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Silk from Crickets: A New Twist on Spinning

  • Andrew A. Walker,

    Affiliations Evolution, Ecology and Genetics, Research School of Biology, Australian National University, Canberra, Australia, Ecosystem Sciences, Commonwealth Scientific and Industrial Research Organisation, Acton, Australia

  • Sarah Weisman,

    Affiliation Ecosystem Sciences, Commonwealth Scientific and Industrial Research Organisation, Acton, Australia

  • Jeffrey S. Church,

    Affiliation Materials Science and Engineering, Commonwealth Scientific and Industrial Research Organisation, Belmont, Australia

  • David J. Merritt,

    Affiliation School of Biological Sciences, University of Queensland, Brisbane, Australia

  • Stephen T. Mudie,

    Affiliation Australian Synchrotron, Clayton, Australia

  • Tara D. Sutherland

    Affiliation Ecosystem Sciences, Commonwealth Scientific and Industrial Research Organisation, Acton, Australia


Raspy crickets (Orthoptera: Gryllacrididae) are unique among the orthopterans in producing silk, which is used to build shelters. This work studied the material composition and the fabrication of cricket silk for the first time. We examined silk-webs produced in captivity, which comprised cylindrical fibers and flat films. Spectra obtained from micro-Raman experiments indicated that the silk is composed of protein, primarily in a beta-sheet conformation, and that fibers and films are almost identical in terms of amino acid composition and secondary structure. The primary sequences of four silk proteins were identified through a mass spectrometry/cDNA library approach. The most abundant silk protein was large in size (300 and 220 kDa variants), rich in alanine, glycine and serine, and contained repetitive sequence motifs; these are features which are shared with several known beta-sheet forming silk proteins. Convergent evolution at the molecular level contrasts with development by crickets of a novel mechanism for silk fabrication. After secretion of cricket silk proteins by the labial glands they are fabricated into mature silk by the labium-hypopharynx, which is modified to allow the controlled formation of either fibers or films. Protein folding into beta-sheet structure during silk fabrication is not driven by shear forces, as is reported for other silks.


The ability to produce silk has evolved in at least 23 groups of insects [1], in spiders [2] and in several other arthropods [3], [4]. Silk research has focused on silkworm cocoon and spider dragline silks, which have independently evolved a number of convergent features. Spider and silkworm silks consist of long, repetitive proteins that fold predominantly into beta-sheets, with the protein backbone parallel to the fiber axis [2]. Highly ordered nanocrystals are embedded in regions of less order and confer high tensile strength to the fibers [5]. The molecular arrangement in spider and silkworm silks is the result of shear forces and controlled dehydration acting on highly concentrated silk protein solutions as they pass through a hardened aperture known as a spinneret [6], [7]. Although less characterised, other silks are dramatically different. For example, protein backbones in silks made by glow-worms and adult lacewings are orientated perpendicular instead of parallel to the fiber axis [8]; the silks of fleas, bees and lacewing larvae contain proteins arranged in alpha-helices instead of beta-sheets [8], [9]; and the fibrous proteins in some silks are an order of magnitude smaller than spider dragline and silkworm cocoon silk proteins [10]. Further characterisation of silks in addition to spider and silkworm silks will allow a comparative approach to understanding the complex molecular arrangements found in silk.

Crickets in the family Gryllacrididae (raspy crickets) produce silk, while only one other insect in the order Orthoptera does so [11], [12]. Raspy crickets use silk fibers to build shelters into which they retreat during the day [12], [13]. The fibers are used variously to sew leaves together, to stabilise burrows in earth or sand, or to restrict access to tree hollows depending on species [12], [14]. The shelters are generally presumed to be a defense against predation, though it has also been suggested that they may limit desiccation in drier environments [12]. Both sexes are capable of producing fibers within hours of hatching and continue to produce shelters throughout their lives [15]. Shelters are highly valued and individuals may label their own shelters with a chemical cue [16] allowing them to return to the same shelter many times.

Very little is known about the method of fabrication of silk fibers by raspy crickets. Rentz and John [12] observed silk production from cricket mouthparts, but the origin of the material is unknown and the internal anatomy of raspy crickets is poorly described. Other insects that generate silk from their mouthparts do so using protein solutions produced in modified labial glands [17]. Wetas and king crickets (Anostostomatidae), the closest relatives of raspy crickets [18], use their labial glands to produce saliva [19]. Anostostomatid labial glands are arranged in grape-like clusters called acini [20]. Acinar cells secrete into the lumen of a branching series of ductules joined to the common duct on each side of the body. The left and right common ducts join at the labium, where they empty into a cavity between the labium and hypopharynx, called the salivarium. An additional organ, the reservoir, is formed by a sack-like outgrowth of the common duct on each side [19], [20].

Nothing is known about the material composition of raspy cricket silk fibers or how they are produced. We investigated the biochemistry and physical structure of raspy cricket fibers and the method of their production. Our motivation in this work was to enhance understanding of which features of different silks that have evolved independently in different arthropod groups are convergent and functional, and which features are historical and accidental.


Raspy cricket build shelters by using silk fibers and films to join other materials

Captive crickets built shelters by binding leaves or plastic card together with webs of silk (figure 1a). The fibers were cylindrical and uniform, with the diameter increasing as the crickets increased in size. For example, the diameter of fibers produced by a 16 mm long early instar Apotrechus were 4.2±1.1 µm. Seven months later, the same animal, measuring 48 mm, was producing fibers with diameter 12.4±2.1 µm.

Figure 1. Raspy cricket silk-webs.

(a) Shelter of silk and dry leaves made by Hyalogryllacris. Scale bar is 2 mm. (b) Scanning electron micrograph of Apotrechus silk-web showing cylindrical fibers and films. Scale bar is 100 µm.

Shelter construction behaviour was similar regardless of whether natural or artificial building materials were supplied. The process is shown in video S1. Silk production began by the cricket touching its labium to the surface of a leaf or piece of plastic, and depositing a film of silk material. As the labium was drawn away from the film, a fiber was produced, which was attached to another piece of building material with another film. Repeating this process resulted in a network of fibers joining the two pieces of building material. As successive layers of fibers were added, films were produced not only to secure fibers to building materials but also where fibers crossed, gluing them together (figure 1b). The end result of the building process was a ‘silk-web’ that served to seal an entry point to the cavity and to hold the building materials together. Most crickets constructed a shelter within 24 hours of being housed, incorporating additional fibers over successive days. To exit the shelter to forage, an access hole was cut through a silk-web using the mandibles. Access holes were sealed with a fresh silk-web after the insect returned. The insect ceased to produce fibers if the shelter was undisturbed for long periods, whilst removal of a shelter resulted in the construction of a replacement.

Fibers and films have a similar molecular structure

We investigated the chemistry of Apotrechus silk using Raman spectroscopy. Spectra obtained from fibers and films indicated a highly proteinaceous material, with no peaks attributed to chitin or other substances (figure 2). Spectra from films were essentially the same as those obtained from fibers indicating that the two materials contained proteins with the same secondary structures. Strong peak vibrations at 1667 cm−1 and 1235 cm−1 indicated that the dominant protein conformation present was beta-sheet [21] and the weak shoulder at 1259 cm−1 was due to disordered protein. Raman spectroscopy is sensitive to a number of specific amino acids and their environments within the protein structure, particularly tyrosine, tryptophan, phenylalanine, histidine, proline, hydroxyproline, methionine and cysteine [22], [23]. No significant difference in the relative strengths of frequencies attributed to any specific amino acid were detected in the spectra obtained from fibers and films, suggesting that the fiber and films had the same amino acid composition.

Figure 2. Raman spectra of raspy cricket fibers and films.

Amide peak positions are indicated.

Fibers were anisotropic, having a birefringence of 0.013±0.002 in the long-slow direction, suggesting that protein chains in fibers are aligned relative to the axis of the fiber. Birefringence was not detected in films, but because films are much thinner than fibers it is unclear if this is because films are isotropic or because our instruments were not sensitive enough to detect their birefringence. Raman spectra obtained from various regions of the film randomly orientated with respect to any adjoining fibers were found to be identical. As the laser is polarized, this infers that there is no protein chain alignment in the films and that films are anisotropic at the molecular level. Silk proteins in beta-sheet conformation can be aligned with the peptide backbone either parallel (extended beta-sheet structure) or perpendicular (cross beta-sheet structure) to the axis of the fiber [8]. The measured WAXS pattern of cricket silk with the fiber axis vertical showed the strong arc attributed to the spacing between beta-strands in the direction of hydrogen bonds on the equator (table 1 and figure S1). This indicates that crystallites in cricket silk are arranged in an extended beta-sheet structure. The placing of another equatorial arc, attributed to the (2 2 0) reflection, depends in part on the inter-sheet spacing between amino acid side groups. Calculations from the position of this reflection suggests that in Apotrechus silk the inter-sheet spacing is 1.27 nm.

Silk is produced from acinar labial glands

Silk made out of protein must have a glandular origin. Visual observations during dissection of Apotrechus and Hyalogryllacris revealed large, acinar labial glands in the thorax and head in a similar arrangement to those of anostostomatids (figure 3a). Ductules from the acini on each side of the body merged into a common duct ending at the labium. Whereas the reservoirs of anostostomatids are connected to the common acinar duct at some distance from the labium [20], raspy cricket reservoirs are joined to the end of the common duct within the labium. At this point the reservoirs and the common ducts from both sides of the body join together and empty into the salivarium through a common aperture. The size of this aperture was much larger than the diameter of silk fibers, measuring in excess of 100 µm across in adult Apotrechus (figure 3b). No structure similar to the hardened, external spinneret of lepidopterans was observed. Instead, the labium was similar to crickets that do not produce silk, with hypopharynx, glossae and paraglossae [24], except that the paraglossae had an unusual shape: the margins of the paraglossae were raised up into ridges that overlap the edges of the hypopharynx, so that the hypopharynx fits ‘hand in glove’ into the labium (figure 3c and 3d). To compare the structure of the labium of crickets that do not produce silk, we dissected a field cricket (Acheta domestica), a katydid (Conocephalus sp.) and an anostostomatid (Penalva flavocalceata). In these species, the surface of the paraglossae is flat or concave, without the distinctive raised margins that occur in raspy crickets.

Figure 3. Raspy cricket labial glands.

(a) Labial glands and labium of Hyalogryllacris; scale bar is 2 mm. (b) Scanning electron micrograph of a preparation from Apotrechus in which the hypopharynx has been pinned back to show the aperture through which the labial glands empty into the salivarium; scale bar is 500 µm. (c) Labium and hypopharynx in open position, showing the raised margins on the labial paraglossae of Hyalogryllacris; scale bar is 500 µm. (d) Same preparation as (c), labium and hypopharynx in closed position, with raised margins of paraglossae overlapping hypopharynx; scale bar is 500 µm. (e) Confocal slice through labial acinus of Apotrechus showing paired arrangement of nuclei (blue) and secretory invaginations (red); scale bar is 50 µm. (f) Projection of 18 images in z-series showing small cuticle-secreting cells lining the acinar labial duct of Apotrechus; scale bar is 50 µm. aci = acini; res = reservoirs; lb = labium; h = hypopharynx; lp = labial palp; a = aperture of labial glands into salivarium; pg = paraglossae of labium.

Secretory cells have a distinctive morphology, with actin-rich secretory membranes and large or multiple nuclei [25]. To investigate the function of the acini and the reservoirs, we stained them with fluorescent dyes that reveal nuclei and the actin cytoskeleton. Acinar cells were organised in pairs, possessed a single large nucleus, to which an actin-rich invagination was closely apposed, consistent with a secretory role (figure 3e). The lobular, actin-rich lumen of each cell conjoined to a common lumen with a stellate, actin-rich periphery that was continuous with the common duct. Each acinus was composed of approximately 20 of these conjoined cells. The ducts were lined with smaller, flattened, cuticle-secreting cells, the luminal surface of which was actin-rich (figure 3f). The reservoirs consisted of a single layer of the same small cuticle-secreting epidermal cells, suggesting a role as a storage organ.

The proteins present in fluid-filled reservoirs were investigated by comparing the sequences of peptides following tryptic digestion to GenBank's non-redundant protein database and in silico translated sequences from our own Apotrechus cDNA library using LC/MS. Apart from house-keeping proteins, the only protein identified was amylase, an enzyme that breaks down the plant polymer starch (table S1). This suggests that reservoirs store saliva. As none of the raspy cricket silk proteins (described below) were detected in the reservoirs, it is unlikely that they store silk proteins.

Raspy cricket silk proteins

The raspy cricket silk proteins were identified using a cDNA library/mass spectroscopy approach that has been applied to other silk-producing species [26]. We constructed a cDNA library of 3.5×105 clones with an average size of 1.1±0.6 kb from Apotrechus acinar labial glands. Analysis of over 100 clones identified 63 putative cDNAs, 22 of which had significant homology to sequences present in GenBank's non-redundant DNA database (E<0.05), mostly to proteins with house-keeping functions (see table S2). A single exception was the sequence encoding an amylase that was detected in fluid-filled reservoirs. Of the other 41 cDNAs that did not have significant homology to known sequences, 17 encoded one of four silk proteins (see below).

Solubilized silk-webs contained major protein bands at approximately 300, 220, and 68 kDa and fainter bands at 120, 30 and 28 kDa (figure 4). Similar protein bands were obtained from silk produced by different individuals and regardless of whether or not protease inhibitors were included during solubilization or reducing agents added during SDS-PAGE. Individual protein bands from SDS-PAGE gels or solid silk-webs were digested with proteases and the resulting peptides were analysed using mass spectroscopy. Comparison of experimentally-derived peptide masses with the predicted masses of in silico digested sequences encoded by the labial gland cDNA library showed that the bands at 300, 220 and 120 kDa all corresponded to a single protein sequence, partially encoded by three cDNA clones (figure 4). We named this protein AilSP1 (Apotrechus illawarra Silk Protein 1; GenBank accessions JF508439 and JF508440). The longest AilSP1 cDNA contained 1282 nucleotides of coding region, a stop codon, a 3′ untranslated region of 315 nucleotides, and a poly-A tail. Two of the AilSP1 cDNAs differed by only by a single silent polymorphism (GenBank accession JF508439) while the third contained 77 single nucleotide polymorphisms (31 silent) and a trinucleotide insert (GenBank accession JF508440). The partial sequence of AilSP1 consisted of an internal repetitive region of 37 or 38 residue repeats followed by a 163 residue C-terminal region that contained a single cysteine residue (figure S2). The small amino acids alanine, glycine and serine made up 24.3%, 23.4%, and 18.5% of residues respectively.

Figure 4. Identification of silk proteins by LC/MS.

Silk proteins separated by SDS-PAGE of solubilized silk-webs are shown on the left. The number of peptides detected and the SpectrumMill score for each band are shown on the right. Spectrum Mill scores higher than 20 indicate a confident identification.

Three other silk proteins that occur at lower abundance were identified. The 68 kDa protein corresponded to a protein sequence we named AilSP2 (figure 4; GenBank accession JF508441). AilSP2 was encoded by 10 cDNA clones, seven of which began with a predicted signal sequence (SignalP 3.0 HMM p = 0.999) and ended with a stop codon, 3′ untranslated region, and poly-A tail, and were presumed to encode the full-length protein sequence. The predicted mature sequence of AilSP2 comprised 404 amino acids and had a predicted mass of 41.4 kDa. The 28 and 30 kDa protein bands both corresponded to a protein sequence we named AilSP3 (figure 4; GenBank accession JF508442), represented by 3 cDNA clones. The longest of these clones encoded a predicted signal sequence (SignalP3.0 HMM P = 0.982), stop codon, 3′ untranslated region and poly-A tail. The predicted mature AilSP3 protein had 203 residues (14 of them cysteine) and a predicted mass of 22.4 kDa. A final silk protein, AilSP4 (GenBank accession JF508443) was identified in silk-webs but not in the solubilized silk (figure 4). AilSP4 was represented by a single partial 1.8 kb cDNA sequence that consisted of a partial coding region, putative stop codon, 3′ untranslated region, and poly-A tail. The partial protein sequence of AilSP4 is high in proline (11.2%) and serine (19.9%) compared to glycine (9.9%) and alanine (7.8%).


The raspy cricket species used in this study produced shelters by joining leaves together with silk-webs. Given that the silk-webs were not air-tight, they are unlikely to be effective in preventing desiccation or the ingress of parasites in the wild, and the most likely function is to reduce predation. Silk-webs were found to be made of protein, and visual observations of crickets fabricating silk suggested that the labial glands might function as silk glands. Our identification of transcripts encoding silk proteins in labial glands confirms this directly.

The silk gland consists of acini connected by a network of ducts to the insect's salivarium. Silk glands from species that make cocoons have large lumens that store silk proteins in preparation for a short period of intense silk production [27]. In contrast, raspy cricket gland lumens are small, the amount of silk required for shelters is low, and animals probably produce silk as required. A reservoir attached to the common duct immediately before it joins the salivarium does not contain silk proteins but does contain amylase, suggesting that it has a salivary function.

The silk-webs consisted of fibers and films, with the fibers providing the mechanical backbone of the webs and the films serving to glue the fibers to other building materials and to each other. Micro-Raman spectra indicated that the proteins present in fibers and films are indistinguishable at the levels of amino acid composition and secondary protein structure, suggesting they are made from the same protein solution. We propose that the anatomical arrangement in raspy crickets is specialised to be able to produce fibers and films interchangeably. Instead of having an external, tubular spinneret like lepidopterans and spiders [28], [29] the labial ducts of raspy crickets end in an aperture that is too large to act as a draw-down taper. Instead, silk dope probably exists in a liquid state in the salivarium chamber created by tucking the hypopharynx under the raised margins of the labial paraglossae. Single fibers are formed by drawing through the taper at the extremity of the labium, between the tips of the two paraglossae and the hypopharynx (figure 5). Opening the labium and hypopharynx by muscular control allows the insect to deposit silk dope in globules, which dry into films. Both films and fibers are found in the silks of other insects, including sawflies, honeybees, hornets and some beetles, e.g. [30].

Figure 5. Fabrication of fibers and films by raspy crickets.

(a) Indicative position of labial gland acini (aci) and reservoirs (res) and associated ducts. (b) Breakdown of raspy cricket mouthparts, including labium (lb), maxillae (mx), mandibles (md) and labrum (lr). (c) Expanded view of the labium with the hypopharynx (h) in closed position, forming a draw-down taper with the labial paraglossae (pg) to allow fiber fabrication. Liquid silk in the salivarium chamber between paraglossae and hypopharynx is shown in purple. lp = labial palp; rd = reservoir duct; ad = acinar duct. (d) Expanded view of the labium with the hypopharynx in open position, allowing silk dope to flow over paraglossae, allowing fabrication of films.

The major silk protein AilSP1 has evolved convergent features to those of the dominant fibrous proteins from silkworm cocoon and spider dragline silks: it is large, contains a high proportion of small amino acids, and adopts primarily an extended beta-sheet conformation in the mature silk. The spacing between beta-sheets in the side group direction is 1.27 nm, within the range reported for lepidopteran beta-sheet silks (0.93–1.57 nm) and slightly greater than that of polyalanine (1.06 nm) [31], [32]. If crystallites are formed from the repeat units of AilSP1 this is a reasonable estimate, since 27% of residues in repeat regions are glycine, 27% are alanine, and 18% are serine, with the remainder larger residues. The presence of a cysteine residue in the C-terminus of AilSP1 and the high frequency of cysteines in AilSP3, as well as the observation that reducing agents are required for silk solubilization, suggest that disulfide bonding plays a structurally important role in cricket silk.

If raspy crickets produce silk films by depositing globules of liquid silk dope which are subsequently allowed to dry, then proteins in films are exposed to minimal shear and compression forces. Since the secondary structure of the proteins in fibres and films is essentially the same, shear forces cannot be a primary mechanism driving beta-sheet formation, as it is for the proteins in silkworm cocoon fibers [33]. Some insect species produce silk consisting of proteins with alpha-helical [34] and cross-beta [35] molecular structures, which shear forces are likely to disrupt rather than favour. In these species, the final molecular structure must form due to some combination of dehydration and the ordering of proteins within the gland prior to silk fabrication. Similar mechanisms may drive beta-sheet formation by raspy cricket silk proteins. Although not required for beta-sheet formation, shear force most likely accounts for the long range alignment of proteins within the fibers demonstrated by WAXS patterns and by birefringence measurements.

A comparison between raspy cricket silk and two unrelated insect silks, produced by silkworms and webspinners, is shown in Table 2. The three types of insects have been faced with a similar problem, the need to produce an insoluble and stable building material, and have independently evolved solutions with some convergent features. All three insect families produce silk proteins, with primary fibroin sizes ranging from moderately large to enormous, and effective size further increased due to cysteine cross-linking. The fibroins are rich in particular amino acids and in repetitive motifs that foster formation of extensive intermolecular beta-sheets, which assemble into insoluble and stable beta-sheet crystallites. On the other hand, there is wide variation between the three silks in regard to the gland of production, form of spinneret, and morphology of the mature silk product. Convergent evolution of silks has occurred at the molecular level; however the silk fabrication process and its anatomical substrates are highly flexible and idiosyncratic.

Table 2. Comparison of silks composed of beta-sheet forming proteins.

Methods and Methods


Raspy crickets of the species Apotrechus illawarra and Hyalogryllacris species 9 were collected from Meroo National Park, NSW, Australia. Crickets were housed in plastic jars with water and orthopteran food mixture [36]. They were supplied either with natural building materials (dry leaves, sticks and sand) or black plastic card. Non-gryllacridid crickets used for comparison were field crickets, Acheta domestica (Pisces Enterprises, Kenmore, Queensland, Australia), katydids of the genus Conocephalus (collected at the Meroo National Park field site) and white-kneed king crickets, Penalva flavocalceata (Minibeast Wildlife, Brisbane, Australia). Species used for comparison were dissected immediately following collection or receipt.


Silk glands and mouthparts from adult insects of either sex were dissected and fixed in phosphate buffered saline (PBS) pH 7.0 containing 2% gluteraldehyde (Invitrogen, Carlsbad, USA) and examined using light microscopy. Silk-webs were mounted onto a stub using conductive tape, sputter-coated with gold and visualised under high vacuum on a Zeiss Evo LS15 scanning electron microscopy (SEM; Zeiss, Jena, Germany). Silk glands from a single large Apotrechus nymph were examined on a confocal scanning laser microscope (Carl Zeiss LSM5, Microimaging GmbH, Jena, Germany). The tissues were dissected and fixed in 4% formaldehyde in PBS for 10 minutes, washed three times for 30 minutes each in PBS, permeabilized for 1 hour in PBT (0.25% BSA, 0.4% Triton X in PBS), blocked for 1 hr in PBT-NGS (2% normal goat serum in PBT). After fixation, tissues were stained overnight at 4°C in 6.6 µM Alexa Fluor 568 phalloidin (Invitrogen, Carlsbad, USA), 1 µg/ml diamidino-2-phenylindole (DAPI; Invitrogen, Carlsbad, USA) in PBS. Before mounting, stained tissue was washed four times for 10 minutes each in PBS and placed in 70% glycerol, 2% propyl gallate in PBS for one hour in the dark.

Fiber birefringence was quantified using a Leica M205C polarising light microscope (Leica, Wetzlar, Germany) with a full wave compensator installed underneath the sample stage, so that its slow axis was aligned at +45° between the crossed polarising filters. Fiber diameters were measured using the Leica LAS software and birefringence calculated with reference to a Leica colour chart.

Raman Spectroscopy

Raman spectra were obtained using an inVia confocal microscope system (Renishaw, Gloucestershire, UK) with 514 nm excitation from an argon ion laser through a ×50 (0.75 na) objective. Incident laser power was 4.5 mW and coaxial backscatter geometry was employed. Spectra were collected over the range 3200 to 100 cm−1 and averaged over at least 5 scans, each with an accumulation time of 20 seconds. Raman shifts were calibrated using the 520 cm−1 line of a silicon wafer. The spectral resolution was ∼1 cm−1. Fibers were analysed aligned parallel to the direction of laser polarisation with the use of a rotating stage. The final spectra used for analysis were averages of spectra collected from 6–10 different areas. All spectra were normalized on the CH2 and CH3 deformation modes at 1452 cm−1 that are attributable to amino acid side chains and thus not sensitive to protein conformation.

X-ray scattering

Parallel bundles of Apotrechus silk fibers were analysed on the SAXS/WAXS beamline of the Australian Synchrotron (Melbourne, Australia). A wavelength of 0.124 nm and a nominal sample to detector distance of 0.559 m provided a q-range (q = 4πsinθ/λ) of approximately 0.01 to 0.22 nm−1, which was calibrated using a silver behenate standard. Samples were mounted in air, perpendicular to the beam, with scattering patterns collected in transmission. An optical microscope alignment system was used to accurately position samples in the X-ray beam. A background profile obtained without a sample was subtracted from experimental profiles to account for air scattering. Peak positions were measured using the Australian Synchrotron's 15ID SAXS/WAXS software.

cDNA library construction

A cDNA library was constructed from the labial glands of four large Apotrechus illawarra nymphs. To induce silk production, the shelters of these crickets were removed and they were supplied with fresh building materials. One day later the insects were dissected in PBS pH 7.0 and the acinar parts of the labial glands removed and stored in RNAlater (Ambion, Austin, USA). Total RNA was prepared using RNAqueous-4PCR (Ambion, Austin, USA) and mRNA isolated using Micro-FastTrack 2.0 (Invitrogen, Carlsbad, USA). The cDNA library was created using Cloneminer II cDNA Library Construction kit (Invitrogen, Carlsbad, USA) according to the manufacturer's instructions. Clones containing cDNA inserts larger than 1 kb were sequenced by Micromon Services (Monash University, Melbourne, Australia) using an Applied Biosystems 3730S Genetic Analyser and Applied Biosystems PRISM BigDye Terminator Mix cycling chemistry. A library of possible protein sequences was generated in silico using EMBOSS Transeq [37]. Putative cDNAs were identified by the presence of a poly-A tail longer than 15 continuous nucleotides and/or the presence of an open reading frame longer than 300 bp. Signal peptides were predicted using the SignalP 3.0 algorithm [38].

Mass spectroscopy

Cricket silk was solubilized in saturated lithium bromide with 5% 2-mercaptoethanol (both from Sigma-Aldrich, St. Louis, USA) at 95°C for one hour, after which no solid material could be observed. Silk-webs were not soluble in guanidinium hydrochloride or sodium dodecyl sulphate (SDS) supplemented with 2-mercaptoethanol, or in saturated lithium bromide in the absence of a reducing agent. In some experiments the denaturing solution included a protease inhibitor cocktail (Complete mini EDTA-free; Roche Applied Science, Penzberg, Germany). Lithium bromide was removed from solubilized samples by repeated concentration and dilution using a Centricon-10 centrifugal filter device (Millipore, Belerica, USA). Solubilized cricket silk proteins were separated by polyacrylamide gel electrophoresis (PAGE) using NuPage 4–12% Bis-Tris gels and 2-(N-morpholino)ethanesulphonic acid running buffer (Invitrogen, Carlsbad, USA) and stained with Coomassie Brilliant Blue (Sigma-Aldrich, St. Louis, USA). Protein gel bands or solid silk samples were digested with either trypsin or α-chymotrypsin (Sigma-Aldrich, St. Louis, USA) and analysed by reversed phase liquid chromatography coupled by electrospray ionisation to ion trap tandem mass spectrometry as previously described [34]. Mass spectral data sets were analysed using Spectrum Mill software (Agilent, Santa Clara, USA).

Supporting Information

Figure S1.

WAXS pattern from Apotrechus silk fiber bundle. Fiber axis is vertical. The d-spacings of scattering peaks are marked.


Figure S2.

Raspy cricket silk protein sequences. Conserved repeats in AilSP1 are highlighted in yellow, and predicted signal peptides for AilSP2 and AilSP3 are highlighted in green. Because disulfide bonding has a structural role in cricket silk, cysteine residues are highlighted in blue.


Video S1.

Hyalogryllacris species 9 fabricating silk. This video appears at 6× actual speed.


Table S1.

Proteins identified in fluid-filled reservoirs by LC/MS.


Table S2.

Assignment of Apotrechus illawarra cDNA non-silk library sequences.



We would like to thank Ros Mourant, Dr. David Rentz, Dr. You Ning Su, Alison Rowell and Dr. John Trueman for advice and assistance in collecting raspy crickets, Dr. Paul Cooper and Dr. Eric Hines for help with dissection and photography, and Dr. Sri Sriskantha and Dr. Xiaoyi Wang for assistance with laboratory procedures.

Author Contributions

Conceived and designed the experiments: AAW TDS SW DJM . Performed the experiments: AAW JSC STM SW DJM. Analyzed the data: AAW SW JSC DJM STM TDS. Contributed reagents/materials/analysis tools: TDS. Wrote the paper: AAW SW TDS.


  1. 1. Sutherland TD, Young JH, Weisman S, Hayashi CY, Merritt DJ (2010) Insect silk: One name, many materials. Annual Review of Entomology 55: 171–188.
  2. 2. Vollrath F, Porter D (2006) Spider silk as archetypal protein elastomer. Soft Matter 2:
  3. 3. Clotuche G, Goff GL, Mailleux AC, Deneubourg JL, Detrain C, et al. (2009) How to visualize the spider mite silk? Microscopy Research and Technique 72: 659–664.
  4. 4. Weygoldt P (1966) Spermatophore web formation in a pseudoscorpion. Science 153: 1647–1649.
  5. 5. Nova A, Keten S, Pugno NM, Redaelli A, Buehler MJ (2010) Molecular and nanostructural mechanisms of deformation, strength and toughness of spider silk fibrils. Nano Letters 10: 2626–2634.
  6. 6. Jin HJ, Kaplan DL (2003) Mechanism of silk processing in insects and spiders. Nature 424: 1057–1061.
  7. 7. Fu C, Shao Z, Vollrath F (2009) Animal silks: their structures, properties and artificial production. Chemical Communications 6515–6529.
  8. 8. Rudall KM (1962) Silk and other cocoon proteins. In: Florkin M, Mason HS, editors. Comparative Biochemistry. New York: Academic Press. pp. 397–433.
  9. 9. Weisman S, Trueman HE, Mudie ST, Church JS, Sutherland TD, et al. (2008) An unlikely silk: the composite material of green lacewing cocoons. Biomacromolecules 9: 3065–3069.
  10. 10. Hayashi CY, Shipley NH, Lewis RV (1999) Hypotheses that correlate the sequence, structure, and mechanical properties of spider silk proteins. International Journal of Biological Macromolecules 24: 271–275.
  11. 11. Rentz DC, Weissman DB (1973) The origins and affinities of the Orthoptera of the channel Islands and adjacent mainland California. Part I. The Genus Cnemotettix. Proceedings of the Academy of Natural Sciences of Philadelphia 125: 89–120.
  12. 12. Rentz DCF, John B (1990) Studies in Australian Gryllacrididae: taxonomy, biology, ecology and cytology. Invertebrate Taxonomy 3: 1053–1210.
  13. 13. Morton SR, Rentz DCF (1983) Ecology and taxonomy of fossorial, granivorous gryllacridids (Orthoptera: Gryllacrididae) from arid central Australia. Australian Journal of Zoology 31: 557–579.
  14. 14. Hale RJ, Rentz DCF (2001) The Gryllacrididae: An overview of the world fauna with emphasis on Australian examples. In: Field LH, editor. The Biology of Wetas, King Crickets and their allies. New York: CABI. pp. 95–110.
  15. 15. Hale RJ (2000) Nest utilisation and recognition by juvenile gryllacridids (Orthoptera: Gryllacrididae). Australian Journal of Zoology 48: 643–652.
  16. 16. Lockwood JA, Rentz DCF (1996) Nest construction and recognition in a gryllacridid: The discovery of pheromonally mediated autorecognition in an insect. Australian Journal of Zoology 44: 129–141.
  17. 17. Sehnal F, Sutherland T (2008) Silks produced by insect labial glands. Prion 2: 145–153.
  18. 18. Jost MC, Shaw KL (2006) Phylogeny of Ensifera (Hexapoda: Orthoptera) using three ribosomal loci, with implications for the evolution of acoustic communication. Molecular Phylogenetics and Evolution 38: 510–530.
  19. 19. Field LH (2001) The biology of wetas, king crickets and their allies. New York: CABI Publishing.
  20. 20. Maskell FG (1927) The anatomy of Hemideina thoracica. Transactions and Proceedings of the New Zealand Institute 57: 637–670.
  21. 21. Frushour BG, Koenig JL (1975) Raman Spectroscopy of Proteins. In: Clark RSH, Hester RE, editors. Advances in Infrared and Raman Spectroscopy. New York: Heyden. pp. 35–97.
  22. 22. Twardowshi J, Anzenbacher P (1994) Raman and IR Spectroscopy in Biology and Biochemistry. New York: Ellis Horwood.
  23. 23. Church JS, Corino G, Woodhead AL (1997) The Raman spectra of cortical and cuticle cells and some protens isolated from wool. Biopolymers 42: 7–17.
  24. 24. Britton EB, Brown WL, Calaby JH, Colless DH, Common IFB, et al. (1970) Insects of Australia: a text book for students and research workers. Carlton: Melbourne University Press.
  25. 25. Akai H (1998) Silk glands. In: Harrison FW, editor. Microscopic anatomy of invertebrates. New York: Wiley-Liss.
  26. 26. Sutherland TD, Weisman S, Trueman HE, Sriskantha A, Trueman JW, et al. (2007) Conservation of essential design features in coiled coil silks. Molecular Biology and Evolution 24: 2424–2432.
  27. 27. Akai H (1983) The structure and ultrastructure of the silk gland. Experientia 39: 443–449.
  28. 28. Sorensen GS, Cribb BW, Merritt DJ, Johnson ML, Zalucki MP (2006) Structure and ultrastructure of the silk glands and spinneret of Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae). Arthropod Structure & Development 35: 3–13.
  29. 29. Coddington JA (1989) Spinneret silk spigot morphology: Evidence for the monophyly of orbweaving spiders, Cyrtophorinae (Araneidae), and the group Theridiidae plus Nesticidae. Journal of Arachnology 17: 71–95.
  30. 30. Ishay JS, Ganor E (1990) Comb cells and puparial silk in the oriental hornet nest: Structure and function. Journal of Morphology 203: 11–19.
  31. 31. Lucas F, Rudall KM (1968) Extracellular fibrous proteins: The silks. In: Florkin M, Stotz EH, editors. Comprehensive Biochemistry. Amsterdam: Elsevier. pp. 475–558.
  32. 32. Colonna-Cesari F, Premilat S, Lotz B (1975) Conformational analysis of the beta-sheet structure of poly-alanine and poly(-alanine-glycine). Journal of Molecular Biology 95: 71–82.
  33. 33. Asakura T, Kaplan DL (1994) Silk production and processing. Arutzen CJ, editor. New York: Academic Press. pp. 1–11.
  34. 34. Sutherland TD, Campbell PM, Weisman S, Trueman HE, Sriskantha A, et al. (2006) A highly divergent gene cluster in honey bees encodes a novel silk family. Genome Research 16: 1414–1421.
  35. 35. Weisman S, Okada S, Mudie ST, Huson MG, Trueman HE, et al. (2009) Fifty years later: The sequence, structure and function of lacewing cross-beta silk. Journal of Structural Biology 168: 467–475.
  36. 36. Rentz DCF (1996) Grasshopper country: the abundant orthopteroid insects of Australia. Sydney: University of New South Wales Press.
  37. 37. Rice P, Longden I, Bleasby A (2000) EMBOSS: the European Molecular Biology Open Software Suite. Trends in Genetics 16: 276–277.
  38. 38. Emanuelsson O, Brunak S, von Heijne G, Nielsen H (2007) Locating proteins in the cell using TargetP, SignalP, and related tools. Nature Protocols 2: 953–971.
  39. 39. Zhou CZ, Confalonieri F, Medina N, Zivanovic Y, Esnault C, et al. (2000) Fine organization of Bombyx mori fibroin heavy chain gene. Nucleic Acids Research 28: 2413–2419.
  40. 40. Collin MA, Garb JE, Edgerly JS, Hayashi CY (2009) Characterization of silk spun by the embiopteran, Antipaluria urichi. Insect Biochemistry and Molecular Biology 39: 75–82.
  41. 41. Okada S, Weisman S, Trueman HE, Mudie ST, Haritos VS, et al. (2008) An Australian webspinner species makes the finest known insect silk fibers. International Journal of Biological Macromolecules 43: 271–275.
  42. 42. Marsh RE, Corey RB, Pauling L (1955) An investigation of the structure of silk fibroin. Biochimica et Biophysica Acta 16: 1–34.
  43. 43. Tanaka K, Kajiyama N, Ishikura K, Waga S, Kikuchi A, et al. (1999) Determination of the site of disulfide linkage between heavy and light chains of silk fibroin produced by Bombyx mori. Biochimica et Biophysica Acta: Protein Structure and Molecular Enzymology 1432: 92–103.
  44. 44. Nagashima T, Niwa N, Okajima S, Nonaka T (1991) Ultrastructure of silk gland of webspinners, Oligotoma japonica (Insecta: Embioptera). Cytologia 56: 679–685.
  45. 45. Moriya M, Ohgo K, Masubuchi Y, Knight DP, Asakura T (2008) Micro-computerized tomographic observation of the spinning apparatus in Bombyx mori silkworms. Polymer 49: 5665–5669.