Different approaches to processing environmental DNA samples in turbid waters have distinct effects for fish, bacterial and archaea communities

10.24072/pcjournal.256 - Peer Community Journal, Volume 3 (2023), article no. e33.

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Coastal lagoons are an important habitat for endemic and threatened species in California that have suffered impacts from urbanization and increased drought. Environmental DNA has been promoted as a way to aid in the monitoring of biological communities, but much remains to be understood on the biases introduced by different protocols meant to overcome challenges presented by unique systems under study. Turbid water is one methodologic challenge to eDNA recovery in these systems as it quickly clogs filters, preventing timely processing of samples. We investigated biases in community composition produced by two solutions to overcome slow filtration due to turbidity: freezing of water prior to filtration (for storage purposes and long-term processing), and use of sediment (as opposed to water samples). Bias assessments of community composition in downstream eDNA analysis was conducted for two sets of primers, 12S (fish) and 16S (bacteria and archaea). Our results show that freezing water prior to filtration had different effects on community composition for each primer, especially for the 16S, when using a filter of larger pore size (3 μm). Nevertheless, pre-freezing water samples can still be a viable alternative for storage and processing of turbid water samples when focusing on fish communities (12S). The use of sediment samples as an alternative to processing water samples should be done with caution, and at minimum the number of biological replicates and/or volume sampled should be increased.

Published online:
DOI: 10.24072/pcjournal.256
Keywords: metabarcoding; turbidity; method comparison; fish
Keywords: metabarcoding, turbidity, method comparison, fish
Turba, Rachel 1; Thai, Glory H. 1; Jacobs, David K. 1

1 Department of Ecology and Evolutionary Biology, University of California, Los Angeles, California 90095 USA
License: CC-BY 4.0
Copyrights: The authors retain unrestricted copyrights and publishing rights
     author = {Turba, Rachel and Thai, Glory H. and Jacobs, David K.},
     title = {Different approaches to processing environmental {DNA} samples in turbid waters have distinct effects for fish, bacterial and archaea communities
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%A Jacobs, David K.
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Turba, Rachel; Thai, Glory H.; Jacobs, David K. Different approaches to processing environmental DNA samples in turbid waters have distinct effects for fish, bacterial and archaea communities
. Peer Community Journal, Volume 3 (2023), article  no. e33. doi : 10.24072/pcjournal.256.

Peer reviewed and recommended by PCI : 10.24072/pci.ecology.100427

Conflict of interest of the recommender and peer reviewers:
The recommender in charge of the evaluation of the article and the reviewers declared that they have no conflict of interest (as defined in the code of conduct of PCI) with the authors or with the content of the article.

[1] Alberdi, A.; Aizpurua, O.; Gilbert, M. T. P.; Bohmann, K. Scrutinizing key steps for reliable metabarcoding of environmental samples, Methods in Ecology and Evolution, Volume 9 (2018) no. 1, pp. 134-147 | DOI

[2] Ballard, J.; Pezda, J.; Spencer, D.; Plantinga, A. An economic valuation of southern California coastal wetlands. , Southern California Wetlands Recovery Project, 2018 (

[3] Bischoff, V.; Zucker, F.; Moraru, C. Marine bacteriophages, Encyclopedia of Virology, Elsevier, 2021, pp. 322-341 | DOI

[4] Bohmann, K.; Chua, P.; Holman, L. E.; Lynggaard, C. DNAqua‐Net conference unites participants from around the world with the quest to standardize and implement DNA‐based aquatic biomonitoring, Environmental DNA, Volume 3 (2021) no. 5, pp. 884-888 | DOI

[5] Bowman, J. P. The family Cryomorphaceae, The Prokaryotes, Springer Berlin Heidelberg, Berlin, Heidelberg, 2014, pp. 539-550 | DOI

[6] Buxton, A. S.; Groombridge, J. J.; Griffiths, R. A. Is the detection of aquatic environmental DNA influenced by substrate type?, PLOS ONE, Volume 12 (2017) no. 8 | DOI

[7] Callahan, B. J.; McMurdie, P. J.; Rosen, M. J.; Han, A. W.; Johnson, A. J. A.; Holmes, S. P. DADA2: High-resolution sample inference from Illumina amplicon data, Nature Methods, Volume 13 (2016) no. 7, pp. 581-583 | DOI

[8] Caporaso, J. G.; Lauber, C. L.; Walters, W. A.; Berg-Lyons, D.; Huntley, J.; Fierer, N.; Owens, S. M.; Betley, J.; Fraser, L.; Bauer, M.; Gormley, N.; Gilbert, J. A.; Smith, G.; Knight, R. Ultra-high-throughput microbial community analysis on the Illumina HiSeq and MiSeq platforms, The ISME Journal, Volume 6 (2012) no. 8, pp. 1621-1624 | DOI

[9] Carøe, C.; Bohmann, K. Tagsteady: a metabarcoding library preparation protocol to avoid false assignment of sequences to samples, bioRxiv, 2020 | DOI

[10] Chen, H. VennDiagram: Generate high-resolution Venn and Euler plots (R Package), 2018 (

[11] Curd, E.; Gomer, J.; Kandlikar, G.; Gold, Z.; Ogden, M.; Shi, B. The Anacapa Toolkit, 2018 (

[12] Curd, E.; Kandlikar, G.; Gomer, J. CRUX: Creating reference libraries using eXisting tools, 2018 (

[13] Deiner, K.; Bik, H. M.; Mächler, E.; Seymour, M.; Lacoursière‐Roussel, A.; Altermatt, F.; Creer, S.; Bista, I.; Lodge, D. M.; Vere, N.; Pfrender, M. E.; Bernatchez, L. Environmental DNA metabarcoding: Transforming how we survey animal and plant communities, Molecular Ecology, Volume 26 (2017) no. 21, pp. 5872-5895 | DOI

[14] Deiner, K.; Walser, J.-C.; Mächler, E.; Altermatt, F. Choice of capture and extraction methods affect detection of freshwater biodiversity from environmental DNA, Biological Conservation, Volume 183 (2015), pp. 53-63 | DOI

[15] Dejean, T.; Valentini, A.; Miquel, C.; Taberlet, P.; Bellemain, E.; Miaud, C. Improved detection of an alien invasive species through environmental DNA barcoding: the example of the American bullfrog Lithobates catesbeianus, Journal of Applied Ecology, Volume 49 (2012) no. 4, pp. 953-959 | DOI

[16] Dell'Anno, A.; Corinaldesi, C. Degradation and turnover of extracellular DNA in marine sediments: ecological and methodological considerations, Applied and Environmental Microbiology, Volume 70 (2004) no. 7, pp. 4384-4386 | DOI

[17] Doi, H.; Uchii, K.; Matsuhashi, S.; Takahara, T.; Yamanaka, H.; Minamoto, T. Isopropanol precipitation method for collecting fish environmental DNA, Limnology and Oceanography: Methods, Volume 15 (2017) no. 2, pp. 212-218 | DOI

[18] Earl, D. A.; Louie, K. D.; Bardeleben, C.; Swift, C. C.; Jacobs, D. K. Rangewide microsatellite phylogeography of the endangered tidewater goby, Eucyclogobius newberryi (Teleostei: Gobiidae), a genetically subdivided coastal fish with limited marine dispersal, Conservation Genetics, Volume 11 (2010) no. 1, pp. 103-114 | DOI

[19] Esling, P.; Lejzerowicz, F.; Pawlowski, J. Accurate multiplexing and filtering for high-throughput amplicon-sequencing, Nucleic Acids Research, Volume 43 (2015) no. 5, pp. 2513-2524 | DOI

[20] Gordon, A.; Hannon, G. J. FASTX-Toolkit, 2010 (

[21] Fernandes, A. D.; Macklaim, J. M.; Linn, T. G.; Reid, G.; Gloor, G. B. ANOVA-like differential expression (ALDEx) analysis for mixed population RNA-Seq, PLoS ONE, Volume 8 (2013) no. 7 | DOI

[22] Ficetola, G. F.; Miaud, C.; Pompanon, F.; Taberlet, P. Species detection using environmental DNA from water samples, Biology Letters, Volume 4 (2008) no. 4, pp. 423-425 | DOI

[23] Gao, X.; Lin, H.; Revanna, K.; Dong, Q. A Bayesian taxonomic classification method for 16S rRNA gene sequences with improved species-level accuracy, BMC Bioinformatics, Volume 18 (2017) no. 1 | DOI

[24] Occurrence Download, The Global Biodiversity Information Facility, 2022 | DOI

[25] Goldberg, C. S.; Turner, C. R.; Deiner, K.; Klymus, K. E.; Thomsen, P. F.; Murphy, M. A.; Spear, S. F.; McKee, A.; Oyler‐McCance, S. J.; Cornman, R. S.; Laramie, M. B.; Mahon, A. R.; Lance, R. F.; Pilliod, D. S.; Strickler, K. M.; Waits, L. P.; Fremier, A. K.; Takahara, T.; Herder, J. E.; Taberlet, P. Critical considerations for the application of environmental DNA methods to detect aquatic species, Methods in Ecology and Evolution, Volume 7 (2016) no. 11, pp. 1299-1307 | DOI

[26] Harper, L. R.; Buxton, A. S.; Rees, H. C.; Bruce, K.; Brys, R.; Halfmaerten, D.; Read, D. S.; Watson, H. V.; Sayer, C. D.; Jones, E. P.; Priestley, V.; Mächler, E.; Múrria, C.; Garcés-Pastor, S.; Medupin, C.; Burgess, K.; Benson, G.; Boonham, N.; Griffiths, R. A.; Lawson Handley, L.; Hänfling, B. Prospects and challenges of environmental DNA (eDNA) monitoring in freshwater ponds, Hydrobiologia, Volume 826 (2019) no. 1, pp. 25-41 | DOI

[27] Hinlo, R.; Gleeson, D.; Lintermans, M.; Furlan, E. Methods to maximise recovery of environmental DNA from water samples, PLOS ONE, Volume 12 (2017) no. 6 | DOI

[28] Historical Weather Summer 2018 at Point Mugu Naval Air Warfare Center (;-California;-United-States)

[29] Jacobs, D. K.; Stein, E. D.; Longcore, T. Classification of California estuaries based on natural closure patterns: Templates for restoration and management (Technical Report), 2011 (

[30] Kandlikar, G. ranacapa: Utility functions and “shiny” app for simple environmental DNA visualizations and analyses (0.1.0), 2020 (

[31] Kelly, R. P.; Shelton, A. O.; Gallego, R. Understanding PCR processes to draw meaningful conclusions from environmental DNA studies, Scientific Reports, Volume 9 (2019) no. 1 | DOI

[32] Kircher, M.; Sawyer, S.; Meyer, M. Double indexing overcomes inaccuracies in multiplex sequencing on the Illumina platform, Nucleic Acids Research, Volume 40 (2012) no. 1 | DOI

[33] Kumar, G.; Farrell, E.; Reaume, A. M.; Eble, J. A.; Gaither, M. R. One size does not fit all: Tuning eDNA protocols for high‐ and low‐turbidity water sampling, Environmental DNA, Volume 4 (2022) no. 1, pp. 167-180 | DOI

[34] Kwambana, B. A.; Mohammed, N. I.; Jeffries, D.; Barer, M.; Adegbola, R. A.; Antonio, M. Differential effects of frozen storage on the molecular detection of bacterial taxa that inhabit the nasopharynx, BMC Clinical Pathology, Volume 11 (2011) no. 1 | DOI

[35] Langmead, B.; Salzberg, S. L. Fast gapped-read alignment with Bowtie 2, Nature Methods, Volume 9 (2012) no. 4, pp. 357-359 | DOI

[36] Laramie, M. B.; Pilliod, D. S.; Goldberg, C. S.; Strickler, K. M. Environmental DNA sampling protocol - filtering water to capture DNA from aquatic organisms, Techniques and Methods, 2015 | DOI

[37] Larsson, A. J. M.; Stanley, G.; Sinha, R.; Weissman, I. L.; Sandberg, R. Computational correction of index switching in multiplexed sequencing libraries, Nature Methods, Volume 15 (2018) no. 5, pp. 305-307 | DOI

[38] Leray, M.; Yang, J. Y.; Meyer, C. P.; Mills, S. C.; Agudelo, N.; Ranwez, V.; Boehm, J. T.; Machida, R. J. A new versatile primer set targeting a short fragment of the mitochondrial COI region for metabarcoding metazoan diversity: application for characterizing coral reef fish gut contents, Frontiers in Zoology, Volume 10 (2013) no. 1 | DOI

[39] Levy-Booth, D. J.; Campbell, R. G.; Gulden, R. H.; Hart, M. M.; Powell, J. R.; Klironomos, J. N.; Pauls, K. P.; Swanton, C. J.; Trevors, J. T.; Dunfield, K. E. Cycling of extracellular DNA in the soil environment, Soil Biology and Biochemistry, Volume 39 (2007) no. 12, pp. 2977-2991 | DOI

[40] Li, J.; Lawson Handley, L.-J.; Read, D. S.; Hänfling, B. The effect of filtration method on the efficiency of environmental DNA capture and quantification via metabarcoding, Molecular Ecology Resources, Volume 18 (2018) no. 5, pp. 1102-1114 | DOI

[41] Liang, Z.; Keeley, A. Filtration recovery of extracellular DNA from environmental water samples, Environmental Science & Technology, Volume 47 (2013) no. 16, pp. 9324-9331 | DOI

[42] Love, M. I.; Huber, W.; Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2, Genome Biology, Volume 15 (2014) no. 12 | DOI

[43] Majaneva, M.; Diserud, O. H.; Eagle, S. H. C.; Boström, E.; Hajibabaei, M.; Ekrem, T. Environmental DNA filtration techniques affect recovered biodiversity, Scientific Reports, Volume 8 (2018) no. 1 | DOI

[44] Martin, M. Cutadapt removes adapter sequences from high-throughput sequencing reads, EMBnet.journal, Volume 17 (2011) no. 1 | DOI

[45] McMurdie, P. J.; Holmes, S. phyloseq: An R package for reproducible interactive analysis and graphics of microbiome census data, PLoS ONE, Volume 8 (2013) no. 4 | DOI

[46] McMurdie, P. J.; Holmes, S. Waste not, want not: Why rarefying microbiome data is inadmissible, PLoS Computational Biology, Volume 10 (2014) no. 4 | DOI

[47] Miya, M.; Sato, Y.; Fukunaga, T.; Sado, T.; Poulsen, J. Y.; Sato, K.; Minamoto, T.; Yamamoto, S.; Yamanaka, H.; Araki, H.; Kondoh, M.; Iwasaki, W. MiFish, a set of universal PCR primers for metabarcoding environmental DNA from fishes: detection of more than 230 subtropical marine species, Royal Society Open Science, Volume 2 (2015) no. 7 | DOI

[48] Nagarajan, R. P.; Bedwell, M.; Holmes, A. E.; Sanches, T.; Acuña, S.; Baerwald, M.; Barnes, M. A.; Blankenship, S.; Connon, R. E.; Deiner, K.; Gille, D.; Goldberg, C. S.; Hunter, M. E.; Jerde, C. L.; Luikart, G.; Meyer, R. S.; Watts, A.; Schreier, A. Environmental DNA methods for ecological monitoring and biodiversity assessment in estuaries, Estuaries and Coasts, Volume 45 (2022) no. 7, pp. 2254-2273 | DOI

[49] Oksanen, J.; Blanchet, F. G.; Friendly, M.; Kindt, R.; Legendre, P.; McGlinn, D.; Minchin, P. R.; O’Hara, R. B.; Simpson, G. L.; Solymos, P.; Stevens, M. H. H.; Szoecs, E.; Wagner, H. vegan: Community Ecology Package (2.5-6), 2019 (

[50] Pawlowski, J.; Bruce, K.; Panksep, K.; Aguirre, F.; Amalfitano, S.; Apothéloz-Perret-Gentil, L.; Baussant, T.; Bouchez, A.; Carugati, L.; Cermakova, K.; Cordier, T.; Corinaldesi, C.; Costa, F.; Danovaro, R.; Dell'Anno, A.; Duarte, S.; Eisendle, U.; Ferrari, B.; Frontalini, F.; Frühe, L.; Haegerbaeumer, A.; Kisand, V.; Krolicka, A.; Lanzén, A.; Leese, F.; Lejzerowicz, F.; Lyautey, E.; Maček, I.; Sagova-Marečková, M.; Pearman, J.; Pochon, X.; Stoeck, T.; Vivien, R.; Weigand, A.; Fazi, S. Environmental DNA metabarcoding for benthic monitoring: A review of sediment sampling and DNA extraction methods, Science of The Total Environment, Volume 818 (2022) | DOI

[51] Perkins, T. L.; Clements, K.; Baas, J. H.; Jago, C. F.; Jones, D. L.; Malham, S. K.; McDonald, J. E. Sediment composition influences spatial variation in the abundance of human pathogen indicator bacteria within an estuarine environment, PLoS ONE, Volume 9 (2014) no. 11 | DOI

[52] Pietramellara, G.; Ascher, J.; Borgogni, F.; Ceccherini, M. T.; Guerri, G.; Nannipieri, P. Extracellular DNA in soil and sediment: Fate and ecological relevance, Biology and Fertility of Soils, Volume 45 (2009) no. 3, pp. 219-235 | DOI

[53] Pilliod, D. S.; Goldberg, C. S.; Arkle, R. S.; Waits, L. P. Estimating occupancy and abundance of stream amphibians using environmental DNA from filtered water samples, Canadian Journal of Fisheries and Aquatic Sciences, Volume 70 (2013) no. 8, pp. 1123-1130 | DOI

[54] Port, J. A.; O'Donnell, J. L.; Romero‐Maraccini, O. C.; Leary, P. R.; Litvin, S. Y.; Nickols, K. J.; Yamahara, K. M.; Kelly, R. P. Assessing vertebrate biodiversity in a kelp forest ecosystem using environmental DNA, Molecular Ecology, Volume 25 (2016) no. 2, pp. 527-541 | DOI

[55] R Core Team R: A language and environment for statistical computing. R Foundation for Statistical Computing , 2018 (

[56] Rees, H. C.; Maddison, B. C.; Middleditch, D. J.; Patmore, J. R.; Gough, K. C. REVIEW: The detection of aquatic animal species using environmental DNA - a review of eDNA as a survey tool in ecology, Journal of Applied Ecology, Volume 51 (2014) no. 5, pp. 1450-1459 | DOI

[57] Robson, H. L. A.; Noble, T. H.; Saunders, R. J.; Robson, S. K. A.; Burrows, D. W.; Jerry, D. R. Fine-tuning for the tropics: Application of eDNA technology for invasive fish detection in tropical freshwater ecosystems, Molecular Ecology Resources, Volume 16 (2016) no. 4, pp. 922-932 | DOI

[58] RStudio Team RStudio: Integrated development for R. RStudio, PBC, 2020 (

[59] Sales, N. G.; Wangensteen, O. S.; Carvalho, D. C.; Mariani, S. Influence of preservation methods, sample medium and sampling time on eDNA recovery in a neotropical river, Environmental DNA, Volume 1 (2019) no. 2 | DOI

[60] Santoro, A. E.; Dupont, C. L.; Richter, R. A.; Craig, M. T.; Carini, P.; McIlvin, M. R.; Yang, Y.; Orsi, W. D.; Moran, D. M.; Saito, M. A. Genomic and proteomic characterization of “Candidatus Nitrosopelagicus brevis”: An ammonia-oxidizing archaeon from the open ocean, Proceedings of the National Academy of Sciences, Volume 112 (2015) no. 4, pp. 1173-1178 | DOI

[61] Sard, N. M.; Herbst, S. J.; Nathan, L.; Uhrig, G.; Kanefsky, J.; Robinson, J. D.; Scribner, K. T. Comparison of fish detections, community diversity, and relative abundance using environmental DNA metabarcoding and traditional gears, Environmental DNA, Volume 1 (2019) no. 4, pp. 368-384 | DOI

[62] Schaarschmidt, F.; Gerhard, D. PairwiseCI: Confidence Intervals for Two Sample Comparisons (0.1-27). , 2019 (

[63] Schnell, I. B.; Bohmann, K.; Gilbert, M. T. P. Tag jumps illuminated - reducing sequence-to-sample misidentifications in metabarcoding studies, Molecular Ecology Resources, Volume 15 (2015) no. 6, pp. 1289-1303 | DOI

[64] Wetlands on the edge: The future of southern California’s wetlands: Regional strategy 2018 (p. 142). California State Coastal Conservancy. , Southern California Wetlands Recovery Project, 2018 (

[65] Sekar, R.; Kaczmarsky, L. T.; Richardson, L. L. Effect of freezing on PCR amplification of 16S rRNA genes from microbes associated with black band disease of corals, Applied and Environmental Microbiology, Volume 75 (2009) no. 8, pp. 2581-2584 | DOI

[66] Shaffer, H. B.; Fellers, G. M.; Randal Voss, S.; Oliver, J. C.; Pauly, G. B. Species boundaries, phylogeography and conservation genetics of the red-legged frog (Rana aurora/draytonii) complex, Molecular Ecology, Volume 13 (2004) no. 9, pp. 2667-2677 | DOI

[67] Shirazi, S.; Meyer, R. S.; Shapiro, B. Revisiting the effect of PCR replication and sequencing depth on biodiversity metrics in environmental DNA metabarcoding, Ecology and Evolution, Volume 11 (2021) no. 22, pp. 15766-15779 | DOI

[68] Smart, A. S.; Weeks, A. R.; Rooyen, A. R.; Moore, A.; McCarthy, M. A.; Tingley, R. Assessing the cost‐efficiency of environmental DNA sampling, Methods in Ecology and Evolution, Volume 7 (2016) no. 11, pp. 1291-1298 | DOI

[69] Stein, E. D.; Cayce, K.; Salomon, M.; Bram, D. L.; De Mello, D.; Grossinger, R.; Dark, S. Wetlands of the southern California coast: Historical extent and change over time (SFEI Report 720; SCCWRP Technical Report 826; p. 58). Southern California Coastal Water Research Project and San Francisco Estuary Institute, 2014 (

[70] Suomalainen, L.-R.; Reunanen, H.; Ijäs, R.; Valtonen, E. T.; Tiirola, M. Freezing induces biased results in the molecular detection of Flavobacterium columnare, Applied and Environmental Microbiology, Volume 72 (2006) no. 2, pp. 1702-1704 | DOI

[71] Swift, C. C.; Haglund, T. R.; Ruiz, M.; Fisher, R. N. The status and distribution of the freshwater fishes of southern California, Bulletin of the Southern California Academy of Sciences, Volume 92 (1993), pp. 101-167 | DOI

[72] Swift, C. C.; Spies, B.; Ellingson, R. A.; Jacobs, D. K. A new species of the bay goby genus Eucyclogobius, endemic to southern California: Evolution, conservation, and decline, PLOS ONE, Volume 11 (2016) no. 7 | DOI

[73] Taberlet, P.; Coissac, E.; Pompanon, F.; Brochmann, C.; Willerslev, E. Towards next-generation biodiversity assessment using DNA metabarcoding: Next-generation DNA metabarcoding, Molecular Ecology, Volume 21 (2012) no. 8, pp. 2045-2050 | DOI

[74] Taberlet, P.; Prud’Homme, S. M.; Campione, E.; Roy, J.; Miquel, C.; Shehzad, W.; Gielly, L.; Rioux, D.; Choler, P.; Clément, J.-C.; Melodelima, C.; Pompanon, F.; Coissac, E. Soil sampling and isolation of extracellular DNA from large amount of starting material suitable for metabarcoding studies: Extraction of extracellular DNA from soil, Molecular Ecology, Volume 21 (2012) no. 8, pp. 1816-1820 | DOI

[75] Takahara, T.; Minamoto, T.; Doi, H. Effects of sample processing on the detection rate of environmental DNA from the Common Carp (Cyprinus carpio), Biological Conservation, Volume 183 (2015), pp. 64-69 | DOI

[76] Thomsen, P. F.; Willerslev, E. Environmental DNA – An emerging tool in conservation for monitoring past and present biodiversity, Biological Conservation, Volume 183 (2015), pp. 4-18 | DOI

[77] Torti, A.; Lever, M. A.; Jørgensen, B. B. Origin, dynamics, and implications of extracellular DNA pools in marine sediments, Marine Genomics, Volume 24 (2015), pp. 185-196 | DOI

[78] Tsuji, S.; Takahara, T.; Doi, H.; Shibata, N.; Yamanaka, H. The detection of aquatic macroorganisms using environmental DNA analysis—A review of methods for collection, extraction, and detection, Environmental DNA, Volume 1 (2019) no. 2, pp. 99-108 | DOI

[79] Turba, R.; Thai, G.; Jacobs, D. Different approaches to processing environmental DNA samples in turbid waters have distinct effects for fish, bacterial and archaea communities (Scripts, Code and Dataset), Dryad, 2023 | DOI

[80] Turba, R.; Thai, G.; Jacobs, D. Different approaches to processing environmental DNA samples in turbid waters have distinct effects for fish, bacterial and archaea communities (Supplementary material), Zenodo, 2023 | DOI

[81] Turner, C. R.; Barnes, M. A.; Xu, C. C. Y.; Jones, S. E.; Jerde, C. L.; Lodge, D. M. Particle size distribution and optimal capture of aqueous macrobial eDNA, Methods in Ecology and Evolution, Volume 5 (2014) no. 7, pp. 676-684 | DOI

[82] Turner, C. R.; Uy, K. L.; Everhart, R. C. Fish environmental DNA is more concentrated in aquatic sediments than surface water, Biological Conservation, Volume 183 (2015), pp. 93-102 | DOI

[83] van der Loos, L. M.; Nijland, R. Biases in bulk: DNA metabarcoding of marine communities and the methodology involved, Molecular Ecology, Volume 30 (2021) no. 13, pp. 3270-3288 | DOI

[84] Williams, K. E.; Huyvaert, K. P.; Piaggio, A. J. Clearing muddied waters: Capture of environmental DNA from turbid waters, PLOS ONE, Volume 12 (2017) no. 7 | DOI

[85] Zinger, L.; Lionnet, C.; Benoiston, A.-S.; Donald, J.; Mercier, C.; Boyer, F. metabaR: an R package for the evaluation and improvement of DNA metabarcoding data quality, bioRxiv (2020) | DOI

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