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Spider Web DNA: A New Spin on Noninvasive Genetics of Predator and Prey

  • Charles C. Y. Xu ,

    Affiliation: Department of Biological Sciences, University of Notre Dame, Notre Dame, Indiana, United States of America

    Current address: Animal Ecology Research Group, Department of Ecology and Genetics, Evolutionary Biology Centre, Uppsala University, Uppsala, Sweden

  • Ivy J. Yen,

    Affiliation: Department of Biological Sciences, University of Notre Dame, Notre Dame, Indiana, United States of America

  • Dean Bowman,

    Affiliation: Potawatomi Zoo, South Bend, Indiana, United States of America

  • Cameron R. Turner

    Affiliation: Department of Biological Sciences, University of Notre Dame, Notre Dame, Indiana, United States of America

    Current address: ecoSystem Genetics LLC, South Bend, Indiana, United States of America

Spider Web DNA: A New Spin on Noninvasive Genetics of Predator and Prey

  • Charles C. Y. Xu, 
  • Ivy J. Yen, 
  • Dean Bowman, 
  • Cameron R. Turner


Noninvasive genetic sampling enables biomonitoring without the need to directly observe or disturb target organisms. This paper describes a novel and promising source of noninvasive spider and insect DNA from spider webs. Using black widow spiders (Latrodectus spp.) fed with house crickets (Acheta domesticus), we successfully extracted, amplified, and sequenced mitochondrial DNA from spider web samples that identified both spider and prey to species. Detectability of spider DNA did not differ between assays with amplicon sizes from 135 to 497 base pairs. Spider and prey DNA remained detectable at least 88 days after living organisms were no longer present on the web. Spider web DNA as a proof-of-concept may open doors to other practical applications in conservation research, pest management, biogeography studies, and biodiversity assessments.


As dominant predators of arthropod communities in natural and agricultural ecosystems, spiders can be important ecological indicators that reflect habitat quality and change [1, 2, 3, 4, 5]. Monitoring the species diversity and abundance of spider assemblages facilitates natural resource management [6]. However, spiders are enormously diverse (~ 45,000 described species) and many can be difficult to identify [7]. Morphological identification of spiders relies primarily on differences in copulatory organs [8] and many complications can prevent identification such as the inability to identify juveniles, extreme sexual dimorphism, size differences between life stages, and genital polymorphisms within species [911]. In recent years, genetic identification methods such as DNA barcoding, the use of a short and standardized fragment of DNA to identify organisms, have been growing in popularity because of decreasing costs and ease of use [12]. In particular, the use of DNA barcodes for species identity and systematics of spiders has proven successful in multiple studies [9, 1315]. The most commonly used genetic marker is the cytochrome c oxidase subunit I (COI) mitochondrial gene because of its designation as the standard DNA barcode [16]. Mitochondrial markers are also ideal for detecting low quantity and quality DNA from environmental or gut samples because each cell contains hundreds to thousands of mitochondrial genomes [17] and there is a positive correlation between gene copy number and detection success [18, 19].

Spiders have a great diversity of life histories and various sampling methods are employed in capturing them including beating, vacuum sampling, sweep netting, pitfall traps, and visual searches. Experiments testing the efficacy of traditional spider sampling methods show high variability in diversity and abundance measurements between methods depending on the habitat and time of sampling. [2022]. Sampling duration is also an important factor as short-term sampling has been found to reduce the number of recorded species by up to 50% [23]. In this paper, we propose a new biomonitoring tool that would complement existing methods: DNA from spider web. While spider web has been found to efficiently collect pollen, fungal spores and agrochemical sprays [24, 25], no study, to our knowledge, has assessed spider web as a potential source of genetic material. We hypothesized that spider web could be a source of noninvasive DNA from both the spider that built the web and spider prey.

Although noninvasive genetic sampling is most common for vertebrates, it has been successfully applied to arthropod exuviae and frass [26, 27]. Webs are an abundant and easily collected spider secretion that may not only provide spider DNA, but may also function as natural biodiversity samplers that contain environmental DNA (eDNA) from captured prey and other local organisms. This idea parallels recent molecular studies using mosquitos, ticks, leeches, and carrion flies to sample local animal biodiversity [2831]. Previous studies have successfully used mitochondrial DNA markers to detect spider prey from gut contents, but this requires physically capturing and killing spiders [32, 33]. One recent advance in noninvasive spider diet analysis is the amplification of prey DNA from fecal pellets [34], but fecal pellets are small and may be hard to locate, especially in the field. Furthermore, traditional taxonomic identification of spider prey items is time-consuming, subject to human error, and often only accurate at high taxonomic levels [35]. Spider webs may provide a unique noninvasive opportunity to study arthropod communities without the need to directly observe spider or insect.

We tested the spider web DNA concept by extracting, amplifying and sequencing DNA of black widow spiders, Latrodectus spp. (Araneae: Theridiidae), and their prey, the house cricket Acheta domesticus (Orthoptera: Gryllidae), from black widow spider webs. Because extraorganismal DNA from spider webs is exposed to environmental degradation and possibly only exists in short fragments, we used nested primer sets to test the effect of amplicon size on detection probability.

Materials and Methods

Web collection

The black widow spider exhibit at the Potawatomi Zoo in South Bend, Indiana was inhabited by a single female western black widow spider (Latrodectus hesperus) before its death on November 19, 2011. The spider was fed 2 medium sized house crickets (A. domesticus), on a weekly basis by zookeepers. The exhibit measured 40 cm by 40 cm by 40 cm and contained a few twigs, a small piece of wood, and wood shavings lining the floor. 88 days after the death of the spider, a web sample was collected from the exhibit on February 15, 2012, hereafter referred to as “Lhes_zoo”. The duration of inhabitance within the exhibit prior to the sample collection date is unknown. Three new individual enclosures measuring 35 cm by 30 cm by 35 cm were constructed with plywood and acrylic sheeting and installed on a wall in the zookeeper access hallway behind the exhibit. The enclosures were decontaminated with 10% bleach and installed at the Potawatomi Zoo in South Bend, Indiana.

Three female southern black widow spiders (Latrodectus mactans) were purchased from Tarantula Spiders ( According to the supplier, these spiders were hatched from egg sacs collected in Marion County, Florida, USA and raised on 2–3 housefly maggots (Musca domestica) twice per week before delivery to the Potawatomi Zoo. A single live L. mactans and a bleach-decontaminated branch for web building were placed into each enclosure on April 26, 2012 (Fig 1). After web construction, each spider was fed two medium-sized crickets by dropping them into the web. Web samples were collected from each enclosure 11 days after spider introduction, hereafter referred to as “Lmac_1”, “Lmac_2”, and “Lmac_3”. All web samples were collected by twisting single-use, sterile plastic applicators to spool silk strands. No organism body parts or exuviae were visible in any web samples but cricket parts and spider feces were clearly evident on the floor of the enclosures. Applicator tips were snipped into 1.5-mL microcentrifuge tubes using bleach-decontaminated scissors and stored at -20°C.

Fig 1. Southern black widow spider (Latrodectus mactans) with its prey house cricket (Acheta domesticus) trapped in spider web.

Image credit Scott Camazine, used with permission.

DNA extraction

DNA extractions from web samples were conducted using a modified extraction protocol for shed reptile skins [36]. A negative control without web was also extracted to test for reagent contamination. 800 μL of cell lysis buffer (10 mM Tris, 10 mM EDTA, 2% sodium dodecyl sulfate [SDS], pH 8.0) and 8 μL of proteinase K (20 mg/L) were added to 1.5 mL microcentrifuge tubes containing web samples followed by 10–20 inversions and incubation at 55°C for 4 hours. Upon reaching room temperature, 4 μL of RNase A (10 mg/mL) were added to each sample followed by 20 inversions. Samples were incubated at 37°C for 15 min and then brought back to room temperature. 300 μL of protein precipitation solution (7.5 M ammonium acetate) were added to each sample and vortexed for 20 seconds followed by incubation on ice for 15 min. Samples were then centrifuged at 14,000 rpm for 3 min. Supernatants were transferred to new 2 mL microcentrifuge tubes containing 750 μL of ice cold isopropanol and inverted 50 times before centrifugation at 14,000 rpm for 2 min. All supernatants were drained and 750 μL of 70% ethanol was added to each sample followed by centrifugation at 14,000 rpm for 3 min. All liquids were removed and samples were air dried. DNA pellets were rehydrated using 100 μL of low TE buffer (10 mM Tris, 0.1 mM EDTA).

Primer design

To detect Latrodectus DNA, we designed four nested primer sets based on an alignment of Latrodectus COI DNA barcoding sequences obtained from the National Center for Biotechnology Information (NCBI) GenBank database [37, 38]. All four assays included the same forward primer but different reverse primers, producing amplicons of 135 bp, 257 bp, 311 bp, and 497 bp respectively (Table 1). To detect prey eDNA, we designed a set of primers that specifically amplifies 248 bp of the DNA barcoding region of the COI gene in A. domesticus using sequences obtained from the NCBI GenBank database (Table 1) [39, 40]. GenBank accession numbers of DNA sequences used to design all primers are provided in S1 Table.

Table 1. PCR primers designed to amplify the cytochrome c oxidase subunit I (COI) gene of target species.

DNA amplification

All DNA samples were amplified in polymerase chain reactions (PCR) of 20 μL containing 13.28 μL of ddH2O, 2 μL of 5 PRIME® 10x Taq Buffer advanced, 2 μL of 5 PRIME® Magnesium Solution at 25 mM, 0.4 μL of dNTPs at 2.5 mM, 0.12 μL of 5 PRIME® Taq DNA polymerase at 5 U/μL, 0.6 μL of forward and reverse primers at 10 μM, and 1.0 μL of DNA template using Eppendorf Mastercycler® pro thermocyclers. Cycling conditions were as follows: 94°C/5 min, 55X (94°C/20 s, 54.4°C/35 s, 72°C/30 s), 72°C/7 m, 4°C/hold. Each Latrodectus spp. primer set was used to amplify all DNA samples with 10 technical replicates to measure detection probability for different amplicon sizes. All web DNA samples were amplified with 2 technical replicates using the A. domesticus primer set. One negative control reaction with ddH2O instead of DNA template was included on every PCR plate to test for contamination. Gel electrophoresis was conducted using 5 μL of PCR product mixed with 3 μL of loading dye and 10 μL of ddH2O. Multiple wells were loaded with 5 μL of 100 bp ladder (Promega) on each gel. Technical replicates showing amplicons of the expected size were pooled and purified using ExoSAP-IT (Affymetrix). Bi-directional Sanger sequencing using ABI BigDye chemistry (Life Technologies) was conducted on an ABI 3730xl 96-capillary sequencer by the University of Notre Dame Genomics Core Facility. Sequencing chromatograms were primer- and quality-trimmed in Sequencher (ver. 5.0; Gene Codes Corp.). Internal ambiguous base calls were denoted as “N”. BLASTn searches of the NCBI GenBank database [41] and Barcode of Life database (BOLD) Identification System (IDS) COI searches of Species Level Barcode Records with default settings were used for taxonomic identification of COI barcode sequences. For each query sequence, the resulting match with the highest percent identity (90–100%) was accepted for taxonomic identification. Accession and IDS numbers of top NCBI and BOLD matches, respectively, are provided in S2 Table.


All extraction and PCR negative controls produced no amplification. Using the nested primer sets, we successfully amplified 135 bp, 257 bp, 311 bp, and 497 bp of Latrodectus spp. COI from web DNA samples (Fig 2). With the exception of zero amplification for the 311 bp PCR assay from two samples, 2–10 technical replicates of each PCR assay successfully amplified from all samples. Web DNA sequences obtained from enclosure samples, “Lmac_1”, “Lmac_2”, and “Lmac_3”, were confirmed by NCBI BLAST and BOLD IDS to be L. mactans and DNA from the zoo exhibit sample, “Lhes_zoo”, was confirmed to be L. hesperus. Two sequences contained short internal runs of ambiguous base calls that did not prevent taxonomic identification (S2 Table). Amplicon size had no effect on PCR success based on the number of successful PCR replicates (ANOVA, F = 1.941, d.f. = 3, P = 0.194). We also successfully amplified 248 bp of Acheta domesticus COI from web DNA samples. Both PCR duplicates from all four web samples amplified successfully and all resulting DNA sequences were confirmed by NCBI BLAST and BOLD IDS to be A. domesticus. The zoo exhibit web sample, “Lhes_zoo”, was collected 88 days after the death and removal of both spider and prey, demonstrating substantial persistence of web DNA. All DNA sequences generated in this study are provided in S2 Table.

Fig 2. Success of different amplicon sizes in detecting the cytochrome c oxidase subunit I (COI) of Latrodectus spp. from web samples.

Percent success calculated from number of successful PCRs out of 10 technical replicates. Error bars represent ± 1 standard error.


The present study represents, to our knowledge, the first demonstration of spider web as a source of noninvasive genetic material. Spider web is an ideal source of noninvasive genetic material for spiders because web can be found and collected without the need to directly observe or capture spiders themselves. Furthermore, unlike most spiders, which are small, mobile, and elusive, webs are relatively large, stationary, and often clearly visible. Spider webs may also remain after spiders move or die, which increases detection probability especially for more elusive species. Webs can also exist in great abundance. For example, web coverage may reach up to more than 50% of land area in agricultural fields [42]. Spider webs have already been utilized by citizen scientists to assess spider biodiversity through visual analysis of web structure [43] and it could be possible to implement similar citizen science initiatives to collect web samples for DNA analysis.

Because black widow spiders are cobweb spiders that generate large three-dimensional cobwebs consisting of sheets dotted with glue droplets [44], they were ideal to use in this experiment. Although spider silk could be considered a form of spider tissue, spider silk fibers are composed of tightly bound β-sheet proteins that exclude water molecules and do not dissolve under the proteinase K treatment of standard DNA extraction protocols [45]. Thus, we hypothesize that most spider web DNA originates either from microscopic pieces of fecal matter, setae, and exuviae adhered to silk strands or directly from the silk gland exudate, which may contain cells shed from silk glands.

Certain black widow spiders like the species used in this study are common venomous pests [46] and spider web DNA could be a particularly useful surveillance tool. Spider web DNA could also help monitor low density populations and determine invasion fronts of invasive widow spiders such as the brown widow, Latrodectus geometricus, in southern California and the Australian redback, Latrodectus hasseltii, in New Zealand and Japan [47, 48]. Besides pest and invasive species, many spiders like the katipo, Latrodectus katipo, are threatened or endangered [49]. The geographic range and abundance of thousands more spider species are unknown but may be declining. Spider web DNA could be particularly useful in rapidly providing occurrence and genetic diversity data for these rare species of concern. As a noninvasive biomonitoring method, spider web DNA could be used for conservation and taxonomy without sacrificing organisms that are already threatened by human disturbance. The collection and genetic analysis of spider webs could also serve spider biogeography studies, which require large-scale sampling across wide geographic ranges [38].

Our proof-of-concept experiment used spider web from indoor enclosures where DNA-degrading conditions such as heat, moisture, and light were likely reduced relative to field conditions. Many spider taxa, including Latrodectus, build webs in protected spaces [50], but further testing of field-collected spider web from more species and habitats is needed to evaluate the generality of our findings. Nevertheless, this first demonstration suggests a promising approach for arthropod monitoring. The ability to target particular species could be useful in monitoring low density populations of pest, invasive, or endangered insects. Future work using massively parallel sequencing on spider web eDNA could reveal entire assemblages of arthropods in a cost-effective manner, especially with the rapid advancement and decreasing costs of such technologies [51]. This method could be used for diet analysis, which would be especially useful in assessing the importance of riparian spiders as links between aquatic and terrestrial food webs [52]. In some environments such as temperate forests, approximately 40% of arthropod biomass is annually consumed by spiders [53]. Although spider predation cannot be concluded from the mere presence of DNA on spider webs and it is unlikely that individual web samples will consistently yield DNA of the full diversity of spider prey, detection of insect eDNA from spider web does at least indicate local proximity. Spider web eDNA may complement traditional assessment methods of local arthropod biodiversity and potentially reveal previously undiscovered biodiversity through improved sensitivity and sampling effort [54]. Such information regarding species diversity is critically important in conservation planning and environmental impact assessments [55, 56]. However, it is crucial to note that DNA barcoding is most valuable in combination with the taxonomic expertise necessary to provide species identities. The successful use of spider web DNA relies heavily on having well-annotated DNA sequences available such as those found in BOLD. Without quality reference databases, species-level identification of rare, endangered, or invasive species is difficult. Generation of new sequence data along with proper annotation will improve the usefulness and efficacy of this new tool.

In conclusion, we provide a proof-of-concept that noninvasive DNA of a spider and its prey can be extracted from spider web and be used for species identification. Spider web DNA appears to be a promising tool with wide applications in biomonitoring, biogeography and biodiversity assessments of spiders and their prey, especially if combined with the power of massively parallel sequencing [57].

Supporting Information

S1 Table. GenBank accession numbers of DNA sequences used to design primers targeting spiders (L. hesperus, L. mactans) and prey (A. domesticus).



S2 Table. Cytochrome c oxidase subunit I sequences of spiders (L. hesperus, L. mactans) and prey (A. domesticus) generated from spider web DNA.




We thank Larry Richey for his help in constructing the enclosures and Scott Camazine for sharing his photograph for Fig 1. We also thank the Potawatomi Zoo for their cooperation as well as all the members of the Lodge lab and especially Dr. David Lodge for their support of this project.

Author Contributions

Conceived and designed the experiments: CCYX CRT. Performed the experiments: CCYX CRT IJY DB. Analyzed the data: CCYX CRT. Contributed reagents/materials/analysis tools: CCYX CRT. Wrote the paper: CCYX CRT IJY DB.


  1. 1. Churchill TB. Spiders as ecological indicators: an overview for Australia. Mem. Natl. Mus. Victoria. 1997;56: 331–337.
  2. 2. Clausen HIS. The use of spiders (Araneae) as ecological indicators. Bull. British Arachn. Soc. 1986;7: 83–86.
  3. 3. Doran NE, Kiernan K, Swain R, Richardson AMM. Hickmania troglodytes, the Tasmanian cave spider, and its potential role in cave management. J. Insect Conserv. 1999;3: 257–262. doi: 10.1017/s0952836901000371
  4. 4. Maelfait J, Hendrickx F. Spiders as bio-indicators of anthropogenic stress in natural and semi-natural habitats in Flanders (Belgium): some recent developments. In: Selden PA, editor. Proceedings of the 17th European Colloquium of Arachnology, Edinburgh. 1997. pp. 293–300.
  5. 5. Scott AG, Oxford GS, Selden PA. Epigeic spiders as ecological indicators of conservation value for peat bogs. Biol. Cons. 2006;127: 420–428. doi: 10.1016/j.biocon.2005.09.001
  6. 6. Pearce JL, Venier LA. The use of ground beetles (Coleoptera: Carabidae) and spiders (Araneae) as bioindicators of sustainable forest management: A review. Ecol. Indic. 2006;6: 780–793. doi: 10.1016/j.ecolind.2005.03.005
  7. 7. World Spider Catalog, Version 16. Natural History Museum Bern, Available: Accessed 03 February 2015.
  8. 8. Huber BA. The significance of copulatory structures in spider systematics. In: Schult J, editor. Biosemiotik—praktische Anwendung und Konsequenzen für die Einzelwissenschaften. Berlin: VWB Verlag; 2004. pp. 89–100.
  9. 9. Barrett RDH, Hebert PDN. Identifying spiders through DNA barcodes. Can. J. Zool. 2005;83: 481–491. doi: 10.1139/z05-024
  10. 10. Brennan KEC, Moir ML, Majer JD. Exhaustive sampling in a Southern Hemisphere global biodiversity hotspot: inventorying species richness and assessing endemicity of the little known jarrah forest spiders. Pac. Conserv. Biol. 2004;10: 241–260.
  11. 11. Huber BA, Gonzalez AP. Female genital dimorphism in a spider (Araneae: Pholcidae). J. Zool. 2001;255: 301–304. doi: 10.1017/s095283690100139x
  12. 12. Jinbo U, Kato T, Ito M. Current progress in DNA barcoding and future implications for entomology. Entomol. Sci. 2011;14: 107–124. doi: 10.1111/j.1479-8298.2011.00449.x
  13. 13. Astrin JJ, Huber BA, Misof B, Klütsch CFC. Molecular taxonomy in pholcid spiders (Pholcidae, Araneae): evaluation of species identification methods using CO1 and 16S rRNA. Zool. Scripta. 2006;35: 441–457. doi: 10.1111/j.1463-6409.2006.00239.x
  14. 14. Blagoev GA, Nikolova NI, Sobel CN, Hebert PDN, Adamowicz SJ. Spiders (Araneae) of Churchill, Manitoba: DNA barcodes and morphology reveal high species diversity and new Canadian records. BMC Ecol. 2013;13: 44. doi: 10.1186/1472-6785-13-44. pmid:24279427
  15. 15. Robinson EA, Blagoev GA, Hebert PDN, Adamowicz SJ. Prospects for using DNA barcoding to identify spiders in species-rich genera. ZooKeys. 2009;16: 27–46. doi: 10.3897/zookeys.16.239
  16. 16. Hebert PDN, Cywinska A, Ball SL, deWaard JR. Biological identifications through DNA barcodes. Proc. R. Soc. B. 2003;270: 313–322. pmid:12614582 doi: 10.1098/rspb.2002.2218
  17. 17. Hoy MA. Insect Molecular Genetics: An Introduction to Principals and Applications. San Diego: Academic Press; 1994.
  18. 18. Agustí N, Unruh TR, Welter SC. Detecting Cacopsylla pyricola (Hemiptera: Psyllidae) in predator guts using COI mitochondrial markers. Bull. Entomol. Res. 2003;93: 179–185. pmid:12762859 doi: 10.1079/ber2003236
  19. 19. Chen Y, Giles KL, Payton ME, Greenstone MH. Identifying key cereal aphid predators by molecular gut analysis. Mol. Ecol. 2000;9: 1887–1898. pmid:11091324 doi: 10.1046/j.1365-294x.2000.01100.x
  20. 20. Churchill TB, Arthur JM. Measuring spider richness: effects of different sampling and spatial and temporal scales. J. Insect Conserv. 1999;3: 287–295.
  21. 21. Green J. Sampling method and time determines composition of spider collections. J. Arachnol. 1999;24: 111–128.
  22. 22. Sørensen LL, Coddington JA, Scharff N. Inventorying and estimating subcanopy spider diversity using semiquantitative sampling methods in an afromontane forest. Environ. Entomol. 2002;31: 319–330. doi: 10.1603/0046-225x-31.2.319
  23. 23. Riecken U. Effects of short-term sampling on ecological characterization and evaluation of epigeic spider communities and their habitats for site assessment studies. J. Arachnol. 1999;27: 189–195.
  24. 24. Eggs B, Sanders D. Herbivory in spiders: The importance of pollen for orb-weavers. PLOS ONE. 2013;8: e82637. doi: 10.1371/journal.pone.0082637. pmid:24312430
  25. 25. Samu F, Matthews GA, Lake D, Vollrath F. Spider webs are efficient collectors of agrochemical spray. Pestic. Sci. 1992;36: 47–51. doi: 10.1002/ps.2780360108
  26. 26. Feinstein J. DNA sequence from butterfly frass and exuviae. Conserv. Genet. 2004;5: 103–104. doi: 10.1023/b:coge.0000014058.34840.94
  27. 27. Petersen SD, Mason T, Akber S, West R, White B, Wilson P. Species identification of tarantulas using exuviae for international wildlife law enforcement. Conserv. Genet. 2006;8: 497–502. doi: 10.1007/s10592-006-9173-2
  28. 28. Calvignac-Spencer S, Merkel K, Kutzner N, Kühl H, Boesch C, Kappeler PM, et al. Carrion fly-derived DNA as a tool for comprehensive and cost-effective assessment of mammalian biodiversity. Mol. Ecol. 2013;22: 915–924. doi: 10.1111/mec.12183. pmid:23298293
  29. 29. Gariepy TD, Lindsay R, Ogden N, Gregory TR. Identifying the last supper: utility of the DNA barcode library for bloodmeal identification in ticks. Mol. Ecol. Resour. 2012;12: 646–652. doi: 10.1111/j.1755-0998.2012.03140.x. pmid:22471892
  30. 30. Schnell IB, Thomsen PF, Wilkinson N, Rasmussen M, Jensen LRD, Willerslev E, et al. Screening mammal biodiversity using DNA from leeches. Curr. Biol. 2012;22: 262–263. doi: 10.1016/j.cub.2012.02.058
  31. 31. Townzen JS, Brower AVZ, Judd DD. Identification of mosquito bloodmeals using mitochondrial cytochrome oxidase subunit I and cytochrome b gene sequences. Med. Vet. Entomol. 2008;22: 386–393. doi: 10.1111/j.1365-2915.2008.00760.x. pmid:19120966
  32. 32. Agustí N, Shayler SP, Harwood JD, Vaughan IP, Sunderland KD, Symondson WOC. Collembola as alternative prey sustaining spiders in arable ecosystems: prey detection within predators using molecular markers. Mol. Ecol. 2003;12: 3467–3475. pmid:14629361 doi: 10.1046/j.1365-294x.2003.02014.x
  33. 33. Sheppard SK, Bell J, Sunderland KD, Fenlon J, Skervin D, Symondson WOC. Detection of secondary predation by PCR analyses of the gut contents of invertebrate generalist predators. Mol. Ecol. 2005;14: 4461–4468. pmid:16313606 doi: 10.1111/j.1365-294x.2005.02742.x
  34. 34. Sint D, Thurner I, Kaufmann R, Traugott M. Sparing spiders: faeces as a non-invasive source of DNA. Frontiers Zool. 2015;12: 3. doi: 10.1186/s12983-015-0096-y
  35. 35. Salomon M. The natural diet of a polyphagous predator, Latrodectus hesperus (Araneae: Theridiidae), over one year. J. Arachnol. 2011;39: 154–160. doi: 10.1636/p10-25.1
  36. 36. Fetzner JW. Extracting high-quality DNA from shed reptile skins: a simplified method. Biotechniques. 1999;26: 1052–1054. pmid:10376138
  37. 37. Arnedo MA, Coddington J, Agnarsson I, Gillespie RG. From a comb to a tree: phylogenetic relationships of the comb-footed spiders (Araneae, Theridiidae) inferred from nuclear and mitochondrial genes. Mol. Phylogenet. Evol. 2004;31: 225–245. pmid:15019622 doi: 10.1016/s1055-7903(03)00261-6
  38. 38. Garb JE, González A, Gillespie RG. The black widow spider genus Latrodectus (Araneae, Theridiidae): phylogeny, biogeography, and invasion history. Mol. Phylogenet. Evol. 2004;31: 1127–1142. pmid:15120405 doi: 10.1016/j.ympev.2003.10.012
  39. 39. Nattier R, Grandcolas P, Elias M, Desutter-Grandcolas L, Jourdan H, Couloux A, et al. Secondary sympatry caused by range expansion informs on the dynamics of microendemism in a biodiversity hotspot. PLOS ONE. 2012;7: e48047. doi: 10.1371/journal.pone.0048047. pmid:23139758
  40. 40. Sint D, Raso L, Kaufmann R, Traugott M. Optimizing methods for PCR-based analysis of predation. Mol. Ecol. Resour. 2011;5: 795–801. doi: 10.1111/j.1755-0998.2011.03018.x
  41. 41. Benson DA, Karsch-Mizrachi I, Clark K, Lipman DJ, Ostell J, Sayers EW. GenBank. Nucleic Acids Res. 2012;40: D48–D53. doi: 10.1093/nar/gkr1202. pmid:22144687
  42. 42. Sunderland KD, Fraser AM, Dixon AFG. Field and laboratory studies on money spiders (Linyphiidae) as predators of cereal aphids. J. Appl. Ecol. 1986;23: 433–447. doi: 10.2307/2404027
  43. 43. Gollan JR, Smith HM, Bulbert M, Donnelly AP, Wilkie L. Using spider web types as a substitute for assessing web-building spider biodiversity and the success of habitat restoration. Biodivers. Conserv. 2010;19: 3141–3155. doi: 10.1007/s10531-010-9882-1
  44. 44. Zevenbergen JM, Schneider NK, Blackledge TA. Fine dining or fortress? Functional shifts in spider web architecture by the western black widow Latrodectus hesperus. Anim. Behav. 2008;76: 823–829. doi: 10.1016/j.anbehav.2008.05.008
  45. 45. Winkler S, Kaplan DL. Molecular biology of spider silk. Rev. Mol. Biotechnol. 2000;74: 85–93. doi: 10.1016/s1389-0352(00)00005-2
  46. 46. Lewitus V. The black widow. Am. J. Nurs. 1935;35: 751–754. doi: 10.1097/00000446-193508000-00011
  47. 47. Vetter RS, Vincent LS, Danielsen DWR, Reinker KI, Clarke DE, Itnyre AA, et al. The prevalence of brown widow and black widow spiders (Araneae: Theridiidae) in urban southern California. J. Med. Entomol. 2012;49: 947–951. pmid:22897057 doi: 10.1603/me11285
  48. 48. Vink CJ, Derraik JGB, Phillips CB, Sirvid PJ. The invasive Australian redback spider, Latrodectus hasseltii Thorell 1870 (Araneae: Theridiidae): current and potential distributions, and likely impacts. Biol. Invasions. 2011;13: 1003–1019. doi: 10.1007/s10530-010-9885-6
  49. 49. Sirvid PJ, Vink CJ, Wakelin MD, Fitzgerald BM, Hitchmough RA, Stringer IAN. The conservation status of New Zealand Araneae. New Zeal. Entomol. 2012;35: 85–90. doi: 10.1080/00779962.2012.686310
  50. 50. Dondale CD. Black widow spiders: An outline of diversity. Biodivers. 2002;3: 17–20. doi: 10.1080/14888386.2002.9712562
  51. 51. Shokralla S, Spall JL, Gibson JF, Hajibabaei M. Next-generation sequencing technologies for environmental DNA research. Mol. Ecol. 2012;21: 1794–1805. doi: 10.1111/j.1365-294X.2012.05538.x. pmid:22486820
  52. 52. Krell B, Röder N, Link M, Gergs R, Entling MH, Schäfer RB. Aquatic prey subsidies to riparian spiders in a stream with different land use types. Limnologica. 2015;51: 1–7. doi: 10.1016/j.limno.2014.10.001
  53. 53. Moulder BC, Reichle DE. Significance of spider predation in the energy dynamics of forest floor arthropods communities. Ecol. Monogr. 1972;42: 473–498. doi: 10.2307/1942168
  54. 54. Nielsen ES, Laurence AM. Global diversity of insects: the problems of estimating numbers. In: Raven PH, editor. Nature and human society: The quest for a sustainable world. Washington DC: National Academy Press; 2000. pp. 213–222.
  55. 55. Kremen C, Colwell RK, Erwin TL, Murphy DD, Noss RF, Sanjayan MA. Terrestrial arthropod assemblages: their use in conservation planning. Conserv. Biol. 1993;7: 796–808. doi: 10.1046/j.1523-1739.1993.740796.x
  56. 56. Rosenberg DM, Danks HV, Lehmkuhl DM. Importance of insects in environmental impact assessment. Environ. Manage. 1986;10: 773–783. doi: 10.1007/bf01867730
  57. 57. Yu DW, Ji Y, Emerson BT, Wang X, Ye C, Yang C, et al. Biodiversity soup: metabarcoding of arthropods for rapid biodiversity assessment and biomonitoring. Methods Ecol. Evol. 2012;3: 613–623. doi: 10.1111/j.2041-210x.2012.00198.x