Molecular data collection
Genomic DNA from freshly preserved material was extracted from the
parapodia and/or foot using the DNeasy Blood and Tissue kit (Qiagen
Inc., Valencia, CA, USA) following the manufacturer’s protocol. In order
to obtain a higher DNA yield for millimetric specimens and a
non-destructive DNA extraction of mostly ‘degraded’ whole museum
individuals we used the protocol described in Tin, Economo & Mikheyev
(2014) using silica-based beads, with some in-lab modifications
(Derkarabetian et al., 2019). Extractions were quantified using the
Qubit 2.0 fluorometer (Life Technologies, Inc.) dsDNA High Sensitivity
kit and visualized on a 2200 Tape Station (Agilent) to assess DNA
degradation. Up to 500 ng of DNA in 130 µL for all fresh specimens was
sonicated for 80 s with a Covaris S220 Focused-ultrasonicator for a
target peak of 500–600 bp, with a Peak Incidence Power of 50, Duty
Factor of 10%, and 200 cycles per burst. Sonication time was adjusted
depending on the Tape Station results for each sample since museum
samples were naturally degraded to the appropriate size for sequence
capture library preparation (see Table 2).
Library preparation followed the general protocol on the UCE website
(https://www.ultraconserved.org/) and some in-lab modifications
(Derkarabetian et al., 2018). Libraries were prepared using the KAPA
Hyper Prep kit following the manufacturer’s protocol and using up to 250
ng of DNA as starting material. For samples with lower DNA yield, we
used it all (down to 5 ng). We used fresh Serapure Speed-beads for all
clean-up steps (Rohland & Reich, 2012) washing with freshly prepared
80% EtOH. After the first clean-up, 25 µL of fragmented,
double-stranded DNA was assembled for end repair and A-tailing, 20 °C
for 30 min and 65 °C for 30 min, respectively. Immediately afterward we
proceeded to adapter ligation, samples with >200 ng of DNA
were run at 20 °C for 30 min using the universal iTru Stubs at 10 µM,
while samples <200 ng were run up to 1h using 5 µM iTru Stubs.
An immediate post-ligation clean-up step was carried at 0.8X for
high-yield, fresh samples, and at up to 3X for degraded, low-yield
samples. Library amplification was conducted using 25 µL of post-ligated
libraries, using individual iTrue i5/i7 dual index adaptors (8 bp
long ; Glenn et al., 2016), with an initial denaturation step at 98 °C
for 45 s, then 6–14 PCR cycles (adjusted to post-ligated Qubit
concentrations) of 98 °C for 15 s, 60 °C for 30 s and 72 °C for 1 min,
and a final extension step of 72 °C for 5 min. Libraries were then
washed, quantified, and 125 ng of each were pooled in batches of eight
samples for a total of 1000 ng. Pools were then speed-vacuumed if
necessary, to a final volume of 14 µL.
Hybridization capture for targeted UCEs was carried following the
myBaits® v.4 user’s manual and the target enrichment for Illumina
standard workflow protocols
(https://www.ultraconserved.org/#protocols).
Pooled libraries were hybridized with the synthesized bait set at 60 °C
for 24 h. Hybrid pools were then bound to streptavidin beads (Dynabeads
MyOne C1, Invitrogen), washed, and eluted in 31 µL of NF Water.
Post-hybridization amplification was carried in a 50 µL reaction using
15 µL of hybridized pools, with same post-ligation PCR conditions but
for a total of 16 cycles. Immediately followed a bead clean up with 70%
EtOH, Qubit 2.0 quantification, and visualization and molarity
calculation using a 2200 Tape Station. Post-hybridized pools were
pooled in equimolar amounts and sequenced in the Illumina NovaSeq 6000
SP platform (paired-end, 150 bp) at the Bauer Core Facility, Harvard
University. New sequences were deposited in the NCBI Sequence Read
Archive (BioProject PRJNA612319); voucher information and assembly
statistics are available in Table 2 and S2.