Species‐level biodiversity assessment using marine environmental DNA metabarcoding requires protocol optimization and standardization

Abstract

DNA extraction from environmental samples (environmental DNA; eDNA) for metabarcoding‐based biodiversity studies is gaining popularity as a noninvasive, time‐efficient, and cost‐effective monitoring tool. The potential benefits are promising for marine conservation, as the marine biome is frequently under‐surveyed due to its inaccessibility and the consequent high costs involved. With increasing numbers of eDNA‐related publications have come a wide array of capture and extraction methods. Without visual species confirmation, inconsistent use of laboratory protocols hinders comparability between studies because the efficiency of target DNA isolation may vary. We determined an optimal protocol (capture and extraction) for marine eDNA research based on total DNA yield measurements by comparing commonly employed methods of seawater filtering and DNA isolation. We compared metabarcoding results of both targeted (small taxonomic group with species‐level assignment) and universal (broad taxonomic group with genus/family‐level assignment) approaches obtained from replicates treated with the optimal and a low‐performance capture and extraction protocol to determine the impact of protocol choice and DNA yield on biodiversity detection. Filtration through cellulose‐nitrate membranes and extraction with Qiagen's DNeasy Blood & Tissue Kit outperformed other combinations of capture and extraction methods, showing a ninefold improvement in DNA yield over the poorest performing methods. Use of optimized protocols resulted in a significant increase in OTU and species richness for targeted metabarcoding assays. However, changing protocols made little difference to the OTU and taxon richness obtained using universal metabarcoding assays. Our results demonstrate an increased risk of false‐negative species detection for targeted eDNA approaches when protocols with poor DNA isolation efficacy are employed. Appropriate optimization is therefore essential for eDNA monitoring to remain a powerful, efficient, and relatively cheap method for biodiversity assessments. For seawater, we advocate filtration through cellulose‐nitrate membranes and extraction with Qiagen's DNeasy Blood & Tissue Kit or phenol‐chloroform‐isoamyl for successful implementation of eDNA multi‐marker metabarcoding surveys.

Considerations for metabarcoding‐based port biological baseline surveys aimed at marine nonindigenous species monitoring and risk assessments

“…Increasing evidences showed that the use of universal and degenerated COI primers is leading to nonspecific amplification of prokaryotic and nontarget eukaryotic DNA (Collins et al, ). Yet, in this context, at least two alternatives are possible to improve eDNA‐based biodiversity assessments: (a) increasing sequencing depth (Grey et al, ) or (b) using group‐specific markers (Jeunen, Knapp, Spencer, Taylor, et al, ). Yet, none of them ensures full biodiversity recovery and both significantly increase costs.…”

Environmental DNA (eDNA) metabarcoding reveals strong discrimination among diverse marine habitats connected by water movement

“…Sample processing followed the recommendations from (Jeunen et al, ). Briefly, water samples were filtered over a 1.2 μm cellulose‐nitrate filter (CN, Whatman™).…”

Maximizing fish detection with eDNA metabarcoding

“…been employed to describe fish assemblages by extracting DNA from the marine environment (Jeunen et al, 2019a;Stat et al, 2017;Thomsen et al, 2012). These environmental DNA (eDNA) techniques remove the need for repeated use of taxonomic expertise as DNA sequences can be taken from samples identified once by an expert and thereafter placed in a reference database of fish DNAso-called DNA barcoding (Taberlet, Bonin, Zinger, & Coissac, 2018).…”

“…Sample volumes for fish studies range widely (400-6,000 ml reported in review by Turner et al, 2014;Wilcox et al, 2013), although both 1,000 ml (Koziol et al, 2019;Stat et al, 2017) and 2,000 ml volumes are frequently sampled (in review by Turner et al, 2014). Sample replication also varies widely, from studies using triplicate 250-ml samples (Singer et al, 2019), eight replicate 1,000-ml samples (Koziol et al, 2019), to five replicate 2,000-ml samples (Jeunen et al, 2019a) per site. For many species, such as amphibians and macroinvertebrates, detection rates are higher with increased sampling volumes (Lopes et al, 2017;Mächler, Deiner, Spahn, & Altermatt, 2016).…”