Experimental set-up
We evaluated the effect of two widely-applied storage solutions (DESS and ethanol) and preservation treatments (heat treatment and storage temperature) on DNA degradation over time (1 day, 1 week, 2 weeks, 1 month and 3 months). Here we consider the long-term storage of samples at temperatures of -20°C or lower, and under dark conditions optimal for sample preservation. Samples were stored under suboptimal storage conditions for 3 months (ambient room temperature of around 20°C and at a cold room set to 5°C). Our goal was to evaluate how different preservation methods effect the initial stages of sample preservation, and choose the right protocol for our study. We extracted DNA from the marine teleost species the Australasian snapper Chrysophrys auratus , which has been observed to experience high rates of DNA degradation within the first 24 hours after sampling (M. Wellenreuther, personal observation). Controlling the initial stage of DNA sample preservation was considered crucial to this study for obtaining good quality genomic data from natural snapper populations.
Methods
Fin clips (approximately 1 cm2) were collected from the anal and caudal fin (avoiding the cartilage tissue as much as possible) from five Australasian snapper (all of the same age and kept under the same environmental conditions) from the Seafood Research Unit in Nelson operated by The New Zealand Institute of Plant and Food Research Limited. We used three storage treatments: a preservation solution, a heat treatment and a set storage temperature (Table 1). Fin clips were stored in 2.0 mL sterile O-ring tubes, with 1.5 mL preservation solution (enough to fully submerge the tissue). Samples were collected within a two-hour time window, cleaning sampling gear with ethanol between individuals. The fin clips were extracted at five different time points: 1 day, 1 week, 2 weeks, 1 month and 3 months. The two preservation solutions used in this study were ethanol (absolute >99.5%, grade: molecular biology) and DESS (20% DMSO, 0.25 M EDTA, sodium chloride saturated solution, see supplementary materials). Ethanol dehydrates cells and causes proteins to coagulate in order to preserve the sample by inhibiting cellular processes that may breakdown the DNA. DESS prevents DNA degradation by deactivating metal-dependent enzymes (i.e. DNase) using EDTA. In addition, our EDTA 0.5 M stock solution was set at pH 8 with NaOH to dissolve the EDTA, creating a pH buffering effect. The solution was salt statured with sodium chloride (NaCl) which stabilizes the DNA (MacFarlane et al., 2017). Finally, DMSO is known to transport compounds (e.g. EDTA and NaCl) across cell membranes and serves as a cryogenic (Seutin et al., 1991). DESS is known to perform well under a wide range of temperatures, including room temperature. Heat treated samples were incubated at 80° C for 5 minutes within an hour of sampling to destroy DNA degrading enzymes. Samples were stored at a “cold” temperature of 5°C and at room temperature (~20°C). Temperatures were chosen to resemble the storage conditions typically encountered in the field, and generally considered suboptimal for storage over multiple days. All samples were stored under dark conditions. Five technical replicates from each individual, were extracted for each combination of preservation treatments over five different time intervals (Table 1), creating a total of 200 DNA extractions.
DNA was extracted using a high-salt extraction protocol adapted from Aljanabi and Martinez (1997) (see supplementary materials). To standardize the amount of DNA extracted from each sample, between 30 and 40 milligram of tissue was used for each extraction. All extractions were performed by the same operator. Wide-bore pipette tips were used every time DNA was pipetted from between tubes to reduce physical stress on the DNA fragments. DNA was eluted overnight at room temperature in 100 µL TE buffer (10 mM Tris-HCl pH 8, 1 mM EDTA). Impurities and DNA concentration were measured using an Implen© NP80 spectrophotometer and Qubit broad range kit (Invitrogen©). Samples were diluted to a concentration of ~3.0 ng/µL and assessed on the Fragment Analyzer (Advanced Analytical) using the high-sensitivity genomic DNA analysis kit, and PROsize V3.0.1.6 to summarize the results.
A multi-factor ANOVA was performed to test which variables had a significant effect on preservation condition of the sample and DNA fragmentation. DNA fragmentation was quantified using GQN quality score implemented in PROsize, based on the running front against known fragment sizes in the HS genomic DNA ladder. We performed three separate multi-factor ANOVAs: One using all samples and two using samples from each of the solution treatments. Visualization of fragment size decay per treatment over time was done by assessing the electropherograms (implemented in PROsize). Electropherograms were merged using a custom Python script. Slight differences between each run resulted in different point estimates for larger fragment sizes. To create overlapping data points between each of the three fragment analyzer runs, fragment sizes above 1 kbp were rounded up to the nearest 10, and relative fluorescence unit (RFU) values were averaged across matching fragment sizes. Similarly, fragment sizes above 10 kbp were rounded to the nearest 100 and again RFU values were averaged. Negative RFU values were clipped to zero. Mean RFU values per treatment were calculated after subsetting and 95% confidence intervals were estimated and plotted in R (R Core Team, 2013).
Finally, the area under each curve was estimated using DescTools (Signorell, 2016). We estimated the area under each curve per treatment to assess the relative abundance of HMW DNA in total amount of DNA. Fragment sizes smaller than 250 bp were not taken into account to prevent biases from RNA.