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.