With the rapid adoption of long-read sequencing and other technologies that require high molecular weight DNA input, there is rising demand for high-quality samples from biorepositories around the world. To accommodate this, scientists at the Smithsonian Institution’s National Museum of Natural History in Washington, DC, developed and published a rapid, inexpensive method for quantifying DNA quality. We spoke with co-authors Jonathan Coddington and Daniel Mulcahy to learn more about this new technique.
Q: Why is there interest in working with high molecular weight DNA?
JC: We work for the Smithsonian Institution, so we’re in the forever business. We want to build a library of the highest-quality genetic resources for biodiversity. There are about 1.9 million species described and many, many millions more out there that haven’t been described — and we’re facing the sixth extinction crisis. So it’s important to collect and preserve samples that would have the maximum utility to science for many decades into the future.
DM: We’re interested in high molecular weight DNA because now it’s very practical to sequence the entire genome of non-model organisms. Even though you shear the DNA up to make the libraries, it’s better to start with big chunks of complete DNA so you’re shearing randomly across the genome, not as it’s been sheared through wear and tear.
JC: We’re also anticipating better technology in the future. Long-read technology is upon us, so everything from what you do in the field until when you extract the DNA needs to be adapted for very high molecular weight DNA.
Q: Before your method was published, how were people assessing sample quality?
DM: A lot of people were putting it on a spectrophotometer and looking at the concentrations. Another common way was to just PCR amplify short fragments, but that doesn’t really tell you if it’s high molecular weight DNA. There are more expensive methods as well, but we were trying to come up with an inexpensive way to quickly assess a lot of tissues based on DNA size.
JC: The concern with DNA quality has been around ever since the beginning of sequencing, but because PCR was so dominant people only measured the quality of their sample based on the marker that they wanted to amplify, not whole genomic DNA. An initial baseline assessment of the size frequency distribution for whole genomic DNA is almost never done in biodiversity biorepositories. We need to move the community to thinking about whole genomes.
Q: Have you seen signs that people are going in that direction?
JC: The sequencing research community has really moved beyond PCR marker-based sequencing. They want to do whole genomes and they’re looking to us to supply that high-quality DNA.
Q: How will having DNA quality information change how genomic scientists work?
DM: Our method can tell you how much of your sample was greater than the band you chose. Ultimately we’ll get to a point where we can sequence an entire chromosome; ideally you would want to be able to run the DNA out to the level where you can see various-sized chromosomes if it’s completely intact DNA. That’s where the future of this is going.
JC: Our paper was designed to enable a really cheap, really fast way for our community to inform people who want to do experiments what sort of quality DNA we might have. It’s a gel electrophoresis-based method because everybody can run gels.
Q: How will these quality assessments affect biorepositories?
JC: Dan and I represent a rapidly growing community of museums and other kinds of biorepositories. The Smithsonian, for example, has publicly about 90,000 samples. We’re also supporting an international network of biorepositories using a common data standard to report the existence and quality of samples that we all hold. The global total is about 615,000 publicly discoverable genetic resources. When we’re loaning out really rare samples, we want to know what you want to do with it. If you need the high-end stuff for a really good reason, we’ll give it to you.
DM: But if they don’t need it, we’ll give them lower-quality DNA to save the better stuff for someone who wants to do whole-genome assemblies.
Q: In the paper, you also looked at preservation methods. What were your key findings?
DM: That was a very preliminary study. If you stick a piece of muscle in the freezer and freeze it, it’s probably as good as it can be. But when you thaw it out, those proteins start to break down and the enzymes start to chew up the DNA.
JC: The DNases can still be active on the thaw cycle. I think the community assumption was that fresh frozen in liquid nitrogen was as good as it gets. What we found that was surprising was that saturating it in some sort of preservation buffer prior to freezing produces results that are better than just freezing. That’s the intriguing implication.
DM: We’re conducting a much longer-term study now. We’d like to see other people test this with other organisms, other buffers, and different storage methods.