Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

DNA from resin-embedded organisms: Past, present and future

  • David Peris ,

    Contributed equally to this work with: David Peris, Kathrin Janssen

    Roles Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Supervision, Validation, Writing – original draft, Writing – review & editing (DP); (KJ)

    Affiliation Section Paleontology, Institute of Geosciences, University of Bonn, Bonn, Germany

  • Kathrin Janssen ,

    Contributed equally to this work with: David Peris, Kathrin Janssen

    Roles Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Validation, Writing – original draft, Writing – review & editing (DP); (KJ)

    Affiliation Institute of Medical Microbiology, Immunology and Parasitology, Medical Faculty, University of Bonn, Bonn, Germany

  • H. Jonas Barthel,

    Roles Formal analysis, Investigation, Writing – review & editing

    Affiliation Section Paleontology, Institute of Geosciences, University of Bonn, Bonn, Germany

  • Gabriele Bierbaum,

    Roles Conceptualization, Funding acquisition, Supervision, Writing – review & editing

    Affiliation Institute of Medical Microbiology, Immunology and Parasitology, Medical Faculty, University of Bonn, Bonn, Germany

  • Xavier Delclòs,

    Roles Conceptualization, Data curation, Funding acquisition, Writing – review & editing

    Affiliation Department of Earth and Ocean Dynamics and Biodiversity Research Institute (IRBio), Faculty of Earth Sciences, Universitat de Barcelona, Barcelona, Spain

  • Enrique Peñalver,

    Roles Conceptualization, Data curation, Investigation, Writing – review & editing

    Affiliation Geological and Mining Institute of Spain (Geominero Museum), Valencia, Spain

  • Mónica M. Solórzano-Kraemer,

    Roles Data curation, Funding acquisition, Methodology, Writing – review & editing

    Affiliation Department of Palaeontology and Historical Geology, Senckenberg Research Institute, Frankfurt am Main, Germany

  • Bjarte H. Jordal,

    Roles Data curation, Funding acquisition, Validation, Writing – review & editing

    Affiliation Museum of Natural History, University Museum of Bergen, University of Bergen, Bergen, Norway

  • Jes Rust

    Roles Conceptualization, Funding acquisition, Methodology, Supervision, Writing – review & editing

    Affiliation Section Paleontology, Institute of Geosciences, University of Bonn, Bonn, Germany

DNA from resin-embedded organisms: Past, present and future

  • David Peris, 
  • Kathrin Janssen, 
  • H. Jonas Barthel, 
  • Gabriele Bierbaum, 
  • Xavier Delclòs, 
  • Enrique Peñalver, 
  • Mónica M. Solórzano-Kraemer, 
  • Bjarte H. Jordal, 
  • Jes Rust


Past claims have been made for fossil DNA recovery from various organisms (bacteria, plants, insects and mammals, including humans) dating back in time from thousands to several million years BP. However, many of these recoveries, especially those described from million-year-old amber (fossil resin), have faced criticism as being the result of modern environmental contamination and for lack of reproducibility. Using modern genomic techniques, DNA can be obtained with confidence from a variety of substrates (e.g. bones, teeth, gum, museum specimens and fossil insects) of different ages, albeit always less than one million years BP, and results can also be obtained from much older materials using palaeoproteomics. Nevertheless, new attempts to determine if ancient DNA (aDNA) is present in insects preserved in 40 000-year old sub-fossilised resin, the precursor of amber, have been unsuccessful or not well documented. Resin-embedded specimens are therefore regarded as unsuitable for genetic studies. However, we demonstrate here, for the first time, that although a labile molecule, DNA is still present in platypodine beetles (Coleoptera: Curculionidae) embedded in six-year-old and two-year-old resin pieces from Hymenaea verrucosa (Angiospermae: Fabaceae) collected in Madagascar. We describe an optimised method which meets all the requirements and precautions for aDNA experiments for our purpose: to explore the DNA preservation limits in resin. Our objective is far from starting an uncontrolled search for aDNA in amber as it was in the past, but to start resolving basic aspects from the DNA preservation in resin and search from the most modern samples to the ancient ones, step by step. We conclude that it is therefore possible to study genomics from resin-embedded organisms, although the time limits remain to be determined.


Deoxyribonucleic acid (DNA) contains the genetic information that allows all life forms to function, grow and reproduce; in addition, it is present in all cells and thus forms part of the tissues. Ideally, fossil—or ancient—DNA (aDNA) might serve as an alternative to morphological analysis and palaeoethology, which for centuries have served as the only tools available for scientists to determine the phylogenetic relationships between past organisms. Genomic sequences also provide insights into molecular evolutionary changes over time, clarify evolutionary relationships among taxa and yield information on mutualism rates [1, 2].

Amber is fossilised tree resin that may preserve deep time fauna in exceptionally clear morphological detail [35], and therefore appeared a promising material for preserving aDNA. It was hoped that DNA could easily be extracted from ancient soft tissue remains that were desiccated and macroscopically well preserved [6], because the complete and rapid engulfment of arthropods in resin and their hypothetically rapid fixation and dehydration—a type of mummification—promised to promote the preservation of DNA [7, 8]. Claims for DNA preserved in amber samples were rapidly made [912] in some cases with samples more than 125 million years old [13]. These studies overshadowed other less well-known studies on younger aDNA [14, 15]. However, amber claims are suspected of being the result of modern environmental DNA contamination because authentication procedures were not followed [1622].

Natural resins are secreted from parenchymal cells in plants and trees and comprise complex mixtures of terpenoid compounds, including acids, alcohols and saccharides, some of which have preservative and antimicrobial properties [5, 23]. It has been observed that modern resin and amber both exhibit extensive chemical variations [24], which may influence preservation processes and therefore the quality of embedded organisms [25, 26]. Furthermore, DNA is a particularly labile macromolecule. In living cells, specific repair mechanisms act on damaged DNA, but these processes cease after cell death and DNA is naturally degraded [27]. Besides active cleavage of DNA by nucleases shortly after cell death [28, 29], long-term DNA degradation by hydrolytic processes (initiating double-strand breaks) or oxidative dinucleotide modification and depurination (removal of the guanine and adenine bases from the sugar-phosphate backbone) highly affect the stability of DNA molecules [16, 3033]. Additionally, it seems that diagenetic events, including overburden pressure and heat generated through orogenesis over millions of years, affect amber permeability [34] and minimise the likelihood of DNA preservation in amber [21, 35].

The resin samples used in this study came from Hymenaea verrucosa Gaertner, 1791 [36] trees (Fabaceae), which produce copious amounts of resin that contains different types of sesquiterpene hydrocarbons and diterpenoid resin acids [23, 37]. Plants from this genus are the source plant of Miocene amber deposits in Mexico, the Dominican Republic, Peru and Ethiopia, and of sub-fossil resin deposits in different parts of the world [38]. Cenozoic amber from India and China was produced by some species of Angiospermae: Dipterocarpaceae [39, 40]. By contrast, the major Mesozoic amber deposits mainly derive from Coniferales [38]. The different plant group origin of the resin implies different sets of compounds in their chemical composition: monosaccharides, alcohols, aldehydes and esters [23]. The participation of esters in resin fossilisation (amberisation) and the biomolecules of the inclusions is still unknown [26]. Resin is a very complex (and not completely studied) preservation source with many variables simultaneously influencing preservation processes.

Distinguishing ancient DNA from recent contamination may be difficult while no strict correlation exists between DNA degradation and the age of the studied organism [41]. Although a comparison of modern and older mitochondrial DNA (mtDNA) showed significant differences in average fragment size, there is no direct correlation between age and fragmentation, and it is assumed that the main processes affecting DNA fragmentation occur rapidly after cell death [41, 42]. Heintzman et al. [43] found a time-dependent decrease in the concentration of amplifiable DNA in museum specimens. As mtDNA degrades more slowly than nuclear DNA (nuDNA), presumably due to additional protection of mtDNA by the double membranes of the mitochondrion, and is present in many more copies per cell, mtDNA may be more useful for aDNA studies [4345]. Many different factors influence the state of preservation [3, 4]; therefore, the conditions under which individual specimens have been preserved are of decisive importance [20].

Attempts in various laboratories to repeat aDNA extraction from amber or from younger resin that will become amber have not been successful [21, 46, 47], which has raised further doubts about claims that aDNA was isolated from various fossilised insects in amber. Penney et al. [21] used next generation sequencing to analyse two Colombian sub-fossil resin bees, dated as less than 60 years and around 10 600 years BP, respectively. The young age of these resins means that they were not subjected to any extreme processes. Nevertheless, no convincing evidence was obtained for the preservation of endogenous DNA in either of the two studied sub-fossil resin inclusions. The clear conclusion of this study was that DNA is not preserved in insect inclusions in sub-fossil resin and amber [21]. After publication of these (and previous) results, the hypothesis that preserved DNA could be extracted and studied from animal remains in amber, or in younger resin, was totally discarded. However, these studies did provide negative evidence, demonstrating that DNA was not preserved in these samples.

Some success with DNA amplification has been reported with pinned museum and permafrost-preserved invertebrates [43, 4851], and some of these samples were older than the oldest specimens analysed by Penney et al. [21]. These studies concluded that museum specimens could serve as a source of molecular information [4850]. Of all recent publications, the sole hypothetical success with aDNA extraction from sub-fossil specimens embedded in resin was reported by Büsse et al. [51], who claimed to have successfully amplified aDNA from 61 species of 46 higher arthropod taxa from ca. 200-year-old dried museum specimens, as well as from two beetles embedded in sub-fossil resin. The two samples of ancient resin (sensu [52]) were analysed via accelerator mass spectrometry (standard-AMS) radiocarbon and dated to 790–700 years BP and 4030–3900 years BP. However, the positive results reported by Büsse et al. [51] should be viewed with caution as the publication does not contain a detailed discussion of the authentication procedures employed. The greatest problem with many studies of insect aDNA is their lack of reproducibility [47], a requirement for scientific evidence. Further limitations of studies of specimens embedded in resin include the implicit destruction of the samples and the small size and often uniqueness of the specimen analysed. Here, we discuss possible ways to solve these questions and limitations.

Fortunately, many studies conducted since 2005 have analysed aDNA using more specific methods, such as next generation sequencing, which not only yields massive amounts of sequencing data and amplification of highly degraded DNA, but also enables more efficient exclusion of modern contamination [53, 54]. Additionally, aDNA research has yielded promising results [e.g. 18, 5457]; positive studies have been reported for aDNA from beetles in museum collections and permafrost [43, 49] and more recently, novel insights have been obtained into biomolecule preservation in amber samples [58] and other fossils [2, 59], including chewed birch pitch [60].

Here, we describe attempts to amplify DNA from ambrosia beetles embedded in six-year-old and two-year-old resins (modern resin, sensu [52]) from Madagascar collected in situ from the producing trees. Authentication and potential contamination with modern DNA were taken carefully into consideration. Our objective was to explore the potential limits of DNA preservation in resins and to develop a standardised protocol for DNA extraction from these samples, which could guarantee unambiguous and independent verification of fossil DNA following the authentication procedures for aDNA research, but applied to a more modern samples by the moment.

Material and methods


We used platypodine beetles of the genus Mitosoma Chapuis, 1865 [61] (Coleoptera: Curculionidae: Platypodinae) embedded in resin (Fig 1). Mitosoma is an endemic genus from Madagascar and is abundantly found embedded in resin drops from Hymenaea verrucosa (Angiospermae: Fabales: Fabaceae). We collected specimens in resins from a lowland forest close to the Pangalanes Canal, in Ambahy (Nosy Varika, Mananjary) (20º46’ S, 48º28’ W) and Andranotsara (Sambava) (14º37’ S, 050º11’ W) on the east coast of Madagascar. The Government of Madagascar authorised sampling (permit no. 160/13 /MEF/SG/DGF/DCB.SAP/SCB and no. 192/17/ MEEF/SG/DGF/DSAP/SCB.Re) and exportation (permit no. 186 N.EA10/MG13 and no. N. 249/17/MEEF/SG/DREEF-SAVA) of samples. The specimens embedded in resin were collected by X.D., E.P. and M.S.K. directly from H. verrucosa trees in Madagascar in October 2013 and September–October 2017, respectively. Resin samples were stored at room temperature until commencing the study in February 2019, which increased the risk of greater DNA degradation [27]. After the first experiments, all resin samples were stored at -20°C.

Fig 1. Resin samples from Hymenaea trees in Madagascar with embedded platypodine beetle specimens of the genus Mitosoma, sampled during fieldtrips in 2013 and 2017.

We used specimens of Mitosoma lobatum Schedl, 1961 [62], M. excisum Schaufuss, 1897 [63] and M. obconiceps Schedl, 1970 [64] as positive controls, collected in 2012 by B.J. when on a fieldtrip in Ranomafana National Park (Madagascar). Specimens were stored in >96% ethanol immediately after collection and preserved at -20°C prior to DNA extraction.

DNA extraction was performed at the Institute of Medical Microbiology, Immunology and Parasitology (University Clinic Bonn, Germany) from ten adult specimens in resin and four adult specimens in ethanol, all representing the same genus and body size, but different localities (see above) and ages (Table 1).

Table 1. Specimens of Mitosoma sp. used for DNA extraction and DNA concentration of the eluted DNA [ng/μl].

Only resin pieces with two or more complete specimens as syninclusions were selected in order to use some of them for analysis and store the other(s) at -20ºC at the Senckenberg Research Institute (Frankfurt, Germany) under the collection numbers SMF Be 13578–13584. We hoped thereby to halt degradation of the hypothetically preserved biomolecules [27] and to preserve the specimen(s) for future controls and experiments. These specimens are available to other researchers/laboratories upon request, for suitable research projects aimed at determining reproducibility. Different specimens from other resin samples collected at the same time (three from 2013 and four from 2017) were treated as independent replicates (Table 1, Fig 1).

Sample preparation and DNA extraction

All samples used in this study were entirely encased within H. verrucosa resin. The resins with insect inclusions were cut into small cubes with a sterile scalpel, leaving a few millimetres around the insect. The surfaces were sterilised with 0.1 M HCl and then washed in sterile water. All the following procedures were carried out under a laminar flow hood in a microbiology laboratory without any contact with entomological experiments. The resin cubes were ground with a micro-pestle in a 1.5 ml reaction tube to which 180 μl ATL-buffer with 20 μl proteinase K (Qiagen, Germany) was added. After incubation at 56°C for 72 h and occasional vortexing, the DNA was extracted using the DNeasy® Blood & Tissue kit (Qiagen, Germany), following the manufacturer’s instructions. DNA was eluted in DNase- and RNase free water and stored at -20°C until further experiments. DNA concentration was measured using the NanoDrop™ One/OneC microvolume-UV/VIS-spectrophotometer (Thermo Scientific, USA). DNA extraction from beetles in ethanol was performed using an identical procedure except for a reduced lysis step of 24 h. To test an alternative methodology some cubes were incubated in 5 ml 100% chloroform (AppliChem GmbH, Germany) for 3 days at 40°C to dissolve the resin. After the chloroform treatment the beetle was washed in ≥ 99% EtOH and further processed as described above. Precautions to eliminate contamination included regular disinfection of all surfaces and working materials with Freka®-NOL AF (Dr. Schumacher GmbH, Germany) and the use of dedicated protective clothing, equipment and reagents. In addition, prior to DNA extraction and amplification experiments, working materials and surfaces were cleaned with DNA-ExitusPlus™ IF (AppliChem GmbH, Germany) to avoid contamination with extrinsic DNA.


Primers and protocols were selected from previously published studies on the weevil subfamily Platypodinae [6567], or newly designed based on Platypodinae DNA sequences. The primer pairs S3690F and A4285R, targeting the D2–D3 domains of the large nuclear ribosomal subunit (28S), and S2442F with A3014R, targeting the 3’ end of the mitochondrial cytochrome oxidase I (COI), efficiently detected DNA from extant Mitosoma samples and were therefore selected for further experiments (Table 2). Two additional COI primers (Table 2; COIRes F and COIRes R2; COInew) were designed based on sequences from the resin-embedded beetles collected in 2013. These enabled amplification of a much smaller fragments than the standardised primers used in previous studies (160 base pairs (bp) vs. more than 600 bp) and were more suitable for new experiments with older (and theoretically more fragmented) material.

Table 2. Primer sequences for DNA extraction selected from the literature and two new sequences tested here.

Polymerase Chain Reaction (PCR) amplification

The PCR reaction mixture used to amplify genes of interest was composed of 12.5 μl OneTaq® 2X Master Mix with Standard Buffer (New England Biolabs, Germany), 0.5 μl of each primer (10 μM) and 50 ng DNA, adding water to a final volume of 25 μl. Addition of 1 μl bovine serum albumin (BSA) (10 mg/ml) to the PCR reaction mixture improved efficiency and enabled a reduction in PCR cycles. In each PCR, a negative (sterile water) and a positive control (beetle DNA from specimens in ethanol) were included. PCR was performed in a Gradient LabCycler (SensoQuest, Germany) with the standard cycle program: initial denaturation step at 95°C for 5 min, followed by 30–50 cycles of denaturation at 95°C for 30 s, annealing at specific temperatures (Table 2) for 30 s and elongation at 68°C for 60 s, and a final elongation step at 68°C for 5 min. Further optimisation included different cycles and gradient PCRs to determine the annealing temperature. The PCR results were analysed by means of agarose gel electrophoresis (1.5%). The PCR products were purified using the GeneJet PCR Purification Kit or GeneJet Gel Extraction Kit (Thermo Scientific, USA), in case that fragments had to be extracted directly from the agarose gel, following the manufacturer’s instructions. Sanger sequencing was performed by Eurofins Genomics GmbH (Germany) with the amplification primers for 28S and COI (Table 2). Alignments between DNA sequences from beetles in EtOH and resin-embedded specimens to analyse the sequence data and investigate potential contamination with modern DNA were performed with Geneious R10 (

Authentication of DNA sequences

We applied the following criteria to authenticate the amplified DNA sequences recovered from the beetles [41, 68]:

  1. Negative controls, extraction blanks and PCR negative controls should be devoid of specific PCR amplification products.
  2. Amplified sequences should be consistently and reproducibly obtained from the same extract(s) from one specimen and from different samples of the same species to guarantee reproducibility.
  3. All procedures should be repeated three times in different laboratory rooms (in our case, at the Institute of Medical Microbiology, Immunology and Parasitology).
  4. All PCR products should be controlled and analysed via sequencing for quality, assembly and specificity.


General pictures of the resin samples were taken using a Nikon D3X with an AF-S Micro-NIKKOR 60 mm 1:2.8 G ED lens. Gel electrophoresis pictures were created using the FastGene FAS-Digi imaging system (NIPPON Genetics Europe). Detailed pictures of the beetle specimen were taken in the laboratory using a Keyence VHX1000 digital microscope under incident light (general body) and using a stereomicroscope in combination with a smartphone adapter. All figures were edited using CorelDraw-X8 software.

Original uncropped and unadjusted images underlying all gel results reported in this work may be found in the S1 Raw images.


Significant differences between DNA concentrations from the analysed samples were calculated by an unpaired t-test using GraphPad Prism version 5.00 for Windows (GraphPad Software, La Jolla, California, USA;


Contrary to some previous experiments using insects in modern resin and sub-fossil resin [21, 47], we successfully amplified DNA sequences from beetles preserved in resins that were six and two years old (Fig 2).

Fig 2. Mean DNA concentration in the different samples analysed.

* = Significant differences P < 0.05; ** = Significant differences P <0.01; n = Non-significant differences.

DNA extraction

DNA was extracted from beetle specimens of Mitosoma sp. and from specimens of the same genus preserved in resin of the same origin (H. verrucosa) but differing in age (2013 and 2017 fieldtrips) and collection site. DNA concentrations differed significantly between recent and resin-embedded beetles (Table 1). We observed a time-correlated decrease, albeit not significant, in the amount of DNA obtained from resin beetles (Table 1, Fig 2). The lowest DNA concentration was detected in resin beetles when the resin encasing the specimens had been dissolved in chloroform prior to DNA extraction (Table 1, Fig 2).

PCR amplification

Initial attempts to amplify DNA from two- and six-year-old resin samples, using the PCR protocol normally employed with recent DNA, were unsuccessful, whereas amplification of recent samples (beetles in ethanol) was positive with the original primer pairs COI and 28S. As DNA extraction from the resin samples was performed with completely ground insect together with remaining resin portions, it was suspected that the resin components might inhibit the PCR. This was tested by the addition of different proportions of resin to beetle samples at different time points during DNA extraction. Neither addition of resin to the sample before DNA extraction nor mixing of extracted DNA from resin and recent samples (Table 3) supported the hypothesis of PCR inhibition by resin components. In further experiments, BSA was added to the PCR mixture to avoid any possible inhibition of the DNA polymerase (Fig 3). The blood plasma protein efficiently binds to inhibitors and thereby prevents a negative impact of these molecules on the DNA polymerase [6971]. After that, two different approaches to amplify the specific DNA fragments were tested: a) two consecutive PCRs of 30 cycles each with a PCR product purification step in between, and b) an increase in the number of PCR cycles up to 50. Both strategies resulted in positive amplifications (Fig 4). Whereas the addition of BSA increased the amplification efficiency of the primer combination targeting the large fragment of COI and enabled successful amplification with a lower number of cycles (≥ 35 cycles), no product could be detected for 28S with less than 45 PCR cycles (Fig 3).

Fig 3. Gel electrophoresis of specific DNA fragments amplified with the original primer combinations COI and 28S with different numbers of cycles.

Comparison of PCR products with and without the addition of BSA to prevent DNA polymerase inhibition. Rb13 = Resin beetle (collected in 2013); Rb17 = Resin beetle (collected in 2017); (+) = positive control, DNA from beetles in ethanol; (-) = negative control, DNase- & RNase-free water; M = 100 bp DNA ladder (New England Biolabs).

Fig 4. Gel electrophoresis of specific DNA fragments amplified with original primer combinations COI and 28S.

A. Positive results with an increase in number of cycles to 50. Rb13 = Resin beetle (collected in 2013); Rb17 = Resin beetle (collected in 2017); (+) = positive control, DNA from beetle in ethanol; (-) = negative control, DNase- & RNase-free water; M = 100 bp DNA ladder (New England Biolabs). B. Two-step PCR with a PCR product purification step in between. Rb172 = Resin beetle (collected in 2017) after two consecutive PCRs of 30 cycles each; (+)1 = positive control, DNA from beetle in ethanol after one PCR of 30 cycles; (+)2 = positive control, DNA from beetle in ethanol after two consecutive PCRs of 30 cycles each; (-)1 = negative control, DNase- & RNase-free water after one PCR of 30 cycles; M = 100 bp DNA ladder (New England Biolabs).

Table 3. Inhibition test performed with the primer combination COI for different proportions of DNA from beetles in ethanol and resin beetles.

Negative amplification was obtained when the DNA from beetles in ethanol was absent, weak positive result when the proportion was low and positive in the remainder of tests, even when resin remains were included.

Control for contamination

Authentication procedures in aDNA studies are aimed at avoiding sample contamination with modern environmental DNA and detecting this should it occur. Therefore, each PCR product was purified and then sequenced by Sanger sequencing. To analyse the sequences several alignments were performed. Besides alignment of the sequences to Platypodinae sequences in the NCBI database BLAST to test for specificity, all products were checked for contamination with modern DNA. We especially examined the risk of inter-sample contamination from the positive control (beetles in ethanol) to the tested samples by multiple alignments of all sequences. Whereas the similarity between the sequences of insects extracted from resin in 2013 and 2017 was ≥ 97.4% with both primer combinations, a comparison between resin beetles and the sequence of beetles in ethanol yielded a similarity of only 73.7% for the original primer pair COI and 52.5% for primer combination 28S, respectively (Table 4), excluding any contamination with DNA from beetles in ethanol.

Table 4. Multiple alignments of all analyzed sequences and the similarity value for the different combinations.


Contamination occurring during analysis

A major handicap in aDNA research, especially when using resin samples, is contamination with modern environmental DNA [1620]. The selection of gene sequences for DNA amplification is a crucial step before working with genetically unknown organisms, as is the case of extinct organisms. Here, the closest living relatives often form the basis for identification of homologous sequences. In this study, primers were selected and specifically designed for beetles of the subfamily Platypodinae [66, 67], to prevent amplification of contaminating modern DNA. Furthermore, our experiments were performed in a microbiology laboratory experienced in molecular methods, but which had never worked with entomological DNA before.

To avoid contamination with modern DNA in the experiments presented here, several control procedures at different stages of the isolation and amplification processes were included in the experimental setup. Besides precautions for working with DNA, all PCR products were purified, sequenced and controlled for sequence specificity and inter-sample contamination. Relatively long fragments of the 28S ribosomal subunit (~ 800 bp) and the mitochondrial gene oxidase (~ 600 bp) (Table 1, Fig 2) were successfully amplified.

To take into account aDNA fragmentation due to decay, the primer used for amplification should encase small sequences. A basic premise is that primers should amplify fragments up to about 160 bp, because longer aDNA sequences become damaged and degraded and have generally proved impossible to amplify [18, 41, 68]. We designed primers for the amplification of small fragments (~ 160 bp) (Table 2; COInew) for future experiments. The new primers were tested successfully in first experiments with resin-embedded platypodine specimens from 2017 (Fig 5) and therefore, build a fundament for future experiments with aDNA using the same insects after confirming the results obtained by longer fragments.

Fig 5. Gelelectrophoresis of the specific DNA fragments amplified with primer combination COInew.

The primers were designed based on specific DNA sequences from resin-embedded beetles (collected in 2017) to amplify smaller DNA fragments. Rb17 = Resin beetle (collected in 2017); (-) = negative control, DNase- & RNase free water; M = 100 bp DNA ladder (New England Biolabs).


Entomological studies can easily implement most recommended aDNA authentication procedures to increase the degree of reliability [1820]. However, other criteria, such as reproducibility or independent replication, are difficult to meet with insect samples due to the small size and frequent uniqueness of the samples. The methodology entailed with samples embedded in resin renders the research even more difficult. The establishment of authentication procedures for aDNA from small samples requires further consideration [27]. Since aDNA isolation in amber insects requires the complete destruction of fossil material, which is obviously undesirable when dealing with rare and very old species stored in museum collections, a more feasible approach would be to analyse several individuals of the species of interest or individuals of any related species of a group [68]. A taxonomic group with relatively abundant representatives would be the perfect target for these experiments; when a specimen is truly unique, samples cannot be used for this purpose because of obvious methodological and ethical problems [27]. The problem of reproducibility is solved here by the selection of insect samples in resin that contain co-specific syninclusions (minimum 2), with a comparable degree of preservation. After cutting the resin sample, the remaining syninclusions were stored at -20°C to maintain their stability as far as possible and thereby preserve the DNA for future tests. Although they do exist, it is difficult to find similar syninclusions in older resins. For rare or unique finds, the approved protocol is to archive independent similar results in two independent laboratories [17, 68]. However, it is extremely challenging to achieve this when using insects from resin or amber deposits because it is difficult to independently replicate DNA extraction from the same fossil insect. In addition, resins of different origin should be treated as independent experiments because of the different nature of their chemical compounds.

Taking into account all the authentication procedures to increase the degree of reliability and avoid contamination in aDNA studies [1720, 27, 68], we summarise here the optimal method that we employed, which we consider should become standard practice in taphonomic research on DNA preservation in modern and ancient resins:

  1. Analyse the sample in a DNA laboratory that has not previously worked with organisms similar to the target specimens, preferably with previous experience in aDNA research.
  2. Select a taxonomic group with abundant representatives as syninclusions in modern resin, sub-fossil resin or amber.
  3. Include negative controls, extraction blanks and PCR negative controls.
  4. Use specific primers for the target specimens instead of more general primers which amplify a broader range of genomes.
  5. Include a sequencing analysis for quality, assembly and specificity control.
  6. Repeat the experiment, ideally with similar specimens from the same sample. Close taxonomic groups from the same sample and corroboration with a sequencing process may be sufficient to demonstrate reliability and discard contamination.
  7. Include a detailed list of the protocols used in the experiments with a clear discussion of the authentication procedures employed.

The use of chloroform in DNA extraction from specimens embedded in resin

The dissolution of resin, sub-fossil resin and amber in chloroform is a typical procedure to release the embedded insects for further analysis [21, 72]. Penney et al. [21] were unable to detect any insect DNA via next generation sequencing after the use of chloroform to dissolve the resin, and thus concluded that DNA was absent. Chloroform:phenol mixtures are frequently used for DNA extraction [7375] and one might assume that the difference in age between the youngest sample analysed in Penney et al. [21] and the oldest one in this study (6 years vs. 60 years) could be a key factor in our successful amplification. Surprisingly, we found that the use of chloroform to dissolve the resin surrounding the specimens compromised DNA concentration (Table 1, Fig 2). During work with DNA samples, the use of 70% ethanol also poses a problem as DNA precipitates at this ethanol concentration [47]. Therefore, only ≥ 80% ethanol should be used to sterilise resin surfaces. Büsse et al. [51] reported the amplification of DNA sequences from specimens preserved inside sub-fossil resin samples, but the lack of information about the methodology used with the specimens embedded in resin precludes comparisons. Further experiments are required to obtain more strongly supported conclusions about the effects of chloroform and ethanol on DNA extraction from resin-embedded material.

Dehydration of specimens

The life-like preservation of many amber inclusions is accompanied by preservation of ultrastructural details such as muscle fibres [76], mitochondria [77], brain tissue [3], and internal genitalia [78]. After the death of an organism, enzymes such as DNases rapidly degrade the DNA, but some repair mechanisms, if active, can revert this process. Under rare circumstances, tissue rapidly desiccates after death [8, 79] and DNA is absorbed by a mineral matrix [80]; in addition, rapid burial may change surrounding conditions, significantly reducing or completely inhibiting enzymatic and microbial degradation [81, 82]. On such occasions, slow but still manifest chemical processes start to affect the DNA [18, 83, 84]. Cano [85] stated that sugars such as arabinose, galactose and sucrose, which are components of natural resins, increase osmotic pressure on cells and thereby draw out the water, which results in tissue dehydration and inhibition of biochemical reactions. Additionally, alcohols and terpenes may act as fixatives to preserve tissues.

Dehydration and preservation in resin or permafrost are the three modes of preservation that retain more life-like organisms than any other kind of fossil, maintaining tissues over time [3]. While promising results have been obtained from permafrost-preserved invertebrates [43], none of the claims of DNA isolated from million-year-old amber samples could be independently replicated [21, 47]. In a recent study, amplified fossil genomes were extracted from birch pitch [60], another organic plant substance like resin, but obtained by heating birch bark. Supposedly, volatile compounds cause thorough and rapid dehydration and fixation of tissue once the trapped organism comes into contact with the resin [3]. This contrasts with our observations of resin-embedded Mitosoma sp. specimens from Madagascar (Fig 6). Two-year-old ground specimens still contained apparently flesh-like tissue without any indication of dehydration or shrinkage (Fig 6C). Therefore, hypothetical dehydration in the resin is not as rapid as was thought and must be regarded as the result of a process lasting for several years. Although the mono- and sesquiterpenoid volatile compounds in resins are known to have antimicrobial and enzyme inhibiting characteristics [23, 86, 87], it is possible that the di- and triterpenoid non-volatile compounds, some of which are also antimicrobial, are crucial to stabilise tissue over longer timescales. The lack of preservation of other molecules, such as chitin and lignin, which are more time-resistant than DNA, in Miocene amber-embedded insects and plants [88] has also been used to argue against the preservation of DNA in these particular fossils [18]. However, ß-1,3 and ß-1,4-linked polysaccharides, and specifically N-acetylglucosamine residues from chitin, have been detected in fungal mycelia in resinicolous fungi from Spanish amber [89]. One explanation for the better preservation of aDNA in bones than in resins may be hydroxyapatite. This mineral, which predominates in bone, is known for its very strong binding affinity to DNA [15, 80, 90]. Our observations indicate that there is still no satisfactory explanation for preservation in amber, but imply that water is available in the system for longer than previously thought, which has a negative effect on DNA stability in specimens embedded in resin.

Fig 6. Different specimens of Mitosoma sp. preserved in resin from Hymenaea verrucosa collected in the Sambava region (Madagascar) in 2017.

A. Lateral view of a complete specimen. B. Dorsal view of a complete specimen. C. Photomicrograph of the sectioned head of one Mitosoma beetle in resin, showing the still fleshy internal tissue after two years inside the resin.

Future approach to determine the time limits of DNA preservation in resin specimens

There are broad areas where further progress in studying aDNA from resin-embedded insects can be expected. Decay experiments provide a useful means to investigate variations in the preservation quality of different resin types [26] and offer a promising subject for future analysis. The preservation of inner structures of insects was highly compromised after only a few months embedded in different kinds of resin [26], but to the best of our knowledge, DNA preservation has never been tested. A comparison of DNA preservation in dead specimens of varying ages (such as those from museums and collections) [91], or specimens fixed in paraffin or embedded epoxy resin from less than one hundred years ago, would be highly interesting. Based on the positive results from this study, future experiments could investigate the limits of the DNA preservation in resin embedded specimens in short-time intervals. For future approaches with samples highly sensitive to contaminations, it will be required further adjustments of the methodology. For instance, the DNA concentration was measured via a microvolume-UV/VIS-spectrophotometer in this study, which lacks specificity, being unable to distinguish between DNA (single- and double-stranded), RNA and nucleotides, and is prone to contamination. Therefore, we recommend to use a fluorometer, which is highly accurate detecting fluorescent nucleic acids. Additionally, DNA of older resin embedded specimens should be analysed with more sensitive next generation sequencing technologies such as shotgun sequencing. This method is more specific, enabling distinction between modern contaminations and ancient DNA and provides further possibilities, e.g. analysis of fragmentation or degradation patterns. Proteins are thought to be more stable than DNA [92, 93] and seem to provide similar phylogenetic information [2]. Some experiments detected biomolecules such as chitin-protein complexes in insect cuticle from sub-fossil resin but failed to do so when Dominican amber samples were tested [88], while other more recent studies found amino acids from fossil feathers in amber from almost 100 million years BP [58]. Therefore, proteins may be a promising target for the study of deep time specimens [94]. However, if the objective is DNA, it may be more effective to amplify mitochondrial DNA since this is present in higher copy numbers than nuclear DNA, rendering it a more promising candidate for genetic analyses of aDNA [4345].

The experiments presented here were performed using the commercial DNeasy® Blood & Tissue Kit (Qiagen, Germany), which was recommended for the extraction of beetle DNA [65, 66]. Other studies have also tested DNeasy® extraction systems, which are based on the binding of DNA to silica membranes, for other types of sample, such as bones or other insects [69, 95]. As a beetle embedded in resin is a mixture of two sample types, tissue (insect) and environmental (resin), selection of the most suitable extraction kit must be carefully assessed. Environmental kits offer the advantage of additional steps to avoid PCR inhibitors [9698], whereas tissue kits are specifically designed for DNA extraction from tissue samples. Other aspects such as choice of DNA polymerase can significantly affect PCR efficiency [69], and should be further investigated to identify the ideal DNA extraction and PCR amplification protocol.

The interaction between a fossil sample and its environment following death of the specimen determines the preservation state of the biomolecules; age per se is not an absolute indicator of quantity or quality, at least over a longer timescale [18]. Storage conditions also play a crucial role in preservation, and dry, cold conditions are best for DNA preservation [27]. Since each case is unique, attending to our results, no biological sample should be disregarded as a potential source for recovery [15].


The existence of aDNA in amber specimens is dependent on the possibility that the resin provides a protected environment for DNA preservation (both protective, through encapsulated conditions, and chemically favourable). In previous studies, no aDNA could be amplified either in amber or in specimens embedded in resin, suggesting that the protective and preservative environment in resin prevents tissue but not DNA degradation. However, modern techniques are more powerful and more sensitive. Recently, Büsse et al. [51] reported successful amplification of aDNA from sub-fossil specimens, but limited information was given about extraction methods, the exclusion of contamination or aspects of reproducibility; therefore, the positive results obtained in that study should be viewed with caution. By contrast, the methodology discussed here is intended for future use as a guide and base for studies of specimens embedded in resin. In future investigations next generation sequencing should be included into the methodology as this technique provides more possibilities when working with aDNA. Our study was designed to clarify fundamental aspects about DNA preservation in resin-embedded insects, including an evident experimental support, that was absent until now. Our positive amplification demonstrates that resin inclusions can also be a resource to explore in aDNA studies, albeit with caution. The risk of contamination demands the design of suitable authentication procedures and the possibility to re-evaluate the inclusion of further analysis once the method is established. Subjects for future studies include the time limits for aDNA detection in resins, the preservation state and assessment of possible taxonomic bias (both entomological and plants) in long-term DNA integrity. With this in mind, a new research project has been launched, moving from newer to older resin samples in order to determine the time limits of DNA preservation in resins.


The first author (DP) thanks Francisco Rodríguez (Vall d’Hebron University Hospital, Barcelona, Spain), Victoria McCoy (University of Wisconsin, Milwaukee, USA), Jesús Gómez-Zurita (Institute of Evolutionary Biology, Barcelona, Spain), and Peter D. Heintzman (Royal Holloway University of London, Egham, UK) for their help and discussion in the early stages of writing this manuscript. We thank Georg Oleschinski (Rheinische Friedrich-Wilhelms-Universität Bonn, Germany) for the pictures of the resin pieces. Finally, we would like to thank the work from the two anonymous reviewers who improved a first version of this manuscript.


  1. 1. Bokelmann L, Hajdinjak M, Peyrégne S, Brace S, Essel E, de Filippo C, et al. A genetic analysis of the Gibraltar Neanderthals. PNAS. 2019; 116: 15610–15615. pmid:31308224
  2. 2. Cappellini E, Welker F, Pandolfi L, Madrigal JR, Fotakis AK, Lyon D, et al. Early Pleistocene enamel proteome sequences from Dmanisi resolve Stephanorhinus phylogeny. Nature. 2019; 574: 103–107. pmid:31511700
  3. 3. Grimaldi DA, Bonwich E, Delannoy M, Doberstein S. Electron microscopic studies of mummified tissues in amber fossils. Am Mus Novit. 1994; 3097: 1–31.
  4. 4. Martínez-Delclòs X, Briggs DEG, Peñalver E. Taphonomy of insects in carbonates and amber. Palaeogeogr Palaeoclimatol Palaeoecol. 2004; 203: 19–64.
  5. 5. Amber Labandeira C. In: Laflamme M, Schiffbauer JD, Darroch SAF, editors. Reading and Writing of the Fossil Record: Preservational Pathways to Exceptional Fossilization. Paleontol Soc Pap. 2014; 20: 163–216.
  6. 6. Pääbo S. Ancient DNA: extraction, characterization, molecular cloning, and enzymatic amplification. PNAS. 1989; 86: 1939–1943. pmid:2928314
  7. 7. Bada JL, Wang XS, Poinar HN, Pääbo S, Poinar GO Jr. Amino acid racemization in amber-entombed insects: Implications for DNA preservation. Geochim Cosmochim Acta. 1994; 58: 3131–3135. pmid:11539553
  8. 8. Mandrioli M. Insect collections and DNA analyses: How to manage collections? Mus Manag Curatorship. 2008; 23: 193–199.
  9. 9. Cano RJ, Poinar HN, Roubik DW, Poinar GO Jr. Enzymatic amplification and nucleotide sequencing of portions of the 18s rRNA gene of the bee Proplebeia dominicana (Apidae: Hymenoptera) isolated from 25–40 million year old Dominican amber. Med Sci Res. 1992; 20: 249–251.
  10. 10. Cano RJ, Poinar HN, Poinar GO Jr. Isolation and partial characterization of DNA from the bee Proplebeia dominicana (Apidae: Hymenoptera) in 25–40 million year old amber. Med Sci Res. 1992; 20: 619–623.
  11. 11. DeSalle R, Gatesy J, Wheeler W, Grimaldi D. DNA sequences from a fossil termite in Oligomiocene amber and their phylogenetic implication. Science. 1992; 257: 1933–1936 pmid:1411508
  12. 12. Poinar HN, Cano RJ, Poinar GO Jr. DNA from an extinct plant. Nature. 1993; 363: 677.
  13. 13. Cano RJ, Poinar HN, Pieniazek NJ, Acra A, Poinar GO Jr. Amplification and sequencing of DNA from a 120–135-million-year-old weevil. Nature. 1993; 363: 536–538. pmid:8505978
  14. 14. Thomas RH, Schaffner W, Wilson AC, Pääbo S. DNA phylogeny of the extinct marsupial wolf. Nature. 1989; 340: 465–467. pmid:2755507
  15. 15. Parsons TJ, Weedn VW. Preservation and recovery of DNA in postmortem specimens and trace samples. In: Haglund W, Sorg M, editors. Advances in Forensic Taphonomy: The Fate of Human Remains. New York: CPR Press; 1996. pp. 109–138.
  16. 16. Instability Lindahl T. and decay of the primary structure of DNA. Nature. 1993; 362: 709–715. pmid:8469282
  17. 17. Lindahl T. Recovery of antediluvian DNA. Nature. 1993; 365: 700. pmid:8413647
  18. 18. Pääbo S, Poinar HN, Serre D, Janicke-Després V, Hebler J, Rohland N, et al. Genetic analyses from ancient DNA. Annu Rev Genet. 2004; 38: 645–679. pmid:15568989
  19. 19. Hebsgaard MB, Phillips MJ, Willerslev E. Geologically ancient DNA: fact or artefact? Trends Microbiol. 2005; 13: 212–220. pmid:15866038
  20. 20. Willerslev E, Cooper A. Ancient DNA. Proc R Soc B. 2005; 272: 3–16. pmid:15875564
  21. 21. Penney D, Wadsworth C, Fox G, Kennedy SL, Preziosi RF, Brown AT. Absence of ancient DNA in sub-fossil insect inclusions preserved in ‘Anthropocene’ Colombian copal. PLoS ONE. 2013; 8: e73150. pmid:24039876
  22. 22. Rosselló JA. The never-ending story of geologically ancient DNA: was the model plant Arabidopsis the source of Miocene Dominican amber? Biol J Linn Soc. 2014; 111: 234–240.
  23. 23. Langenheim JH. Plant resins: chemistry, evolution, ecology and ethnobotany. Cambridge: Timber Press; 2003.
  24. 24. Lambert JB, Santiago-Blay JA, Wu Y, Levy AJ. Examination of amber and 490 related materials by NMR spectroscopy. Magn Reson Chem. 2015; 53: 2–8. pmid:25176402
  25. 25. McCoy VE, Soriano C, Gabbott SE. A review of preservational variation of fossil inclusions in amber of different chemical groups. Earth Environ Sci Trans R Soc Edinb. 2018; 107: 203–211.
  26. 26. McCoy VE, Soriano C, Pegoraro M, Luo T, Boom A, Foxman B, et al. Unlocking preservation bias in the amber insect fossil record through experimental decay. PLoS ONE. 2018; 13: e0195482. pmid:29621345
  27. 27. Reiss RA. Ancient DNA from ice age insects: proceed with caution. Quat Sci Rev. 2006; 25: 1877–1893.
  28. 28. Darzynkiewicz Z, Juan G, Li X, Gorczyca W, Murakami T, Traganos F. Cytometry in cell necrobiology: Analysis of apoptosis and accidental cell death (necrosis). Cytometry.1997; 27: 1–20. pmid:9000580
  29. 29. Nagata S. Apoptotic DNA fragmentation. Exp Cell Res. 2000; 256: 12–18. pmid:10739646
  30. 30. Lindahl T, Andersson A. Rate of chain breakage at apurinic sites double-stranded deoxyribonucleic acid. Biochemistry. 1972; 11: 3618–3623. pmid:4559796
  31. 31. Lindahl T, Nyberg B. Rate of depurination of native deoxyribonucleic acid. Biochemistry 1972; 11: 3610–3618. pmid:4626532
  32. 32. Briggs AW, Stenzel U, Johnson PLF, Green RE, Kelso J, Prüfer K, et al. Patterns of damage in genomic DNA sequences from a Neandertal. PNAS. 2007; 104; 14616–14621. pmid:17715061
  33. 33. Dabney J, Meyer M, Pääbo S. Ancient DNA damage. Cold Spring Harb Perspect Biol. 2013; 5: a012567. pmid:23729639
  34. 34. Hopfenberg HB, Witchey LC, Poinar GO, Beck CW, Chave KE, Smith SV, et al. Is the air in amber ancient? Science. 1988; 241: 717–721. pmid:17839082
  35. 35. Smith CI, Chamberlain AT, Riley MS, Stringer C, Collins MJ. The thermal history of human fossils and the likelihood of successful DNA amplification. J Hum Evol. 2003; 45: 203–217. pmid:14580590
  36. 36. Gaertner J. De Fructibus et Seminibus Plantarum. Stuttgart: Stutgardie Typis Academie Caroline; 1791.
  37. 37. Anderson KB, Winans RE, Botto RB. The nature and fate of natural resins in the geosphere—II. Identification, classification and nomenclature of resinites. Org Geochem. 1992; 18: 829–841.
  38. 38. Penney D. Biodiversity of fossils in amber from the major world deposits. Manchester: Siri Scientific Press; 2010.
  39. 39. Rust J, Singh H, Rana RS, McCann T, Singh L, Anderson K, et al. Biogeographic and evolutionary implications of a diverse paleobiota in amber from the early Eocene of India. PNAS. 2010; 107: 18360–18365. pmid:20974929
  40. 40. Shi G, Dutta S, Paul S, Wang B, Jacques FMB. Terpenoid compositions and botanical origins of Late Cretaceous and Miocene amber from China. PLoS ONE. 2014; 9: e111303. pmid:25354364
  41. 41. Pääbo S, Higuchi RG, Wilson AC. Ancient DNA and the polymerase chain reaction. J Biol Chem. 1989; 264: 9709–9712. pmid:2656708
  42. 42. Sawyer S, Krause J, Guschanski K, Savolainen V, Pääbo S. Temporal patterns of nucleotide misincorporations and DNA fragmentation in ancient DNA. PLoS ONE. 2012; 7: e34131. pmid:22479540
  43. 43. Heintzman PD, Elias SA, Moore K, Paszkiewicz K, Barnes I. Characterizing DNA preservation in degraded specimens of Amara alpina (Carabidae: Coleoptera). Mol Ecol Resour. 2014; 14: 606–615. pmid:24266987
  44. 44. Schwarz C, Debruyne R, Kuch M, Mcnally E, Schwarcz H, Aubrey AD, et al. New insights from old bones: DNA preservation and degradation in permafrost preserved mammoth remains. Nucleic Acids Res. 2009; 37: 3215–3229. pmid:19321502
  45. 45. Allentoft ME, Collins M, Harker D, Haile J, Oskam CL, Hale ML, et al. The half-life of DNA in bone: Measuring decay kinetics in 158 dated fossils. Proc R Soc B. 2012; 279: 4724–4733. pmid:23055061
  46. 46. Pawlowski J, Kmieciak D, Szadziewski R, Burkiewicz A. Attempted isolation of DNA from insects embedded in Baltic amber. Inclusion-Wrostek. 1996; 26: 12–13.
  47. 47. Austin JJ, Ross AJ, Smith AB, Fortey RA, Thomas RH. Problems of reproducibility–does geologically ancient DNA survive in amber-preserved insects? Proc R Soc B. 1997; 264: 467–474. pmid:9149422
  48. 48. Gilbert MTP, Moore W, Melchior L, Worobey M. DNA extraction from dry museum beetles without conferring external morphological damage. PLoS ONE. 2007; 3: e272.
  49. 49. Thomsen PF, Elias S, Gilbert MT, Haile J, Munch K, Svetlana K, et al. Non-destructive sampling of ancient insect DNA. PLoS ONE. 2009; 4: e5048. pmid:19337382
  50. 50. Tagliavia M, Massa B, Albanese I, La Farina M. DNA extraction from Orthoptera museum specimens. Anal Lett. 2011; 44: 1058–1062.
  51. 51. Büsse S, von Grumbkow P, Mazanec J, Tröster G, Hummel S, Hörnschemeyer T. Note on using nuclear 28S rDNA for sequencing ancient and strongly degraded insect DNA. Entomol Sci. 2017; 20: 137–141.
  52. 52. Anderson KB. The nature and fate of natural resins in the geosphere. VII. A radiocarbon (14C) age scale for description of immature natural resins. An invitation to scientific debate. Org Geochem. 1996; 25: 251–253.
  53. 53. Bunning SL, Jones G, Brown TA. Next generation sequencing of DNA in 3300-year-old charred cereal grains. J Archaeol Sci. 2012; 39: 2780–2784.
  54. 54. Hofreiter M, Paijmans JLA, Goodchild H, Speller CF, Barlow A, Fortes GG, et al. The future of ancient DNA: Technical advances and conceptual shifts. BioEssays. 2014; 37: 284–293. pmid:25413709
  55. 55. Pääbo S. Neanderthal man: In search of lost genomes. New York: Basic Books; 2014.
  56. 56. Slon V, Mafessoni F, Vernot B, de Filippo C, Grote S, Viola TB, et al. The genome of the offspring of a Neanderthal mother and a Denisovan father. Nature. 2018; 561: 113–116. pmid:30135579
  57. 57. Bailleul AM, Zheng W, Horner JR, Hall BK, Holliday CM, Schweitzer MH. Natl Sci Rev. 2020; nwz206.
  58. 58. McCoy VE, Gabott SE, Penkman K, Collins MJ, Presslee S, Holt J, et al. Ancient amino acids from fossil feathers in amber. Scientific Reports. 2019; 9: 6420. pmid:31015542
  59. 59. Boatman EM, Goodwin MB, Holman H-Y, Fakra S, Zheng W, Gronsky R, et al. Mechanisms of soft tissue and protein preservation in Tyrannosaurus rex. Sci Rep. 2019; 9: 15678. pmid:31666554
  60. 60. Jensen TZT, Niemann J, Iversen KH, Fotakis AK, Gopalakrishnan S, Vågene ÅJ, et al. A 5700 year-old human genome and oral microbiome from chewed birch pitch. Nat Commun. 2019; 10: 5520. pmid:31848342
  61. 61. Chapuis F. Monographie des Platypides. Liège: H. Dessain; 1865.
  62. 62. Schedl KE. Fauna Madagascariensis IV: 188. Contribution à la morphologie et à la systématique des coléoptères scolytoidea. Mém Inst Sci Madag.1961; 12: 127–170.
  63. 63. Schaufuss CFC. Beitrag zur Kaferfauna Madagascars. III. Missions scientifiques de M. Ch. Alluaud auxîles Séchelles (1892) et a Diego-Suarez, Madagascar (1893) (Scolytidae et Platypodidae). Tijdschr. Entomol. 1897; 40: 209–225.
  64. 64. Schedl KE. Fauna Madagascariensis, V, Coleopteres, Scolytoidea. Ann Soc Entomol Fr. 1970; 6: 233–241.
  65. 65. Pistone D, Mugu S, Jordal BH. Genomic mining of phylogenetically informative nuclear markers in bark and ambrosia beetles. PLoS ONE. 2016; 11: e0163529. pmid:27668729
  66. 66. Mugu S, Pistone D, Jordal BH. New molecular markers resolve the phylogenetic position of the enigmatic wood-boring weevils Platypodinae (Coleoptera: Curculionidae). Arthropod Syst Phylo. 2018; 76: 45–58.
  67. 67. McKenna DD, Farrell BD. Molecular phylogenetics and evolution of host plant use in the Neotropical rolled leaf ‘hispine’ beetle genus Cephaloleia (Chevrolat) (Chrysomelidae: Cassidinae). Mol Phylogenet Evol. 2005; 37: 117–131. pmid:16054400
  68. 68. Handt O, Höss M, Krings M, Pääbo S. Ancient DNA: Methodological challenges. Experientia. 1994; 50: 524–529. pmid:8020612
  69. 69. Rohland N, Hofreiter M. Comparison and optimization of ancient DNA extraction. BioTechniques. 2007; 42: 343–352. pmid:17390541
  70. 70. Farell EM, Alexandre G. Bovine serum albumin further enhances the effects of organic solvents on increased yield of polymerase chain reaction of GC-rich templates. BMC Res Notes. 2012; 5: 257. pmid:22624992
  71. 71. Sidstedt M, Steffen CR, Kiesler KM, Vallone PM, Rådström P, Hedman J. The impact of common PCR inhibitors on forensic MPS analysis. Forensic Sci Int. 2019; 40: 182–191.
  72. 72. Mazur N, Nagel M, Leppin U, Bierbaum G, Rust J. The extraction of fossil arthropods from Lower Eocene Cambay amber. Acta Palaeontol Pol. 2014; 59: 455–459.
  73. 73. Gaikwad AB. DNA extraction: Comparison of methodologies. PLoS Biol. 2002; 20: 162–173.
  74. 74. Köchl S, Niederstätter H, Parson W. DNA extraction and quantitation of forensic samples using the phenol-chloroform method and real-time PCR. Methods Mol Biol. 2005; 297: 13–29. pmid:15570097
  75. 75. Barnett R, Larson G. A phenol-chloroform protocol for extracting DNA from ancient samples. In: Shapiro B, Hofreiter M, editors. Ancient DNA. Totowa: Humana Press; 2012. pp. 13–19.
  76. 76. Henwood AA. Exceptional preservation of Dipteran flight muscle and the taphonomy of insects in amber. Palaios. 1992; 7: 203–212.
  77. 77. Poinar HN, Melzer RR, Poinar GO Jr. Ultrastructure of 30–40 million year old leaflets from Dominican amber (Hymenaea protera, Fabaceae: Angiospermae). Experientia. 1996; 52: 387–390.
  78. 78. Perreau M, Tafforeau P. Virtual dissection using phase-contrast X-ray synchrotron microtomography: reducing the gap between fossils and extant species. Syst Entomol. 2011; 36: 573–580.
  79. 79. Shirkey B, McMaster NJ, Smith SC, Wright DJ, Rodriguez H, Jaruga P, et al. Genomic DNA of Nostoc commune (Cyanobacteria) becomes covalently modified during long-term (decades) desiccation but is protected from oxidative damage and degradation. Nucleic Acids Res. 2003; 31: 2995–3005. pmid:12799425
  80. 80. Brundin M, Figdor D, Sundqvist G, Sjögren, U. DNA binding to hydroxyapatite: A potential mechanism for preservation of microbial DNA. J Endod. 2013; 39: 211–216. pmid:23321233
  81. 81. Seilacher A, Reif WE, Westphal F. Sedimentological, ecological and temporal patterns of fossil Lagerstätten. Philos T R Soc B. 1985; 311: 5–24.
  82. 82. Briggs DEG. The role of decay and mineralization in the preservation of soft-bodied fossils. Annu Rev Earth Planet Sci. 2003; 31: 275–301.
  83. 83. Hofreiter M, Serre D, Pääbo S. Ancient DNA. Nat Rev. 2001; 2: 353–359.
  84. 84. Willerslev E, Hansen AJ, Poinar HN. Isolation of nucleic acids and cultures from fossil ice and permafrost. Trends Ecol Evol. 2004; 19: 141–147. pmid:16701245
  85. 85. Cano RJ. Analysing ancient DNA. Endeavour. 1996; 20: 162–167. pmid:9022353
  86. 86. Deans SG, Ritchie G. Antibacterial properties of plant essential oils. Int J Food Microbiol. 1987; 5: 165–180.
  87. 87. Ryan MF, Byrne O. Plant-insect coevolution and inhibition of acetylcholinesterase. J Chem Ecol. 1988; 14: 1965–1975. pmid:24277106
  88. 88. Stankiewicz BA, Poinar HN, Briggs DEG, Evershed RP, Poinar GO Jr. Chemical preservation of plants and insects in natural resins. Proc R Soc B. 1998; 265: 641–647.
  89. 89. Speranza M, Ascaso C, Delclòs X, Peñalver E. Cretaceous mycelia preserving fungal polysaccharides: Taphonomic and paleoecological potential of microorganisms preserved in fossil resins. Geol Acta. 2015; 13: 363–385.
  90. 90. Adegoke JA, Ighavini BO, Onuigbo RO. Characteristic features of the sonicated DNA of Agama agama agama L. (Reptilia, Agamidae) on hydroxyapatite columns, using mouse DNA as a reference. Generica. 1991; 83: 171–80.
  91. 91. Goldstein PZ, DeSalle R. Calibrating phylogenetic species formation in a threatened insect using DNA from historical specimens. Mol Ecol. 2003; 12: 1993–1998. pmid:12803647
  92. 92. Nielsen-Marsh C. Biomolecules in fossil remains: Multidisciplinary approach to endurance. Biochemist. 2002; 24: 12–14.
  93. 93. Wadsworth C, Procopio N, Anderung C, Carretero J-M, Iriarte E, Valdiosera C, et al. Comparing ancient DNA survival and proteome content in 69 archaeological cattle tooth and bone samples from multiple European sites. J Proteom. 2017; 158: 1–8.
  94. 94. Schweitzer MH, Schroeter ER, Cleland TP, Zheng W. Paleoproteomics of Mesozoic dinosaurs and other Mesozoic fossils. Proteomics. 2019; 19: 1800251.
  95. 95. Chen H, Rangasamy M, Tan SY, Wang H, Siegfried BD. Evaluation of five methods for total DNA extraction from western corn rootworm beetles. PLoS ONE. 2010; 5: e11963. pmid:20730102
  96. 96. Whitehouse CA, Hottel HE. Comparison of five commercial DNA extraction kits for the recovery of Francisella tularensis DNA from spiked soil samples. Mol Cell Probe. 2007; 21: 92–96.
  97. 97. Dineen SM, Aranda R, Anders DL, Robertson JM. An evaluation of commercial DNA extraction kits for the isolation of bacterial spore DNA from soil. J Appl Microbiol. 2010; 109: 1886–1896. pmid:20666869
  98. 98. Faber KL, Person EC, Hudlow WR. PCR inhibitor removal using the NucleoSpin® DNA Clean-Up XS kit. Forensic Sci Int Genet. 2013; 7: 209–213. pmid:22784879