Current and emerging tools for the recovery of genetic information from post mortem samples: New directions for disaster victim identification

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

Contents lists available at ScienceDirect

Forensic Science International: Genetics journal homepage: www.elsevier.com/locate/fsigen

Review article

Current and emerging tools for the recovery of genetic information from post mortem samples: New directions for disaster victim identification J. Watherstona,b,

⁎,1

, D. McNevina,c, M.E. Gahana, D. Brucea,b, J. Warda,b

a

National Centre for Forensic Studies, Faculty of Science and Technology, University of Canberra, Bruce, ACT, 2601, Australia NSW Health Pathology, Forensic & Analytical Science Service, Lidcombe, NSW, 2141, Australia c Centre for Forensic Science, University of Technology Sydney, Broadway, NSW, 2007, Australia b

A R T I C LE I N FO

A B S T R A C T

Keywords: Disaster victim identification (DVI) DNA profiling Unidentified human remains Compromised samples Emerging DNA technologies Post mortem

DNA profiling has emerged as the gold standard for the identification of victims in mass disaster events providing an ability to identify victims, reassociate remains and provide investigative leads at a relatively low cost, and with a high degree of discrimination. For the majority of samples, DNA-based identification can be achieved in a fast, streamlined and high-throughput manner. However, a large number of remains will be extremely compromised, characteristic of mass disasters. Advances in technology and in the field of forensic biology have increased the options for the collection, sampling, preservation and processing of samples for DNA profiling. Furthermore, recent developments now allow a vast array of new genetic markers and genotyping techniques to extract as much genetic information from a sample as possible, ensuring that identification is not only accurate but also possible where material is degraded, or limited. Where historically DNA profiling has involved comparison with ante mortem samples or relatives, now DNA profiling can direct investigators towards putative victims or relatives, for comparison through the determination of externally visible characteristics, or biogeographical ancestry. This paper reviews the current and emerging tools available for maximising the recovery of genetic information from post mortem samples in a disaster victim identification context.

1. Introduction

2. International standards in disaster victim identification

The primary and most reliable means of identification for disaster victim identification (DVI) are fingerprint analysis, dental comparison and DNA analysis [1]. DNA profiling has become the gold standard for the identification of victims in both mass casualty incidents and forensic cases where human remains are highly fragmented and/or degraded [2]. This is due to the relatively low cost and high degree of discrimination DNA-based identification can provide. In addition to identifying victims, DNA profiling also offers the ability to reassociate body parts and can aid in the identification of offenders where human activity has led to a mass casualty event [2]. Challenges associated with the sampling of remains can include the number of victims, mechanisms of body destruction, extent of body fragmentation and body accessibility [3]. Disaster locations considered hostile environments can also pose additional challenges for the recovery effort.

The 2004 tsunami in South East Asia and subsequent DVI effort highlight the necessity of standards in the DVI process; it was during this mass disaster that forensic scientists and police organisations started to develop standards for the identification process based on their practical experience [4]. The Tsunami Evaluation Report [5] documents the workflow, responsibilities and other significant issues influencing the decision making process, as well as the aspects of the operation that would influence the efficiency of the identification process. The INTERPOL Standing Committee on DVI [6] would go on to develop guidelines for all aspects of the DVI process, with the inclusion of three working groups: forensic pathology, forensic odontology and police. International standards have continued to develop [4,7–9] with strong evidence following disaster events that local structures should adopt international standards and recommendations, as well as be provided with more detailed guidance regarding appropriate DVI responders [10]. Goodwin [9] highlights two International Organisation for

⁎

Corresponding author at: 480 Weeroona Road, Lidcombe, NSW, 2141, Australia. E-mail address: [email protected] (J. Watherston). 1 Postal address: PO Box 162, Lidcombe, Sydney, NSW, 1825, Australia. https://doi.org/10.1016/j.fsigen.2018.08.016 Received 29 March 2018; Received in revised form 27 August 2018; Accepted 27 August 2018 1872-4973/ © 2018 Elsevier B.V. All rights reserved.

Please cite this article as: Watherston, J., Forensic Science International: Genetics (2018), https://doi.org/10.1016/j.fsigen.2018.08.016

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

quantities of DNA from the skeletal samples [24]. Due to the poor quality and/or quantity of nuclear DNA (nDNA) in samples such as bone and teeth, the analysis of nDNA markers may fail to yield a reportable profile [25–27]. In these instances, mitochondrial DNA (mtDNA) is an alternative target due to its higher resistance to degradation and high copy number per cell [2]. Structurally the major proportion of bone is matrix, consisting of both an inorganic (principally hydroxyapatite) and an organic fraction, which is composed chiefly of type I collagen and extracellular matrix proteins, such as glycosaminoglycans and osteocalcin [28,29]. The collagen provides a soft framework and the minerals add strength and harden the framework. Approximately 70% of bone consists of the inorganic mineral hydroxyapatite which includes calcium phosphate, calcium carbonate, calcium fluoride, calcium hydroxide and citrate [30]. Different skeletal elements have been found to vary in the way they preserve DNA and consequently, yield different amounts of DNA [15,31]. Historically, sampling advice suggested that although spongy and cancellous bone can be rich in DNA, preservation is not reliable and the dense cortical bone (preferably weight bearing long leg bones) should be collected preferentially [7]. This is due to the DNA being protected by the physical and chemical structure of compact bone within the calcium (Ca2+) matrix, which is not present in spongy bones [23]. During the identification efforts of the World Trade Center disaster (9/11) in 2001, it was determined that skeletal samples from femur and metatarsal bones offered more DNA, while skull bones were less suitable [32]. The International Commission on Missing Persons [31] has also identified weight bearing bones such as femur, tibia, pelvis, metatarsal and talus as some of the most suitable skeletal elements for sample collection. The unique composition of teeth and their location in the jawbone provide additional protection from environmental and physical conditions that accelerate post mortem (PM) decomposition and DNA decay [33,34]. The total DNA content of teeth varies considerably between individuals and also within the same individual [35–37]. Pulp and cementum are the most valuable sources of nDNA in the tooth, as well as being good sources of mtDNA [38]. The pulp provides the richest source of DNA in teeth due to the relatively high cellularity [39]; however, pulp may be limited or even absent in aged and/or diseased teeth [38]. Teeth with the largest pulp volume provide the best source of DNA [40,41] and Higgins and Austin [38] suggest teeth with the largest root surface area (i.e. molars) should be targeted. Factors such as tooth type, tooth health and chronological age of the donor will have an effect on the relative proportions of DNA present in the tooth [38].

Standardisation (ISO) standards having high relevance to the identification of human remains. These include ISO/IEC 17020:2012 'Conformity assessment - Requirements for the operation of various types of bodies performing inspection'; ISO/IEC 17025:2018 'General requirements for the competence of testing and calibration laboratories' and; the Forensic Science ISO/IEC 17025 Application Document and ISO 18385:2017 ‘Minimising the risk of human DNA contamination in products used to collect, store and analyse biological material for forensic purposes – Requirements’. These standards have applicability usually in the recovery of evidence at a crime scene and as a technical standard for forensic genetics. The supplementary publication ILAC G19:08/2014 Modules in a Forensic Science Process, by the International Laboratory Accreditation Cooperation Organisation (ILAC) helps to bring a specific forensic application to the broad scope of the ISO standards. Additionally, both practical and simulated quality exercises are becoming more common in an attempt to standardise DVI procedures internationally [11,12]. 3. Sources of DNA DNA can be recovered from a range of biological sources. Depending on the circumstances of a mass casualty incident, some sources may be more ideal for the purposes of DNA-based identification than others. Factors such as resistance of the source to degradation or damage due to its natural structure can also affect DNA recovery. 3.1. Current approaches 3.1.1. Blood and saliva Current INTERPOL Guidelines [1] recommend the collection of blood or saliva on Flinders Technology Australia (FTA®) paper or a swab in complete, non-decomposed bodies. It can be difficult to collect blood and saliva from deceased individuals once the blood ceases to circulate, and saliva ceases to be produced. The collection of blood from a body cavity becomes labour intensive and may require surgical recovery [2]. Blood is the recommended sample if the body is a complete or mutilated non-decomposed body [1], with DNA from blood usually being less degraded than saliva [13]. Blood vessels have been found to yield better short tandem repeat (STR) typing results than muscle samples in decomposed, dismemberment cases [14]. Buccal smears on FTA® cards are recommended only if the body condition is complete and non-decomposed [15]. 3.1.2. Skin and muscle Deep-seated red muscle tissue is currently recommended if the remains are mutilated and incomplete [1]. During decomposition the soft tissues of the body will begin to decompose much earlier than hard tissues. Consequently, DNA in soft tissues tends to degrade faster than in hard tissues [2,7,15,16]. When sampling soft tissue, skeletal muscle is recommended [3,17–19]. In severely burnt corpses, smears from the bladder have been shown to be a particularly effective alternate source of DNA [20].

3.1.4. Hair Hair is associated with use as an ante mortem (AM) sample rather than as a good source of DNA for the purposes of identification of a PM sample [1,7]. Because of its nature, hair has limited value for a conclusive identification, especially if separated from the body. This is further complicated in decomposed remains, particularly following hair loss during the bloat stage [42]. The difficulty with hair samples is being able to recover enough nDNA for a useable DNA profile. This is due to the structure of hair itself which is made up mostly of keratinised proteins, with little or no undegraded source of DNA [43]. In nDNA testing, the success of DNA profiling using hair samples is attributed to the presence of the root, and adhering epithelial cells [44,45]. As for bone, mtDNA can provide an alternative target.

3.1.3. Bone and teeth DNA is well preserved in bone cells and teeth [15], making them reliable sources of DNA, particularly in adverse environmental conditions and for long-term sampling [7,21]. Current recommendations suggest the collection of bone is most appropriate for compromised remains due to a higher success rate of DNA recovery from femur shafts and teeth as compared to blood, buccal and tissue samples [2,7,15,16,22,23]. Specifically, where bodies are complete and decomposed or mutilated, INTERPOL Guidelines [1] recommend the collection of a sample from long, compact bones (4–6 cm window section without shaft separation), healthy teeth (preferably molars) or any other available bones (∼10 g if possible; preferably cortical bones with dense tissue). This is limited, however, by the ability to isolate sufficient

3.1.5. Nails Nails can be a valuable source of DNA because they have been shown to be resistant to decay and preserve their DNA content well [46–48]. DNA in fingernails is assumed to adhere to the underside of the nails but DNA is also preserved within the keratin structure of the nail [49]. STRs were successfully recovered from nails after one month 2

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

collected material to an FTA® card. The FTA® card is then left to air dry, before DNA profiling. This approach offers a simple and effective method for post mortem sampling with sufficient DNA recovery across several PMIs, seasons and environmental conditions for timeframes usually associated with DVI efforts.

when left dry, in wet soil, water (river, sea and distilled water) and air at room temperature. Nails were found to be more fragile in wet soil compared to all other environmental conditions [47]. It has also been suggested that the preservation of DNA in nails (exhumed remains cases) may be attributed to a positive microenvironment surrounding the hands after burial (e.g. cross-positioned onto the abdomen and above the body with no or little contact with other putrefying tissues) [50]. DNA (both endogenous and exogenous) has also been found to persist under the fingernails even after submersion in salt and fresh water [51]. In addition, it has been shown that no differences exist in the DNA content of nails across sex, young versus elderly individuals, or even between each finger of the same person [47]. The use of nails as a source of DNA-based identification on decomposed cadavers has been assessed with post mortem intervals (PMIs) ranging from 5 days to longer than 6 months [48], with full profiles obtained using the PowerPlex® 16 System (Promega). The added benefit of nails (finger or toes) is the simplicity involved in their collection, which makes them an attractive option for routine and/or urgent DNA-based identification [48]. Nails taken from exhumed and partially skeletonised bodies and clippings stored at room temperature for 10–12 years have been successfully typed using STRs, even when no results were obtained from soft tissue or bone [50]. DNA profiles have also been obtained from fingernails added directly to polymerase chain reaction (PCR) (i.e. without DNA extraction) [49]. A limitation with nails is the negative impact of exogenous DNA contaminants [48]; fingernails are sometimes collected specifically for the identification of these exogenous contributors. This complication may be overcome in an identification context by sampling toenails over fingernails, which may be expected to have less extraneous DNA [48]. A DNA extraction method utilising toenails has been optimised and implemented at the Victorian Institute of Forensic Medicine as the primary sample type for collection of decomposed remains when blood sampling is not available [52].

3.2.2. Bones Mundorff and Davoren [24] found that small cancellous bones yield more DNA and STR loci than the cortical bones at increasing PMIs. A comparison of DNA yields from within an individual also showed that, on average, the small cancellous bones (e.g. finger, toe and ankle bones) have much higher concentrations of DNA per unit mass, than dense cortical bones such as femur [24]. The phalanges of the hallux have been used for DNA-based identification in the Brazilian floods and mudslides in January 2011, mostly resulting in DNA yields of > 1.0 ng/ μL with the added benefit of a simple collection method [56]. High load bearing bones, particularly the bones of the feet, have been reported to be a preferable source of DNA in submersion cases [57]. Additionally, small bones can be sampled intact with a disposable scalpel instead of using a bone saw to cut out a bone wedge [24]. Easier sampling associated with collection of the small cancellous bones also justifies pursuing them as a source of DNA. The direct swabbing of exposed bone marrow after the removal of a bone tissue wedge has also proven successful, having recently been applied in the MH17 DVI operation [58]. The auditory ossicles have been shown to provide DNA of sufficient quality and quantity for identification in highly putrefied cadavers [59]. The success and advantages of targeting and DNA profiling costal cartilage attached to rib has been reported, offering a rapid and cheap collection method with no requirement of special equipment [60]. For cases where the cranium is available, it has been found that optimal results are obtained from the petrous portion of the temporal bone [61,62]. 4. DNA profiling

3.1.6. Connective tissue Different types of connective tissues are an alternative DNA source for decomposing remains. Trindade-Filho and colleagues [19] found hyaline cartilage was superior to skeletal muscle in obtaining STR profiles and more reliable when dealing with compromised remains in tropical environments. The skeletal muscle decomposed faster than the hyaline cartilage from joints and it is believed that the DNA degrades in a similar fashion [19]. Achilles tendon has been shown to be a stable tissue and can also be used as a source of DNA in highly decomposed bodies, yielding twice as much DNA than non-decayed kidney and muscle samples [53].

4.1. Current approaches 4.1.1. Preservative solutions Preservative solutions work by inactivating nucleases, slowing microbial growth, denaturing proteins, protecting against bacterial growth and lysing cells [63–68]. Traditionally, there has been an archival focus with long-term tissue storage; however, forensic situations such as mass disasters are mostly field-based and only short-term storage is required. Samples collected for identification are usually analysed relatively soon after an event, or placed in refrigeration following transportation to the laboratory which could take up to a month [69]. A range of preservative solutions have been trialled and recommended for maintaining the integrity of samples [69,70]. INTERPOL DVI Guidelines [1] recommend the use of a preservative (e.g. ethanol) for tissue and muscle samples collected for the purposes of DVI, but formaldehyde and formalin should not be used as they have been found to degrade DNA [1,7]. Sodium chloride (NaCl) is a historically and commonly used preservative. It desiccates the sample as a solid, which inactivates nucleases and slows microbial growth. Conversely, in aqueous solution, it denatures proteins [68]. Solid NaCl has been shown to effectively preserve DNA in human muscle tissue for up to a year [71]. Chelating agents like ethylenediaminetetraacetic acid (EDTA) bind to metal ions which slow or halt nuclease activity on DNA; these metal ions are required by nucleases for normal function [63,64]. Dimethyl sulphoxide (DMSO) promotes the dermal absorption of chemicals [72] and consequently enhances the absorption of other preservatives across membranes and into the cell [73]. Buffering the pH will decrease the rate of acid-catalysed depurination and aid in stabilising the DNA [74,75]. A recent study by Connell et al. [76] has shown NaCl and 70% ethanol preserved fresh and decomposed muscle samples, and 70% ethanol and 20% DMSO preserved fresh bone samples up

3.2. Emerging approaches Recent studies have highlighted the availability of DNA from tissues and organs not previously considered for the purposes of genetic identification, with promising results. Furthermore, such DNA sources can offer time and resource savings for DVI, in contrast to the lengthy sample collection, preparation and extraction of some traditionally targeted samples (e.g. femur). 3.2.1. Tissues including internal organs DNA and RNA have been shown to remain stable in certain organ tissues post mortem [54]. van den Berge et al. [54] examined brain, lung, liver, skeletal muscle, heart, kidney and skin samples from excavated graves with burial times ranging from 4 to 42 years. By comparing the average percentage of detected alleles for each organ type, they found that brain followed by heart were the most promising organ types for DNA profiling of long-buried human remains, and yielded the least degraded profiles. Mundorff et al. [55] describe the direct swabbing of muscle tissue following an incision through the skin, and then the transfer of 3

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

to 65 ͦC, with results comparable with refrigeration.

profile being detected [83]. While some investigators recommend cleaning samples with sodium hypochlorite prior to DNA extraction to remove contaminants, others caution that such treatment can destroy the majority of endogenous DNA [102]. Sodium hypochlorite treatment can erroneously cause exogenous contaminating DNA to take on characteristics of endogenous DNA that is of poor quality and/or quantity [102]. Sodium hypochlorite destroys DNA through oxidative damage such as base modifications and the production of chlorinated base products [107–110]. However, the ability to amplify DNA from a bone or tooth after treatment with sodium hypochlorite suggests that the endogenous DNA is protected from the heavy oxidant through its adsorption to hydroxyapatite, a bond not afforded to the contaminating DNA [95]. Endogenous DNA has been shown to be stable even when exposed to higher sodium hypochlorite treatments (6% for 21 h) facilitating the preservation of DNA in ancient skeletal remains [88].

4.1.2. Sample preparation Contamination can occur in the field or prior to sample receipt in the laboratory. Various studies have shown that bones and teeth may easily become contaminated prior to DNA extraction [77–81]. The susceptibility of bones and teeth to contamination has been postulated to be largely due to the ultra-structural organisation of bones and teeth and in particular, sample permeability [82] and porosity [81], meaning old bones are highly vulnerable to contamination from modern sources [83]. Human teeth lack many of the cell bodies found in bone and seal their roots with age [84] making them less porous. Additionally, the upper surface is protected by impermeable enamel which may explain why they are less susceptible to contamination [82]. Bone and teeth samples are prone to water-borne sources of contaminant DNA, again due to the porous nature of bone and teeth, facilitating the contaminant’s deep penetration [82]. Even dry soils are likely to hold some water, and due to the surface tension of water, skeletal samples with characteristically small pores will contain water even below the soil wilting point [85]. Consequently, contamination with foreign DNA usually originates from individuals who were involved in the initial washing and cleaning of the specimens rather than from those who collected the samples [86]. A study examining resistance to further contamination suggests that samples are most susceptible to contamination just after excavation when they are still damp from the burial environment [82]. The risk of contamination of remains with DNA from other sources including handling of the sample (i.e. by the collector), contaminated instruments, equipment and/or work spaces, or from comingled remains needs to be minimised [15]. The best ways of minimising and eliminating contamination are: use of personal protective equipment, clean sampling areas, use of clean instruments, and the appropriate storage of collected samples (containers and conditions) [87]. Additionally, contamination can be excluded by extracting material from the interior of a bone or tooth sample [88]. Elimination samples can also assist in the identification and interpretation of contamination events [89]. Prior to DNA testing, chemical and physical decontamination procedures can be applied to hard tissues such as nail and bone to remove exogenous DNA. Nail samples can be decontaminated using water with detergents such as sodium dodecyl sulfate (SDS) and 5% Terg-a-zyme™ (Sigma-Aldrich®), or SDS with proteinase K however, a one-hour SDS/ proteinase K soak may be required for removing all exogenous material [90]. Laboratories use a variety of techniques to clean the exterior surface of skeletal remains, including but not limited to, complete removal of the exterior of the bone [91–94], washing of the sample using a diluted bleach solution [88,95], acid wash [96–98] and ultra-violet (UV) irradiation [99,100]. Mundorff and Davoren [24] report cleaning bone by wiping with 10% bleach, sterile water, and 70% ethanol, followed by at least 12 h of drying in a hood. More recently, a combination of physical cleaning (i.e. mechanical removal of the surface of the bone by sanding), chemical cleaning (i.e. bleach, water and ethanol) and UV irradiation has been recommended as the best way to decontaminate hard tissues prior to powdering using a freezer mill, commercial blender or drill [9,101]. While sodium hypochlorite is highly efficient (∼81–99%) at contamination removal, no treatment is capable of completely removing contaminants [102], nor is there consensus on the optimum concentration of sodium hypochlorite, or the amount of time the bone or tooth should be exposed [88]. Moreover, there is disagreement in the literature regarding the ability to completely remove contaminants, with some believing it is difficult to entirely remove the contaminants [82,83,102–104], and others claiming complete decontamination is possible [88,105,106]. Decontamination is even more problematic for mtDNA testing with up to 15% of samples potentially retaining detectable contaminant mtDNA after cleaning, despite no autosomal DNA

4.1.3. DNA extraction It is imperative that tissue samples thought to contain inhibitors are either diluted to a point where inhibitor concentration is too low for inhibition to occur or are subject to DNA extraction such that the inhibitors are removed. This is important as PCR inhibition is the most common cause of PCR failure when sufficient DNA is recovered [111]. This is particularly the case for DNA extraction from bone where a demineralisation procedure has been employed, involving concentrated EDTA to both dissolve the bone and sequester Ca2+ from solution [30,112]. PCR inhibitors from bone, such as Ca2+ and collagen [113], have been routinely reported [114,115] and EDTA treatment of bone samples can result in co-extraction of an excess of proteins and other biomolecules, possibly through release of extracellular matrix components [116]. PCR inhibition from bone exposed to water and/or soil is thought to be due to the presence of humic and fulvic acids, tannins, iron, cobalt and other materials that can enter the bone after long periods of exposure [117–120]. DNA extraction efficiency and the removal of potential PCR inhibitors is crucial for the subsequent steps in DNA processing [121]. There are a number of different extraction methods and commercial extraction kits available. Silica-based extraction methods (e.g. QIAamp® DNA Mini Extraction kit (QIAGEN), EZ1® DNA Investigator kit (QIAGEN), DNA IQ™ System (Promega)) provide acceptable results in terms of DNA yield and inhibitor removal but have been reported to be inferior to the PrepFiler™ Forensic DNA Extraction kit (Thermo Fisher Scientific: TFS) [122]. PrepFiler® has also been shown to be as effective as the organic method for extracting DNA from fresh bone [123]. However, the QIAamp® DNA Investigator kit (QIAGEN) was successfully used to extract DNA from bone, tissue, buccal swabs and blood stains during the 2009 Victorian bushfire identification effort using the QIAcube (QIAGEN) platform [124] and subsequently from toenail samples for decomposed remains cases [52]. The DNA IQ™ System has been proven to be an effective technology in the separation of DNA from PCR inhibitors and contaminants from most casework samples [125–128] and has been applied to an automated workflow with enhanced throughput and capacity [125,129,130]. DNA purification kits based on magnetic beads (e.g. PrepFiler™, ChargeSwitch® (TFS)) have been shown to extract DNA from very small quantities of material including blood, saliva and semen [2]. Further, they have been extensively applied to automated DNA processing platforms. The combination of uniquely structured magnetic particles and an optimised multi-component surface chemistry provides a very efficient DNA binding capacity, removal of inhibitors and maximum recovery of DNA [131]. Demineralisation is a DNA extraction protocol for bones and teeth involving submersion of powder material in an EDTA-based buffer. Incubation in the buffer aids in the breakdown of calcium phosphate leaving decalcified material for extraction [9]. The EDTA demineralises the bone and inactivates DNAses by chelating bivalent cations such as Mg2+ or Ca2+ [30,132]. Huel et al. [133] highlight the ratio of bone 4

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

concentration of total DNA and male DNA. Quantifiler™ Trio includes a ‘Quality Index’ which helps to identify degraded DNA and the presence of inhibitors [151]. Specifically, the ‘Degradation Index’ can indicate the extent of DNA degradation by comparison of large and small autosomal targets.

powder to 0.5 M EDTA is important for optimum digestion. For example, they recommend that 1 g of bone powder should be demineralised in 15 mL of 0.5 M EDTA. While an overnight dissolution is usually applied (for convenience), in most cases 5–6 h is sufficient to completely demineralise 0.5 g of bone powder [112,133]. Increasing the EDTA incubation time to 48–120 h can induce DNA damage or degradation [134–136]. Current recommendations suggest bone and tooth powder should be subject to total demineralisation [30,101]. This method, coupled with proteinase digestion, significantly increases DNA yields and DNA typing results by completely breaking down the hard material [30,112,132,137,138]. Complete dissolution of the bone powder is thought to permit access to larger, high-quality fragments of endogenous DNA that are held in dense crystal aggregates of the bone matrix [95]. This also eliminates the possibility that DNA is left behind in the solid bone matrix, which is important for smaller quantities of starting material [30,112]. Amory et al. [112] suggests total demineralisation success might be related not only to ‘releasing’ more nucleic acids but also targeting well preserved DNA bound to the hydroxyapatite crystal aggregates as suggested by Gotherstrom et al. [139] and Salamon et al. [95]. A total demineralisation protocol has also been automated but may offer increased capacity at the cost of sacrificing DNA yields [112]. However, an automated total demineralisation protocol has recently been shown to extract DNA more efficiently on both contemporary and World War II skeletal extracts [140]. Total demineralisation of thin slices of bone have also been compared to that of bone powder. Caputo et al. [141] determined that better quality STR profiles were obtained when DNA was extracted from bone slices, with optimal yields obtained from 150 to 200 mg of starting material [141]. Following demineralisation, samples can be extracted using either an organic or inorganic DNA extraction method. Organic extraction (phenol:chloroform) is effective at removing proteins and lipids from DNA extracts but tends to be ineffective for the removal of hydrophilic compounds, which is particularly problematic for skeletal remains exposed to water and/or soil [136,137]. Additionally, solvents used in the extraction pose known health hazards requiring handling in a fume hood. In comparison, most non-organic bone extraction methods use the ability to reversibly bind DNA to silica via salt bridging [112,142–147], and use ultrafiltration membranes to remove contaminants [30,93,148,149]. The total demineralisation method of Loreille et al. [30] and the silica-based cleanup of Yang et al. [142] using QIAquick™ spin columns has been combined for the processing of more challenging skeletal samples [112,132,145]. The approach has been reported to recover more DNA with less PCR inhibitors and with improved STR typing results for aged samples [132,145]. In degraded bone samples, large variations have been observed in both quantitative data and in the recovered DNA profiles among separate extractions of the same aliquot of bone powder. This reflects variability in quantity and quality of the DNA in such samples. This variability is thought to be attributed to differential distribution of DNA within the sample, or to uncharacterised variables that affect the extraction process [112,141]. This is further complicated by an inability to predict DNA yield from bone macroscopic appearance [28,29,95,137]. Such uncertainty highlights the importance of using an extraction protocol which has been optimised for a smaller starting amount of bone powder, facilitating re-extraction of challenging samples following failure to recover a sufficient profile [23,112].

4.2. Emerging approaches 4.2.1. Rapid DNA testing Rapid DNA analysis systems are suitable for mobile and non-expert operation [152]. Such technologies include the RapidHIT™ 200 (IntegenX, recently acquired by TFS), ParaDNA (LGC forensics) and DNAscan (NetBio) [153]. The RapidHIT™ 200 Human DNA Identification System is a fully integrated system capable of generating STR profiles in approximately 90 min [154]. The early RapidHIT™ system reliably allowed the rapid DNA profiling of buccal swabs, with some profiling success also obtained for saliva, semen, skin and hair samples [152]. The RapidHIT™ 200 has been assessed for reproducibility, contamination, storage, and sensitivity, and has been determined to perform well when compared to traditional capillary electrophoresis (CE)based methods in producing useable DNA profile information [155]. While the RapidHIT™ 200 system incorporates extraction, amplification, CE separation and a large proportion of software, it is unable to quantify the amount of DNA added to PCR [155]. Moreover, rapid DNA devices cannot yet compete with the sensitivity of standard laboratory analyses. Mapes et al. [153] highlight the dilemma of getting suboptimal results within two hours, or analysing the sample in the laboratory with higher sensitivity, but with a much longer processing time. This will prove especially important when considering degraded and compromised human remains. 4.2.2. Direct-to-PCR The direct-to-PCR preparations involve directly adding the samples to the PCR tube. The success of the direct-to-PCR approach has been reported for blood [156,157], hairs [158,159], fingernails [49] and more recently, a single hair follicle [158]. Habib et al. [160] report a successful direct-to-PCR application for reference and/or post mortem samples including toothbrush bristles, muscle tissue and bone shavings, following a Proteinase K lysis step. Samples have been directly amplified using several STR multiplex kits, including both Promega (e.g. PowerPlex® 21 System [161–163]) and TFS (e.g. Identifiler® Plus (TFS) [164], GlobalFiler® [163]) kits. Direct-to-PCR methodology has also been applied to Y-STRs using the PowerPlex® Y23 (Promega) and AmpFlSTR® Y-filer® (TFS) kits [164]. Direct-to-PCR has been found to generate a significant increase in the height of electropherogram peaks, as a result of more DNA template, compared to traditional extraction methodologies [165]. Only 17 cells are required to obtain a full STR profile using direct-to-PCR, compared to 250 cells required for standard extraction methodology as a result of losses from DNA extraction [159]. The success and quality of the DNA profiles recovered using the direct-to-PCR approach is dependent on the nature of the material sampled and the presence of PCR inhibitors, which are usually removed during the DNA extraction process [166]. Optimisation options include diluting PCR product prior to CE and reducing the number of cycles. The implications on the routine operation of the laboratory will also need to be considered [159]. 4.2.3. DNA leeching preservative solutions Some preservatives leech DNA from the preserved tissue and into solution which makes subsequent handling easier and less prone to contamination as an aliquot of the preservative can be withdrawn (by pipette) without having to disturb the tissue itself. These include DMSO-EDTA salt-saturated (DESS) solution, TENT buffer (Tris, EDTA, salt and detergent) and some proprietary preservatives (e.g. DNAgard® (Biomatrica®)). Sorensen et al. [167] added these solutions to tissue samples obtained from decomposing human remains and extracted the

4.1.4. DNA quantification Following DNA extraction, extracts are quantified to determine the quantity and quality of DNA in a sample. Amplifying samples with suboptimal target DNA concentration can result in substrate inhibition or insufficient signal intensities [150]. A common approach in the forensic science community is the use of multiple target Taqman assays in real-time polymerase chain reactions (RT-PCRs). For example, the Quantifiler™ Trio DNA Quantification kit (TFS) provides the 5

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

‘free DNA’ in solution. There is evidence to suggest that these aliquots can be added directly to PCR although the presence of inhibitors requires careful optimisation of preservative concentrations and aliquot volumes [69,163]. Sorensen et al. [163] obtained DNA profiles without DNA extraction, by adding aliquots of preservative solutions surrounding fresh and decomposing human muscle and skin tissue samples directly to PCR. A Tecan Freedom EVO® 150 robotic platform amplification workstation was used to setup the PowerPlex® 21 reactions [161–163]. DESS solution, TENT buffer and two proprietary preservatives (DNAgard® and DNA Genotek) produced full and partial profiles from skin and muscle samples procured before full bloat of two cadavers, although dilution was required to relieve inhibition for DESS and the two proprietary preservatives. This work has continued with the trial of a modified TENT buffer and FTA® Elute Card on decomposed tissue for up to six months at room temperature [168]. A successful application of leeching preservative solution on aged blood stain samples has also been reported [162].

samples to be collected from distant maternal relatives (with the expectation that mtDNA should remain largely the same) when reference samples from direct relatives may be difficult or impossible to obtain. 5.2. Emerging approaches The forensic community has looked beyond the use of STRs for DNA identification. Often challenges to identification efforts include the degraded and compromised nature of the DNA remaining. Some genetic markers are less susceptible to degradation, and this can aid in the genotyping of the available DNA. Where traditionally DNA in a forensic, mass disaster or missing person context has relied strictly on comparative grounds, recent advances in genetics are allowing highly informative DNA markers to be used for the prediction of externally visible characteristics (EVCs) and biogeographical ancestry (BGA) [184]. 5.2.1. Single nucleotide polymorphisms Single nucleotide polymorphisms (SNPs) are single base sequence variations between individuals at a particular point in the genome. Amplicons which contain target SNPs are usually short, making them useful for degraded samples [15,185]. Panels of SNPs can provide individual discrimination comparable to the standard Combined DNA Index System (CODIS) panel of STRs if a suitable number are typed [186–188]. SNPs are classified according to their forensic application: identity-informative SNPs (IISNPs) for human identification, lineageinformative SNPs (LISNPs) for inferring genealogies, phenotype informative SNPs (PISNPs) for inferring EVCs and ancestry informative SNPs (AISNPs) for inferring BGA [189]. PISNPs can reveal forensically useful physical traits [184,190–192] while AISNPs can provide information as to the BGA of the contributor of the DNA [193–197]. These DNA intelligence tools, referred to as forensic DNA phenotyping (FDP), show promise for missing person and DVI applications, offering scope to provide leads to putative victims, or relatives of victims. Assays for predicting EVCs such as eye, hair and skin colour have been developed [192,198–201]. For the DNA-based prediction of eye colour, the developmentally validated IrisPlex system can be used, consisting of a multiplex genotyping assay and a prediction model based on genotype and phenotype data from multiple European populations [202]. A newer developed HIrisPlex system has also been introduced, capable of simultaneously predicting both hair and eye colour [192,200]. The IrisPlex system has been vigorously assessed at the international level in the context of a collaborative exercise involving 21 laboratories overseen by the European DNA Profiling (EDNAP) Group of the International Society of Forensic Genetics (ISFG), which demonstrated the reproducibility and robustness of the IrisPlex assay, and accuracy of the IrisPlex model, with simple implementation [203]. Some limitations exist when dealing with some non-European populations [204]. Success of the HIrisPlex system has been demonstrated in the application to World War II skeletal remains, and highlights the potential value in missing person and DVI cases, especially in the ability to provide leads for locating putative relatives [205]. EVCs panels have continued to develop to also provide information on skin colour [204,206] and pigmentation, culminating in the newly introduced HIrisPlex-S system [201]. AISNPs are genetic polymorphisms that exhibit large allele frequency divergences among major global populations which capture the genetic distances between them [186,207,208]. There are many custom assays that have been developed for predicting BGA and SNPs from some of these have been assembled into two proprietary massively parallel sequencing (MPS) panels: the ForenSeq™ Signature Prep Kit (Illumina®) [209] which utilises the Kidd panel [210] together with STRs, identity SNPs and the IrisPlex panel; and the Precision ID Ancestry Panel (TFS) [211–213] which is made up of both the Kidd and Seldin [193] panels with 13 overlapping SNPs. Both of these panels will require the use of appropriate reference populations to accurately infer

5. Genetic markers 5.1. Current approaches 5.1.1. Autosomal short tandem repeats Autosomal STRs are the most frequently employed markers for genetic identification and are characterised by high values of polymorphic information content [2] and have a high power of discrimination [169]. As a result, STR analysis has proven to be a powerful tool in many mass fatality incidents since the 1990s; often in isolation [2]. While the shortcomings of nine-loci STR multiplexes (e.g. AmpFlSTR® Profiler Plus™ (TFS)) for kinship matching have been highlighted in DVI [170], modern STR multiplexes such as the PowerPlex® 21 System (Promega) and GlobalFiler® (TFS), continue to increase the power of discrimination with the addition of more STR loci. Furthermore, multiplexes can be applied to automated platforms and can be modified to alter the number of cycles, reaction volumes and input amount of DNA depending on the equipment used and type of samples encountered in the laboratory [159,171,172]. 5.1.2. Y chromosome STRs and X chromosome STRs Y chromosome STRs (Y-STRs) are useful when establishing paternal lineages as Y-STR profiles are expected to remain the same along a patrilineage for slowly mutating Y-STRs [173]. It is generally necessary to supplement the analysis of autosomal STRs with X chromosome STRs (X-STRs) or Y-STRs to achieve extra discrimination when using siblings as reference samples for DNA identification. X-STRs can be useful as they are highly polymorphic, and can meet Hardy Weinberg and linkage expectations if not within the same linkage group [174–176]. 5.1.3. Mitochondrial DNA hypervariable regions I and II mtDNA is of great advantage in forensic testing due to its high copy number per cell [177] making sequencing of mitochondrial DNA hypervariable regions I and II (HVI/HVII) particularly useful in missing persons and unidentified remains cases when nDNA is not available [178]. While nDNA is present in two copies per diploid cell, a mature oocyte is estimated to have thousands of mitochondria and greater than 100,000 copies of mtDNA [179,180]. The circular nature of the molecules also makes mtDNA more resistant to degradation [181]. In highly degraded specimens, the use of mini-primer sets can improve the chances of typing success; the amplification strategy consisting of eight overlapping short amplicons that span the HVI/II regions and range from 126 to 170 base pairs (bp) [182]. mtDNA molecules are inherited maternally [177,183]. This characteristic is valuable in closed disasters where victims are thought to be unrelated [15]. Here mtDNA testing can confirm or refute maternal relatedness between individuals. This application also allows reference 6

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

degraded DNA samples. As neutral genetic loci that are poorly transcribed and identical by descent only (with no mechanism for parallel independent insertions to occur), they are ideal for kinship analysis of degraded human remains. Bi-allelic INNULs in the InnoTyper® 21 Kit offer a sensitivity and power of discrimination that makes them useful for human identification of extremely degraded samples [238].

ancestry in an operational context [210,214]. 5.2.2. Microhaplotypes Multiple haplotypes exist as a consequence of the origins of variants at different sites, rare recombinants, and the unpredictability of genetic drift and/or selection [215]. Clusters of tightly linked SNPs as novel forensic markers have been proposed including haploblocks [216], mini-haplotypes [217] and microhaplotypes [185,215]. The latter consist of regions encompassing two or more SNPs within an extent of < 200 bp that define multiallelic haplotype loci [185,215]. A panel of microhaplotype loci with high regional or global FST (a measure of genetic distance between population groups) should be able to provide information on global ancestry and on lineage-familial relationships [185]. As well as ancestry assignment, microhaplotype loci could also be used for individual identification, kinship analyses, determining family/clan relationships and the identification and deconvolution of mixtures [185,215,218,219]. They offer the advantages of SNPs for degraded DNA but overcome one of the disadvantages of SNPs: the inability to detect mixtures as a result of their generally bi-allelic nature. Haplotyped SNPs allow more efficient inference of family relationships on a per locus basis because they constitute multiallelic loci [215,217]. These closely linked haplotype systems could offer an optimal marker for family or lineage inference [185,217,220,221]. As more microhaplotypes are identified and appropriate population databases constructed, a microhaplotype panel could potentially outperform existing SNP panels for individual identification, ancestry inference, familial information and the ability to detect and deconvolute mixtures [185]. Approximately 130 microhaplotypes have now been identified as well as their estimated haplotype (allele) frequencies in 83 different population samples [222].

6. Detection methods 6.1. Current approach 6.1.1. Capillary electrophoresis Current DNA profiling is achieved by CE. The selectivity of the DNA fragments is based on charge-to-size ratio. Only a relatively small amount of sample is required for injection, and separation in the capillaries is achieved in minutes due to the high voltages that are permitted with efficient heat dissipation from capillaries [239]. CE platforms can be used for both fragment length analysis (e.g. STR genotyping) and sequencing applications (e.g. Sanger sequencing of mtDNA HVI and HVII and SNaPshot® (TFS) SNP assays). SNaPshot® is a single-base extension (SBE) assay minisequencing method that is commonly applied to forensic DNA due to its sensitivity and high multiplexing ability [189,240]. SNaPshot® assays incorporate all SNP classes and are also expected to be able to incorporate other EVCs of potential forensic value in the future. Application of SNP panels to MPS platforms will aid this endeavour yet SNaPshot® is a low-cost and time-efficient alternative to MPS for smaller scale genotyping requirements [189]. Analysis of mtDNA has focused on HVI and HVII due to a high concentration of variants in these regions. However, > 70% of the total variation within the whole mitochondrial DNA genome (mtGenome) has been reported to exist outside of HVI and HVII [241], meaning sequencing the whole mitochondrial genome can provide far greater resolving power for human identification [242,243]. PCR-based sequencing approaches for mitochondrial HVI and HVII typically amplify two to 12 overlapping fragments of approximately 150 bp to 600 bp in length [244–247] as well as consuming large amounts of DNA extract and being template-length dependent [248]. To sequence the whole mtGenome using PCR-based sequencing would require hundreds of overlapping amplicons. Moreover, Sanger sequencing beyond the control region is laborious, expensive and time consuming [249,250].

5.2.3. Insertion-deletion markers Insertion-deletion length polymorphisms (InDels) are a type of biallelic short DNA length variation [223–227]. InDels are the second most abundant polymorphism in humans and have been shown to be a useful alternative to STRs [226,228–232]. InDels share many of the favourable characteristics of SNPs that make them ideal for the analysis of degraded DNA including short amplicon ranges, high multiplexing capability and low mutation rates [233,234]. In highly multiplexed assays, where dye-labelled PCR products can be separated and detected by CE, InDels offer a promising development for the analysis of degraded DNA [233,234]. When typing severely compromised remains (with the expectation of recovering incomplete STR profiles), Fondevila and colleagues [233,235] suggest the typing of approximately 60 InDels in two simple and robust multiplexes has considerable potential for the genetic identification of human remains. The Investigator DIPplex® kit (QIAGEN) has been shown to provide a powerful stand-alone or supplement capability for identity testing [230].

6.2. Emerging approach 6.2.1. Massively parallel sequencing MPS or next generation sequencing (NGS) refers to the next generation of post-Sanger sequencing technologies. MPS allows for a substantial increase in throughput of a large number of genetic targets and samples with significant depth of coverage at a relatively affordable price [243]. As the next generation of sequencing, these new methods rely on the preparation of MPS libraries in a cell free system. Many millions of sequencing reactions are produced in parallel (instead of up to 96 with CE), and the sequencing output is detected directly where base interrogation is performed cyclically and in parallel [251]. With its ability to type large batteries of markers in multiple samples simultaneously, MPS is increasingly being applied within a forensic context [234,243,252–258]. Existing forensic STR polymorphisms are more informative when genotyping is carried out by MPS because previously undetectable SNPs can be identified both within STR regions and in flanking regions. MPS also makes it possible to sequence a cluster of SNPs in phase, thus enabling the genotyping of microhaplotypes [185,215,218,219]. STR polymorphisms and SNP panels can be multiplexed to add information on both ancestry and phenotype to the identification information from STR markers when typed by MPS [185,189,215,218]. InDels can also be analysed on an MPS platform where they can be defined better and

5.2.4. Retrotransposable elements/short interspersed nuclear elements/ insertion and null alleles Retrotransposable elements (REs), consisting of long interspersed nuclear elements (LINEs) and short interspersed nuclear elements (SINEs), are a group of markers that can be useful for human identity testing [236,237]. SINEs are non-coding genomic DNA repeat sequences, or mobile insertion elements, comprising approximately 40% of the human genome [238]. Due to inherent size differences associated with insertion and null alleles (INNULs), the use of REs for facilitated population studies had not been pursued until a new novel primer design facilitated INNUL marker testing [236]. Markers included in the InnoTyper® 21 Kit (InnoGenomics®) are bi-allelic, having two possible allelic states (insertion or null). Alu elements are primate specific SINEs that have reached a copy number in excess of one million in the human genome, which makes them highly sensitive and desirable for extremely 7

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

offer the potential to identify proximal SNPs that can increase the power of discrimination of currently defined InDels [259]. MPS offers a practical solution for the routine processing of mtGenomes, providing the highest level of maternal lineage discrimination. MPS of high quality mtDNA can be achieved by using longrange PCR to generate two amplicons of approximately 8 kilobases (kb) in length [243,260]. The primer pairs generate overlapping amplicons of ∼8.3 and ∼8.6 kb [261] to allow sequencing of the whole 16,569 bp mtGenome. The general workflow for library preparation includes tagmentation (enzymatic fragmentation) of long-range PCR products, PCR amplification, clean up, quantification, pooling and sequencing [243,262]. For degraded samples, PCR amplification of large fragments (several kb) may fail due to mtDNA fragmentation. An MPS tiling approach for simultaneous mtGenome sequencing using 161 short overlapping amplicons (average 200 bp) is available for such samples [263]. Commercial kits are now available using this approach, namely the Precision ID mtDNA Whole Genome Panel (TFS), which is currently undergoing developmental validation [264]. mtGenome sequencing using MPS also offers a number of other advantages for forensic testing. Just et al. [265] describe the use of MPS technologies to detect mtDNA point heteroplasmy. Resolution of mtDNA heteroplasmy is now possible when applying MPS to minor components down to 1% [266,267]. MPS can also detect the presence of damage-induced lesions [267] and resolve length heteroplasmy [268–270]. This also allows the resolution of mixture components to be vastly improved [268,271,272]. Multiple rounds of in-solution hybridisation-based DNA capture can retrieve whole mtGenome sequences from the most challenging samples, including those of a highly degraded nature [248,273]. Originating from ancient DNA studies to routinely generate complete mtGenomes [241,274–276], this DNA capture strategy is based on the hybridisation of target DNA sequences to probes that are immobilised in solution or on a surface [277,278]. Primer extension capture methods also target smaller mtDNA fragment sizes and have been found to produce reliable and plausible mtDNA haplotypes unable to be recovered with Sanger-type sequencing and other MPS techniques [279]. In a forensic context, repeating the enrichment process on samples with very low amounts of mtDNA can substantially improve the overall coverage of the mtDNA genome by more than doubling the number of unique reads and average coverage [248]. The methodology has the ability to capture DNA templates that are damaged and fragmented (< 100 bp in length) and that are difficult to recover using traditional methods [280]. DNA capture approaches have been shown to selectively enrich short endogenous DNA templates over longer exogenous contaminant DNA [281], an obvious advantage for compromised samples contaminated with large amounts of exogenous DNA [248]. A probe capture assay targeting forensically relevant nuclear SNP markers (375 SNPs and 36 microhaplotype markers) has also been designed with potential application to degraded and mixed DNA samples [282].

because pigs take a quadrupedal stance, it is not possible to bury pigs supine [289]. Pigs have been used extensively in entomological and thanatochemistry studies [290,291] as well as taphonomic investigations regarding different burial environments and climates [292–296]. The first human taphonomic facility was opened in 1980 in Tennessee and was colloquially known as the ‘body farm’. It was not until 2007 that the second such facility was established, with a total of six facilities currently being operated in the United States [297]. These scientific research facilities are traditionally anthropological and use human cadavers to focus on taphonomy with several forensic viewpoints including anthropology, entomology, pathology and biology. In 2016, the Australian Facility for Taphonomic Experimental Research opened, which is the first taphonomic facility in the southern hemisphere. Not only do such facilities negate the need for human analogues, they provide an accurate and precise mechanism by which to expose bodies to real life environmental insults. Where historically human analogues have been used, and biological samples were degraded artificially (e.g. heat, UV, etc.), a more realistic alternative is now offered. A plethora of studies in the forensic community are using samples from such facilities with an invaluable ability to produce the most accurate and appropriate data [24,163,167]. As more facilities open across the world, the possibility of test environments with the appropriate climate, regional and seasonal differences, and faunal assemblages, can be applied to a range of DVI scenarios.

7. Taphonomic facilities for conducting DVI research

Declarations of interest

8. Conclusions Current developments have changed traditional thinking with regards to components implicit to DNA-based identification. For example, following success of DNA recovery from a new range of biological samples that can be collected with minimally or non-invasive procedures, perhaps now is the time to move past the traditional sample types and collection techniques. Additionally, in the case of degraded samples, the forensic scientist now has a number of genetic markers available and means by which to recover available DNA. DNA intelligence tools also means DVI does not rely strictly on comparative grounds with AM data and a candidate pool of victims can be refined. The exploration of new sources of DNA, and the development of genetic markers and technologies continues to offer an increasing number of tools for DNA-based DVI. At this time, it would be pertinent to revise international DVI standards and guidelines to reflect current thinking about sample selection, collection, preservation and DNA profiling procedures to ensure best practice is being applied globally for DNAbased DVI efforts. Will the future of DNA-based DVI resemble a toenail clipping being submitted for direct-to-PCR processing on an automated STR platform with subsequent comparison to AM samples, proceeded by the option to conduct FDP on an MPS platform to generate a physical profile of the deceased for investigative purposes?

None.

Current research of relevance for DVI covers the different stages of the DVI process including sample selection, collection and preservation, through to successful profiling of severely compromised remains. Even more recently, research has emerged into DNA intelligence that can be offered in the identification of victims or relatives of victims. One development that ensures the most relevant samples are used for this work is the establishment of taphonomic facilities. Prior to taphonomic facilities using human subjects, pig carcasses were used as standard human body analogues [283,284]. Pig carcasses are largely hairless and have a similar body mass, skin structure, fat: muscle ratio and physiology to humans [285–287]. However, the bones of pigs have a different microstructure to that of humans [288]. The limbs of pigs are proportionately much shorter than human limbs and,

Acknowledgements The authors thank two anonymous reviewers for their constructive criticisms which contributed to the final manuscript. References [1] INTERPOL, Disaster Victim Identification (Amended) Guide, Interpol, Lyon, 2014. [2] E. Ziętkiewicz, M. Witt, P. Daca, J. Zebracka-Gala, M. Goniewicz, B. Jarząb, M. Witt, Current genetic methodologies in the identification of disaster victims and in forensic analysis, J. Appl. Genet. 53 (1) (2012) 41–60.

8

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

[33] T.R. Schwartz, E.A. Schwartz, L. Mieszerski, L. McNally, L. Kobilinsky, Characterization of deoxyribonucleic acid (DNA) obtained from teeth subjected to various environmental conditions, J. Forensic Sci. 36 (4) (1991) 979–990. [34] A. Alvarez García, I. Muñoz, C. Pestoni, M.V. Lareu, M.S. Rodríguez-Calvo, A. Carracedo, Effect of environmental factors on PCR-DNA analysis from dental pulp, Int. J. Leg. Med. 109 (3) (1996) 125–129. [35] R. Gaytmenn, D. Sweet, Quantification of forensic DNA from various regions of human teeth, J. Forensic Sci. 48 (3) (2003) 622–625. [36] R.C. Dobberstein, J. Huppertz, N. von Wurmb-Schwark, S. Ritz-Timme, Degradation of biomolecules in artificially and naturally aged teeth: implications for age estimation based on aspartic acid racemization and DNA analysis, Forensic Sci. Int. 179 (2-3) (2008) 181–191. [37] D. Higgins, J. Kaidonis, J. Austin, G. Townsend, H. James, T. Hughes, Dentine and cementum as sources of nuclear DNA for use in human identification, Aust. J. Forensic Sci. 43 (2011) 287–295. [38] D. Higgins, J.J. Austin, Teeth as a source of DNA for forensic identification of human remains: a review, Sci. Justice 53 (4) (2013) 433–441. [39] P.C. Malaver, J.J. Yunis, Different dental tissues as source of DNA for human identification in forensic cases, Croat. Med. J. 44 (3) (2003) 306–309. [40] D. De Leo, S. Turrina, M. Marigo, Effects of individual dental factors on genomic DNA analysis, Am. J. Forensic Med. Pathol. 21 (4) (2000) 411–415. [41] L. Rubio, L.J. Martinez, E. Martinez, S. Martin de las Heras, Study of short- and long-term storage of teeth and its influence on DNA, J. Forensic Sci. 54 (6) (2009) 1411–1413. [42] C.L. Parks, A study of the human decomposition sequence in central Texas, J. Forensic Sci. 56 (1) (2011) 19–22. [43] D. McNevin, L. Wilson-Wilde, J. Robertson, J. Kyd, C. Lennard, Short tandem repeat (STR) genotyping of keratinized hair: part 1. Review of current status and knowledge gaps, Forensic Sci. Int. 153 (2) (2005) 237–246. [44] D. McNevin, L. Wilson-Wilde, J. Robertson, J. Kyd, C. Lennard, Short tandem repeat (STR) genotyping of keratinized hair: part 2. An optimized genomic DNA extraction procedure reveals donor dependence of STR profiles, Forensic Sci. Int. 153 (2) (2005) 247–259. [45] E.A. Graham, DNA reviews: hair, Forensic Sci. Med. Pathol. 3 (2) (2007) 133–137. [46] M.A. Tahir, N. Watson, Typing of DNA HLA-DQ alpha alleles extracted from human nail material using polymerase chain reaction, J. Forensic Sci. 40 (4) (1995) 634–636. [47] A. Nakanishi, F. Moriya, Y. Hashimoto, Effects of environmental conditions to which nails are exposed on DNA analysis of them, Leg. Med. (Tokyo) 5 (Suppl. 1) (2003) S194–7. [48] M. Allouche, M. Hamdoum, P. Mangin, V. Castella, Genetic identification of decomposed cadavers using nails as DNA source, Forensic Sci. Int. Genet. 3 (1) (2008) 46–49. [49] R. Ottens, D. Taylor, A. Linacre, DNA profiles from fingernails using direct PCR, Forensic Sci. Med. Pathol. 11 (1) (2015) 99–103. [50] A. Piccinini, N. Cucurachi, F. Betti, M. Capra, S. Coco, et al., Forensic DNA typing of human nails at various stages of decomposition, Int. Congr. Ser. 1288 (2006) 586–588. [51] S. Harbison, S. Petricevic, S. Vintiner, The persistence of DNA under fingernails following submersion in water, Int. Congr. Ser. 1239 (2003) 809–813. [52] A. Schlenker, K. Grimble, A. Azim, R. Owen, D. Hartman, Toenails as an alternative source material for the extraction of DNA from decomposed human remains, Forensic Sci. Int. 258 (2016) 1–10. [53] A. Roeper, W. Reichert, R. Mattern, The Achilles tendon as a DNA source for STR typing of highly decayed corpses, Forensic Sci. Int. 173 (2–3) (2007) 103–106. [54] M. van den Berge, D. Wiskerke, R.R. Gerretsen, J. Tabak, T. Sijen, DNA and RNA profiling of excavated human remains with varying postmortem intervals, Int. J. Leg. Med. 130 (6) (2016) 1471–1480. [55] A.Z. Mundorff, S. Amory, R. Huel, A. Bilić, A.L. Scott, T.J. Parsons, An economical and efficient method for postmortem DNA sampling in mass fatalities, Forensic Sci. Int. Genet. 36 (2018) 167–175. [56] S. Ferreira, R. Garrido, K. Paula, R. Nogueira, E. Oliveira, A. Moraes, Cartilage and phalanges from hallux: alternative sources of samples for DNA typing in disaster victim identification (DVI). A comparative study, Forensic Sci. Int. Genet. Suppl. Ser. 4 (2013) e366–e367. [57] J. Fredericks, K. Brown, A. Williams, P. Bennett, DNA analysis of skeletal tissue recovered from the English Channel, J. Forensic Leg. Med. 20 (6) (2013) 757–759. [58] H.H. de Boer, G.J.R. Maat, D.A. Kadarmo, P.T. Widodo, A.D. Kloosterman, A.J. Kal, DNA identification of human remains in Disaster Victim Identification (DVI): an efficient sampling method for muscle, bone, bone marrow and teeth, Forensic Sci. Int. 289 (2018) 253–259. [59] T. Schwark, J.H. Modrow, E. Steinmeier, M. Poetsch, J. Hasse, H. Fischer, N. von Wurmb-Schwark, The auditory ossicles as a DNA source for genetic identification of highly putrefied cadavers, Int. J. Leg. Med. 129 (3) (2015) 457–462. [60] T. Siriboonpiputtana, T. Rinthachai, J. Shotivaranon, V. Peonim, B. Rerkamnuaychoke, Forensic genetic analysis of bone remain samples, Forensic Sci. Int. 284 (2018) 167–175. [61] S. Edson, A. Christensen, S. Barritt, A. Meehan, M. Leney, L. Finelli, Sampling of the cranium for mitochondrial DNA analysis of human skeletal remains, Forensic Sci. Int. Genet. Suppl. Ser. 2 (2009) 269–270. [62] G. Kulstein, T. Hadrys, P. Wiegand, As solid as a rock-comparison of CE- and MPSbased analyses of the petrosal bone as a source of DNA for forensic identification of challenging cranial bones, Int. J. Leg. Med. 132 (1) (2018) 13–24. [63] C. Giannakis, I.J. Forbes, P.D. Zalewski, Ca2+/Mg(2+)-dependent nuclease: tissue distribution, relationship to inter-nucleosomal DNA fragmentation and inhibition by Zn2+, Biochem. Biophys. Res. Commun. 181 (2) (1991) 915–920.

[3] A. Alonso, P. Martin, C. Albarrán, P. Garcia, L. Fernandez de Simon, M. Jesús Iturralde, et al., Challenges of DNA profiling in mass disaster investigations, Croat. Med. J. 46 (4) (2005) 540–548. [4] R. Lessig, M. Rothschild, International standards in cases of mass Disaster Victim Identification (DVI), Forensic Sci. Med. Pathol. 8 (2) (2012) 197–199. [5] I.T.E.W. Group, The DVI Response to the South East Asian Tsunami between December 2004 and February 2006, (2010). [6] INTERPOL, 2018. [7] M. Prinz, A. Carracedo, W.R. Mayr, N. Morling, T.J. Parsons, A. Sajantila, et al., DNA Commission of the International Society for Forensic Genetics (ISFG): recommendations regarding the role of forensic genetics for disaster victim identification (DVI), Forensic Sci. Int. Genet. 1 (1) (2007) 3–12. [8] D. Sweet, INTERPOL DVI best-practice standards–an overview, Forensic Sci. Int. 201 (1–3) (2010) 18–21. [9] W.H. Goodwin, The use of forensic DNA analysis in humanitarian forensic action: the development of a set of international standards, Forensic Sci. Int. 278 (2017) 221–227. [10] J. Lee, P. Scott, D. Carroll, C. Eckhoff, S. Harbison, V. Ientile, et al., Recommendations for DNA laboratories supporting disaster victim identification (DVI) operations–Australian and New Zealand consensus on ISFG recommendations, Forensic Sci. Int. Genet. 3 (1) (2008) 54–56. [11] U.D. Immel, S. Lutz-Bonengel, R. Lessig, Quality exercise in disaster victim identification, Forensic Sci. Int. Genet. Suppl. Ser. 5 (2015) e202–e203. [12] C.M. Vullo, M. Romero, L. Catelli, M. Šakić, V.G. Saragoni, M.J. Jimenez Pleguezuelos, et al., GHEP-ISFG collaborative simulated exercise for DVI/MPI: lessons learned about large-scale profile database comparisons, Forensic Sci. Int. Genet. 21 (2016) 45–53. [13] L.A. Dixon, A.E. Dobbins, H.K. Pulker, J.M. Butler, P.M. Vallone, M.D. Coble, et al., Analysis of artificially degraded DNA using STRs and SNPs–results of a collaborative European (EDNAP) exercise, Forensic Sci. Int. 164 (1) (2006) 33–44. [14] K. Shintani-Ishida, K. Harada, M. Nakajima, K. Yoshida, Usefulness of blood vessels as a DNA source for PCR-based genotyping based on two cases of corpse dismemberment, Leg. Med. (Tokyo) 12 (1) (2010) 8–12. [15] K. Montelius, B. Lindblom, DNA analysis in disaster victim identification, Forensic Sci. Med. Pathol. 8 (2) (2012) 140–147. [16] G.C. Calacal, D.L.T. Apaga, J.M. Salvador, J.A.D. Jimenez, L.J. Lagat, R.P.F. Villacorta, et al., Comparing different post-mortem human samples as DNA sources for downstream genotyping and identification, Forensic Sci. Int. Genet. 19 (2015) 212–220. [17] B. Rerkamnuaychoke, W. Chantratita, U. Jomsawat, J. Thanakitgosate, N. Pattanasak, P. Rojanasunan, Comparison of DNA extraction from blood stain and decomposed muscle in STR polymorphism analysis, J. Med. Assoc. Thai. 83 (Suppl. 1) (2000) S82–88. [18] H.J. Meyer, The Kaprun cable car fire disaster–aspects of forensic organisation following a mass fatality with 155 victims, Forensic Sci. Int. 138 (1–3) (2003) 1–7. [19] A. Trindade-Filho, C. Mendes, S. Ferreira, S. Oliveira, A. Vasconcelos, F. Maia, et al., DNA obtained from decomposed corpses cartilage: a comparison with skeleton muscle source, Forensic Sci. Int. Genet. Suppl. Ser. 1 (2008) 459–461. [20] R. Owen, P. Bedford, J. Leditschke, A. Schlenker, D. Hartman, Post mortem sampling of the bladder for the identification of victims of fire related deaths, Forensic Sci. Int. 233 (1–3) (2013) 14–20. [21] B.C. Manjunath, B.R. Chandrashekar, M. Mahesh, R.M. Vatchala Rani, DNA profiling and forensic dentistry–a review of the recent concepts and trends, J. Forensic Leg. Med. 18 (5) (2011) 191–197. [22] S. Andelinović, D. Sutlović, I. Erceg Ivkosić, V. Skaro, A. Ivkosić, F. Paić, et al., Twelve-year experience in identification of skeletal remains from mass graves, Croat. Med. J. 46 (4) (2005) 530–539. [23] A. Milos, A. Selmanović, L. Smajlović, R.L. Huel, C. Katzmarzyk, A. Rizvić, T.J. Parsons, Success rates of nuclear short tandem repeat typing from different skeletal elements, Croat. Med. J. 48 (4) (2007) 486–493. [24] A. Mundorff, J.M. Davoren, Examination of DNA yield rates for different skeletal elements at increasing post mortem intervals, Forensic Sci. Int. Genet. 8 (1) (2014) 55–63. [25] W. Parson, A. Brandstätter, A. Alonso, N. Brandt, B. Brinkmann, A. Carracedo, et al., The EDNAP mitochondrial DNA population database (EMPOP) collaborative exercises: organisation, results and perspectives, Forensic Sci. Int. 139 (2–3) (2004) 215–226. [26] W. Parson, H.J. Bandelt, Extended guidelines for mtDNA typing of population data in forensic science, Forensic Sci. Int. Genet. 1 (1) (2007) 13–19. [27] M. Nilsson, H. Andréasson-Jansson, M. Ingman, M. Allen, Evaluation of mitochondrial DNA coding region assays for increased discrimination in forensic analysis, Forensic Sci. Int. Genet. 2 (1) (2008) 1–8. [28] J.P. Orgel, A. Miller, T.C. Irving, R.F. Fischetti, A.P. Hammersley, T.J. Wess, The in situ supermolecular structure of type I collagen, Structure 9 (11) (2001) 1061–1069. [29] P.F. Campos, O.E. Craig, G. Turner-Walker, E. Peacock, E. Willerslev, M.T. Gilbert, DNA in ancient bone – where is it located and how should we extract it? Ann. Anat. 194 (1) (2012) 7–16. [30] O.M. Loreille, T.M. Diegoli, J.A. Irwin, M.D. Coble, T.J. Parsons, High efficiency DNA extraction from bone by total demineralization, Forensic Sci. Int. Genet. 1 (2) (2007) 191–195. [31] ICMP, Standard Operating Procedure for Sampling Bone and Tooth Specimens From Human Remains for DNA Testing at the ICMP, ICMP, 2015. [32] A.Z. Mundorff, E.J. Bartelink, E. Mar-Cash, DNA preservation in skeletal elements from the World Trade Center disaster: recommendations for mass fatality management, J. Forensic Sci. 54 (4) (2009) 739–745.

9

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

[97] C. Lalueza, A. Pérez-Pérez, E. Prats, L. Cornudella, D. Turbón, Lack of founding Amerindian mitochondrial DNA lineages in extinct aborigines from Tierra del Fuego-Patagonia, Hum. Mol. Genet. 6 (1) (1997) 41–46. [98] R. Montiel, A. Malgosa, P. Francalacci, Authenticating ancient human mitochondrial DNA, Hum. Biol. 73 (5) (2001) 689–713. [99] J. Burger, S. Hummel, B. Hermann, W. Henke, DNA preservation: a microsatelliteDNA study on ancient skeletal remains, Electrophoresis 20 (8) (1999) 1722–1728. [100] A. González-Oliver, L. Márquez-Morfín, J.C. Jiménez, A. Torre-Blanco, Founding Amerindian mitochondrial DNA lineages in ancient maya from Xcaret, Quintana roo, Am. J. Phys. Anthropol. 116 (3) (2001) 230–235. [101] J. Ward, Best practice recommendations for the establishment of a national DNA identification program for missing persons: a global perspective, Forensic Sci. Int. Genet. Suppl. Ser. 6 (2017) e43–e45. [102] J.L. Barta, C. Monroe, B.M. Kemp, Further evaluation of the efficacy of contamination removal from bone surfaces, Forensic Sci. Int. 231 (1–3) (2013) 340–348. [103] H. Malmström, J. Storå, L. Dalén, G. Holmlund, A. Götherström, Extensive human DNA contamination in extracts from ancient dog bones and teeth, Mol. Biol. Evol. 22 (10) (2005) 2040–2047. [104] H. Malmström, E.M. Svensson, M.T. Gilbert, E. Willerslev, A. Götherström, G. Holmlund, More on contamination: the use of asymmetric molecular behavior to identify authentic ancient human DNA, Mol. Biol. Evol. 24 (4) (2007) 998–1004. [105] K. Watt, Decontamination Techniques in Ancient DNA Analysis, Simon Fraser University, British Columbia, 2005. [106] J. Dissing, M. Kristinsdottir, C. Friis, On the elimination of extraneous DNA in fossil human teeth with hypochlorite, J. Archaeol. Sci. 35 (6) (2008) 1445–1452. [107] H. Hayatsu, S. Pan, T. Ukita, Reaction of sodium hypochlorite with nucleic acids and their constituents, Chem. Pharm. Bull. (Tokyo) 19 (10) (1971) 2189–2192. [108] M. Whiteman, A. Jenner, B. Halliwell, Hypochlorous acid-induced base modifications in isolated calf thymus DNA, Chem. Res. Toxicol. 10 (11) (1997) 1240–1246. [109] S. Ohnishi, M. Murata, S. Kawanishi, DNA damage induced by hypochlorite and hypobromite with reference to inflammation-associated carcinogenesis, Cancer Lett. 178 (1) (2002) 37–42. [110] M. Whiteman, H. Hong, A. Jenner, B. Halliwell, Loss of oxidized and chlorinated bases in DNA treated with reaction oxygen species: implications for assessment of oxidative damage in vivo, Biochem. Biophys. Res. Commun. 296 (2002) 883–889. [111] R. Alaeddini, Forensic implications of PCR inhibition–a review, Forensic Sci. Int. Genet. 6 (3) (2012) 297–305. [112] S. Amory, R. Huel, A. Bilić, O. Loreille, T.J. Parsons, Automatable full demineralization DNA extraction procedure from degraded skeletal remains, Forensic Sci. Int. Genet. 6 (3) (2012) 398–406. [113] M. Scholz, I. Giddings, C.M. Pusch, A polymerase chain reaction inhibitor of ancient hard and soft tissue DNA extracts is determined as human collagen type I, Anal. Biochem. 259 (2) (1998) 283–286. [114] D.L. Fisher, M.M. Holland, L. Mitchell, P.S. Sledzik, A.W. Wilcox, M. Wadhams, V.W. Weedn, Extraction, evaluation, and amplification of DNA from decalcified and undecalcified United States Civil War bone, J. Forensic Sci. 38 (1) (1993) 60–68. [115] E. Sørensen, S.H. Hansen, B. Eriksen, N. Morling, Application of thiopropyl sepharose 6B for removal of PCR inhibitors from DNA extracts of a thigh bone recovered from the sea, Int. J. Leg. Med. 117 (4) (2003) 245–247. [116] M. Evison, D. Smillie, A. Chamberlain, Extraction of single-copy nuclear DNA from forensic specimens with a variety of post-mortem histories, J. Forensic Sci. 42 (6) (1997) 1032–1038. [117] C.C. Tebbe, W. Vahjen, Interference of humic acids and DNA extracted directly from soil in detection and transformation of recombinant DNA from bacteria and a yeast, Appl. Environ. Microbiol. 59 (8) (1993) 2657–2665. [118] Y.L. Tsai, B.H. Olson, Rapid method for separation of bacterial DNA from humic substances in sediments for polymerase chain reaction, Appl. Environ. Microbiol. 58 (7) (1992) 2292–2295. [119] C.A. Kreader, Relief of amplification inhibition in PCR with bovine serum albumin or T4 gene 32 protein, Appl. Environ. Microbiol. 62 (3) (1996) 1102–1106. [120] D. Sutlovic, S. Gamulin, M. Definis-Gojanovic, D. Gugic, S. Andjelinovic, Interaction of humic acids with human DNA: proposed mechanisms and kinetics, Electrophoresis 29 (7) (2008) 1467–1472. [121] P. Zimmermann, K. Vollack, B. Haak, M. Bretthauer, A. Jelinski, M. Kugler, et al., Adaptation and evaluation of the PrepFiler™ DNA extraction technology in an automated forensic DNA analysis process with emphasis on DNA yield, inhibitor removal and contamination security, Forensic Sci. Int. Genet. Suppl. Ser. 2 (1) (2009) 62–63. [122] K. Sturk-Andreaggi, T. Diegoli, R. Just, J. Irwin, Evaluation of automatable silicabased extraction methods for low quantity samples, Forensic Sci. Int. Genet. 3 (1) (2011) e504–e505. [123] M. Kuś, A. Ossowski, G. Zielińska, Comparison of three different DNA extraction methods from a highly degraded biological material, J. Forensic Leg. Med. 40 (2016) 47–53. [124] D. Hartman, O. Drummer, C. Eckhoff, J.W. Scheffer, P. Stringer, The contribution of DNA to the disaster victim identification (DVI) effort, Forensic Sci. Int. 205 (1-3) (2011) 52–58. [125] N. Laurin, F. Célestin, M. Clark, D. Wilkinson, B. Yamashita, C. Frégeau, New incompatibilities uncovered using the Promega DNA IQ™ chemistry, Forensic Sci. Int. 257 (2015) 134–141. [126] P.V. Mandrekar, L. Flanagan, A. Tereba, Forensic extraction and isolation of DNA from hair, tissue and bone, Profiles in DNA, (2002).

[64] G. Seutin, B. White, P. Boag, Preservation of avian blood and tissue samples for DNA analysis, Can. J. Zool. 69 (1991) 82–90. [65] K. Muralidharan, C. Wemmer, Transporting and storing field-collected specimens for DNA without refrigeration for subsequent DNA extraction and analysis, Biotechniques 17 (3) (1994) 420–422. [66] L.E. Flournoy, R.P. Adams, R.N. Pandy, Interim and archival preservation of plant specimens in alcohols for DNA studies, Biotechniques 20 (4) (1996) 657–660. [67] M. Grassberger, C. Stein, S. Hanslik, M. Hochmeister, Evalution of a novel tagging and tissue preservation system for potential use in forensic sample collection, Forensic Sci. Int. 151 (2–3) (2005) 233–237. [68] Z. Nagy, A hands-on overview of tissue preservation methods for molecular genetic analyses, Org. Divers. Evol. 10 (1) (2010) 91–105. [69] A. Allen-Hall, D. McNevin, Human tissue preservation for disaster victim identification (DVI) in tropical climates, Forensic Sci. Int. Genet. 6 (5) (2012) 653–657. [70] A. Allen-Hall, D. McNevin, Non-cryogenic forensic tissue preservation in the field: a review, Aust. J. Forensic Sci. 45 (4) (2013) 450–460. [71] M. Caputo, L.A. Bosio, D. Corach, Long-term room temperature preservation of corpse soft tissue: an approach for tissue sample storage, Invest. Genet. 2 (2011) 17. [72] R.B. Stoughton, W. Fritsch, Influence of dimethylsulfoxide (DMSO) on human percutaneous absorption, Arch. Dermatol. 90 (1964) 512–517. [73] C.W. Kilpatrick, Noncryogenic preservation of mammalian tissues for DNA extraction: an assessment of storage methods, Biochem. Genet. 40 (1–2) (2002) 53–62. [74] T. Lindahl, B. Nyberg, Rate of depurination of native deoxyribonucleic acid, Biochemistry 11 (19) (1972) 3610–3618. [75] T. Suzuki, S. Ohsumi, K. Makino, Mechanistic studies on depurination and apurinic site chain breakage in oligodeoxyribonucleotides, Nucleic Acids Res. 22 (23) (1994) 4997–5003. [76] J. Connell, J. Chaseling, M. Page, K. Wright, Tissue preservation in extreme temperatures for rapid response to military deaths, Forensic Sci. Int. Genet. 36 (2018) 86–94. [77] M. Richards, B. Sykes, R. Hedges, Authenticating DNA extracted from ancient skeletal remains, J. Archaeol. Sci. 22 (1995) 291–299. [78] O. Handt, M. Krings, R.H. Ward, S. Pääbo, The retrieval of ancient human DNA sequences, Am. J. Hum. Genet. 59 (2) (1996) 368–376. [79] M. Hofreiter, D. Serre, H.N. Poinar, M. Kuch, S. Pääbo, Ancient DNA, Nat. Rev. Genet. 2 (5) (2001) 353–359. [80] P. Wandeler, S. Smith, P.A. Morin, R.A. Pettifor, S.M. Funk, Patterns of nuclear DNA degeneration over time–a case study in historic teeth samples, Mol. Ecol. 12 (4) (2003) 1087–1093. [81] M. Gilbert, L. Rudbeck, E. Willerslev, A. Hansen, C. Smith, K.E.H. Penkmane, et al., Biochemical and physical correlates of DNA contamination in archaeological human bones and teeth excavated at Matera, Italy, J. Archaeol. Sci. 32 (2005) 783–795. [82] M. Gilbert, A. Hansen, E. Willerslev, G. Turner-Walker, M. Collins, Insights into the processes behind the contamination of degraded human teeth and bone samples with exogenous sources of DNA, Int. J. Ostoarchaeol. 16 (2006) 156–164. [83] N. von Wurmb-Schwark, A. Heinrich, M. Freudenberg, M. Gebührand, T. Schwark, The impact of DNA contamination of bone samples in forensic case analysis and anthropological research, Leg. Med. (Tokyo) 10 (3) (2008) 125–130. [84] J. Currey, Bones: Structure and Mechanics, Princeton University Press, Princeton, 2002. [85] R. Hedges, Bone diagenesis: an overview of processes, Archaeometry 44 (2002) 319–328. [86] M.L. Sampietro, M.T. Gilbert, O. Lao, D. Caramelli, M. Lari, J. Bertranpetit, C. Lalueza-Fox, Tracking down human contamination in ancient human teeth, Mol. Biol. Evol. 23 (9) (2006) 1801–1807. [87] A.A. Westen, R.R. Gerretsen, G.J. Maat, Femur, rib, and tooth sample collection for DNA analysis in disaster victim identification (DVI) : a method to minimize contamination risk, Forensic Sci. Med. Pathol. 4 (1) (2008) 15–21. [88] B.M. Kemp, D.G. Smith, Use of bleach to eliminate contaminating DNA from the surface of bones and teeth, Forensic Sci. Int. 154 (1) (2005) 53–61. [89] S.M. Edson, A.F. Christensen, Field contamination of skeletonized human remains with exogenous DNA, J. Forensic Sci. 58 (1) (2013) 206–209. [90] R.E. Cline, N.M. Laurent, D.R. Foran, The fingernails of Mary Sullivan: developing reliable methods for selectively isolating endogenous and exogenous DNA from evidence, J. Forensic Sci. 48 (2) (2003) 328–333. [91] T. Kalmár, C.Z. Bachrati, A. Marcsik, I. Raskó, A simple and efficient method for PCR amplifiable DNA extraction from ancient bones, Nucleic Acids Res. 28 (12) (2000) E67. [92] E. Meyer, M. Wiese, H. Bruchhaus, M. Claussen, A. Klein, Extraction and amplification of authentic DNA from ancient human remains, Forensic Sci. Int. 113 (1–3) (2000) 87–90. [93] A. Alonso, S. Andelinović, P. Martín, D. Sutlović, I. Erceg, E. Huffine, et al., DNA typing from skeletal remains: evaluation of multiplex and megaplex STR systems on DNA isolated from bone and teeth samples, Croat. Med. J. 42 (3) (2001) 260–266. [94] S.M. Edson, J.P. Ross, M.D. Coble, T.J. Parsons, S.M. Barritt, Naming the dead – confronting the realities of rapid identification of degraded skeletal remains, Forensic Sci. Rev. 16 (1) (2004) 63–90. [95] M. Salamon, N. Tuross, B. Arensburg, S. Weiner, Relatively well preserved DNA is present in the crystal aggregates of fossil bones, Proc. Natl. Acad. Sci. U. S. A. 102 (39) (2005) 13783–13788. [96] C. Lalueza-Fox, Analysis of ancient mitochondrial DNA from extinct Aborigines from Tierra del Fuego-Patagonia, Ancient Biomol. 1 (1996) 43–54.

10

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al. [127] P.V. Mandrekar, B.E. Krenkeand, A. Tereba, DNA IQTM: the intelligent way to purify DNA, Profiles in DNA, (2001). [128] A.M. Tereba, R.M. Bitner, S.C. Koller, C.E. Smith, D.D. Kephartand, S.J. Ekenberg, Simultaneous Isolation and Quantitation of DNA, (2004). [129] C.J. Frégeau, A. De Moors, Competition for DNA binding sites using Promega DNA IQ™ paramagnetic beads, Forensic Sci. Int. Genet. 6 (5) (2012) 511–522. [130] C.J. Frégeau, C.M. Lett, R.M. Fourney, Validation of a DNA IQ-based extraction method for TECAN robotic liquid handling workstations for processing casework, Forensic Sci. Int. Genet. 4 (5) (2010) 292–304. [131] A. Barbaro, P. Cormaci, A. Agostino, Validation of a new magnetic particle-based method (Prepfiler™ Forensic DNA Extraction Kit) for rapid extraction of high quality DNA from a wide variety of forensic samples, Forensic Sci. Int. Genet. Suppl. Ser. 2 (2009) 176–177. [132] S.B. Seo, A. Zhang, H.Y. Kim, J.A. Yi, H.Y. Lee, D.H. Shin, S.D. Lee, Technical note: efficiency of total demineralization and ion-exchange column for DNA extraction from bone, Am. J. Phys. Anthropol. 141 (1) (2010) 158–162. [133] R. Huel, S. Amory, A. Bilić, S. Vidović, E. Jasaragić, T.J. Parsons, DNA extraction from aged skeletal samples for STR typing by capillary electrophoresis, Methods Mol. Biol. 830 (2012) 185–198. [134] E. Hagelberg, J.B. Clegg, Isolation and characterization of DNA from archaeological bone, Proc. Biol. Sci. 244 (1309) (1991) 45–50. [135] N. Rohland, M. Hofreiter, Comparison and optimization of ancient DNA extraction, Biotechniques 42 (3) (2007) 343–352. [136] J. Jakubowska, A. Maciejewska, R. Pawłowski, Comparison of three methods of DNA extraction from human bones with different degrees of degradation, Int. J. Leg. Med. 126 (1) (2012) 173–178. [137] J. Davoren, D. Vanek, R. Konjhodzić, J. Crews, E. Huffine, T.J. Parsons, Highly effective DNA extraction method for nuclear short tandem repeat testing of skeletal remains from mass graves, Croat. Med. J. 48 (4) (2007) 478–485. [138] S.M. Edson, T.P. McMahon, Extraction of DNA from skeletal remains, Methods Mol. Biol. 1420 (2016) 69–87. [139] A. Gotherstrom, M. Collins, A. Angerbjorn, K. Liden, Bone preservation and DNA amplification, Archaeometry 44 (3) (2002) 395–404. [140] I. Zupanič Pajnič, M. Debska, B. Gornjak Pogorelc, K. Vodopivec Mohorčič, J. Balažic, T. Zupanc, et al., Highly efficient automated extraction of DNA from old and contemporary skeletal remains, J. Forensic Leg. Med. 37 (2016) 78–86. [141] M. Caputo, M. Irisarri, E. Alechine, D. Corach, A DNA extraction method of small quantities of bone for high-quality genotyping, Forensic Sci. Int. Genet. 7 (5) (2013) 488–493. [142] D.Y. Yang, B. Eng, J.S. Waye, J.C. Dudar, S.R. Saunders, Technical note: improved DNA extraction from ancient bones using silica-based spin columns, Am. J. Phys. Anthropol. 105 (4) (1998) 539–543. [143] J. Ye, A. Ji, E.J. Parra, X. Zheng, C. Jiang, X. Zhao, et al., A simple and efficient method for extracting DNA from old and burned bone, J. Forensic Sci. 49 (4) (2004) 754–759. [144] B. Kemp, C. Monroe, D. Smith, Repeat silica extraction: a simple technique for the removal of PCR inhibitors from DNA extracts, J. Archaeol. Sci. 33 (12) (2006) 1680–1689. [145] H.Y. Lee, M.J. Park, N.Y. Kim, J.E. Sim, W.I. Yang, K.J. Shin, Simple and highly effective DNA extraction methods from old skeletal remains using silica columns, Forensic Sci. Int. Genet. 4 (5) (2010) 275–280. [146] N. Rohland, H. Siedel, M. Hofreiter, A rapid column-based ancient DNA extraction method for increased sample throughput, Mol. Ecol. Resour. 10 (4) (2010) 677–683. [147] P.L. Marshall, M. Stoljarova, S.E. Schmedes, J.L. King, B. Budowle, A high volume extraction and purification method for recovering DNA from human bone, Forensic Sci. Int. Genet. 12 (2014) 155–160. [148] A.D. Kloosterman, P. Kersbergen, Efficacy and limits of genotyping low copy number (LCN) DNA samples by multiplex PCR of STR loci, J. Soc. Biol. 197 (4) (2003) 351–359. [149] Y.J. Deng, Y.Z. Li, X.G. Yu, L. Li, D.Y. Wu, J. Zhou, et al., Preliminary DNA identification for the tsunami victims in Thailand, Genomics Proteomics Bioinf. 3 (3) (2005) 143–157. [150] R. Schwenzer, A. Schoneberg, W. Pflug, Implementation of a robotized real-time PCR setup for the use of the Quantifiler™ Human DNA Quantification Kit, Forensic Sci. Int. Genet. Suppl. Ser. 1 (2008) 68–70. [151] C. Vieira-Silva, H. Afonso-Costa, T. Ribeiro, M. Porto, M. Dias, A. Amorim, Quantifiler® Trio DNA validation and usefulness in casework samples, Forensic Sci. Int. Genet. Suppl. Ser. 5 (2015) e246–e247. [152] S. Verheij, L. Clarisse, M. van den Bergeand, T. Sijen, RapidHITTM 200, a promising system for rapid DNA analysis, Forensic Sci. Int. Genet. Suppl. Ser. 4 (2013) e254–e255. [153] A.A. Mapes, A.D. Kloosterman, C.J. Poot, V. van Marion, Objective data on DNA success rates can aid the selection process of crime samples for analysis by rapid mobile DNA technologies, Forensic Sci. Int. 264 (2016) 28–33. [154] L.K. Hennessy, H. Franklin, Y. Li, J. Buscaino, K. Chear, J. Gass, et al., Dvelopmental validation studies on the RapidHITTM human DNA identification system, Forensic Sci. Int. Genet. Suppl. Ser. 4 (2013) e7–e8. [155] M. Holland, F. Wendt, Evaluation of the RapidHIT™ 200, an automated human identification system for STR analysis of single source samples, Forensic Sci. Int. Genet. 14 (2015) 76–85. [156] B. Mercier, C. Gaucher, O. Feugeas, C. Mazurier, Direct PCR from whole blood, without DNA extraction, Nucleic Acids Res. 18 (19) (1990) 5908. [157] K. Gray, D. Crowle, P. Scott, Direct amplification of casework bloodstains using the Promega PowerPlex(®) 21 PCR amplification system, Forensic Sci. Int. Genet. 12 (2014) 86–92.

[158] R. Ottens, D. Taylor, D. Abarno, A. Linacre, Successful direct amplification of nuclear markers from a single hair follicle, Forensic Sci. Med. Pathol. 9 (2) (2013) 238–243. [159] R. Ottens, D. Taylor, D. Abarno, A. Linacre, Optimising direct PCR from anagen hair samples, Forensic Sci. Int. Genet. Suppl. Ser. 4 (1) (2013) e109–110. [160] M. Habib, A. Pierre-Noel, F. Fogt, Z. Budimlija, M. Prinz, Direct amplification of biological evidence and DVI samples using the Qiagen Investigator 24plex GO! Kit, Forensic Sci. Int. Genet. Suppl. Ser. 6 (2017) e208–e210. [161] C. Berry, The Use of a Direct to PCR Analysis Method for Disaster Victim Identification, in National Centre for Forensic Science, University of Canberra, Bruce, 2014. [162] J. Kingston, DNA Recovery from Aged Blood Stains, in National Centre for Forensic Science, University of Canberra, Bruce, 2014. [163] A. Sorensen, C. Berry, D. Bruce, M.E. Gahan, S. Hughes-Stamm, D. McNevin, Direct-to-PCR tissue preservation for DNA profiling, Int. J. Leg. Med. 130 (3) (2016) 607–613. [164] H. Altshuler, R. Roy, Evaluation of direct PCR amplification using various swabs and washing reagents, J. Forensic Sci. 60 (6) (2015) 1542–1552. [165] J.E. Templeton, D. Taylor, O. Handt, P. Skuza, A. Linacre, Direct PCR improves the recovery of DNA from various substrates, J. Forensic Sci. 60 (6) (2015) 1558–1562. [166] J. Templeton, R. Ottens, V. Paradiso, O. Handt, D. Taylor, A. Linacre, Genetic profiling from challenging samples: direct PCR of touch DNA, Forensic Sci. Int. Genet. Suppl. Ser. 4 (2013) e224–e225. [167] A. Sorensen, E. Rahman, C. Canela, D. Gangitano, S. Hughes-Stamm, Preservation and rapid purification of DNA from decomposing human tissue samples, Forensic Sci. Int. Genet. 25 (2016) 182–190. [168] A.S. Holmes, M.G. Roman, S. Hughes-Stamm, In-field collection and preservation of decomposing human tissues to facilitate rapid purification and STR typing, Forensic Sci. Int. Genet. 36 (2018) 124–129. [169] R. Chakraborty, K.K. Kidd, The utility of DNA typing in forensic work, Science 254 (5039) (1991) 1735–1739. [170] D. Hartman, L. Benton, L. Morenos, J. Beyer, M. Spiden, A. Stock, Examples of kinship analysis where Profiler Plus™ was not discriminatory enough for the identification of victims using DNA identification, Forensic Sci. Int. 205 (2011) 64–68. [171] J. Edson, E.M. Brooks, C. McLaren, J. Robertson, D. McNevin, A. Cooper, J.J. Austin, A quantitative assessment of a reliable screening technique for the STR analysis of telogen hair roots, Forensic Sci. Int. Genet. 7 (1) (2013) 180–188. [172] D. McNevin, J. Edson, J. Robertson, J.J. Austin, Reduced reaction volumes and increased Taq DNA polymerase concentration improve STR profiling outcomes from a real-world low template DNA source: telogen hairs, Forensic Sci. Med. Pathol. 11 (3) (2015) 326–338. [173] P. Gill, C. Brenner, B. Brinkmann, B. Budowle, A. Carracedo, M.A. Jobling, et al., DNA commission of the international society of forensic genetics: recommendations on forensic analysis using Y-chromosome STRs, Forensic Sci. Int. 124 (1) (2001) 5–10. [174] L. Alzate, N. Agudelo, J. Builes, X-STRs as a tool for missing persons identification using only siblings as reference, Forensic Sci. Int. Genet. Suppl. Ser. 5 (2015) e636–e637. [175] R. Szibor, M. Krawczak, S. Hering, J. Edelmann, E. Kuhlisch, D. Krause, Use of Xlinked markers for forensic purposes, Int. J. Leg. Med. 117 (2) (2003) 67–74. [176] T.M. Diegoli, Forensic typing of short tandem repeat markers on the X and Y chromosomes, Forensic Sci. Int. Genet. 18 (2015) 140–151. [177] M.M. Holland, T.J. Parsons, Mitochondrial DNA sequence analysis - validation and use for forensic casework, Forensic Sci. Rev. 11 (1) (1999) 21–50. [178] V. Bourdon, C. Ng, J. Harris, M. Prinz, E. Shapiro, Optimization of human mtDNA control region sequencing for forensic applications, J. Forensic Sci. 59 (4) (2014) 1057–1063. [179] L. Pikó, L. Matsumoto, Number of mitochondria and some properties of mitochondrial DNA in the mouse egg, Dev. Biol. (Basel) 49 (1) (1976) 1–10. [180] G.S. Michaels, W.W. Hauswirth, P.J. Laipis, Mitochondrial DNA copy number in bovine oocytes and somatic cells, Dev. Biol. (Basel) 94 (1) (1982) 246–251. [181] M.M. Holland, D.L. Fisher, L.G. Mitchell, W.C. Rodriquez, J.J. Canik, C.R. Merril, V.W. Weedn, Mitochondrial DNA sequence analysis of human skeletal remains: identification of remains from the Vietnam War, J. Forensic Sci. 38 (3) (1993) 542–553. [182] M.N. Gabriel, E.F. Huffine, J.H. Ryan, M.M. Holland, T.J. Parsons, Improved MtDNA sequence analysis of forensic remains using a "mini-primer set" amplification strategy, J. Forensic Sci. 46 (2) (2001) 247–253. [183] X. Chen, R. Prosser, S. Simonetti, J. Sadlock, G. Jagiello, E.A. Schon, Rearranged mitochondrial genomes are present in human oocytes, Am. J. Hum. Genet. 57 (2) (1995) 239–247. [184] M. Kayser, P.M. Schneider, DNA-based prediction of human externally visible characteristics in forensics: motivations, scientific challenges, and ethical considerations, Forensic Sci. Int. Genet. 3 (3) (2009) 154–161. [185] K. Kidd, A. Pakstis, W. Speed, R. Lagace, J. Chang, S. Wootton, N. Ihuegbu, Microhaplotype loc are a powerful new type of forensic marker, Forensic Sci. Int. Genet. Suppl. Ser. 4 (2013) 123–124. [186] C. Phillips, A. Salas, J.J. Sánchez, M. Fondevila, A. Gómez-Tato, J. Alvarez-Dios, et al., Inferring ancestral origin using a single multiplex assay of ancestry-informative marker SNPs, Forensic Sci. Int. Genet. 1 (3–4) (2007) 273–280. [187] A.J. Pakstis, W.C. Speed, R. Fang, F.C. Hyland, M.R. Furtado, J.R. Kidd, K.K. Kidd, SNPs for a universal individual identification panel, Hum. Genet. 127 (3) (2010) 315–324. [188] K.K. Kidd, J.R. Kidd, W.C. Speed, R. Fang, M.R. Furtado, F.C. Hyland, A.J. Pakstis,

11

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

[189] [190]

[191]

[192]

[193]

[194]

[195]

[196]

[197] [198]

[199]

[200]

[201]

[202]

[203]

[204]

[205]

[206]

[207]

[208]

[209] [210]

[211]

[212]

[213]

[214]

[215]

[216] J. Ge, B. Budowle, J.V. Planz, R. Chakraborty, Haplotype block: a new type of forensic DNA markers, Int. J. Leg. Med. 124 (5) (2010) 353–361. [217] A.J. Pakstis, R. Fang, M.R. Furtado, J.R. Kidd, K.K. Kidd, Mini-haplotypes as lineage informative SNPs and ancestry inference SNPs, Eur. J. Hum. Genet. 20 (11) (2012) 1148–1154. [218] K. Kidd, W. Speed, S. Wootton, R. Lagace, R. Langit, E. Haigh, et al., Genetic markers for massively parallel sequencing in forensics, Forensic Sci. Int. Genet. Suppl. Ser. 5 (2015) 677–679. [219] H. Wang, J. Zhu, N. Zhou, Y. Jiang, L. Wang, et al., NGS technology makes microhaplotype a potential forensic marker, Forensic Sci. Int. Genet. Suppl. Ser. 5 (2015) 233–234. [220] A. Pakstis, W. Speed, J. Kiddand, K. Kidd, An expanded, nearly universal, panel of SNPs for individual identification, Annual Meeting of the National Institute of Justice, (2007). [221] J. Butler, B. Budowle, P. Gill, K. Kidd, C. Phillips, P. Schneider, et al., Report on ISFG SNP panel discussion, Forensic Sci. Int. Genet. Suppl. Ser. 1 (2008) 471–472. [222] K.K. Kidd, W.C. Speed, A.J. Pakstis, D.S. Podini, R. Lagacé, J. Chang, et al., Evaluating 130 microhaplotypes across a global set of 83 populations, Forensic Sci. Int. Genet. 29 (2017) 29–37. [223] J.L. Weber, D. David, J. Heil, Y. Fan, C. Zhao, G. Marth, Human diallelic insertion/ deletion polymorphisms, Am. J. Hum. Genet. 71 (4) (2002) 854–862. [224] R.E. Mills, C.T. Luttig, C.E. Larkins, A. Beauchamp, C. Tsui, W.S. Pittard, S.E. Devine, An initial map of insertion and deletion (INDEL) variation in the human genome, Genome Res. 16 (9) (2006) 1182–1190. [225] U. Väli, M. Brandström, M. Johansson, H. Ellegren, Insertion-deletion polymorphisms (indels) as genetic markers in natural populations, BMC Genet. 9 (2008) 8. [226] J.M. Mullaney, R.E. Mills, W.S. Pittard, S.E. Devine, Small insertions and deletions (INDELs) in human genomes, Hum. Mol. Genet. 19 (R2) (2010) R131–136. [227] R.E. Mills, W.S. Pittard, J.M. Mullaney, U. Farooq, T.H. Creasy, A.A. Mahurkar, et al., Natural genetic variation caused by small insertions and deletions in the human genome, Genome Res. 21 (6) (2011) 830–839. [228] B. Budowle, A. van Daal, Forensically relevant SNP classes, Biotechniques 44 (5) (2008) 603–608 610. [229] R. Pereira, C. Phillips, C. Alves, A. Amorim, A. Carracedo, L. Gusmão, A new multiplex for human identification using insertion/deletion polymorphisms, Electrophoresis 30 (21) (2009) 3682–3690. [230] B.L. LaRue, J. Ge, J.L. King, B. Budowle, A validation study of the Qiagen Investigator DIPplex® kit; an INDEL-based assay for human identification, Int. J. Leg. Med. 126 (4) (2012) 533–540. [231] H.B. Pena, S.D. Pena, Automated genotyping of a highly informative panel of 40 short insertion-deletion polymorphisms resolved in polyacrylamide gels for forensic identification and kinship analysis, Transfus. Med. Hemother. 39 (3) (2012) 211–216. [232] B.L. LaRue, R. Lagacé, C.W. Chang, A. Holt, L. Hennessy, J. Ge, et al., Characterization of 114 insertion/deletion (INDEL) polymorphisms, and selection for a global INDEL panel for human identification, Leg. Med. (Tokyo) 16 (1) (2014) 26–32. [233] M. Fondevila, C. Phillips, C. Santos, R. Pereira, L. Gusmão, A. Carracedo, et al., Forensic performance of two insertion-deletion marker assays, Int. J. Leg. Med. 126 (5) (2012) 725–737. [234] F.R. Wendt, X. Zeng, J.D. Churchill, J.L. King, B. Budowle, Analysis of short tandem repeat and single nucleotide polymorphism loci from single-source samples using a custom HaloPlex target enrichment system panel, Am. J. Forensic Med. Pathol. 37 (2) (2016) 99–107. [235] M. Fondevila, C. Phillips, N. Naveran, L. Fernandez, M. Cerezo, A. Salas, et al., Case report: identification of skeletal remains using short-amplicon marker analysis of severely degraded DNA extracted from a decomposed and charred femur, Forensic Sci. Int. Genet. 2 (3) (2008) 212–218. [236] B.L. LaRue, S.K. Sinha, A.H. Montgomery, R. Thompson, L. Klaskala, J. Ge, et al., INNULs: a novel design amplification strategy for retrotransposable elements for studying population variation, Hum. Hered. 74 (1) (2012) 27–35. [237] S. Sinha, G. Murphy, H. Brown, A. Green, A. Montgomery, M. Carrol, J. Tabak, Retrotransposable elements: novel and sensitive DNA markers and their application in human identity, Forensic Sci. Int. Genet. Suppl. Ser. 5 (2015) e627–e629. [238] H. Brown, R. Thompson, G. Murphy, D. Peters, B. La Rue, J. King, et al., Development and validation of a novel multiplexed DNA analysis system, InnoTyper, Forensic Sci. Int. Genet. 29 (2017) 80–99. [239] J. Butler, Forensic DNA Typing: Biology, Technology and Genetics of STR Markers, 2nd ed., Elsevier Academic Press, New York, 2005. [240] Q. Wang, L. Fu, X. Zhang, X. Dai, M. Bai, G. Fu, et al., Expansion of a SNaPshot assay to a 55-SNP multiplex: assay enhancements, va lidation, and power in forensic science, Electrophoresis 37 (10) (2016) 1310–1317. [241] P. Brotherton, W. Haak, J. Templeton, G. Brandt, J. Soubrier, C. Jane Adler, et al., Neolithic mitochondrial haplogroup H genomes and the genetic origins of Europeans, Nat. Commun. 4 (2013) 1764. [242] T. Melton, S. Clifford, M. Kayser, I. Nasidze, M. Batzer, M. Stoneking, Diversity and heterogeneity in mitochondrial DNA of North American populations, J. Forensic Sci. 46 (1) (2001) 46–52. [243] J.L. King, B.L. LaRue, N.M. Novroski, M. Stoljarova, S.B. Seo, X. Zeng, et al., Highquality and high-throughput massively parallel sequencing of the human mitochondrial genome using the Illumina MiSeq, Forensic Sci. Int. Genet. 12 (2014) 128–135. [244] C. Ginther, L. Issel-Tarver, M.C. King, Identifying individuals by sequencing mitochondrial DNA from teeth, Nat. Genet. 2 (2) (1992) 135–138. [245] P. Gill, P.L. Ivanov, C. Kimpton, R. Piercy, N. Benson, G. Tully, et al., Identification

Expanding data and resources for forensic use of SNPs in individual identification, Forensic Sci. Int. Genet. 6 (5) (2012) 646–652. B. Mehta, R. Daniel, C. Phillips, D. McNevin, Forensically relevant SNaPshot, Int. J. Leg. Med. 131 (1) (2017) 21–37. R.K. Valenzuela, M.S. Henderson, M.H. Walsh, N.A. Garrison, J.T. Kelch, O. Cohen-Barak, et al., Predicting phenotype from genotype: normal pigmentation, J. Forensic Sci. 55 (2) (2010) 315–322. V.A. Canfield, A. Berg, S. Peckins, S.M. Wentzel, K.C. Ang, S. Oppenheimer, K.C. Cheng, Molecular phylogeography of a human autosomal skin color locus under natural selection, G3 (Bethesda) 3 (11) (2013) 2059–2067. S. Walsh, F. Liu, A. Wollstein, L. Kovatsi, A. Ralf, A. Kosiniak-Kamysz, et al., The HIrisPlex system for simultaneous prediction of hair and eye colour from DNA, Forensic Sci. Int. Genet. 7 (1) (2013) 98–115. R. Kosoy, R. Nassir, C. Tian, P.A. White, L.M. Butler, G. Silva, et al., Ancestry informative marker sets for determining continental origin and admixture proportions in common populations in America, Hum. Mutat. 30 (1) (2009) 69–78. J.R. Kidd, F.R. Friedlaender, W.C. Speed, A.J. Pakstis, F.M. De La Vega, K.K. Kidd, Analyses of a set of 128 ancestry informative single-nucleotide polymorphisms in a global set of 119 population samples, Invest. Genet. 2 (1) (2011) 1. M. Fondevila, C. Phillips, C. Santos, A. Freire Aradas, P.M. Vallone, J.M. Butler, et al., Revision of the SNPforID 34-plex forensic ancestry test: assay enhancements, standard reference sample genotypes and extended population studies, Forensic Sci. Int. Genet. 7 (1) (2013) 63–74. B. Keating, A.T. Bansal, S. Walsh, J. Millman, J. Newman, K. Kidd, et al., First allin-one diagnostic tool for DNA intelligence: genome-wide inference of biogeographic ancestry, appearance, relatedness, and sex with the Identitas v1 Forensic Chip, Int. J. Leg. Med. 127 (3) (2013) 559–572. C. Phillips, Forensic genetic analysis of bio-geographical ancestry, Forensic Sci. Int. Genet. 18 (2015) 49–65. S. Walsh, A. Lindenbergh, S.B. Zuniga, T. Sijen, P. de Knijff, M. Kayser, K.N. Ballantyne, Developmental validation of the IrisPlex system: determination of blue and brown iris colour for forensic intelligence, Forensic Sci. Int. Genet. 5 (5) (2011) 464–471. S. Walsh, F. Liu, K.N. Ballantyne, M. van Oven, O. Lao, M. Kayser, IrisPlex: a sensitive DNA tool for accurate prediction of blue and brown eye colour in the absence of ancestry information, Forensic Sci. Int. Genet. 5 (3) (2011) 170–180. S. Walsh, L. Chaitanya, L. Clarisse, L. Wirken, J. Draus-Barini, L. Kovatsi, et al., Developmental validation of the HIrisPlex system: DNA-based eye and hair colour prediction for forensic and anthropological usage, Forensic Sci. Int. Genet. 9 (2014) 150–161. L. Chaitanya, K. Breslin, S. Zuñiga, L. Wirken, E. Pośpiech, M. Kukla-Bartoszek, et al., The HIrisPlex-S system for eye, hair and skin colour prediction from DNA: introduction and forensic developmental validation, Forensic Sci. Int. Genet. 35 (2018) 123–135. S. Walsh, A. Wollstein, F. Liu, U. Chakravarthy, M. Rahu, J.H. Seland, et al., DNAbased eye colour prediction across Europe with the IrisPlex system, Forensic Sci. Int. Genet. 6 (3) (2012) 330–340. L. Chaitanya, S. Walsh, J.D. Andersen, R. Ansell, K. Ballantyne, D. Ballard, et al., Collaborative EDNAP exercise on the IrisPlex system for DNA-based prediction of human eye colour, Forensic Sci. Int. Genet. 11 (2014) 241–251. P. Dario, H. Mourino, A. Oliveira, I. Lucas, T. Ribeiro, M. Porto, et al., Assessment of IrisPlex-based multiplex for eye and skin color prediction with application to a Portugese population, Int. J. Leg. Med. 129 (2015) 1191–1200. L. Chaitanya, I.Z. Pajnič, S. Walsh, J. Balažic, T. Zupanc, M. Kayser, Bringing colour back after 70 years: predicting eye and hair colour from skeletal remains of World War II victims using the HIrisPlex system, Forensic Sci. Int. Genet. 26 (2017) 48–57. S. Walsh, L. Chaitanya, K. Breslin, C. Muralidharan, A. Bronikowska, E. Pospiech, et al., Global skin colour prediction from DNA, Hum. Genet. 136 (7) (2017) 847–863. I. Halder, M. Shriver, M. Thomas, J.R. Fernandez, T. Frudakis, A panel of ancestry informative markers for estimating individual biogeographical ancestry and admixture from four continents: utility and applications, Hum. Mutat. 29 (5) (2008) 648–658. R. Pereira, C. Phillips, N. Pinto, C. Santos, S.E. dos Santos, A. Amorim, et al., Straightforward inference of ancestry and admixture proportions through ancestry-informative insertion deletion multiplexing, PLoS One 7 (1) (2012) e29684. Illumina, ForenSeq™ DNA Signature Prep Reference Guide, Illumina, Inc., 2015. K.K. Kidd, W.C. Speed, A.J. Pakstis, M.R. Furtado, R. Fang, A. Madbouly, et al., Progress toward an efficient panel of SNPs for ancestry inference, Forensic Sci. Int. Genet. 10 (2014) 23–32. AppliedBiosystems, Precision ID Panels with the Ion PGM™ System, in: E. Life Technologies Corporation (Ed.), Application Guide. Carlsbad, CA: TFS SCIENTIFIC, 2017. C. Hollard, C. Keyser, T. Delabarde, A. Gonzalez, C. Vilela Lamego, V. Zvénigorosky, B. Ludes, Case report: on the use of the HID-Ion AmpliSeq™ Ancestry Panel in a real forensic case, Int. J. Leg. Med. 131 (2) (2017) 351–358. M. Al-Asfi, D. McNevin, B. Mehta, D. Power, M.E. Gahan, R. Daniel, Assessment of the precision ID ancestry panel, Int. J. Leg. Med. 19 (Mar) (2018), https://doi.org/ 10.1007/s00414-018-1785-9 [Epub ahead of print]. V. Pereira, H.S. Mogensen, C. Børsting, N. Morling, Evaluation of the Precision ID Ancestry Panel for crime case work: a SNP typing assay developed for typing of 165 ancestral informative markers, Forensic Sci. Int. Genet. 28 (2017) 138–145. K.K. Kidd, A.J. Pakstis, W.C. Speed, R. Lagacé, J. Chang, S. Wootton, et al., Current sequencing technology makes microhaplotypes a powerful new type of genetic marker for forensics, Forensic Sci. Int. Genet. 12 (2014) 215–224.

12

Forensic Science International: Genetics xxx (xxxx) xxx–xxx

J. Watherston et al.

[246] [247]

[248]

[249]

[250]

[251] [252]

[253]

[254]

[255]

[256]

[257]

[258]

[259]

[260]

[261]

[262]

[263]

[264]

[265]

[266] [267]

[268]

[269]

[270]

sequencing, Forensic Sci. Int. Genet. 30 (2017) 127–133. [271] M.M. Holland, L.A. Wilson, S. Copeland, G. Dimick, C.A. Holland, R. Bever, J.A. McElhoe, MPS analysis of the mtDNA hypervariable regions on the MiSeq with improved enrichment, Int. J. Leg. Med. 131 (4) (2017) 919–931. [272] S.H. Vohr, R. Gordon, J.M. Eizenga, H.A. Erlich, C.D. Calloway, R.E. Green, A phylogenetic approach for haplotype analysis of sequence data from complex mitochondrial mixtures, Forensic Sci. Int. Genet. 30 (2017) 93–105. [273] C. Marshall, K. Sturk-Andreaggi, J. Daniels-Higginbotham, R.S. Oliver, S. BarrittRoss, T.P. McMahon, Performance evaluation of a mitogenome capture and Illumina sequencing protocol using non-probative, case-type skeletal samples: implications for the use of a positive control in a next-generation sequencing procedure, Forensic Sci. Int. Genet. 31 (2017) 198–206. [274] T. Maricic, M. Whitten, S. Pääbo, Multiplexed DNA sequence capture of mitochondrial genomes using PCR products, PLoS One 5 (11) (2010) e14004. [275] M. Knapp, M. Stiller, M. Meyer, Generating barcoded libraries for multiplex highthroughput sequencing, Methods Mol. Biol. 840 (2012) 155–170. [276] Q. Fu, A. Mittnik, P.L.F. Johnson, K. Bos, M. Lari, R. Bollongino, et al., A revised timescale for human evolution based on ancient mitochondrial genomes, Curr. Biol. 23 (7) (2013) 553–559. [277] E. Hodges, Z. Xuan, V. Balija, M. Kramer, M.N. Molla, S.W. Smith, et al., Genomewide in situ exon capture for selective resequencing, Nat. Genet. 39 (12) (2007) 1522–1527. [278] A. Gnirke, A. Melnikov, J. Maguire, P. Rogov, E.M. LeProust, W. Brockman, et al., Solution hybrid selection with ultra-long oligonucleotides for massively parallel targeted sequencing, Nat. Biotechnol. 27 (2) (2009) 182–189. [279] M. Eduardoff, C. Xavier, C. Strobl, A. Casas-Vargas, W. Parson, Optimized mtDNA control region primer extension capture analysis for forensically relevant samples and highly compromised mtDNA of different age and origin, Genes (Basel) 8 (10) (2017). [280] S. Köhnemann, H. Pfeiffer, Application of mtDNA SNP analysis in forensic casework, Forensic Sci. Int. Genet. 5 (3) (2011) 216–221. [281] P. Brotherton, J.J. Sanchez, A. Cooper, P. Endicott, Preferential access to genetic information from endogenous hominin ancient DNA and accurate quantitative SNP-typing via SPEX, Nucleic Acids Res. 38 (2) (2010) e7. [282] N. Bose, K. Carlberg, G. Sensabaugh, H. Erlich, C. Calloway, Target capture enrichment of nuclear SNP markers for massively parallel sequencing of degraded and mixed samples, Forensic Sci. Int. Genet. 34 (2018) 186–196. [283] K. Schoenly, A statistical analysis of successional patterns in carrion-arthropod assemblages: implications for forensic entomology and determination of the postmortem interval, J. Forensic Sci. 37 (6) (1992) 1489–1513. [284] N. Haskell, Testing reliability of animal models in forensic entomology with 50200lb. Pigs vs. humans in Tennessee, XXI International Conference of Entomology (2000). [285] W.C. Rodriguez, W.M. Bass, Decomposition of buried bodies and methods that may aid in their location, J. Forensic Sci. 30 (3) (1985) 836–852. [286] K. Schoenly, K. Griest, S. Rhine, An experimental field protocol for investigation of postmortem interval using multidisciplinary indicators, J. Forensic Sci. 36 (1991) 1395–1415. [287] D. France, T. Griffin, J. Swanburg, J. Lindemann, G. Davenport, V. Trammell, et al., A multidisciplinary approach to the detection of clandestine graves, J. Forensic Sci. 37 (1992) 1445–1458. [288] L. Harsanyi, Differential diagnosis of human and animal bone, Histology of Ancient Human Bone: Methods and Diagnosis, Springer-Verlag, Berlin, 1993. [289] E.M. Schotsmans, J. Denton, J. Dekeirsschieter, T. Ivaneanu, S. Leentjes, R.C. Janaway, A.S. Wilson, Effects of hydrated lime and quicklime on the decay of buried human remains using pig cadavers as human body analogues, Forensic Sci. Int. 217 (1–3) (2012) 50–59. [290] M. Grassberger, C. Frank, Initial study of arthropod succession on pig carrion in a central European urban habitat, J. Med. Entomol. 41 (3) (2004) 511–523. [291] J. Dekeirsschieter, F.J. Verheggen, M. Gohy, F. Hubrecht, L. Bourguignon, G. Lognay, E. Haubruge, Cadaveric volatile organic compounds released by decaying pig carcasses (Sus domesticus L.) in different biotopes, Forensic Sci. Int. 189 (1–3) (2009) 46–53. [292] S.L. Forbes, B.B. Dent, B.H. Stuart, The effect of soil type on adipocere formation, Forensic Sci. Int. 154 (1) (2005) 35–43. [293] S.L. Forbes, B.H. Stuart, B.B. Dent, The effect of the method of burial on adipocere formation, Forensic Sci. Int. 154 (1) (2005) 44–52. [294] S.L. Forbes, B.H. Stuart, B.B. Dent, The effect of the burial environment on adipocere formation, Forensic Sci. Int. 154 (1) (2005) 24–34. [295] A.S. Wilson, R.C. Janaway, A.D. Holland, H.I. Dodson, E. Baran, A.M. Pollard, D.J. Tobin, Modelling the buried human body environment in upland climes using three contrasting field sites, Forensic Sci. Int. 169 (1) (2007) 6–18. [296] R. Janaway, A. Wilson, G. Caprio Diazand, S. Guillen, Taphonomic changes to the buried body in arid environments: an experimental case study in Peru, in: K. Ritz, L. Dawson, D. Miller (Eds.), Criminal and Environmental Soil Forensics, Springer, London, 2009, pp. 341–356. [297] J. Bytheway, M. Connor, G. Dabbs, C. Johnston, M. Sunkel, The ethics and best practices of human decomposition facilities in the United States, Forensic Sci. Policy Manag. 6 (3–4) (2015) 59–68.

of the remains of the Romanov family by DNA analysis, Nat. Genet. 6 (2) (1994) 130–135. T.C. Boles, C.C. Snow, E. Stover, Forensic DNA testing on skeletal remains from mass graves: a pilot project in Guatemala, J. Forensic Sci. 40 (3) (1995) 349–355. M. Allen, A.S. Engström, S. Meyers, O. Handt, T. Saldeen, A. von Haeseler, et al., Mitochondrial DNA sequencing of shed hairs and saliva on robbery caps: sensitivity and matching probabilities, J. Forensic Sci. 43 (3) (1998) 453–464. J.E. Templeton, P.M. Brotherton, B. Llamas, J. Soubrier, W. Haak, A. Cooper, J.J. Austin, DNA capture and next-generation sequencing can recover whole mitochondrial genomes from highly degraded samples for human identification, Invest. Genet. 4 (1) (2013) 26. E.A. Lyons, M.K. Scheible, K. Sturk-Andreaggi, J.A. Irwin, R.S. Just, A highthroughput Sanger strategy for human mitochondrial genome sequencing, BMC Genomics 14 (2013) 881. M. Stoljarova, J.L. King, M. Takahashi, A. Aaspõllu, B. Budowle, Whole mitochondrial genome genetic diversity in an Estonian population sample, Int. J. Leg. Med. 130 (1) (2016) 67–71. E.L. van Dijk, H. Auger, Y. Jaszczyszyn, C. Thermes, Ten years of next-generation sequencing technology, Trends Genet. 30 (9) (2014) 418–426. M.A. Quail, M. Smith, P. Coupland, T.D. Otto, S.R. Harris, T.R. Connor, et al., A tale of three next generation sequencing platforms: comparison of Ion Torrent, Pacific Biosciences and Illumina MiSeq sequencers, BMC Genomics 13 (2012) 341. J. Churchill, J. Chang, J. Ge, N. Rajagopalan, S. Wootton, et al., Blind study evaluation illustrates utility of the Ion PGM™ system for use in human identity DNA typing, Croat. Med. J. 56 (2015) 218–229. R. Daniel, C. Santos, C. Phillips, M. Fondevila, R.A. van Oorschot, A. Carracedo, et al., A SNaPshot of next generation sequencing for forensic SNP analysis, Forensic Sci. Int. Genet. 14 (2015) 50–60. S. Fordyce, H. Mogensen, C. Borsting, R. Lagace, C. Chang, et al., Second-generation sequencing of forensic STRs using the ion torrent™ HID STR 10-plex and the ion PGM™, Forensic Sci. Int. Genet. 14 (2015) 132–140. X. Zeng, J.L. King, M. Stoljarova, D.H. Warshauer, B.L. LaRue, A. Sajantila, et al., High sensitivity multiplex short tandem repeat loci analyses with massively parallel sequencing, Forensic Sci. Int. Genet. 16 (2015) 38–47. B. Mehta, R. Daniel, C. Phillips, S. Doyle, G. Elvidge, D. McNevin, Massively parallel sequencing of customized forensically informative SNP panels on the MiSeq, Electrophoresis 37 (21) (2016) 2832–2840. F.R. Wendt, J.D. Churchill, N.M.M. Novroski, J.L. King, J. Ng, R.F. Oldt, et al., Genetic analysis of the Yavapai Native Americans from West-Central Arizona using the Illumina MiSeq FGx™ forensic genomics system, Forensic Sci. Int. Genet. 24 (2016) 18–23. F.R. Wendt, D.H. Warshauer, X. Zeng, J.D. Churchill, N.M.M. Novroski, B. Song, et al., Massively parallel sequencing of 68 insertion/deletion markers identifies novel microhaplotypes for utility in human identity testing, Forensic Sci. Int. Genet. 25 (2016) 198–209. W. Parson, G. Huber, L. Moreno, M. Madel, M. Brandhagen, S. Nagl, et al., Massively parallel sequencing of complete mitochondrial genomes form hair shaft samples, Forensic Sci. Int. Genet. 15 (2015) 8–15. E.D. Gunnarsdóttir, M. Li, M. Bauchet, K. Finstermeier, M. Stoneking, Highthroughput sequencing of complete human mtDNA genomes from the Philippines, Genome Res. 21 (1) (2011) 1–11. J.A. McElhoe, M.M. Holland, K.D. Makova, M.S. Su, I.M. Paul, C.H. Baker, et al., Development and assessment of an optimized next-generation DNA sequencing approach for the mtgenome using the Illumina MiSeq, Forensic Sci. Int. Genet. 13 (2014) 20–29. L. Chaitanya, A. Ralf, M. van Oven, T. Kupiec, J. Chang, R. Lagacé, M. Kayser, Simultaneous whole mitochondrial genome sequencing with short overlapping amplicons suitable for degraded DNA using the ion torrent personal genome machine, Hum. Mutat. 36 (12) (2015) 1236–1247. J. Churchill, D. Peters, C. Capt, C. Strobl, W. Parson, B. Budowle, Working towards implementation of whole genome mitochondrial DNA sequencing into routine casework, Forensic Sci. Int. Genet. Suppl. Ser. 6 (2017) e388–e389. R. Just, M. Scheible, S. Fast, K. Sturk-Andreaggi, A. Rock, J. Bush, et al., Full mtGenome reference data: development and characterization of 588 forensicquality haplotypes representing three US populations, Forensic Sci. Int. Genet. 14 (2015) 141–155. S. Tang, T. Huang, Characterization of mitochondrial DNA heteroplasmy using a parallel sequencing system, Biotechniques 48 (4) (2010) 287–296. M.M. Rathbun, J.A. McElhoe, W. Parson, M.M. Holland, Considering DNA damage when interpreting mtDNA heteroplasmy in deep sequencing data, Forensic Sci. Int. Genet. 26 (2017) 1–11. M.M. Holland, M.R. McQuillan, K.A. O’Hanlon, Second generation sequencing allows for mtDNA mixture deconvolution and high resolution detection of heteroplasmy, Croat. Med. J. 52 (3) (2011) 299–313. C. Davis, D. Peters, D. Warshauer, J. King, B. Budowle, Sequencing the hypervariable regions of human mitochondrial DNA using massively parallel sequencing: enhanced data acquisition for DNA samples encountered in forensic testing, Leg. Med. (Tokyo) 17 (2) (2015) 123–127. C.Y. Lin, L.C. Tsai, H.M. Hsieh, C.H. Huang, Y.J. Yu, B. Tseng, et al., Investigation of length heteroplasmy in mitochondrial DNA control region by massively parallel

13