Optimising Extraction of DNA From Museum Insect Specimens

DNA technologies have many advantages for biomonitoring and biodiversity analyses, but these depend on the availability of relevant reference DNA barcodes. To be most useful, a DNA barcode should be linked to a taxonomic name, which can in turn be connected to ecological information. This linking can be achieved by DNA barcoding of taxonomically identified specimens. Museums are a promising source of such specimens, but the DNA in museum specimens is often degraded, necessitating carefully optimised DNA extraction methods. In this issue of Molecular Ecology Resources, Holmquist et al. (2025) present a DNA extraction protocol for museum insect specimens, using in‐house formulated Solid Phase Reversible Immobilisation (SPRI) beads. The authors carried out several experiments with statistical evaluation to determine optimal DNA extraction parameters, before testing the protocol on a large and diverse pool of museum‐held insect specimens. The result is a low‐cost and effective DNA extraction protocol for diverse museum insect specimens.

Insects are vitally important components of Earth's biodiversity, but monitoring these communities is challenging due to the huge diversity of species that exist. DNA sequencing technologies enable efficient molecular characterisation of insect diversity, but the resulting molecular taxonomic units are typically disconnected from species or functional information (Meier et al. 2024). This makes ecological insights difficult to achieve for wide swathes of biodiversity. Reference DNA barcodes from taxonomically identified species can bridge this gap (Kress et al. 2015), but the process of taxonomically identifying insect specimens is very difficult due to a scarcity of suitable taxonomic expertise. New workflows that combine machine learning with mass DNA barcoding of trapped insect samples have the potential to resolve this challenge over time (Meier et al. 2024). On the other hand, it is important to consider existing resources such as museum collections as sources of reference DNA barcodes.

Museums often hold rich collections of biological specimens, usually with taxonomic identifications, accumulated over long periods of time (Figure 1). In theory, these collections represent a compelling source of DNA barcodes (Raxworthy and Smith 2021). The DNA in museum specimens is often degraded due to specimen age and suboptimal preservation, however; this makes it difficult to recover DNA barcode sequences from some specimens (Hebert et al. 2013). Assembly of multiple shorter amplicons can be effective in cases where DNA fragmentation makes PCR amplification of standard barcode regions unfeasible (D'Ercole et al. 2021; Prosser et al. 2016), but this is more complex and costly than conventional DNA barcode generation. Therefore, it is important to develop optimal methods of DNA extraction from museum insect collections to maximise the chance that conventional DNA barcoding can succeed.

FIGURE 1.

Museums are a promising source of DNA barcodes from taxonomically identified insect specimens.

Holmquist et al. (2025) address this issue by developing a low‐cost and effective DNA extraction approach for museum insect specimens using SPRI beads. Their work has two main strengths. Firstly, they carried out a fine‐grained optimisation and statistical evaluation of DNA extraction parameters. While many other studies have compared different DNA extraction protocols for museum specimens, these have usually focused on vertebrate specimens rather than insects (Raxworthy and Smith 2021). Furthermore, consideration of parameters within a protocol using statistical approaches is rare and gives additional confidence in the conclusions. Secondly, the optimised protocol was tested on a large and taxonomically diverse set of museum‐held insect specimens. This provides confidence that the method has broad application, and results in practical recommendations for adjusting the method for groups with differing characteristics (e.g., size and toughness).

Importantly, SPRI beads, more commonly used in sequencing library preparation, can be formulated in house at much lower cost than those purchased from commercial suppliers, making this approach economical to apply to large numbers of specimens. This is a critical point because developing large‐scale DNA barcode reference libraries will be expensive and so optimising costs is important. Nevertheless, the SPRI beads still represent the most expensive DNA extraction component, so the authors first varied the PEG to SPRI bead ratio to identify the point that balanced retention of DNA with a low concentration of beads.

In a second experiment, the authors optimised the amounts of NaCl, PEG and SPRI beads to maximise the purity of the extracted DNA. This experiment included specimens from a wide range of insect orders to determine how generalisable these findings are. In a third experiment, they compared the performance of their optimised SPRI bead protocol with two commonly used DNA extraction kits and the HotSHOT method (Truett et al. 2000), using one beetle and one fly species. This experiment showed that the performance of the SPRI bead method was close to that of the best‐performing (and more expensive) Qiagen DNeasy kit, while the HotSHOT protocol was the least effective. One factor likely contributing to this outcome is the gentle nature of the SPRI bead process, especially compared with the hot NaOH of the HotSHOT method.

Finally, the authors applied an optimised SPRI bead protocol (based on findings of the first three experiments) to 3728 museum‐held insect specimens belonging to 14 taxonomic orders, with collection dates ranging from 1972 to 2024 (mean age 19 years). Based on this experiment, the cost per specimen ranged from four to 11.6 cents, depending on extraction constituent proportions, and took 45 min per 96 well plate after lysis (using a semi‐automated 96‐well pipettor), indicating that multiple plates of specimens could be processed within 1 day. The cost and efficiency of the protocol thus compare favourably with other approaches. A trade‐off between DNA yield and PCR success was identified, most likely because improved DNA purity correlates with lower yield but higher PCR success. The inclusion of a wide range of different taxa was important, as there is variation in how well DNA is preserved and how recoverable it is due to differing body sizes and degrees of sclerotisation, and different collection and preservation methods typically used for different taxa. This experiment also enabled the authors to provide some useful guidelines for adjusting the protocol for different taxa. PEG and NaCl were the most important components to optimise, and it may be worthwhile doing so for bespoke projects in other insect collections.

Surprisingly, no correlation between DNA concentration and specimen age was observed, but specimen age is still likely to be an important factor. The oldest specimens considered were collected in 1972. Many museum specimens are much older than this, and it seems likely that alternative approaches will be needed at some point, although this may vary by taxon and other factors. It is also important to remember the importance of verifying taxonomic names in collections and digitisation of specimen metadata (if not already done), which is likely to become the largest cost of the DNA barcoding process.

The research delivered a useful tool for DNA barcoding of museum‐held insect specimens. The authors have achieved an impressive reduction in operating costs, which are complemented by low costs of modern DNA sequencing technologies (Srivathsan et al. 2018). This study is an important advance towards the goals of developing complete DNA barcode libraries for local insect faunas, and enabling DNA‐based biomonitoring of insect communities (Pawlowski et al. 2021).

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

The authors have nothing to report.