Comparing sediment preservation methods for genomic biomonitoring of coastal marine ecosystems

Highlights

• We assessed effects of sample preservation for eDNA-based biomonitoring.

• We compared freezing versus preservation-solution treatment.

• DNA extraction efficiency was highly congruent among treatments.

• Bacterial community profiles as bioindicators were highly similar among treatments.

• Standard Operating Procedures (SOPs) for biomonitoring can use both treatments.

Abstract

To avoid loss of genetic information in environmental DNA (eDNA) field samples, the preservation of nucleic acids during field sampling is a critical step. In the development of standard operating procedures (SOPs) for eDNA-based compliance monitoring, the effect of different routinely used sediment preservations on biological community structures serving as bioindicators has gone untested. We compared eDNA metabarcoding results of marine bacterial communities from sample aliquots that were treated with a nucleic acid preservation solution (treated samples) and aliquots that were frozen without further treatment (non-treated samples). Sediment samples were obtained from coastal locations subjected to different stressors (aquaculture, urbanization, industry). DNA extraction efficiency, bacterial community profiles, and measures of alpha- and beta-diversity were highly congruent between treated and non-treated samples. As both preservation methods provide the same relevant information to environmental managers and regulators, we recommend the inclusion of both methods into SOPs for biomonitoring in marine coastal environments.

Introduction

The advent of efficient, inexpensive, fast, up-scalable and fully automatable DNA metabarcoding protocols (Aylagas et al., 2016; Cordier et al., 2017; Keeley et al., 2018; Lanzén et al., 2020) and data analyses packages (Aylagas and Rodriguez-Ezpeleta, 2016; Macher et al., 2021; Zinger et al., 2021) have heralded in a new era in the field of environmental biomonitoring. Traditional methods for assessing marine ecosystem health usually rely on the microscopic identification of macroinvertebrates, which can be used as bioindicators due to their specific response to individual stressors or a combination of multiple stressors. Such stressors include for example aquaculture associated organic enrichment (Brown et al., 1987; Carroll et al., 2003), hydrocarbon pollution at production sites (Cordier et al., 2019; Lanzén et al., 2016; Laroche et al., 2016), or toxic chemicals originating in industry or urban wastewater (Chariton et al., 2010; Vaalgamaa et al., 2013; Yoon et al., 2020). Traditional taxonomic monitoring approaches rely on the observations of organisms that can be identified morphologically, such as macroinvertebrates (Bonada et al., 2006; Magurran et al., 2010) or algae (Reavie et al., 2010), and do not allow the exploitation of strong biomarkers that are not readily identifiable and quantifiable using microscopy, such as bacteria. Furthermore, they are limited by high costs, lengthy sample analysis time, often unverifiable taxonomic precision, and cannot easily be scaled up (Baird and Hajibabaei, 2012). The necessity to increase the frequency and scale of environmental biomonitoring due to an increasing human impact on our planet in general (Chiang et al., 2021; Myers and Smith, 2018; Orr et al., 2005; Wu et al., 2013) and coastal ecosystems in particular (Harley et al., 2006; Puritz and Toonen, 2011; van de Velde et al., 2018), has fueled the development of alternative biomonitoring strategies. The solution that is widely accepted as having the highest potential for assessing the ecological status of marine ecosystems is the interrogation of DNA extracted from environmental samples. In these protocols, environmental DNA (eDNA) is extracted from e.g. sediment or water samples and specific taxonomic target genes are amplified using PCR prior to high-throughput sequencing (Taberlet et al., 2018). For marine biomonitoring, bacteria have emerged as a potentially powerful indicator group (Dowle et al., 2015; Keeley et al., 2018; Stoeck et al., 2018a; Verhoeven et al., 2018), alongside other microscopic organisms such as ciliates (Forster et al., 2019; Stoeck et al., 2018b), foraminifera (Pawlowski et al., 2016; Pochon et al., 2015) and diatoms (Apothéloz-Perret-Gentil et al., 2017; Rivera et al., 2018). Current efforts of the scientific, industrial and regulatory communities are to develop eDNA-standard protocols to exploit bacteria as bioindicators, which can be implemented in routine biomonitoring regulations and practice (Cordier et al., 2017; Laroche et al., 2018; Stoeck et al., 2018a). The establishment of standard protocols for sample collection, treatment and analysis, are critical for enabling results that can be compared between different monitoring studies, including in relation to sampling (Hestetun et al., 2021a), DNA extraction (Hestetun et al., 2021b; Pearman et al., 2021) and reproducibility of PCR and Illumina sequencing (Dully et al., 2021). One aspect that has so far gone untested, to the best of our knowledge, is the treatment of samples between collection and DNA extraction. In the first step after sample collection, it is necessary to preserve the sediment samples after sampling. In the ideal case, DNA should be extracted from sediment samples immediately after sample collection. However, this is often times not possible and samples will have to be stored several days to months until DNA extraction. Poorly controlled storage condition may then result in DNA degradation or microbial growth and severely bias downstream analyses of the DNA-derived biological community profiles (Rubin et al., 2013). The two most widely used options are preservation of samples using a nucleic acid preservation solution (Cordier et al., 2019; Laroche et al., 2017; Lejzerowicz et al., 2015) or freezing of the sample without any further sample treatment (Aylagas et al., 2014; Lanzén et al., 2020; Polinski et al., 2019; Steyaert et al., 2020). Commercially available preservation solutions such as Lifeguard® prevent RNAse and DNAse activities, allowing for 16S rRNA profiling (metabarcoding) of bacterial communities on samples collected in the field under any conditions (MoBio, 2011). The advantage of using a preservation solution is that no additional equipment is required to freeze the sample after collection (such as a freezer, dry ice or a liquid nitrogen cooler), which is often challenging in remote locations or aboard small sampling vessels. Furthermore, an expensive cooled transport to the sample-processing laboratory by a courier service is not required because microbial community profiles in environmental samples are maintained with such a solution for at least one week and RNA integrity even for 30 days at room temperature (MoBio, 2011). On the downside, commercially available standardized nucleic acid preservation solutions can be relatively expensive (currently up to ca. €2000 per liter), which adds substantially to the costs of environmental monitoring. Furthermore, adding preservation buffer to each sediment sample is more time consuming, adding further increasing costs compared to freezing samples. Thus, freezing samples within a few hours of collection is a faster and less-expensive option for sample preservation. However, in this case the additional transportation costs of the frozen samples on blue or dry ice from the location of the sampling to the sample processing laboratory should also be considered. The major question, however, is the influence of the sediment preservation method on the resultant data.

To address this question, we here collected sediment samples from different marine coastal locations, namely two different locations at the west coast of Scotland subjected to organic enrichment resulting from aquafarming installations and estuarine sites at the Basque coast subjected to different degrees of urban and industrial impacts. We split all samples in two aliquots, one set of which was preserved with a commonly used nucleic acid preservation solution, and one set of which was preserved by freezing. We then extracted DNA from all samples and used a standard eDNA metabarcoding protocol to analyze the bacterial community composition. Statistical analyses were then conducted to compare the results obtained from frozen and solution-preserved samples to inform the development of standardized operating protocols for eDNA-based compliance monitoring.