How to store a beetle larva? Comparing temporal effects of common fluid preservation methods on color, shape, and DNA quality

Abstract

Proper fixing and long-term preservation of entomological evidence are essential in collections and research and crucial in applied fields such as forensic entomology. Incorrectly stored samples may lose important morphological features over time, rendering molecular analyses exceedingly difficult. The most effective method for preserving soft samples such as larvae is fluid preservation. It uses a combination of a wide range of fixatives and storage fluids. However, very little comparative work has been done to determine the effects of long-term storage on sample quality in terms of color, shape, and DNA stability. Moreover, the current golden standard in forensic entomology has been tailored for age estimation of larvae of Diptera, which differ from larvae of Coleoptera in morphology and subsequently in applied methods. We compared the effects of combinations of 6 commonly used fixatives and 6 commonly used storage fluids on midsized larvae of the forensically important beetle, Necrodes littoralis (Linnaeus, 1758), in terms of color, shape, and suitability for DNA analyses over a 2-yr period. We were looking for combinations that can preserve specimens in a satisfactory state, can be used on a regular basis, do not require advanced protection or skills of the personnel, and are not toxic or too harmful to the environment. We found not only several methods that scored significantly better in the tested parameters compared with the golden standard but also several common methods that should be avoided. The effects of agents on each tested category are discussed in detail.

Introduction

Forensic entomology utilizes immature and adult stages of necrophagous arthropods found on human remains to determine the time since the first colonization on the carcass, defined as the minimum postmortem interval (PMImin) (Amendt et al. 2007, Byrd and Tomberlin 2020). Historically, the flies (order Diptera) have been the most commonly used insect group for PMImin estimations as they are the first colonizers arriving on a body within minutes after death. They are later followed by a succession of other insect groups preferring different stages of decomposition (Byrd and Tomberlin 2020). In recent years, beetles (order Coleoptera) have been given increasing attention due to their later arrival times after the files have already departed and longer development on the corpse, which makes them indispensable for PMImin estimations in case a body is found in later stages of decay (Midgley and Villet 2009, Ridgeway et al. 2014).

The most important insect evidence on the crime scene is represented by the developmental stages of various necrophagous species found on the body or in its vicinity (Byrd and Tomberlin 2020). As a regular procedure, some part of the evidence should always be stored and archived in case it needs to be revisited or reviewed (Amendt et al. 2007). Depending on the amount and the state of the evidence, the rest can be used for rearing for age determination or can be discarded if redundant or damaged based on internal regulations of each institution (H. Šuláková, personal communication, head forensic entomologist of the Institute of Criminalistics, Prague). Archived samples should ideally be kept under such conditions that will allow morphological and molecular analysis for the duration of the statute of limitations period, i.e., at least for up to several decades (e.g., up to 20 yr according to the Czech filing and shredding regulations, H. Šuláková, personal communication).

Larvae and pupae generally represent the soft-bodied stages in the development of insects. They either entirely lack the hardened plates that form the exoskeleton in adults or possess them in smaller amounts (Gillot 2005). The classic method of dry preservation and pinning the specimen thus cannot be used, as their bodies would collapse and distort, damaging the morphological features needed for their determination. Although there are other methods like freezing or supercritical drying, the most practical method for preserving immature stages of insects is still considered to be fluid preservation (Simmons 2014). It does not require specialized machinery, and if sealed well, the samples can simply be stored for years without additional energy requirements (i.e., running a freezer).

Modern-day literature offers a wide range of recommended chemicals to use in fluid preservation that vary depending on the author (Riley 1892, Cox et al. 2006, Eymann et al. 2010, Mourek and Lišková 2010, Simmons 2014, Hammond et al. 2019). The contemporary process of fluid preservation of insects usually consists of 2 steps. First, the specimen is submerged in a fixative (Eymann et al. 2010, Mourek and Lišková 2010, Simmons 2014). Its purpose is to kill it as fast as possible, without unnecessary suffering and struggle during which the individual could damage its morphological features. The other purpose of the fixative is to quickly penetrate the body, arrest the decomposition processes that start almost immediately after death, and prepare the tissue for the second step (Eymann et al. 2010, Mourek and Lišková 2010, Simmons 2014, Hammond et al. 2019). The specimen is then moved into a storage solution that should secure its stability for the longest time period possible (Simmons 2014, Hammond et al. 2019).

The fluids can consist of specific chemicals in various concentrations or are mixtures that sometimes bear the name of their inventor or manufacturer (i.e., Pampel, Carnoy’s, Kahle’s, Hood’s, Bouin’s, San Veino, etc.). Here, we mention only the most common ones (Table 1). Finding a suitable chemical or mix of chemicals is rather a difficult task, as comparative works are often missing, and various authors present their favorite recipes often based on their personal preferences and experience. Moreover, some commonly used practices are based solely on “oral traditions” but otherwise unfounded. For example, putting a few drops of glycerin in ethanol preservatives has been believed to form a protective film on the specimen’s surface, which should slow down the evaporation of the alcohol (Simmons 2014) or increase the viscosity of the solution, so if the stopper leaks, it will not dry completely, but a denser solution with a smaller volume will remain at the bottom of the bottle (Jan Růžička, personal communication). However, such practices have never been appropriately tested and may have only minimal effects (Simmons 2014).

In forensic entomological practice, preservation methods differ depending on the body types and sizes of the insects collected, which require different approaches (Schauff 2001, Amendt et al. 2007, Eymann et al. 2010). The bodies of fly larvae are very soft, almost completely lacking sclerotization, and are devoid of many morphological features, and their age is thus determined by changes in the length of their body (Adams and Hall 2003, Bugelli et al. 2016). On the other hand, beetle larvae are usually at least partly sclerotized, can be larger, and their age can be traditionally estimated only to specific intervals defined as instars (Novák et al. 2017, Díaz et al. 2018, Jakubec et al. 2018). The specific instars are characterized by morphological (including color) or morphometrical features (of rigid structures) because the total body length in beetle larvae can dramatically change based on various factors, e.g., the amount of food ingested (often observed by the authors). A strong agent well preserving the morphological characteristics of a large beetle larva can thus distort fly larval length, and vice versa, a more “gentle” agent suitable for soft-bodied fly larvae can be insufficient for beetle larva and can cause its degradation.

Furthermore, some beetle species can be so similar that the only differences are represented by different pigmentation (Díaz et al. 2018, Jakubec et al. 2018). For example, Thanatophilus rugosus (Linnaeus, 1758), T. sinuatus (Fabricius, 1775), and Necrodes littoralis (Linnaeus, 1758) (Coleoptera: Staphylinidae: Silphinae) larvae look alike in terms of bodily form but are different in terms of coloration, especially on their dorsal plates and legs (Byzova 1964, Díaz et al. 2018, Jakubec et al. 2018). They, however, differ in habitat preferences and times of activity (Růžička and Jakubec 2016). Even different instars within a single species can look alike. The second- and third-instar larvae of T. sinuatus may be similar in size but are best differentiated by the color patterns on their thorax, abdomen, and legs (Jakubec et al. 2018). However, incorrectly stored larvae tend to completely darken, losing these important features. A misidentified species can lead to under- or overestimation of PMImin due to the possible differences in developmental lengths and/or make it difficult to determine whether a body was moved between habitats due to possible differences in ecology. A lower (younger) instar mistaken for a higher (older) instar causes overestimation of PMImin and vice versa higher instar mistaken for a lower instar causes its underestimation. Preserving color in the long-term storage of beetle larvae is thus crucial to avoid incorrect conclusions or make revisiting old evidence impossible.

While the effect on shape is very important in preserving fly larvae (due to determination by total length; Adams and Hall 2003, Amendt et al. 2007, Day and Wallman 2008, Rosilawati et al. 2014), it does have an auxiliary role rather than a crucial role in beetle larval determination. However, it is generally convenient to have specimens preserved in a straightened position, as it enables full access to all potential morphological features, often placed on the venter (Novák et al. 2017, Jakubec et al. 2018). Larvae in certain preservatives tend to curve ventrally, obstructing access to this part of the body (Midgley and Villet 2009). Any extra manipulation, especially in stiffened specimens or even the process of preservation itself, can lead to breakage or destruction of fine extremities, for example, urogomphi, which are often the only feature for instar determination in many Silphini species (Novák et al. 2017, 2023).

With the development of techniques enabling the identification of insect species based on molecular methods, special consideration should also be given to the effect of a fluid preservative on the DNA quality of a sample (Amendt et al. 2007, Byrd and Tomberlin 2020). In species identification through barcoding, the mitochondrial COI plays an important role (e.g., Hebert et al. 2003, Serjeant and Beebee 2013), and correct preservation and storage of material for molecular analysis are thus crucial. The destructive effects of formaldehyde, chloroform, and methanol on DNA are known (Fukatsu 1999, Srinivasan et al. 2002, Day and Wallman 2008) and immediately disqualify these chemicals from use if the collected evidence cannot be determined morphologically, and genetic barcoding is needed.

The current recommended general preservation practice in entomology is to fix the specimen in heated water over 80 °C, but not boiling, between 30 s (Hot Water Kill, HWK) and 5 min (Schauff 2001, Haglund and Sorg 2002). The former is recommended specifically in forensic entomology as the “golden standard” tailored specifically to fully extend the body of Dipteran larvae for age estimation (Amendt et al. 2007, Gennard 2014, Byrd and Tomberlin 2020). The fixative effect is reached by the denaturation of proteins at high temperatures rather than by a chemical reaction. The sample should then be stored in 70–95% ethanol depending on the author. However, in the field, it is not always manageable nor practical to have heated water at hand, and since putting specimens directly into alcohol will cause their rapid decay, finding a corresponding easy-to-use substitute represents a crucial problem in forensic practice.

Most forensic entomology-related studies have researched the effects of chemicals on fly larvae only due to their historically given importance (Haglund and Sorg 2002, Adams and Hall 2003, Day and Wallman 2007, Rosilawati et al. 2014, etc.). In this article, we compared 36 different combinations of 6 fixative and 6 storage fluids on mature third (last) instar larvae of a common forensically important (Matuszewski et al. 2010, Charabidze et al. 2015) necrophagous beetle, N. littoralis. The species is predominantly reported to be found on cadavers in later stages of decomposition in outdoor locations, especially forests and rural areas from May to November, with the most cases from July to September (Charabidze et al. 2015). Our goal was to find the ideal combination of fixative and storage fluids that would be nontoxic for the technical personnel, practical, and easy to use (i.e., no other equipment would be needed other than the fluid itself) and would preserve the specimens in a good state for various potential morphological and molecular analyses for an extended period of time. Furthermore, we aimed to find a method that preserves the material in such a manner that it will allow fast, simple, and cheap DNA analyses without the need for additional optimization in order to minimize costs and thus can be utilized across the world.

Materials and Methods

The specimens used in this experiment comprised of third-instar larvae of N. littoralis (Fig. 1a), obtained from breeding colonies in the laboratories of the Czech University of Life Sciences Prague and specimens freshly collected from a wild boar carcass on the outskirts of Prague in summer 2019 and reared to the third instar the same year in the same laboratory. We used sonly third-instar larvae, so the effects of the preservation methods are analyzed on the larval stage with the largest body mass, which should be the hardest to preserve. The duration of the experiment was approximately 24 months, beginning in the first half of September 2019 and ending in the second half of August 2021.

We analyzed the effects of the unique combinations of 6 fixative and 6 storage fluids (Table 2), which amounts to 36 different combinations of treatments in a randomized factorial design. Temperature in treatments using heat was not kept stable throughout the whole immersion time but was allowed to cool down after the insertion of the specimens. Ten specimens were designated for each treatment, of which 8 specimens were selected for visual and morphological evaluation and 2 for molecular analysis. We used only 2 specimens for molecular analysis because only a small part of their body was needed for each DNA extraction. Overall, 360 larval specimens were used in morphological and molecular analyses.

The living larvae were divided into 6 groups of 60 specimens (based on the different fixative solutions), inserted into containers with the respective fixatives, and left inside for 6 h (Fig. 1b). Six hours were chosen arbitrarily by the authors and should represent the extended time taken between collecting the specimen in the field (e.g., a crime scene) and the transport into the laboratory for further processing and transfer into a storage fluid. The only exception was one of the HWK in distilled water, where the specimens were put into the fixative for 5 min only. This represents one of the generally recommended techniques in entomology (Schauff 2001, Hammond et al. 2019) and the current gold standard in forensic practice (Amendt et al. 2007). We slightly modified it for larvae of larger sizes by prolonging the time in HWK to 5 min and selecting storage fluid to be 70% EtOH. This treatment combination was used as the baseline to which all other combinations were compared. Subsequently, the specimens were transferred into 6 types of storage fluids for long-term storage (Table 2; Fig. 1b). The larvae were stored individually in polypropylene tubes (8 ml) with a screw cap with EPDM isolation O-ring and properly labeled. The tubes with samples were stored in closed carton boxes at room temperature for the duration of the experiment.

The visual state of the stored larvae was documented by photographing them individually each month (starting the first month after storage). Possible changes in specimen coloration, a telltale sign of decay, were analyzed based on measuring RGB (Red Green Blue) values from the images in graphical software following Novák et al. (2020). We measured the luminosity of the darkest part and lightest part, and from these values, we calculated the difference in luminosity (see Fig. 2), which is the crucial metric for spotting the variability in color patterns.

To find differences between the treatments, we created a generalized additive model (GAM) with asbsolute contrast between the lightest and darkest part of the larval body using RGB luminosity as a response variable. The model was chosen because the contrast luminosity values did not behave linearly when plotted over time. REML (restricted maximum likelihood) smoothing was applied to 2 factors; the first being the fixed effect factor of time for different treatments, which consisted of a combination of 6 methods for fixation and 6 methods for storage (36 in total). The second was the random factor of larval identity, as each sample was measured multiple times during the experiment. We also included a varying intercept based on the treatment factor, creating smooth-factor interactions as we presumed each treatment affects RGB’s values differently from the start. The fit of the model quality was evaluated based on convergence and diagnostic plots created by the function “gam.check” (package mgvc) to ensure that the residuals of the model do not violate assumptions of normality and homoskedasticity. The significance level was set at 5%. The data management and analyses were conducted using “R v4.1.1.” with packages “tidyr” and “mgcv,” respectively. The packages “gratia” and “ggplot2” were used to visualize the results.

The shape of the larvae was evaluated from the photographs taken using Olympus cellSens Entry software. We measured the specimen’s shape from the lateral side, defined as an angle between the anterior edge of the pronotum and the base of the urogomphi, with its vertex in the middle of the tergite on the abdominal segment III (Fig. 1a). Average values and standard deviations were calculated for each batch of specimens (8 samples) per treatment. We were looking for treatments that had minimal effect on the curvature of the specimens, i.e., with values around 180° or at least making an obtuse angle.

To estimate the effects of the treatments on the preservation of the genetic material, DNA was extracted in the first month after the storage and then every 2 months (7 events of DNA isolations in total), using Tissue/Blood DNA Mini Kit (Geneaid, New Taipei City, Taiwan). The genetic material was extracted by leg clippings from the specimens designated solely for this purpose (2 individuals per treatment combination) from all treatment combinations. The isolation was followed by amplification of cytochrome oxidase I (COI). The PCR reactions were carried out at 25 µl based on the provided PPP Master Mix protocol (Top Bio, s.r.o., Czech Republic) under the following conditions: initial denaturation at 95 °C for 5 min; 40 cycles of denaturation at 95 °C for 30 s; annealing at 50 °C for 40 s; extension at 72 °C for 2 min; and final extension at 72 °C for 8 min; using previously published specific primers for Coleoptera C1-J-2183 (alias Jerry) and TL2-N-3014 (alias Pat) (Simon et al. 1994). The PCR products were visually checked using electrophoreses (90 V, 25 min, 1% agarose gel). The succession rate of amplification was evaluated based on the presence or absence of the amplicon. If the amplicon was present, the sample was treated with the ExoSAP-IT (Applied Biosystems, USA) according to the protocol and sequenced in the BIOCEV laboratory (Vestec, Czech Republic) using one-sided Sanger sequencing. The sequences obtained were then trimmed and compared against the NCBI (https://www.ncbi.nlm.nih.gov/) database using BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi). The database contains 51 hits for (“Necrodes littoralis” and “COI”) or (“Necrodes littoralis” and “COX1”) of individuals collected in various localities across the area of distribution (Greece, Germany, Finland, South Kora, Japan, China, and unknown).

Results

In total, 288 of 360 larvae were visually analyzed for changes in color and effects on shape. The GAM model confirmed that the luminosity had a significantly nonlinear response over time in all treatments (Fig. 2). We found the effect on preserving color quality in combinations of 80 °C HWK 6 h and acetone, and KAA and 80% EtOH significantly better than the control treatment (80 °C HWK 5 min and 70% EtOH). Significantly worse results were obtained in all treatments using cold Abs. EtOH as a fixative; boiling Abs. EtOH in combination with 70% EtOH, 80% EtOH, and AGA; acetic alcohol when both fixative and storage fluids or in combination with Acetone or AGA; and 80 °C HWK 6 h in combination with 70% EtOH.

The simplified additive model was able to quantify the differences in contrast values produced by individual fixative and storage fluids (Table 3; Figs. 3 and 4). Considering the fixative only (Table 3), we did not find any significantly better treatment than the control (80 °C HWK 5 min) Fixing specimens in 80 °C HWK 6 h proved to be significantly worse than the control as well as all the other fixatives. From the storage fluids only (Table 3), significantly better results than the control (70% EtOH) were achieved using (in descending order) absolute EtOH, acetic alcohol, acetone, and 80% EtOH. The control scored the worst of all the storage fluids, together with AGA, which did not score significantly better or worse (Table 3). Furthermore, we noticed that the legs of the larvae stored in acetic alcohol obtained a slight greenish-blue hue.

The effects of fixatives, storage fluids, and their combinations on the shape of the specimens in terms of curving expressed in angular size are shown in Table 4 (per individual combinations) and Table 5 (per fixative or storage fluid only). The angle values averaged per storage fluid do not seem to differ greatly from each other. However, substantial differences can be seen among the averaged values based on the fixative type and even more among specific treatments. Fixing specimens in cold alcohol or KAA had the smallest effect on the curving of the larvae. The largest distortion was caused by fixing the specimens in boiling alcohol, followed by 80 °C HWK for 5 min.

DNA was extracted and amplified from specimens from all 36 treatment combinations. All the generated sequences of COI were identically trimmed using Chromas (Version 2.6.6; Trim low quality). Sequences longer than 500 bp (553–794 bp) were obtained from 20 treatments after 24 months, all of which were identified as N. littoralis (Table 6). Six treatments produced only short sequences (105–317 bp), although the extraction was sufficient for species identification after 24 months. Eight treatments failed to generate sufficient barcode sequences for species identification: 80 °C HWK for 6 h and acetic alcohol (the last successful extraction was after 6 months), followed by acetic alcohol and AGA; cold Abs. EtOH and AGA; and cold Abs. EtOH and acetic alcohol (for all the last successful extraction was after 8 months; 754–803 bp). The last successful extraction from the acetic alcohol-only treatment was after 20 months, while in 80 °C HWK for 5 min and acetic alcohol; 80 °C HWK for 5 min and AGA; and KAA and acetic alcohol treatments, the last successful extraction was 22 months after their storage.s

Discussion

Effect on Color

Our results show that the color quality of beetle larval specimens (here represented by the difference in contrast between naturally dark and light parts of the larva) is primarily influenced by the fixative used (Table 7; Fig. 5). However, a correct combination with storage fluid is crucial to achieving an ideal outcome. The use of fixatives should never be omitted or underestimated in specimen preservation as it can significantly prolong their longevity in a satisfactory state.

We found 2 significantly better methods than the gold standard utilized in forensic practice: 80 °C HWK 6 hours (fixative) and acetone (storage fluid), and KAA (fixative) and 80% EtOH (storage fluid). The positive effects of acetone on color preservation could be explained by its stronger dehydrating properties compared with ethanol. It penetrates insect tissues more rapidly by faster removal of water content (Moreira et al. 2013), thus stopping the decomposition processes and consequent darkening of the specimens (Fig. 5). Also, KAA as a fixative yielded better results in preserving color than our control. It did so in 2 treatments; in combination with 80% EtOH and in combination with absolute EtOH, albeit the p-value, in this case, did not reach a standard minimum level of significancy (p = 0.051). Moreover, from fixatives alone, KAA was the only one that did not score significantly worse than the control. In fact, the mean estimated effect on contrast value was positive compared to the gold standard, albeit on a lower significance level (p = 0.098), which indicates it is comparable to it in terms of color quality preservation (Table 3; Fig. 5). These superior fixing properties of KAA may come from the combination of its components. Kerosene is a nonpolar solvent, which means it can facilitate penetration of the remaining polar agents into fat tissues and thus improve the preservation efficacy. Glacial acetic acid helps the penetration of ethanol into the tissues (Rosilawati et al. 2014) and swells them, acting as a counter to the shrinkage effects (Hammond et al. 2019) caused mainly by ethanol (Simmons 1999). The combination of these agents thus seems to make KAA a powerful fixative where the individual chemicals act in synergy. In fact, none of the combinations employing KAA scored significantly worse than the control (Table 7). The only treatment with a negative estimate value was in combination with Acetone as a storage fluid. Perhaps the strong dehydrating properties of acetone overpowered the swelling effect of the glacial acetic acid in the fixative.

The method with the most negative impact on the color quality appears to be fixing in cold ethanol and storing in low-concentration ethanol (Table 7; Fig. 5), which is, unfortunately, a very common entomological practice used for its simplicity. However, the specimens treated in this manner quickly darken, which is a telltale sign of decay processes taking place. Furthermore, it can hinder morphological identification in cases where species differ only in pigmentation of specific small areas on the body. This includes different instars of the same species as well, as their sizes may sometimes overlap (Jakubec et al. 2018). Storing or collecting samples directly in alcohol in the form of various liquors is well known in history (Simmons 2014). Still, due to their low ethanol concentrations (mostly way below 70%), the preservation qualities of these fluids are highly inferior, in addition to their inability to kill specimens quickly. It is thus recommended in extreme cases only when completely lacking any other equipment (Niederegger 2021). Our results show that even absolute EtOH used cold and on its own is, despite its high concentration, an inferior fixative when compared with other methods (Tables 3 and 7; Fig. 5). That includes methods using heat for fixing (even by just hot water) or even some cold methods such as KAA.

Effect on Shape

The shape of the specimens is determined by the fixative used, which is apparent when comparing average values (including standard deviation) per fixative only against values per storage fluid only (Table 5; Fig. 5). The specimens with the largest curvature can be found in treatments with boiling EtOH (avg = 45°) and 80 °C HWK 5 min (avg = 53°) as fixatives. The extreme contortion of certain larvae, in some cases reaching a circular shape, is probably a combined result of behavioral (curling up) and physiological (protein denaturation) reactions to the extreme heat. In fact, the smaller angle in specimens treated in boiling EtOH could be explained by an even stronger behavioral response to not only heat but also ethanol (in contrast to distilled water), which is a strong irritant on its own. The difference in average shape between 80 °C HWK 5 min and 80 °C HWK 6 h (avg = 72°) can be explained by the fact that in the latter case, the larvae were left in the medium for a prolonged period of time. Distilled water does not have any preserving qualities on its own, and the decay processes most likely started during the time, facilitating the “relaxing” of the specimens. The least curved larvae were found among specimens fixed in cold EtOH and KAA. In both cases, the spastic position could not be immediately set due to the lack of heat in the fixative, which likely resulted in more straight, “relaxed” samples.

Effect on Genetic Material

The obtained barcodes of cytochrome oxidases I were compared to the open database of NCBI (https://www.ncbi.nlm.nih.gov/). The database contains 51 accessions of N. littoralis and contains its closest related species and genera. None of the produced sequences of sufficient quality showed any indication of misidentification with these or any other taxa within the tribe Silphini. The positive identification of our samples is thus indicative of a well-preserved specimen and cannot be interpreted as a lack of other possible matches.

We found remarkable differences in the quality of the genetic material depending on the treatments used (Table 6). Contrary to the effect on the shape of the specimen, the DNA quality appears to be dependent predominantly on the storage fluid used. We did not find any treatments using AGA or acetic alcohol suitable for DNA preservation since they appear to degrade the DNA, and the obtained sequences were generally short (<216 bp). However, the fixative may also have a particular effect, as we found 5 of 6 treatments fixed in 80 °C HWK 5 min unsuitable, primarily due to the short sequences produced. This was not observed in 80 °C HWK 6 h, and it may be likely that, similar to the effect on the visual quality, the time period of 5 min HWK is too short to fix the specimen sufficiently, and the results could be improved by prolonging it. In fixatives using heat (80 °C HWK 5 min, 6 h, and boiling EtOH), we observed that the generated sequences were degraded, and only short fragments were of sufficient quality, which is in accordance with the fact that heat stress damages DNA (Kantidze et al. 2016), albeit in half of these treatments, species determination based on COI barcode was still successful. Regarding the storage fluids, on average, the shortest sequences were produced by 70% EtOH, in addition to the already mentioned acetic alcohol and AGA.

The genetic material was best preserved in samples fixed in absolute EtOH, KAA, and acetic alcohol, which were subsequently stored in either 80% EtOH, acetone, or absolute EtOH. These treatments produced sequences of standard COI fragments of sufficient length for barcode determination. Although acetic alcohol yielded good results when used as a fixative alone, it appeared to cause long-term DNA degradation as a storage fluid; hence, we do not recommend using it even as a fixative, especially when other more suitable alternatives are at hand. Preserving samples in absolute EtOH for genetic analysis and storing them in temperatures well below zero is already a well-established practice. Still, not much is known about other suitable fluid alternatives. Haskell and Williams (2008) successfully analyzed DNA from specimens preserved in KAA, although with a warning that the matter requires further studies. Fukatsu (1999) and Moreira et al. (2013) showed that the genetic material of various insect specimens could be preserved in acetone at room temperature for several years. Our work confirms the results of these previous studies and demonstrates that all 3 fluids are suitable for DNA preservation even when stored at room temperature.

Effectiveness of Alternative Procedures

Modified versions of 2 solutions were used in this study. First, we opted for using KAA (Byrd and Castner 2001) instead of KAAD (Schauff 2001), which includes highly toxic dioxane as an extra agent in the mix. We proved that the effect of KAA on samples was more than satisfactory, thus making it a completely valid and safer alternative to KAAD for long-term storage of midsized beetle larvae. Second, we used absolute EtOH instead of 75% EtOH in acetic alcohol. Our motive was to ensure the proper conservation of genetic information, which we feared would be less likely with low-concentration ethanol. However, the mixture had anyway proven to be highly detrimental to DNA, which leads us to conclude that the cause for this was glacial acetic acid, and both versions would score equally bad. From a visual point of view, the results were average, and the extremities of the larvae seemed to gain an unnatural color. In this case, our modification did not seem to have any positive or negative effect.

Using HWK for fixing specimens is commonly recommended in entomology (Schauff 2001, Hammond et al. 2019) as well as in forensic practice (Haglund and Sorg 2002, Amendt et al. 2007). However, obtaining or keeping hot water at the right temperature in the field conditions of a crime scene or some remote biodiversity hot spot may be rather tricky. Our study shows that there is a suitable “cold” alternative to this fixative represented by KAA. It preserves the visual state of the specimens well (Tables 3 and 7; Fig. 5), does not distort their shape (Tables 4 and 5; Fig. 5), nor does it have a detrimental effect on the quality of the genetic material (Table 6).

Conclusion

We were looking for a method of preserving larger beetle larval specimens that would be universally effective for all categories of specimen quality that we analyzed in this work (color, shape, DNA quality), even though specific methods might have scored better in a single category. We prioritized the effects on the color and the obtainability of the DNA of the specimens over the impact on the shape. We believe that they represent better markers of the level of tissue decay and are crucial for correct sample identification. In conclusion, we recommend using KAA and absolute EtOH as the ideal method for fixing and long-term preservation of midsized beetle larval specimens as evidence in criminal investigations. We also recommend 80 °C HWK and absolute EtOH or 80 °C HWK and acetone as suitable alternatives. All 3 methods preserve specimens very well both in terms of visual and genetic quality. However, using KAA does not employ hot water, which makes its utilization exceedingly more practical in the field and does not damage specimens in terms of shape distortion or genetic sequence length. Moreover, we propose considering our findings for implementation in both forensic and general entomology practice regarding the preservation of Coleoptera larvae specifically. All of these methods enable storing specimens at room temperature for a minimum of 2 yr and likely even longer at lower temperatures. We highly suggest that follow-up research on beetle morphology and developmental biology in forensic science utilize the proposed methods to standardize the protocols and achieve comparability of results.

Acknowledgments

We thank the Molecular Laboratory of the Faculty of Environmental Sciences (CZU Prague) for providing research equipment for DNA analyses. We also thank Chris Harding (Prague) for the language corrections and to 2 anonymous reviewers for many valuable comments.

Author Contributions

Martin Novák (Conceptualization [Equal], Formal analysis [Equal], Investigation [Equal], Writing—original draft [Lead], Writing—review & editing [Equal]), Pavel Jakubec (Conceptualization [Equal], Funding acquisition [Lead], Investigation [Equal], Methodology [Lead], Supervision [Equal], Writing—review & editing [Equal]), Karolina Mahlerová (Formal analysis [Equal], Investigation [Equal], Methodology [Equal], Visualization [Equal], Writing—review & editing [Equal]), Santiago Montoya-Molina (Investigation [Equal], Writing—review & editing [Equal]), and Jarin Qubaiová (Investigation [Equal], Writing—review & editing [Equal])

Data Availability

All collected data are provided in the online repository https://github.com/jakubecp/Larval-storage-experiment.

References