Comprehensive study of bee phylogeny (using UCEs), age, and biogeography. What is pretty cool is that it serves as a comparative treatment to Charles Michener's classic 1979 paper on bee biogeography. Michener posited that bees originated in Western Gondwana, but it hasn't been adequately tested using molecular phylogenies and new methodologies. Another thing that I think is amazing -- the molecular dating. I'm talking 185 bee fossils used -- and not just node calibrations! Silas did the dating analysis, using a fossilized birth-death model (in the program MrBayes), meaning he assigned fossils to clades and not to a specific node or branch.

We had some media attention on this, too! Kind of interesting to see how that process worked. Our College of Ag, CAHNRS, put out an article and then WSU Insider published it a couple of days later. Several internet sites picked up the story put out by WSU, such as Popular Science. Silas and I did a radio interview for Northwest Newsradio. Silas was interviewed by NPR! 

WSU distributes their news articles (like ours) on EurekAlert, so that other sites can access them for content. Some internet sites put their own 'creative' spin on it -- like the one that was titled "A team of paleontologists find fossils that could radically change what is known about bees" on Crast.net. Hm.... not exactly. This paper didn't deal with describing new fossils, but it was the most extensive use of fossil data on a bee molecular phylogeny. 

The week was fun and info-packed! Participants were provided with comprehensive training and hands-on experience in the latest methodologies and techniques relevant to obtaining and analyzing ultraconserved elements. Through a combination of lectures, lab experiences, and practical tutorials and exercises, attendees gained an understanding of the UCE workflow – from DNA extraction and steps in library preparation, to the bioinformatic pipeline and preparing data for a comparative analysis in R.
 
The instructors think that we were successful in having an environment of active learning, engagement, and knowledge exchange. Participants had the opportunity to meet scientific peers, allowing for valuable networking and collaboration opportunities. We surveyed the attendees on the first and last days to gauge impact of the workshop, and the feedback was positive. Participants reported an increase in skills and knowledge, and some of the highest impact areas were: 1) understanding of the Phyluce pipeline (used to process UCE data), 2) describing the purpose of a bead clean step, and 3) knowing the difference between a concatenated and a coalescent phylogenetic analysis. Stay tuned for 2025, when we host the UCE Phylogenomics Workshop in Pullman, WA! Also great news - there are NSF funds to help support travel and meals.

We are excited to welcome Lexi Menth to the lab. She is starting at WSU this summer so that she can get a jump on fieldwork. Lexi is undertaking a cool project  -- an insect survey of the Fairchild Air Force Base, located west of Spokane. She'll be using various techniques to collect insects, such as light trapping, pitfall traps, and sweep netting. We are working with the US Fish and Wildlife Service, and Silas Bossert is also a PI on the grant. Everyone is anticipating the results of the next two years of sampling!

The lab has been getting the molecular protocols for next-gen sequencing up and running. One of the primary tools we have for our phylogenomics work is to do low-coverage genome sequencing on an Illumina sequencer. For the short-read sequencing, we actually NEED fragmented DNA (we're targeting 250-600 basepairs total to submit to sequencing (with adapters, etc.). We need to ascertain the quality of the DNA extract then make an educated guess as to how long to shear the DNA to get it broken into the ideal size pieces. We have a lot of old specimens with degraded DNA, and then we may choose not to shear at all. We're using a Bioruptor here at WSU.  After getting our sheared sample, we go through a Kapa kit-based library preparation in the lab. 

Although I think it's stressful to purposely fragment the DNA, as you'll see below, it is critical that you really commit to it and often do a few cycles in order to obtain smaller pieces -- the long fragments will not be useful downstream anyway!

The first step of preparing libraries for Illumina sequencing is fragmenting the input DNA into a desired range of ~300-600 bp. Typically, there are two ways for doing that: mechanic fragmentation and enzymatic fragmentation. Mechanical fragmentation has been more widely used because of many factors, but mainly because its unbiased and can be more consistent about the fragment size obtained. Besides these factors, despite requiring a higher initial investment (buying the equipment, which is generally a sonicator), throughout time it turns cheaper because you don’t need anything else than tubes to accommodate your DNA, and electricity and water for the machine to work.

We are starting to setup our lab for preparing libraries to sequence Low Coverage Genome using Illumina technology. And, once we have a sonicator (Dianogene Bioruptor plus) available at WSU we decided to perform some tests to establish the best parameters to fragment the DNA we will be using for our projects. Most of our projects rely on museum specimens, which have, generally, DNA degraded in some levels.

After consulting the available manual of the equipment (https://www.diagenode.com/en/protocols/dna-shearing-for-standard-and-plus-protocol), and a blog post (https://igatechnology.com/blog/exome-seq-dealing-with-degraded-samples/) about the same topic and having the same questions – samples with some level of fragmentation, we prepared an experiment to certify that our setup would produce the results we are expecting considering the kind of samples we have.

For our experiment we selected two samples: one with a less degraded or more contiguous DNA, extracted from a fresh specimen, and another one with degraded DNA, extracted from a museum sample. it was expected that these samples would give us two different scenarios or levels of fragmentation, so we would be able to evaluate how different configurations of the sonicator will influence the different samples. We then undertook a TapeStation analysis for these two samples to have a sense of the distribution of fragment sizes in both samples before the sonication. The results show that our expectations were right, the fresh specimen had DNA with fragment sizes mainly longer than we were able to read with the screen tape we had currently available (Figure 1.A), which can detect fragments between 25 and 1500 bp. Conversely, the museum sample had a fragment distribution ranging between 100-1500bp, with a peak around 620bp (Figure 1.B), which is slightly above the mean fragment size we were targeting.

We then used two configurations of the sonicator to fragment the DNA: the first one had two cycles with the machine on for 30 seconds and off for 60 seconds. While the second one had three cycles with the same pattern 30/60.