Soil extracellular DNA fragments show variable degradation rates among sequences and environmental conditions

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This valuable study introduces an innovative experimental design to address a crucial and timely issue in microbial ecology: the potential bias in soil microbial community analyses caused by extracellular DNA degradation. The work contributes to the field by providing convincing evidence that variable extracellular DNA degradation rates impact microbial ecology studies. This research will appeal to microbial ecologists and researchers interested in using molecular techniques to evaluate microbial community structure.

https://doi.org/10.7554/eLife.110251.3.sa0

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Abstract

While extracellular DNA (eDNA) persistence substantially influences soil microbiome investigations, its degradation kinetics remain poorly quantified. Here, we developed a primer-labeled DNA approach coupled with microcosm incubation to determine the overall and sequence-specific degradation rates of eDNA amplicon fragments across China. We observed substantial variations in the overall degradation rates of extracellular 16S rRNA gene amplicon fragments among the study sites, with degradation rate constants ranging from 0.05 to 0.16 day⁻¹. The overall degradation rate constants showed significant correlations with soil moisture content, prokaryotic abundance, prokaryotic community profiles, and mean annual precipitation. The significant influences of moisture content on the overall degradation rates were further verified by a moisture gradient microcosm experiment. The sequence-specific degradation rate constant profiles were additionally correlated with pH, nitrogen content, and mean annual temperature. Furthermore, propidium monoazide-based exclusion of eDNA signals significantly altered soil prokaryotic abundance, richness, and prokaryotic community profiles, and the pool sizes of sequence-specific extracellular 16S rRNA gene amplicon fragments were significantly correlated with their respective degradation rates. This study developed a methodology for determining the overall and sequence-specific degradation rates of eDNA amplicon fragments, highlighting the profound influences of eDNA on soil microbial research and informing the optimization of environmental DNA technologies.

Introduction

The investigation of soil microbial abundance and diversity heavily relies on DNA-based technologies, such as real-time PCR, high-throughput amplicon sequencing, and metagenomic analysis (Che et al., 2018; Du et al., 2020; Yang et al., 2023). Soil DNA originates from both living and relic microbial cells, with the DNA from deceased microbial cells commonly referred to as relic or extracellular DNA (eDNA) (Ye et al., 2022). eDNA serves as a critical vector for horizontal gene transfer, facilitating the uptake of genetic material by competent microorganisms and promoting the spread of functional traits such as antibiotic resistance (Liu et al., 2024). In addition, eDNA participates in soil biogeochemical cycling because its enzymatic degradation releases bioavailable nutrients, particularly phosphorus and nitrogen, which can be reused by soil microorganisms (Ye et al., 2022). Moreover, the prevailing paradigm of total DNA extraction in soil microbiome studies introduces eDNA as a critical noise factor. Generally, its environmental persistence can lead to overestimation of microbial diversity in amplicon sequencing (Barnes et al., 2014; Wang et al., 2025; Xue et al., 2025) and distort qPCR-based quantification of marker genes (Carini et al., 2016; Sun and Ge, 2023; Wang et al., 2024b). A model simulation study showed that the extent of this influence is primarily determined by the abundance and sequence-specific degradation rates of eDNA (Lennon et al., 2018). Therefore, determining the overall and sequence-specific degradation rates of soil eDNA can provide crucial insights into evaluating the effects of eDNA on soil microbial analyses and the reliability of research based on environmental DNA technologies.

The overall degradation of soil eDNA has received significant attention due to its crucial roles in nutrient cycling and horizontal gene transfer (Nagler et al., 2018). Early studies mainly assessed eDNA persistence and degradation using PCR, DNA hybridization, radioisotope labeling, and competent cell transformation techniques (Paget et al., 1992; Zhang et al., 2020; Samuels et al., 2025). These investigations consistently demonstrated that eDNA can persist in soils for months to years (Barnes et al., 2014; Pathan et al., 2020), maintaining its capacity to transform competent cells (Levy-Booth et al., 2007; Pietramellara et al., 2009). Most of these studies primarily concentrated on assessing the risks associated with transgenic technologies, narrowly examining the degradation dynamics of DNA sequences related to such technologies. Consequently, the insights they provided were usually qualitative or semiquantitative (Morrissey et al., 2015), limiting the comprehensive understanding of eDNA degradation dynamics. More recently, there has been a notable shift towards quantification, and the dynamics of soil eDNA were quantified in many studies (Eichmiller et al., 2016; Wei et al., 2018). For instance, the degradation rates of soil eDNA were quantified using real-time PCR with specific plasmid labels such as T7 and SP6 promoters (Ceccherini et al., 2009). Another study simultaneously determined the dynamics of both soil microbial community and eDNA by adding 16S rRNA gene primer labels to exogenous DNA (Sirois and Buckley, 2019). Moreover, stable isotopic probing was also utilized for determining the degradation rates of soil eDNA (Morrissey et al., 2015). However, most quantitative approaches have remained restricted to single or highly specific DNA targets (e.g., transgenic sequences), thereby overlooking sequence-specific variation in soil eDNA degradation rates (Pietramellara et al., 2009; Sirois and Buckley, 2019; Wang et al., 2019).

In this study, ‘sequence-specific degradation’ refers to statistically significant differences in first-order degradation rate constants (k, day⁻¹) among distinct 16S rRNA gene amplicon sequence variants (ASVs) under identical soil and incubation conditions. The potential variations in sequence-specific eDNA degradation rates can be attributed to several factors. First, sequence-dependent degradation can arise from differences in nucleotide composition, particularly GC content. This influences the thermodynamic stability and base-stacking interactions of the DNA duplex, thereby altering its accessibility to extracellular nucleases (Marrone and Ballantyne, 2008; Wolpe and Guertin, 2022). Second, local conformational features and the formation of potential secondary structures, such as stem-loops or hairpins, can create steric hindrance that protects the phosphodiester backbone. Differences in base composition also alter the elemental stoichiometry (e.g., C:N ratio) of DNA molecules, potentially affecting microbial preference for recycling specific sequences as nutrient sources (Cai et al., 2006b; Buitrago et al., 2021). Third, the persistence of soil DNA can be mainly attributed to its adsorption and protection by minerals and humus in soils (Cai et al., 2006a; Vuillemin et al., 2017; McKinney and Dungan, 2020). Thus, sequence-dependent differences in the physicochemical behavior of DNA molecules, including their affinity for soil minerals and organic matter, may also contribute to variation in degradation rates among sequences (Levy-Booth et al., 2007; Morrissey et al., 2015). Consequently, we proposed three central hypotheses. (1) The degradation rates of eDNA amplicon fragments were expected to be highly sequence-specific. (2) The rates and patterns of eDNA fragments degradation would be influenced by environmental factors such as temperature and moisture content. (3) The sequence-specific degradation of extracellular 16S rRNA gene amplicon fragments would significantly influence estimates of soil prokaryotic abundance and diversity.

To test these hypotheses, we investigated the overall and sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments and the factors influencing these rates. Additionally, the effects of extracellular 16S rRNA genes on soil prokaryotic community analysis and their relationships with the overall and sequence-specific degradation rates were determined. Soil samples were collected from 30 representative ecosystems across China. A new methodology, involving the addition of exogenous 16S rRNA gene amplicon fragments tagged with specific primers, microcosm incubation, real-time PCR, and amplicon sequencing, was developed to determine the overall and sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments. We employed 16S rRNA gene amplicon fragments as the standardized and trackable model substrate because the 16S rRNA gene is one of the most extensively used molecular markers in microbiome research (Knight et al., 2018; Du et al., 2025).

Results

The overall degradation rate constants of soil extracellular 16S rRNA gene amplicon fragments

The glyceraldehyde-3-phosphate dehydrogenase (GAPDH F)-tagged 16S rRNA gene amplicon fragments were consistently detectable throughout the 48-day incubation period, but their abundance rapidly declined as the incubation progressed (Figure 1a, p < 0.05). After 48 days of incubation, 0.2–3.1% of the initially spiked GAPDH F-tagged 16S rRNA gene amplicon fragments persisted in the soils (Figure 1a). The degradation rate constants of the spiked extracellular 16S rRNA gene amplicon fragments displayed considerable variability among the study sites, ranging from 0.05 to 0.16 day^–1 (Figure 1b). Furthermore, we found that degradation rate constants differed significantly among ecosystem types (Figure 1c, p < 0.05). Specifically, cropland and forest soils exhibited significantly higher degradation rates than grassland soils (p < 0.05). Random forest modeling revealed that soil moisture content, prokaryotic abundance, and prokaryotic community profiles were critical predictors for the overall degradation rate constants of extracellular 16S rRNA gene amplicon fragments, and they all showed significant positive correlations (Figure 1d, Figure 1—figure supplement 1a–d). However, we did not observe significant correlations between soil texture and the degradation rate constants (Figure 1—figure supplement 1e, f). Structural equation modeling (SEM) analysis indicated that the overall degradation rate constants of 16S rRNA gene amplicon fragments were mainly directly affected by prokaryotic abundance, which was indirectly influenced by soil moisture (Figure 1e). In the moisture gradient microcosm experiment, we also observed a strong positive correlation between soil moisture content and the overall degradation rates of extracellular 16S rRNA gene amplicon fragments (Figure 1f).

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The overall degradation rates of soil extracellular 16S rRNA gene amplicon fragments and their influential factors.

(a) The GAPDH F-tagged 16S rRNA gene amplicon fragment abundance at different incubation time points. (b, c) The degradation rate constants of soil extracellular 16S rRNA gene amplicon fragments across the study sites and different ecosystem types. (d) The factors influencing extracellular 16S rRNA gene degradation rates. (e) The influencing factors for the overall degradation rates of soil extracellular 16S rRNA gene amplicon fragments revealed by structural equation modeling. Orange and green lines indicate positive and negative relationships, respectively. Solid and dashed lines indicate significant and non-significant relationships, respectively. Path coefficients are denoted by numbers adjacent to the arrows, with arrow width reflecting their strength. (f) The influence of soil moisture on the degradation rates of soil microbial eDNA. Significance levels are indicated as follows: *p < 0.05, **p < 0.01, and ***p < 0.001. Moisture: soil moisture content; Abundance: prokaryotic abundance; NMDS1: the scores at the first axis of the NMDS ordination of prokaryotic community profiles; MAP: mean annual precipitation; Richness: soil prokaryotic richness; NMDS2: the scores at the second axis of the NMDS ordination of prokaryotic community profile; TK: soil total potassium contents; MAT: mean annual temperature; AP: soil available phosphorus content; TN: soil total nitrogen contents; and TOC: soil total organic carbon content.

The sequence-specific degradation rate constants of the extracellular 16S rRNA gene amplicon fragments

The richness of the GAPDH F-tagged 16S rRNA gene amplicon fragments significantly decreased during the incubation period, declining to approximately 60% of the initial values after 48 days (Figure 2a). The prokaryotic community profiles based on the GAPDH F-tagged 16S rRNA gene amplicon fragments also exhibited significant variations across different incubation time points (Figure 2b, Figure 2—figure supplement 1), and community-profile similarities declined more strongly with increasing incubation intervals (Figure 2—figure supplement 2). These findings suggest sequence-specific differences in degradation patterns among ASVs. Indeed, the degradation rate constants for different extracellular 16S rRNA gene amplicon fragments were within 0.40 day^–1, with most ranging between 0.06 and 0.12 day^–1 (Figure 2c). For example, many sequences assigned to Proteobacteria had significantly higher degradation rate constants than those assigned to Acidobacteriota, whereas Methylomirabilota showed intermediate degradation rate constants (Figure 3a). The sequence-specific degradation rate constant profiles of the extracellular 16S rRNA gene amplicon fragments also showed significant correlations with multiple environmental factors, including soil moisture, mean annual temperature (MAT), mean annual precipitation (MAP), soil pH, as well as the contents of soil ammonium nitrogen (NH4⁺-N) and nitrate nitrogen (NO₃⁻-N) (Figure 3b). However, no significant relationships were found between the degradation rates and the GC content of the representative sequences (Figure 3—figure supplement 1). Further analysis suggested that the degradation rate constants of extracellular 16S rRNA gene amplicon fragments of the dominant phyla showed similar correlations with the environmental factors, and MAT was identified as the strongest predictor for them (Figure 3—figure supplement 2).

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The sequence-specific degradation rates of soil exogenous extracellular 16S rRNA gene amplicon fragments.

(a) Soil GAPDH F-tagged 16S rRNA gene richness. (b) The nonmetric multidimensional scaling (NMDS) ordination of the community profiles based on GAPDH F-tagged 16S rRNA gene amplicon fragments at different incubation time points. (c) The number of GAPDH F-tagged 16S rRNA gene ASVs within different degradation rate ranges.

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The difference in sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments and their influencing factors.

(a) The paired comparison of degradation rates among different ASVs. In the heatmap, each cell represents a pairwise comparison between two ASVs. Blue indicates that the degradation rate of the ASVs listed in the row (row ASVs) is significantly lower than that of the ASVs listed in the column (column ASVs); red indicates that the row ASVs have a significantly higher degradation rate than the column ASV. A positive t value indicates that the row ASVs degrade significantly faster than the column ASVs; a negative t value indicates the opposite. Significance levels are indicated as follows: *p < 0.05, **p < 0.01, and ***p < 0.001. (b) The relationships between the sequence-specific degradation rate profiles and environmental factors. NO₃⁻-N: soil NO₃⁻-N contents; NH₄⁺-N: soil NH₄⁺-N contents; TN: soil total N contents; TP: soil total P contents; TK: soil total K contents; TOC: soil total organic carbon content; MAT: mean annual temperature; and MAP: mean annual precipitation.

The influences of extracellular 16S rRNA genes on soil prokaryotic community analysis and their links with the degradation rates

Based on the propidium monoazide (PMA) treatment, we observed significant effects of extracellular 16S rRNA genes on the analysis of soil prokaryotic abundance and diversity (Figures 4 and 5). The PMA treatment revealed that intact cells accounted for approximately 40% (range: 9–73%) of the total 16S rRNA gene copies. In contrast, over 80% (range: 27–97%) of the observed ASV richness was associated with sequences originating from intact cells (Figure 4a, b). Meanwhile, the Shannon index of the PMA-treated prokaryotic communities was significantly lower than that of the total prokaryotic community (Figure 4c). Furthermore, significant differences were found between the profiles of total and PMA-treated prokaryotic communities, especially in the forest and cropland ecosystems (Figure 4d–f, Figure 4—figure supplement 1). The Bray-Curtis dissimilarity between the intact cells and the total prokaryotic community was approximately 52.8% (Figure 4f). For instance, Abditibacteriota, Bacteroidota, Nitrospirota, Fibrobacterota, Entotheonellaeota, Elusimicrobiota, and Armatimonadota were significantly enriched in the total prokaryotic community, whereas Actinobacteriota and Planctomycetota showed opposite trends (Figure 5a). However, many other microbial taxa, such as Proteobacteria, Acidobacteriota, and Chloroflexi, showed no significant differences between the total and PMA-treated prokaryotic communities (Figure 5a). Additionally, the correlations with environmental factors were stronger for the total soil prokaryotic community structure than the PMA-treated community structure (Figure 5b). Total prokaryotic community structure was significantly correlated with soil pH, total potassium (TK), NH₄⁺-N, MAT, and MAP. However, the PMA-treated prokaryotic community structure was only significantly correlated with soil pH, MAT, and MAP (Figure 5b).

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The differences in the abundance, richness, Shannon, and community composition between total and PMA-treated soil prokaryotes.

(a) The abundance of total and PMA-treated soil prokaryotes. (b, c) The richness and Shannon index of total and PMA-treated soil prokaryotes. (d) The differences between the relative abundance of total and intracellular ASVs. The red points represent the prokaryotic taxa exhibiting statistically significant differences. (e) The nonmetric multidimensional scaling (NMDS) ordination of total and PMA-treated soil prokaryotes. Different colors represent samples from different sites. (f) The Bray–Curtis dissimilarity between total and PMA-treated soil prokaryotes.

Figure 5

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Effects of eDNA exclusion on soil prokaryotic community composition and its relationships with environmental factors.

(a) The taxa with significant differences between total and intact cell prokaryotic communities. P1: Thermoplasmatota; P2: Nitrospirota; P3: Fibrobacterota; P4: Entotheonellaeota; P5: Elusimicrobiota; P6: Armatimonadota; P7: Myxococcota; C1: Thermoplasmata; C2: Nitrospiria; C3: Vampirivibrionia; C4: Clostridia; C5: Bacilli; C6: Armatimonadia; C7: Chthonomonadetes; C8: Polyangia; C9: Blastocatellia; C10: Parcubacteria; C11: Bacteroidia; C12: Thermoleophilia; O1: Nitrospirales; O2: Abditibacteriales; O3: Clostridiales; O4: Bacillales; O5: Armatimonadales; O6: Pyrinomonadales; O7: Bryobacterales; O8: Chitinophagales; O9: Rubrobacterales; O10: Propionibacteriales; and O11: Corynebacteriales. (b) The relationships between soil prokaryotic community profiles and environmental factors. NO₃⁻-N: soil NO₃⁻-N contents; NH₄⁺-N: soil NH₄⁺-N contents; Moisture: soil moisture content; MAT: mean annual temperature; MAP: mean annual precipitation; TP: soil total P contents; TK: soil total K contents; and TOC: soil total organic carbon contents.

Interestingly, a significant negative correlation was observed between the total prokaryotic community structure and the degradation rates of extracellular 16S rRNA gene amplicon fragments. However, no significant relationship was observed for the PMA-treated prokaryotic communities (Figure 5b). The relationships between extracellular 16S rRNA gene pool sizes and degradation rates were further explored. We found a significant positive correlation between the overall degradation rates and the copies of soil extracellular 16S rRNA genes (Figure 1—figure supplement 1c). Moreover, most study sites exhibited significant positive correlations between sequence-specific degradation rates and the pool sizes of soil extracellular 16S rRNA genes (Figure 1—figure supplement 1g). Additionally, there were significant correlations between the differences in relative abundance of taxa in the total and PMA-treated prokaryotic community and the sequence-specific degradation rates; but these relationships varied across the study sites (Figure 1—figure supplement 1g).

Discussion

In this study, we found that extracellular 16S rRNA gene amplicon fragments persisted in soils for at least several weeks, and the degradation rate constants ranged from 0.05 to 0.16 day^–1. The degradation rate constants of soil eDNA based on plasmid and stable isotope labeling microbial genomes were usually 0.03–0.14 day^–1 (Morrissey et al., 2015; Sirois and Buckley, 2019; Wang et al., 2019), which is consistent with our results. Therefore, the PCR-generated DNA fragments provide a standardized substrate for quantifying the degradation kinetics of added DNA under controlled conditions, offering a useful proxy for eDNA turnover. Additionally, substantial variations in the overall degradation rates of added exogenous eDNA amplicon fragments were also observed in this study, which can be mainly elucidated in the following ways (Figure 1a and b). First, the degradation of soil eDNA is strongly influenced by the availability of enzymes (Nihemaiti et al., 2020). As a large proportion of soil enzymes originate from microbes (Bhardwaj et al., 2024; Tan et al., 2025), differences in microbial abundance among study sites could make a substantial contribution to the variations in the overall degradation rates. Second, different soil microbial taxa play diverse roles in eDNA degradation. In this study, there were considerable discrepancies in soil community profiles across study sites, providing another plausible explanation for the differing degradation rates of eDNA. Third, environmental factors, including soil moisture, pH, and temperature, can predominantly govern enzymatic reaction rates (He et al., 2024; Shah et al., 2024). Indeed, strong positive correlations were observed between moisture content and eDNA degradation rates in both the survey and microcosm experiments (Figure 1d–f). These results support our second hypothesis that environmental factors regulate the degradation rates and persistence patterns of eDNA fragments, with soil moisture emerging as a particularly important driver. This finding further suggests that dryland soils may be particularly susceptible to eDNA persistence, potentially increasing the risk of overestimating microbial abundance and diversity in arid ecosystems (Carini et al., 2016; Lennon et al., 2018).

Consistent with our first hypothesis, we observed that the degradation of soil eDNA amplicon fragments showed strong sequence-specific patterns (Figure 2c). In particular, many sequences belonging to Proteobacteria exhibited a significantly faster DNA degradation rate than those of Acidobacteriota (Figure 3a). The sequence-specific degradation of the eDNA amplicon fragments can be interpreted from multiple perspectives. First, the variation in the number and position of restriction enzyme cutting sites for different sequences could be a critical reason for the sequence-specific degradation rates of eDNA (Brown, 2020). Second, sequence differences could also influence the spatial structure and mineral adsorption of DNA fragments (Cleaves et al., 2011; Buitrago et al., 2021), indirectly affecting the sequence-specific degradation rates. Third, microbes might preferentially degrade and recycle the abundant DNA sequences present in their habitats. Thus, the sequence-specific degradation rates could also be ascribed to the varied abundance of different eDNA amplicon fragment sequences (Finkel and Kolter, 2001). This assertion is further evidenced by the generally observed positive correlations between eDNA abundance and degradation rates (Figure 1—figure supplement 1). We also examined whether GC content could explain the observed sequence-specific patterns, but no significant correlation was found (Figure 3—figure supplement 1), suggesting that simple base composition is not the primary driver in this study. However, this does not exclude the possibility that higher-order structural features (e.g., hairpin loops) or sequence-specific nuclease recognition motifs contribute to differential degradation (Wang et al., 2007). This should be tested in future studies using synthetic DNA constructs with controlled structural elements. As observed in this study, the differential sequence-specific degradation rates also led to significant alterations in the community profiles based on the exogenous extracellular 16S rRNA genes (Figure 2a and b). Additionally, this study essentially determined the effects of DNA sequences on the degradation rates of extracellular 16S rRNA gene amplicon fragments. However, actual sequence-specific degradation rates of extracellular 16S rRNA genes can be additionally influenced by the varying degrees of protection offered by cell residuals from different microbial taxa (Shi et al., 2024). Thus, the differences in the actual sequence-specific degradation rates of extracellular 16S rRNA genes should be even more significant than those observed in this study. This can be one of the main reasons for the influences of eDNA on the analysis of soil microbial community profiles (Lennon et al., 2018).

Accordingly, we further explored how eDNA may influence prokaryotic community analyses using PMA treatment, and significant disparities were observed between the profiles of the total and PMA-treated soil prokaryotic communities (Figure 4). The differences between total and PMA-treated community profiles can arise from multiple factors. As mentioned above, the disparities can be ascribed to the differential sequence-specific degradation rates of eDNA. Additionally, these differences are also related to several other factors. The primary reason should be the distinct source of extracellular and intracellular DNA, which are derived from dead and live microbes, respectively. The death of microorganisms is mainly caused by environmental selection or stochastic processes (Zhang et al., 2016). In terms of environmental selection, the dead microbial taxa should be less competitive than PMA-treated ones (Upton et al., 2019; Chu et al., 2020). The stochastic death should be correlated with the population size of each microbial species, but the varied stochastic mortality among different microbial taxa can still lead to different community profiles between dead and live microbes (Blazewicz et al., 2020). This study also revealed that extracellular 16S rRNA genes led to substantial overestimation of soil prokaryotic abundance and diversity (Figures 4 and 5). These findings were supported by many recent studies (Carini et al., 2016; Sun and Ge, 2023; Du et al., 2025) and can be explained as follows. The overestimated prokaryotic abundance can be attributed to the long-term persistence of eDNA (Sun and Ge, 2023). However, as DNA extraction efficiency may differ between intact cells and eDNA, the actual differences between total and living prokaryotic abundance could be smaller than those observed in this study. Similarly, the overestimated prokaryotic richness may arise from historically accumulated microbial taxonomic information stored in eDNA pools (Deshpande and Fahrenfeld, 2023; Wang et al., 2024a).

We observed a significant correlation between eDNA degradation rates and the overall structure of the prokaryotic community, but this relationship was absent in PMA-treated communities (Figure 5b). This discrepancy highlights the divergent ecological roles of extracellular and intracellular DNA. Analyses of the total community integrate intracellular DNA from metabolically active cells with eDNA, which primarily originates from historical microbial residues (Lennon et al., 2018). eDNA incorporates signals that likely reflect the legacy effects of past environmental conditions (Wang et al., 2021). In contrast, the PMA-treated community reflects transient microbial activity driven by current selective pressures. Additionally, eDNA can serve as a nutrient source and facilitate horizontal gene transfer, which may further shape its interactions with contemporary microbial communities (Levy-Booth et al., 2007).

Notably, we also observed significant positive correlations between sequence-specific degradation rates and the pool sizes of extracellular 16S rRNA gene amplicon fragments at most of the study sites (Figure 1—figure supplement 1g). This finding suggests that abundant eDNA degrades at a faster rate compared to rare eDNA. As mentioned earlier, this could be explained by several mechanisms. First, as soil eDNA is subject to enzymatic degradation and microbial recycling, abundant DNA sequences may be more likely to be encountered and degraded by extracellular nucleases simply due to their higher copy numbers (Levy-Booth et al., 2007; Nagler et al., 2018). Similarly, if microbes preferentially take up DNA as a nutrient source, they may degrade abundant sequences more frequently as a stochastic consequence of higher encounter rates (Finkel and Kolter, 2001). However, we also found that the relationships between the sequence-specific degradation rates and the effect sizes of extracellular 16S rRNA gene amplicon fragments varied across the study sites (Figure 1—figure supplement 1g). The sequence-specific effect sizes of extracellular 16S rRNA gene amplicon fragments are mainly determined by both their production and degradation rates (Pietramellara et al., 2009; Sirois and Buckley, 2019). These inconsistent correlations emphasize the critical role played by the production rates of extracellular 16S rRNA genes in influencing the analysis of prokaryotic communities. Therefore, future studies should systematically determine both the production and degradation rates of eDNA.

Despite the high-resolution insights afforded by our methodology, several limitations should be considered. First, utilizing PCR-amplified 16S rRNA gene fragments as proxies oversimplifies the structural and sequence complexity of natural soil eDNA pools. In natural environments, eDNA varies widely in fragment length and conformation and exhibits complex interactions with mineral surfaces, all of which fundamentally affect degradation dynamics (Levy-Booth et al., 2007; McKinney and Dungan, 2020). Additionally, the highly conserved nature of the 16S rRNA gene means that the nucleotide variability explored here (e.g., GC content gradients) does not fully capture the genomic heterogeneity of entire metagenomes (Knight et al., 2018). Consequently, our reported degradation rates indicate the decay potential of highly accessible linear eDNA rather than a universal rate for all soil DNA fractions. Future studies incorporating diverse metagenomic DNA, especially DNA with extremely high AT or GC content, are essential for building a more generalizable predictive framework for eDNA persistence (Morrissey et al., 2015). Second, methodological biases inherent in quantifying the intracellular community must be acknowledged (Du et al., 2025). Although PMA treatment is widely used to exclude eDNA, its efficiency in complex soil matrices can be compromised by limited light penetration in turbid suspensions and competitive adsorption to soil particles (Nocker et al., 2007; Carini et al., 2016; Heise et al., 2016). Compounding this issue, downstream DNA recovery is subject to differential cell lysis, as taxa with robust cell walls (e.g., Gram-positive bacteria) may resist extraction (Frostegård et al., 1999; Albertsen et al., 2015). While our standardized bead-beating protocol and calculation of degradation rate constants (k) minimize systematic biases, future studies should integrate complementary viability markers (e.g., RNA-based analyses or protein synthesis activity probes) and multi-extraction comparisons to robustly validate these ecological patterns (Emerson et al., 2017).

Materials and methods

Study sites and soil sampling

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Soil samples were collected from 30 sites across China (Figure 1—figure supplement 2). The selection of sampling sites was mainly based on the National Soil Fertility and Fertilizer Effect Long-term Monitoring Network and the Chinese Ecosystem Research Network (CERN). Several other sampling sites were included based on the systematic consideration of ecosystem typicality, soil types, and climates. The 30 sites spanned major climatic zones and land-use types in China, including 10 grasslands, 10 forests, 6 croplands, and 4 deserts, ensuring broad applicability of the results across diverse environmental scenarios. The longitude of the sites ranged from 80.724 °E to 124.817 °E, while the latitude ranged from 21.917 °N to 50.168 °N. The altitude of the study sites varied from 13 m to 4793 m. The ranges of MAT and MAP were –3.35 to 22.53°C and 39 to 1809 mm, respectively. The geographic and climate information of study sites is detailed in Supplementary file 1A. No specific permits or ethical approvals were required for soil sampling at the study sites.

At each sampling site, 10 subsampling points were randomly selected, with a minimum distance of 10 m between adjacent subsampling points. Soil samples (0–20 cm) from 10 subsampling points at each study site were collected, thoroughly homogenized, and sieved to ≤2 mm to form a composite sample. Subsequently, all the composite soil samples were divided into two sub-samples. The first sub-sample was air-dried for the determination of soil pH values, as well as measurements of total organic carbon (TOC), total nitrogen (TN), total phosphorus (TP), and available phosphorus (AP). The second sub-sample was preserved at 4°C for the determination of moisture content, inorganic nitrogen content, as well as for microcosm experiment and PMA treatment.

Analysis of soil physicochemical properties

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Soil moisture content was determined by drying the soils at 105°C for 48 hr (Che et al., 2019). Soil pH values were measured using a pH meter (Mettler Toledo, Switzerland) with a soil-to-water ratio of 1:2.5 (Zhang et al., 2023a). TOC content of the soils was examined via a TOC analyzer (GB/T 30740-2014). The TN content of the soils was determined using an automatic Kjeldahl apparatus (Liu et al., 2023). Additionally, the determination of soil TP concentration was performed using the Mo-Sb colorimetric method. The soil NH₄⁺-N and NO₃⁻-N contents were measured by indophenol blue colorimetry and vanadium chloride spectrophotometry with a potassium chloride (KCl) extraction method, respectively (Zhang et al., 2019; Zhang et al., 2023b). The soil AP content was measured following the methods of Olsen, 1954. Detailed physical and chemical properties of the soils are listed in Supplementary file 1B.

Determination of 16S rRNA gene amplicon fragments overall and sequence-specific degradation rate constants

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The overall and sequence-specific degradation rate constants of added exogenous extracellular 16S rRNA gene amplicon fragments were determined using a primer labeling method, combined with real-time PCR and amplicon sequencing (Figure 6). Briefly, exogenous eDNA was prepared by PCR amplification using a modified forward primer consisting of a GAPDH F tag fused to the 16S rRNA gene primer 515F, together with the reverse primer 806R. The full primer sequences were as follows: GAPDH-F-515F: 5′-CAT TGG CAA TGA GCG GTT C-GTG CCA GCM GCC GCG GTA A-3′, in which CAT TGG CAA TGA GCG GTT C represents the GAPDH F tag and GTG CCA GCM GCC GCG GTA A represents the 16S rRNA gene forward primer sequence (515F); and 806R: 5′-GGA CTA CHV GGG TWT CTA AT-3′ (Caporaso et al., 2011; Walters et al., 2016). GAPDH is a primer for a human housekeeping gene and it has no homologous sequences in soils. Subsequently, GAPDH was selected as the label primer based on two criteria. First, this primer was selected to avoid interference from the original soil sequences (Huang et al., 2014; Yang et al., 2021; Arvizu-Hernandez et al., 2025), and no detectable PCR amplification was observed for the primer set GAPDH F-806R across all the soil DNA samples included in this study. Second, the melting temperature (Tm) value of GAPDH F approximately matched that of 806R. The GAPDH was incorporated only into the forward primer for several reasons. Methodologically, adding a long linker to the degenerate reverse primer (806R) could reduce amplification efficiency or introduce bias. Economically, single-end labeling allowed us to use the standard reverse primer already carrying sample-specific barcodes, avoiding the costly synthesis of a full set of dual-labeled barcoded primers. This design minimized the risk of secondary structure and primer–dimer artifacts while maintaining sufficient specificity and compatibility with downstream qPCR and sequencing. Subsequently, the exogenous eDNA was separately added to the corresponding fresh soils collected from the 30 study sites, and incubated for 0, 3, 6, 12, 24, and 48 days. Finally, the overall and sequence-specific degradation rate constants of 16S rRNA gene amplicon fragments were calculated based on amplicon sequencing and real-time PCR. Detailed procedures are described as follows.

Figure 6

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Schematic illustration of the experimental workflow for determining the overall and sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments.

Preparation of exogenous eDNA

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Soil DNA was extracted from 0.5 g of soil using the DNeasy PowerSoil kit (QIAGEN, Hilden, Germany) following the manufacturer’s protocols (Li et al., 2023). The GAPDH F-tagged PCR products were generated using DNA extracted from the original soil sample as templates and amplified with GAPDH F-tagged 515F and 806R. The PCR mixture (50 μl) consisted of ExTaq buffer (10×, TaKaRa; 5 μl), dNTP Mix (2.5 mM; 4 μl), forward primer (10 μM; 1 μl), reverse primer (10 μM; 1 μl), template DNA (1 μl), ExTaq (0.25 μl), and DNase-free water (38 μl). The PCR protocol involved an initial denaturation at 95°C for 10 min, followed by 32 PCR cycles consisting of 30 s at 95°C, 30 s at 56°C, and 40 s at 72°C. Additionally, a final extension was performed at 72°C for 4 min. Three technical replicates were conducted to amplify each DNA sample, followed by PCR product purification using the GeneJET Gel Extraction Kit (Thermo Scientific, Lithuania). Finally, the purified GAPDH F-tagged 16S rRNA gene fragments were utilized as exogenous extracellular 16S rRNA gene amplicon fragments for subsequent experiments.

Microcosm experiment

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The microcosm experiment was conducted using 30 g of soil for each sample. After pre-incubation at 20°C for 1 week, each soil was thoroughly mixed with the GAPDH F-tagged 16S rRNA gene amplicon fragments and incubated further at 20°C (Figure 1—figure supplement 3). The amount of exogenous GAPDH F-tagged 16S rRNA gene amplicon fragments added to each soil sample was equivalent to 1% of the total DNA concentration naturally present in that soil, as determined fluorometrically prior to the experiment. This concentration was chosen to approximate natural eDNA fluxes resulting from microbial lysis, ensuring experimental relevance to in situ conditions (Supplementary file 1B). Weekly water additions were performed to maintain the original moisture content of the soils. Soil samples (5 g) were collected after incubation periods of 0, 3, 6, 12, 24, and 48 days with complete mixing prior to each collection. A total of 180 soil samples (30 study sites × 6 incubation times) were included in this study.

We further explored the influence of soil moisture content on the degradation rates of microbial extracellular 16S rRNA gene amplicon fragments based on soils collected from Kaiyuan (KY) and Dashanbao (DSB) in southwest China. The moisture content gradient included 100%, 75%, 50%, 25%, and 10% of soil water holding capacity. GAPDH F-tagged 16S rRNA gene amplicon fragments originating from the corresponding soils were added to the soils based on the natural soil DNA concentration. The soils were collected after 0, 1, 3, 6, 12, and 24 days of incubation. This complementary experiment included two sites, five moisture levels, six incubation time points, and two replicates per treatment combination, resulting in a total of 120 soil samples.

Soil DNA extraction, real-time PCR, and amplicon sequencing

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DNA was extracted from the microcosm experiment soils using the DNeasy PowerSoil kit (QIAGEN, Hilden, Germany). The copy numbers of the GAPDH F-tagged 16S rRNA gene amplicon fragments persisting in the soils were determined by employing a LightCycler real-time PCR System (Roche, Mannheim, Germany) with primers GAPDH F and 806R. For each real-time PCR reaction mixture, 1.0 μl of template DNA, 10.0 μl of TB Green Premix Ex Taq II (Takara, Japan), 0.5 μl of forward primer (20 μM), and 0.5 μl of reverse primer (20 μM) were mixed with 8.0 μl of DNase-free water. The real-time PCR protocol commenced with an initial denaturation at 95°C for 40 s, followed by 40 cycles (5 s at 95°C, 30 s at 56°C, and 40 s at 72°C). The standard curves exhibited fitted curve R² values higher than 0.99, with amplification efficiencies of 85%. Each DNA sample quantification was performed in triplicate.

The community profiles of the GAPDH F-tagged 16S rRNA gene amplicon fragments were determined using high-throughput amplicon sequencing. Briefly, GAPDH F-tagged 16S rRNA gene amplicon fragments from the microcosm soils were first amplified from individual samples using GAPDH F and barcode-labeled 806R primers. The reverse primer 806R carried a 12-bp sample-specific barcode, whereas the GAPDH F primer did not contain a barcode. Therefore, each sample was assigned a unique barcode during PCR, which allowed sample demultiplexing after sequencing. The PCR reaction system and thermal cycling conditions were similar to those described above, except that the number of amplification cycles was increased to 35 to obtain sufficient amplicon products for sequencing. The barcoded PCR products from individual samples were purified using a GeneJET Gel Extraction Kit (Thermo Scientific, Lithuania), quantified, and then pooled in equimolar amounts for subsequent library construction. Sequencing libraries were prepared from the pooled barcoded amplicons using the ALFA-SEQ DNA Library Prep Kit according to the manufacturer’s protocol. Universal Illumina-compatible adapters were first ligated to the pooled amplicon products, followed by bead-based purification. An indexing PCR was then performed using the index primer mix, which introduced the complete P5/P7 flow-cell binding sequences and a library-level Illumina index into the pooled library molecules. The indexed library was purified, quantified, and subjected to paired-end sequencing on the NovaSeq platform at Magigene Biotechnology Co, Ltd (Guangdong, China).

The USEARCH (v11) with denoising algorithm was used to analyze the raw sequences (Edgar, 2013). Briefly, paired-end reads were merged using USEARCH, and primer sequences (GAPDH-F-515F and 806R) were removed using the search_pcr2 script. Reads with more than two primer mismatches were discarded. Quality filtering was performed using the fastq_filter script, and sequences with quality scores below 20 were removed. Redundant sequences were dereplicated using the fastx_uniques script. ASVs were generated using the UNOISE3 non-clustering denoising algorithm, which infers 100% exact sequence variants by distinguishing biological sequences from PCR/sequencing errors. ASVs with total sequence counts fewer than 9 across all samples were removed to reduce noise. To quantify the abundance of each ASV, an ASV table was generated by mapping the quality-filtered raw reads back to the ASV set using the otutab command. A 97% similarity threshold was applied for this recruitment to accommodate stochastic sequencing noise while maintaining biological resolution. Crucially, the mapping followed a best-hit priority rule, where each read was assigned to the ASV with the highest percent identity within the 97% radius. This approach ensures that reads derived from the same biological template are accurately counted toward their respective ASV, preventing the underestimation of abundances that would occur with exact matching while strictly preserving the single-nucleotide resolution of the ASV framework. Taxonomic annotation of the ASVs was performed in QIIME2 with the Silva v138 database. A total of 89322 prokaryotic ASVs were obtained. To standardize sequencing depth across samples, the read number of each sample was rarefied to 53251 using the rarefy function in the vegan package in R. The soil prokaryotic diversity (i.e., richness) was assessed via the vegan package in R (Oksanen et al., 2021). All the raw sequencing data and analysis codes have been deposited in the NCBI Sequence Read Archive under BioProject PRJNA1141901 and GitHub repository (https://github.com/lt916/16S-rRNA-genes-rates, copy archived at Li, 2026), respectively. The BioSample accession numbers for the 16S rRNA gene amplicon sequencing data are SAMN42909669–SAMN42909908.

Determination of soil extracellular 16S rRNA gene amplicon fragments degradation rate constants

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The overall degradation rate constants of the extracellular 16S rRNA gene amplicon fragments were determined by examining the relationships between the copies of labeled 16S rRNA gene amplicon fragments and incubation time, and they were mainly indicated by degradation rate constants. The relationships were fitted using first-order enzyme-catalyzed reaction kinetics, as described by the following equations:

l n c = - k t + l n c_{0}

t: incubation time, day;

c: the copies of GAPDH F-tagged 16S rRNA gene amplicon fragments at time t;

c₀: the initial copies of GAPDH F-tagged 16S rRNA gene amplicon fragments;

k: the degradation rate constants of soil extracellular 16S rRNA gene amplicon fragments, day^–1.

The sequence-specific degradation rate constants of the exogenous extracellular 16S rRNA gene amplicon fragments were determined using an approach similar to that used for the overall degradation rates of the extracellular 16S rRNA gene amplicon fragments. The absolute abundance of each prokaryotic taxa was estimated by multiplying its relative abundance with the copies of total GAPDH F-tagged 16S rRNA gene amplicon fragments. Subsequently, the reaction rate constants (k) were calculated by fitting the relationships between the sequence-specific 16S rRNA gene amplicon fragment copies and incubation time. When comparing the rates of first-order reactions, the reaction rate constant (k) usually serves as the core indicator. Therefore, we used the degradation rate constant (k) to characterize the degradation dynamics of the added exogenous DNA, which serves as a proxy for how eDNA may be degraded in soils. Because these estimates combine relative abundance profiles with total qPCR abundance, the resulting sequence-specific degradation rates should be interpreted as apparent ASV-level degradation patterns within this analytical framework.

Impacts of eDNA on prokaryotic abundance and diversity analysis

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To further explore the implications of the overall and sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments, we also determined the impacts of extracellular 16S rRNA gene amplicon fragments on soil prokaryotic community analysis and their links with the overall and sequence-specific degradation rates. To inhibit amplification of eDNA, soils were incubated with PMA, as described previously (Carini et al., 2016). Upon photoactivation, eDNA can form covalent bonds through cross-linking, leading to the inhibition of its PCR amplification. In contrast, microbes with intact cell membranes exclude PMA, and their DNA is not cross-linked with PMA, and remains amenable to PCR amplification (Cangelosi and Meschke, 2014). Because cells with intact membranes are less permeable to PMA than eDNA or DNA from membrane-compromised cells, the PMA-treated fraction is expected to be enriched in DNA derived from intact cells. Currently, PMA treatment is widely used to suppress PCR amplification of eDNA (Xue et al., 2023; Canini et al., 2024).

In this study, 0.50 g of soil was mixed with PMA in a total volume of 0.5 ml (40 µM PMA in phosphate-buffered saline, PBS), while the control soil samples were mixed with PBS without PMA. Both the PMA-treated and control soil samples (PMA-untreated) were gently vortexed for 10 min in the dark at room temperature. Following the incubation, both sets of samples were exposed to a 650 W halogen lamp placed at a distance of 20 cm from the tube, undergoing four consecutive cycles of alternating light (30 s) and darkness (30 s). Subsequently, the tubes were centrifuged at 10,000 × g for 2 min, and the pellets were retained. Finally, DNA was extracted from these pellets using the DNeasy PowerSoil kit (QIAGEN, Hilden, Germany) following manufacturer’s protocols.

The abundance of 16S rRNA gene amplicon fragments was determined using quantitative PCR employing a LightCycler real-time PCR System (Roche, Germany). The universal primers set for the 16S rRNA gene amplification were 515F and 806R. The real-time PCR mixture and procedure followed the aforementioned protocol. The 20.0 μl real-time PCR reaction mixture included 1.0 μl of template DNA, 10.0 μl of TB Green Premix Ex Taq II (Takara, Japan), 0.5 μl of forward primer (20 μM), and 0.5 μl of reverse primer (20 μM) were mixed with 8.0 μl of DNase-free water. The real-time PCR protocol commenced with an initial denaturation at 95°C for 40 s, followed by 40 cycles (5 s at 95°C, 30 s at 56°C, and 40 s at 72°C). Additionally, prokaryotic 16S rRNA gene fragments were amplified via PCR with universal primers 515F-806R, following the same PCR system, procedure, and bioinformatics analysis as described above. Ultimately, discrepancies in soil microbial properties between the control and PMA-treated samples were utilized to indicate the impact of extracellular 16S rRNA gene amplicon fragments on soil prokaryotic community analysis.

Statistical analysis

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The changes in GAPDH F-tagged 16S rRNA gene copies and richness across different incubation time points were determined using repeated-measures analysis of variance. One-way analysis of variance (ANOVA) followed by Tukey’s honestly significant difference (HSD) post hoc test was used to compare degradation rate constants among ecosystem types. Prokaryotic community structure differences among the study sites and incubation time points were examined through non-metric multidimensional scaling analysis (NMDS), permutational multivariate analysis of variance (PERMANOVA), and permutational analysis of multivariate dispersion (PERMDISP) (Kruskal, 1964; Anderson, 2001). Random forest modeling was conducted to assess the importance of environmental variables in predicting the overall degradation rates of extracellular 16S rRNA gene amplicon fragments. SEM was employed to further evaluate the direct and indirect effects of soil moisture, soil pH, MAP, and prokaryotic abundance on the overall degradation rates of extracellular 16S rRNA gene amplicon fragments (Grace, 2006). The sequence-specific degradation rate constants of the exogenous extracellular 16S rRNA gene amplicon fragments were visualized using a heatmap with corrplot packages. To reduce noise arising from unstable detection and unreliable model fitting, only ASVs with relative abundances >0.01%, present in more than 90% of the study sites, and exhibiting a fitted degradation curve with R² > 0.5 were retained for pairwise sequence-specific degradation rate comparisons. As for the analysis, we performed paired t-tests across all the study sites. Thus, the degradation rates were essentially compared within each site, with both values originating from a same soil sample under identical incubation conditions. A positive t value indicates that the first ASV has a significantly higher degradation rate than the second one, and a negative t value indicates the opposite. The p values were adjusted for multiple comparisons using the FDR method. The relationships between the sequence-specific degradation rate profiles of soil extracellular 16S rRNA gene amplicon fragments and environmental factors were analyzed by Mantel tests (Mantel, 1967).

The impacts of extracellular 16S rRNA gene on the abundance and richness of soil prokaryotic community were determined using the paired t-test. Differences in the relative abundance of microbial taxa between the total and PMA-treated prokaryotic communities were assessed using Wilcoxon test, and the taxa with significant differences were visualized with Graphlan (v1.1.3). Mantel test was employed to further investigate the relationships of PMA-treated or total prokaryotic communities and environmental factors. Pearson correlation test was employed to examine the relationships between the pool sizes and degradation rates of sequence-specific soil extracellular 16S rRNA gene amplicon fragments. Only the abundant taxa, with the proportions exceeding 0.01%, were included in the analysis. For each prokaryotic taxon, its extracellular 16S rRNA gene pool size was calculated as the total 16S rRNA gene copies ×the relative abundance of the taxon × (1 – intracellular 16S rRNA gene copies of the taxon/total 16S rRNA gene copies of the taxon). Most of the aforementioned statistical analyses were conducted in R software with a range of packages including vegan, ggplot2, RandomForest, piecewiseSEM, corrplot, matlab, igraph, ggcor, ggpubr, Rmisc, and Hmisc (Breiman, 2001; Cutler et al., 2012; Lefcheck, 2016; Harrell and Dupont, 2020; Oksanen et al., 2021).

Data availability

All the raw sequencing data have been deposited in the NCBI Sequence Read Archive under BioProject PRJNA1141901. The BioSample accession numbers for the 16 S16S rRNA gene amplicon sequencing data are SAMN42909669–SAMN42909908.

The following data sets were generated

1. Li T
2. Zhang S
3. Wang Z
4. Zhang Z
5. Wang F
6. Liu D
7. Cui X
8. Che R
(2025) NCBI BioProject
ID PRJNA1141901. The taxon-specific degradation of soil extracellular 16S rRNA genes: rates, influential factors, and implications.

https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1141901

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Article and author information

Author details

Ting Li
1. Yunnan Key Laboratory of Soil Erosion Prevention and Green Development, Institute of International Rivers and Ecosecurity, Yunnan University, Kunming, China
2. State Key Laboratory for Vegetation Structure, Function and Construction (VegLab), Ministry of Education Key Laboratory for Ecosecurity of Southwest China, Yunnan University, Kunming, China
3. College of Life Sciences, University of Chinese Academy of Sciences, Beijing, China
Contribution
Conceptualization, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0009-0007-8064-1615
Song Zhang

State Key Laboratory of Pollution Control and Resource Reuse, School of the Environment, Nanjing University, Nanjing, China

Contribution
Data curation, Formal analysis, Visualization

Competing interests
No competing interests declared
Zelin Wang
1. Yunnan Key Laboratory of Soil Erosion Prevention and Green Development, Institute of International Rivers and Ecosecurity, Yunnan University, Kunming, China
2. State Key Laboratory for Vegetation Structure, Function and Construction (VegLab), Ministry of Education Key Laboratory for Ecosecurity of Southwest China, Yunnan University, Kunming, China
Contribution
Data curation, Visualization, Methodology

Competing interests
No competing interests declared
Wei Huang
1. Yunnan Key Laboratory of Soil Erosion Prevention and Green Development, Institute of International Rivers and Ecosecurity, Yunnan University, Kunming, China
2. State Key Laboratory for Vegetation Structure, Function and Construction (VegLab), Ministry of Education Key Laboratory for Ecosecurity of Southwest China, Yunnan University, Kunming, China
Contribution
Investigation, Methodology

Competing interests
No competing interests declared
Zejin Zhang

College of Life Sciences, University of Chinese Academy of Sciences, Beijing, China

Contribution
Data curation, Investigation, Methodology

Competing interests
No competing interests declared
Fang Wang

College of Life Sciences, University of Chinese Academy of Sciences, Beijing, China

Contribution
Investigation, Methodology, Writing – original draft

Competing interests
No competing interests declared
Dong Liu

School of Life Sciences, Yunnan University, Kunming, China

Contribution
Funding acquisition, Investigation, Methodology, Writing – original draft

Competing interests
No competing interests declared
Xiaoyong Cui

College of Life Sciences, University of Chinese Academy of Sciences, Beijing, China

Contribution
Conceptualization, Methodology, Writing – original draft

Competing interests
No competing interests declared
Rongxiao Che
1. Yunnan Key Laboratory of Soil Erosion Prevention and Green Development, Institute of International Rivers and Ecosecurity, Yunnan University, Kunming, China
2. State Key Laboratory for Vegetation Structure, Function and Construction (VegLab), Ministry of Education Key Laboratory for Ecosecurity of Southwest China, Yunnan University, Kunming, China
Contribution
Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Writing – original draft, Writing – review and editing

For correspondence
cherongxiao@ynu.edu.cn

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0001-7907-478X

Funding

National Natural Science Foundation of China (42007035)

Rongxiao Che

National Natural Science Foundation of China (42261012)

Rongxiao Che

National Natural Science Foundation of China (32560035)

Dong Liu

Yunnan Fundamental Research Projects (202301AW070004)

Rongxiao Che

Yunnan Fundamental Research Projects (202301BF070001-006)

Rongxiao Che

Yunnan Fundamental Research Projects (202301AT070211)

Dong Liu

Xingdian Youth Talent Support Program of Yunnan Province (YNQR-QNRC-2018-024)

Rongxiao Che

Xingdian Youth Talent Support Program of Yunnan Province (YNQR-QNRC-2020-087)

Dong Liu

Science and Technology Project of Yunnan University for Serving Local Development (YDFWDF202509)

Rongxiao Che

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by the National Natural Science Foundation of China (42007035, 42261012, and 32560035), Yunnan Fundamental Research Projects (202301AW070004, 202301BF070001-006, and 202301AT070211), Xingdian Youth Talent Support Program of Yunnan Province (YNQR-QNRC-2018-024 and YNQR-QNRC-2020-087), and Science and Technology Project of Yunnan University for Serving Local Development (YDFWDF202509).

Version history

Sent for peer review: January 12, 2026
Preprint posted: January 14, 2026
Reviewed Preprint version 1: April 15, 2026
Reviewed Preprint version 2: August 6, 2026
Version of Record published: September 4, 2026

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You can cite all versions using the DOI https://doi.org/10.7554/eLife.110251. This DOI represents all versions, and will always resolve to the latest one.

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Ting Li
Song Zhang
Zelin Wang
Wei Huang
Zejin Zhang
Fang Wang
Dong Liu
Xiaoyong Cui
Rongxiao Che

(2026)

Soil extracellular DNA fragments show variable degradation rates among sequences and environmental conditions

eLife 15:RP110251.

https://doi.org/10.7554/eLife.110251.3

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The overall degradation rates of soil extracellular 16S rRNA gene amplicon fragments and their influential factors.

The sequence-specific degradation rates of soil exogenous extracellular 16S rRNA gene amplicon fragments.

The difference in sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments and their influencing factors.

The differences in the abundance, richness, Shannon, and community composition between total and PMA-treated soil prokaryotes.

Effects of eDNA exclusion on soil prokaryotic community composition and its relationships with environmental factors.

Schematic illustration of the experimental workflow for determining the overall and sequence-specific degradation rates of soil extracellular 16S rRNA gene amplicon fragments.

Author details

Ting Li

Contribution

Competing interests

Song Zhang

Contribution

Competing interests

Zelin Wang

Contribution

Competing interests

Wei Huang

Contribution

Competing interests

Zejin Zhang

Contribution

Competing interests

Fang Wang

Contribution

Competing interests

Dong Liu

Contribution

Competing interests

Xiaoyong Cui

Contribution

Competing interests

Rongxiao Che

Contribution

For correspondence

Competing interests

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