Ethanol curation plays a crucial role in preserving museum specimens, especially soft-bodied arthropods. Dehydration happens as a result of less than ideal storage and preservation conditions. If rehydration is desired for a specimen, then effective rehydration methods must be investigated and compared. The impact different rehydration techniques have on specimen tissues has been studied in vertebrates but is less explored in arthropods. Additionally, how these techniques impact the DNA of dehydrated arthropod specimens has not been assessed. In this study, we investigated the impact of two rehydration approaches on dehydrated spiders from the same donation. We used a gradual ethanol rehydration “Step-Up” method, and a heat-accelerated, or Heat-Rehydration, method. To determine which approach was most effective for restoring dehydrated spiders in ethanol, we assessed spider morphology for damage, DNA yield, and DNA integrity. We found that all desiccated and rehydrated spiders, regardless of treatment choice, displayed varying levels of tissue desiccation and separation of the abdomen and legs. The Step-Up method was more consistent than the Heat-Rehydration method in rehydrating tissues and reducing the separation of tissues from the abdomen. Overall, we found a significant difference in abdominal tissue separation between our controls and treatments but no significant difference when assessing damage to pedipalps. Additionally, DNA capture from all specimens was low and significantly degraded compared to a positive control group. Our recommendations for museum collections managers are to consider the fragility of dehydrated materials in deciding what rehydration technique to use and to preferentially extract DNA from other material if possible.

Fluid preservation, museum collections, specimen preservation, spiders

The proper management of natural history museum collections is integral to preserving the integrity of research materials, and doing so requires trained personnel who oversee the maintenance and preservation of specimens (Singer 2014; Neisskenwirth 2020). Collections’ staff watch over extensive numbers of specimens to maintain their integrity for generations to come. Despite the important and irreplaceable role that natural history specimens play in answering research questions, specimens are always at risk of decay and data loss from various preservation threats. Choices of preservation to prevent specimen degradation can differ vastly based on the taxonomic group (Van Cleave and Ross 1947; Singer 2014). The method of specimen preservation is based primarily on the type of specimen (e.g., hard-bodied insects are traditionally pinned; non-insect arthropods are generally preserved in ethanol or other fluids) (Kumar et al. 2022). When museum specimens become degraded, as when ethanol specimens dehydrate, the treatment choice for rejuvenating these degraded specimens may be dependent upon the potential value the specimen has in terms of its historical significance, its fragility, and/or the importance of the associated data for addressing particular scientific questions (Neisskenwirth 2020).

Fluid preservation is a centuries old technique developed to permanently store specimens in liquids, such as ethanol (Simmons 2014). The practice of using ethanol as a fluid preservative is common for soft bodied arthropods such as arachnids (Simmons 2014; Marquina et al. 2021). A particular challenge that arises with ethanol collections is fluid evaporation and dehydration (Cushing and Slowik 2007; Neisskenwirth 2020). If a container experiences less than optimal preservation conditions, the ethanol may experience evaporation, leading to specimen desiccation. This may cause damage to the specimens’ morphology, may promote the growth of mold and microbes, and may degrade DNA (Singer 2014; Simmons 2014). The potential for desiccation to occur increases when containers are not sealed properly, especially if the specimens in such containers are not regularly monitored (Neisskenwirth 2020). If desiccated specimens are identified, collections staff may choose to rehydrate them, ideally selecting the most restorative method possible.

Much of the historical literature for rehydration practices is based on vertebrate taxa and prioritizes morphological integrity, but new approaches have been developed and specialized for various other taxa as well (Simmons 2014; Neisskenwirth 2020). Rehydration done on specimens does not reverse the damage caused by the initial desiccation and attempts to rehydrate should reduce the possibility of additional damage. Different approaches should then be considered for restoration as they will irreversibly impact the specimens. Several approaches have been developed, including submersion in wetting agents (Jocque 2008; Beccaloni 2012); alkaline solutions to help rehydrate specimens (Vogt 1991); and using a heated surfactant or heated rehydration fluid to accelerate rehydration times (Banks and Williams 1972; Waterhouse and Graner 2009). While heated rehydration is rapid, it may potentially damage already fragile material (Neisskenwirth 2020). Directly heating specimens in ethanol has been used in arachnid collections because it rehydrates specimens rapidly, but there is no published literature assessing this technique. While directly transferring spider specimens into higher concentrations of ethanol has not been observed to negatively impact the morphology of preserved spiders (Cushing and Slowik 2007), the technique has not been assessed for completely desiccated spiders. Heating may also cause damage to specimens, but this has not been assessed. Singer (2014) recommended softening tissues of desiccated specimens by putting them in a humid environment, instead of subjecting specimens to potentially damaging approaches such as heat-accelerated rehydration (Neisskenwirth 2020). More recently developed water vapor rehydration methods may reduce damage to fragile specimens, as they reintroduce fluids into tissues more slowly. If rehydration is pursued, one must consider that it is an irreversible process that alters the specimen and may degrade DNA (Neumann et al. 2022). Rehydrated specimens can provide invaluable morphological data, but their potential to provide genetic data needs to be tested.

In the last 20 years, ethanol-preserved specimens housed in collections have been increasingly sought after for genetic information because of the rise of new DNA extraction and sequencing techniques (McCormack et al. 2017). Specimens housed in collections can provide key information to the biologists who study them, especially as access to newer technology, methodologies, and information progresses (Short et al. 2018; Webster 2019; Derkarabetian et al. 2019). Storing invertebrates in ethanol at concentrations of 70% or higher is frequently used to preserve specimens’ DNA, and the genetic data may then be used in various studies (Szinwelski et al. 2012). However, a tradeoff exists between genetic and morphological preservation. Higher ethanol concentrations (95–100%) are better for DNA preservation (Nagy 2010) but cause specimens to become brittle as the water content in tissues further decreases. At lower ethanol concentrations, DNA will naturally degrade over time, but the specimen’s morphology will retain its flexibility (Baird et al. 2011). Even for museum specimens stored at lower concentrations, protocols have been specifically developed for the recovery of degraded DNA (Tin et al. 2014). This has allowed for the capture of genomic and sub-genomic data using, for example, Ultra-Conserved Elements (UCEs) and double digest Restriction-site Associated DNA (ddRAD) (Faircloth et al. 2012), for various studies focused on phylogenetics, systematics, and population genetics (Miller et al. 2013; Burrell et al. 2015; Starrett et al. 2017; Derkarabetian et al. 2019; Garcia et al. 2024).

Desiccated specimens are often ignored when searching for specimens to sample, since the assumption is that the DNA may be too degraded for capturing. While ethanol adequately preserves DNA for long periods (Anchordoquy and Molina 2007), when an ethanol preserved specimen is dehydrated, DNA degradation will accelerate, and the tissues of specimens may desiccate which can introduce risk for physical damage. These dehydrated specimens could have important data associated with them and provide both morphological and genetic data to researchers, but the effects of different rehydration treatments have not been investigated.

This project seeks to compare the restoration of desiccated spider specimens housed in the arachnology collection at the Denver Museum of Nature and Science (DMNS) between a rapid heated rehydration (Heat-Rehydration) treatment and water vapor (Step-Up) rehydration treatment to evaluate the impact on their morphology, the potential for DNA capture, and quality of the captured DNA. We assessed the potential for DNA capture, as well as damage to the tissues of the abdomen and genitalia. The treatment spiders were compared to desiccated spiders (negative control group) as well as a positive control group that had not experienced dehydration. For the abdomen, we looked for separation of the tissues from the exoskeleton, as this has been observed in heated ethanol rehydration. For the genitalia, we looked for signs of damage such as fracturing and breaks, as the genitalia in spiders is often crucial for taxonomic identification. If the structures are damaged, this may reduce the ability to positively identify the spiders. The present study seeks to propose recommendations for rehydrating fluid preserved arthropod specimens in order to maximize their usefulness for both morphological and molecular analyses.

We found a significant relationship between rehydration method and tissue separation (Table 1; df = 9, Chi Square = 63.91, p < 0.05). All of the experimental specimens and dehydrated controls showed varying levels of tissues separation from their exoskeleton (Figs 1, 2, 3). When we assessed the dehydrated control group, a majority of the samples displayed severe tissue separation. For the Heat-Rehydration treatment, two spiders fractured into multiple pieces, one of which could not be sampled further for DNA (Fig. 3). A majority of the samples in that treatment showed moderate tissue separation. In the Step-Up treatment, only one of the spiders had broken into pieces, and most displayed minimal tissue separation. In comparison, none of our positive controls showed tissue pulling.

Figure 1. 

Dorsal view of a spider specimen (Voucher 20) rehydrated using the Step-Up method. Tissue separation was assessed as a 1 (mild to moderate), and pedipalps were intact. Scale bar: 2.5 mm.

Figure 2. 

Dorsal view of a spider (voucher 13) rehydrated using the Heat-Rehydration method. Tissue separation was assessed as a 2 (moderate to severe), and pedipalps were intact. Scale bar: 2.5 mm.

Figure 3. 

Dorsal view of a spider (voucher 2) that was rehydrated using the Heat-Rehydration method and destroyed in the process. The level of tissue separation was assessed as a 3 (severe tissue separation) and the pedipalps could not be examined. Scale bar: 5 mm.

When assessing damage to pedipalps, we found that all eleven specimens of the positive control had intact pedipalps as well as all eleven of the dehydrated control; the Step-Up process damaged the pedipalps of one (out of eleven) males; the Heat-Rehydration process damaged the pedipalps of four (out of eleven) males (df = 3; Chi-Square = 7.16, p = 0.07). Thus, neither rehydration process significantly damaged the genitalia.

After the DNA extracts were quantified, we found almost all of the specimens had a low DNA content (Table 2). In the Step-Up, Dehydrated (negative control), and Hydrated groups (positive control), we found an outlier that had a much greater DNA content than the rest of the samples and was removed. We performed a one-way ANOVA to explain differences in variation in DNA content by treatment choice. The ANOVA showed a significant difference for the given data. We then performed a follow-up Tukey HSD test (p < 0.05) (Table 3, Fig. 4). The results of the Tukey HSD test show that most of the group comparisons do not show a significant difference with the exception of Step-Up and Hydrated (positive control) (P < 0.05) (Table 4).

Figure 4. 

Boxplot showing differences for DNA content in the legs of spiders sampled by treatment. Final eluted DNA had concentrations estimated using the Qubit fluorometer. Boxes represent 25–75% quartiles bisected by the median with outliers represented in circles. SU = Step-Up rehydration technique; HE = Heated-Rehydration technique; Dehydrated = negative control; Hydrated = positive control.

To determine if the quality of DNA was different between treatments and controls, we used an Ethidium Bromide gel run and found only one of our groups had visible smears indicating some level of DNA. The positive control, negative control, Heated-Rehydration, and Step-Up samples were run and only the positive control had a bright visible smear; no visible DNA smear was evident in the negative control or either of the experimental rehydrated samples (Fig. 5). Regardless of treatment choice, all desiccated spiders had heavily degraded DNA.

Figure 5. 

DNA integrity of control and rehydrated spiders. The DNA for the samples was loaded onto an Ethidium Bromide (EtBr) 1.5% gel plate and run for 20 minutes.

After assessing damage to the specimens, we found that the desiccated specimens displayed more tissue separation than our positive controls (Table 1). Our positive controls displayed no tissue separation, and the specimens were overall in good condition. The dehydrated spiders displayed moderate to severe separation of the tissues, and several were beginning to break apart while handling. As the tissue separation in specimens became more severe, we noticed that specimens were more fragile and susceptible to breaking during rehydration. All of the desiccated spiders had tissues that separated from the exoskeletons, but the Step-Up rehydration method was more consistent in rehydrating the tissues and reducing separation when compared to the Heat-Rehydration method (Table 1). The pedipalps of spiders in both control groups did not display damage. But when we looked at the treatment spiders, we found occasional breaks and fracturing that had occurred after rehydrating (Fig. 3). However, the damage incurred was not statistically significant and did not differ much between the two rehydration treatments. When we looked at the spiders that did have breakage, they were generally smaller in size. Size and fragility of the specimen may influence the amount of damage that occurs during a rehydration treatment.

The Step-Up rehydration process was developed to reduce morphological damage to specimens (Neisskenwirth 2020) and is longer and more logistically demanding. This approach has been successful for various vertebrates but needed testing for arthropods, such as arachnids. For our samples, this approach was more successful in rehydrating the tissues and did reduce tissue separation. The Heat-Rehydration method took between 1 to 3 minutes to perform compared to the 4 weeks required for the Step-Up method, but we observed more separation and a bit more damage to the pedipalps. This treatment choice also comes with the risk of destroying fragile specimens. Two of the heat accelerated specimens were separated into smaller pieces, making them unidentifiable after treatment. We had to remove one of those two specimens from DNA sampling, as we could not properly sample tissues for extraction. Overall, if the adult specimen is intact and the genitalia can be evaluated, the spider specimen is of value for morphological studies. If rehydration of the specimens is desired, then collections staff must consider the fragility of the desiccated specimen before proceeding. If damage to the material is a concern, we suggest that the curation or collections team considers the potential historical or research value the specimens may have when deciding whether to rehydrate specimens, and what technique to use. This may include who the specimens were collected by, when and where they were collected, or if they were designated as type series for a species. The Step-Up method may not rehydrate all desiccated material depending on their initial condition, but it is gentler than our Heat-Rehydration method and more adequately rehydrates tissues in fragile specimens.

In this study, we performed DNA extraction and isolation using a commercially available extraction kit that is accessible and affordable for labs. The DNEasy Blood and Tissue Kit (Qiagen) captured DNA but may not have been the most effective approach for such degraded material. However, using specialized protocols is costlier and requires additional considerations for safety due to the health hazards of several reagents (Tin et al. 2014). Our sampled materials had a low amount of DNA captured for every treatment, but we were able to successfully capture DNA in our positive control group for every specimen. For both rehydration treatments and the dehydrated control, we were unable to quantify DNA from the extracts for several specimens (Table 2). While we were able to extract DNA from each group, this does not fully address if the quality of the DNA is usable for sequencing technologies and downstream analysis. If the DNA quantity is of concern for ethanol preserved materials in collections, then museum personnel must be careful to ensure that specimens do not dehydrate in the first place.

When we ran the DNA on gels, we found that all desiccated or rehydrated specimens were too degraded (i.e. the DNA too fractured) to show up on our gels (Fig. 5). The positive controls did, however, have a smear present on the plate (Fig. 5). The DNA capture potential for these specimens exists, but the quality of these specimens’ DNA may not provide much success if attempting to sequence more than a few DNA loci. McCormack et al. (2015) were able to successfully capture UCEs from museum specimens, but there were limitations to consider for older, lower quality specimens. For this scenario, where our spiders had been desiccated for up to ten years, the DNA may have degraded to such a degree that any DNA detected in a fluorometer may be too fragmented even for Next Generation Sequencing.

DNA will begin degrading once the specimen dies, but high concentrations of ethanol will slow the degradation process (Guo et al. 2018; Oosting et al. 2020). Over time multiple processes including oxidative agents and enzymatic action can induce damage to preserved DNA (Oosting et al. 2020). Once the spider specimens were dehydrated, those processes were accelerated. Seeing no difference in DNA capture between most of the rehydration treatments except for one comparison (Tables 3, 4, Fig. 4), our assumption is that, while we may be capturing similar amounts of DNA from the sampled tissues, the capture is too low and the quality too degraded. The quality of the DNA from both rehydration treatments was almost comparable to those that remained desiccated (Fig. 5). Thus, neither treatment greatly reduced our ability to capture DNA compared to the dehydrated controls, and we did not observe a decrease in quality (Fig. 5), though our study shows that recovering DNA from dehydrated arachnids in general remains a challenge.

While higher temperatures can denature or damage DNA, in this study, heat may not have been a significant factor in further damaging the DNA in the tissues of the samples. It is difficult, if not impossible, to isolate every potential confounding variable impacting the degradation of DNA in dehydrated museum specimens. Neither rehydration treatment will improve yield or reduce degradation of the DNA if the intention is to restore them for genetics-focused work. These specimens can still provide valuable morphological data, and so a collections team should assess the integrity of the specimens and determine if a rapid approach, such as our Heat-Rehydration method, should be used at the potential cost of more fragile material, or if a slower Step-Up approach should be used. The Step-Up rehydration is labor and time intensive and may not successfully rehydrate all desiccated arthropod samples, but this approach is less damaging to fragile specimens. The heat accelerated rehydration is not labor intensive and is faster to conduct, but it poses more risk for damaging fragile specimens. Rehydrating museum specimens is an irreversible process and multiple methods should be considered before proceeding (Neumann et al. 2022). The goal for any rehydration effort should be to reduce morphological damage and reduce the degradation of DNA imposed on specimens by rehydration. Ultimately, specimens can never be fully restored after experiencing desiccation; therefore, resources should be devoted to keeping specimens hydrated so that desiccation and its impacts on specimens do not become a concern.

This project was supported by funding from the National Science Foundation grant DEB-1754587 awarded to Dr. Paula E. Cushing. It was part of the master’s thesis work of the first author. Dr. Bridget Chalifour and Tiffany Nuessle from the DMNS Genetics Lab contributed to the design of the methods for gel plating small inputs of degraded DNA.