymerase activation followed by 42 cycles of denaturation at 95\u2009\u00b0C for 5\u2009s and combined extension\/annealing at 60\u2009\u00b0C for 30\u2009s.
\nEach plate of 48 reactions comprised: eight extracted water samples of the experimental tanks, with three technical replicates per sample (8\u2009\u00d7\u20093\u2009=\u200924 reactions); a six-fold standard-curve serial dilution of 1\u20131 million copies\/\u03bcL, in triplicate (=21 reactions); and three no-template controls (=three reactions). To allow low-copy-number templates, an increased opportunity to amplify, PCRs were repeated a further three times for each sample when there was no amplification in any of the three initial technical replicates (excluding negative controls). The standard curve stock solutions were generated by PCR-amplifying and purifying tissue extractions of genomic DNA in a standard PCR using the primers in Supplementary Table 1, and were subsequently diluted and quantified using a Qubit assay, with the number of copies estimated at a standard dsDNA molar mass of 650\u2009g 28. We tested for PCR inhibitors by performing triplicate qPCRs on three serial dilutions of the 0 h replicates from three treatments (synthetic, inshore, offshore) over both seasons (total 24 samples). If inhibitors were co-extracted, the cycle threshold (Ct) values at each tenfold dilution point would deviate from the expected increase of 3.3 PCR cycles, and therefore the expected efficiency values of 90\u2013110% 28.<\/p>\n

Analysis<\/span><\/strong><\/span>
\nCycle threshold values and target DNA concentrations were calculated on the Eco48 machine software using the default settings, and converted from copies per reaction (=copies\/\u03bcL given a 1-\u03bcL template volume) to copies\/L of initial sample water (given a 600 mL filtration volume and a 50 \u03bcL elution volume). All amplifications were checked manually in the log plot view and any amplifications that crossed the baseline threshold, but that did not represent a clean, obviously exponentially increasing reaction, were excluded. The final eDNA concentrations for each sample were averaged over the technical replicates, with non-amplifications included as an arbitrarily low but non-zero value of 13.7 copies per litre of sample water (Ct\u2009=\u200938.5; the lowest concentration that the assay reliably detected).<\/p>\n

Statistical analyses were conducted in R v3.5.159. Decay of eDNA was modelled using a linear mixed-effects model as implemented in the lme function of nlme v3.1-13760. The response variable was natural loge transformed eDNA concentration normalised as a proportion of starting concentration, i.e. the value at time t\u2009=\u2009x divided by the value at time t\u2009=\u20090. We specified time, treatment, season, assay, and the natural log of eDNA starting concentration as predictor variables (our fixed effects), while the individual tank used in each biological replicate was treated as a random effect. To minimise heteroscedasticity\u2014i.e. the increasing variance of regression residuals over time\u2014we excluded the normalised zero-hour (t\u2009=\u20090) data, which had no variance. The synthetic water control was also excluded from the main model\u2014this was a control for reference rather than to investigate its biological effect\u2014and decay rates for this subset were calculated separately (following the same procedure as outlined below).<\/p>\n

We determined the optimal model to fit our data according to the procedure of Zuur et al.61. We started with a full model containing all fixed effects and their interactions, and determined the optimal variance weighting for different treatment\u2013season\u2013assay combinations by AIC comparison (given by the form weights\u2009=\u2009varIdent (form\u2009=\u20091|treatment*season)). We then determined the optimal random structure for the full model with this variance weighting by AIC comparison (given by the form random\u2009=\u20091\u2009+\u2009time|tank). Finally, we determined the optimal fixed effects structure using the ‘drop1’ approach and specifying method\u2009=\u2009\u2019ML\u2019 until all terms in the model were statistically significant. We switched to method\u2009=\u2009\u2019REML\u2019 and performed model validation to ensure that the model residuals were approximately normal and homogeneously distributed (see Supplementary Fig. 3). The fixed effects structure and output for the full model and the optimal model are also presented in Supplementary Note 1<\/a><\/strong>.<\/p>\n

The first-order decay-rate constant k for each treatment\u2013season\u2013assay combination was calculated from the estimated marginal mean of regression slopes using the emtrends function of emmeans v1.2.362. To test the importance of predictor variables on the degradation rate, pairwise post hoc Tukey tests were carried out on the marginal mean regression slopes, again using emmeans. To explore the environmental covariates we constructed a simple lme model with time, assay, pH, conductivity, natural log transformed starting concentration, and background DNA concentration as fixed effects, and tank as a random effect. For this model, we excluded the treatment and season predictors\u2014which were deliberately chosen for their heterogeneity\u2014as we assumed these to be correlated with the environmental covariates. We additionally included them in a combined model to estimate the degree of correlation between the predictors and covariates.<\/p>\n

 <\/p>\n

 <\/p>\nLINK TO PART I<\/span><\/a>\n

 <\/p>\n

Original study Persistence of environmental DNA in marine systems | Communications Biology (nature.com)<\/a><\/p>\n

 <\/p>\n

 <\/p>\n

Code availability<\/span><\/strong>
\nThe code generated during and\/or analysed during the current study is available in the Figshare repository 65<\/a><\/p>\n

Data availability<\/span><\/strong>
\nThe datasets generated during and\/or analysed during the current study are available in the Figshare repository 65<\/a>. New sequence data generated here were deposited in the GenBank nucleotide archive under the accessions MH931374:MH931388.<\/p>\n

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tempo di lettura: <\/span> 19<\/span> minuti<\/span><\/span>. . ARGOMENTO: BIOLOGIA PERIODO: XXI SECOLO AREA: RICERCA parole chiave: DNA, marine systems   Discussion Our results show evidence for a strong spatial effect of eDNA degradation in the natural marine environment, with eDNA degrading around 1.6 times faster in the terrestrially influenced inshore environment than the ocean-influenced offshore environment. 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