Terebra steering in chalcidoid wasps

Eggs, Benjamin; Fischer, Stefan; Csader, Michael; Mikó, István; Rack, Alexander; Betz, Oliver

doi:10.1186/s12983-023-00503-1

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Published: 08 August 2023

Terebra steering in chalcidoid wasps

Frontiers in Zoology volume 20, Article number: 26 (2023) Cite this article

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Abstract

Various chalcidoid wasps can actively steer their terebra (= ovipositor shaft) in diverse directions, despite the lack of terebral intrinsic musculature. To investigate the mechanisms of these bending and rotational movements, we combined microscopical and microtomographical techniques, together with videography, to analyse the musculoskeletal ovipositor system of the ectoparasitoid pteromalid wasp Lariophagus distinguendus (Förster, 1841) and the employment of its terebra during oviposition. The ovipositor consists of three pairs of valvulae, two pairs of valvifers and the female T9 (9th abdominal tergum). The paired 1st and the 2nd valvulae are interlocked via the olistheter system, which allows the three parts to slide longitudinally relative to each other, and form the terebra. The various ovipositor movements are actuated by a set of nine paired muscles, three of which (i.e. 1st valvifer-genital membrane muscle, ventral 2nd valvifer-venom gland reservoir muscle, T9-genital membrane muscle) are described here for the first time in chalcidoids. The anterior and posterior 2nd valvifer-2nd valvula muscles are adapted in function. (1) In the active probing position, they enable the wasps to pull the base of each of the longitudinally split and asymmetrically overlapping halves of the 2nd valvula that are fused at the apex dorsally, thus enabling lateral bending of the terebra. Concurrently, the 1st valvulae can be pro- and retracted regardless of this bending. (2) These muscles can also rotate the 2nd valvula and therefore the whole terebra at the basal articulation, allowing bending in various directions. The position of the terebra is anchored at the puncture site in hard substrates (in which drilling is extremely energy- and time-consuming). A freely steerable terebra increases the chance of contacting a potential host within a concealed cavity. The evolution of the ability actively to steer the terebra can be considered a key innovation that has putatively contributed to the acquisition of new hosts to a parasitoid’s host range. Such shifts in host exploitation, each followed by rapid radiations, have probably aided the evolutionary success of Chalcidoidea (with more than 500,000 species estimated).

Background

The evolution of parasitoidism in Hymenoptera has led to one of the largest species radiations in insects [1,2,3,4,5]. A large proportion of parasitoids belong to the Chalcidoidea, an extremely diverse and ecologically important group (nearly 27,000 species described, over 500,000 species estimated) of mainly minute wasps (average body size range from 1–2 mm) that are omnipresent in almost all terrestrial habitats [6,7,8,9,10,11]. Most chalcidoids are ectoparasitoids of other insects, usually attacking enclosed host stages with reduced mobility (i.e. egg or larval stages of wood and stem borers, leaf-miners or inhabitants of galls, seeds and fruits) [12], although other life stages are also targeted [13]. The parasitization of hosts living deep within substrates allows the ectoparasitoid larvae to develop within the protection of a concealed environment and without exposure to the host immune system as occurs in endoparasitoids. An evolutionary novelty and presumably a strong driver of diversification in Chalcidoidea is the secondary reversal to monocondylic mandibles (reduction of the posterior condyle accompanied with modified musculature with functional separation), which allow the emerging wasp to bite through a hard substrate by precise cutting movements that overcome the limitations of a single degree of freedom [14]. However, the use of hosts living concealed within hard substrates poses challenges not only for the emerging wasp (i.e. in leaving the substrate), but also for females attempting to parasitize them (i.e. entering the substrate to find a potential host) [15]. In this context, the ovipositor has to fulfil several functional demands: penetration or navigation through the substrate or the target egg/puparium, assessment of the host, discrimination between suitable and previously parasitized hosts, piercing of the host, injection of venom, formation of a feeding tube for host feeding, ovicide or larvicide of the competitors’ eggs or larvae, respectively, marking of the host and find a suitable place for egg laying and oviposition [16]. However, putative evolutionary novelties of the chalcidoid ovipositor system, such as morphological and behavioural adaptations that enable the steering of the terebra (= ovipositor (shaft) sensu [17,18,19,20,21,22,23,24,25,26,27]) and its underlying mechanisms have not been thoroughly investigated hitherto.

As in all hymenopterans, the chalcidoid ovipositor consists of the female T9 (9th abdominal tergum; = outer ovipositor plates sensu [17,18,19,20,21,22,23,24,25]), two pairs of valvifers and three pairs of valvulae derived from the 8th and 9th abdominal segments (7th and 8th metasomal segments). The basally situated valvifers accommodate the operating musculature, whereas all the valvulae are devoid of intrinsic musculature [28,29,30,31,32]. The 1st valvifers (8th gonocoxites [33, 34] or the fusion of the same with the gonangula [30]; = fulcral plates sensu [17,18,19,20,21,22,23,24,25, 35,36,37]; = gonangulum, gonangula sensu [26, 27]) are anteriorly continuous with the rami of the 1st valvulae (8th gonapophyses; = stylets sensu [17,18,19,20,21,22,23,24,25, 35,36,37,38]; = lower valves sensu [26, 27]), and their posterior angles articulate dorsally with the female T9 via the tergo-valvifer articulation and ventrally with the 2nd valvifers via the intervalvifer articulation. The 2nd valvifers (9th gonocoxites; = inner ovipositor plates sensu [17,18,19,20,21,22,23,24,25]) extend as the 3rd valvulae (9th gonostyli; = (articulating/terminal) palps sensu [19, 20, 22, 23, 36]; = ovipositor sheaths sensu [26, 27]) and are ventrally articulated with the 2nd valvula (fusion of the 9th gonapophyses; = (stylet) sheath sensu [17,18,19,20,21,22,23,24,25, 36,37,38]; = upper valve sensu [26, 27]) [28, 29], which is asymmetrically split except at the apex in all chalcidoid families [39]. The two overlapping asymmetric halves of the 2nd valvula are connected dorsally by the notal membrane, which extends almost to the apex [17, 19,20,21,22,23,24,25, 40]. The interlocked 1st and 2nd valvulae enclose the egg canal and form the terebra, which is embraced by the 3rd valvulae when not in use. The ventral surface of the 2nd valvula is interlocked with both of the 1st valvulae by a sublateral longitudinal tongue called the rhachis, which runs within a corresponding groove called the aulax along the dorsal surface of each of the 1st valvulae. This so-called olistheter system allows the three elements of the terebra to slide longitudinally relative to each other and simultaneously prevents their unwanted separation [29, 39].

In order to reach their hosts and permit greater control over egg placement, several parasitoid wasps are able actively to bend and rotate their terebra in any direction relative to their body axis [41,42,43,44], despite the lack of intrinsic terebral musculature. Such terebra movements have also been reported in chalcidoid wasps of the family of Pteromalidae [40, 45,46,47], a polyphyletic group sensu lato [6, 10, 48, 49] (over 3500 species described [8]). However, little is known about the actuation of the various ovipositor movements, with the mechanisms involved in terebra steering (i.e. bending and rotating) remaining unclear. In this study, we investigated the working mechanisms of the terebra steering movements of Lariophagus distinguendus (Förster, 1841) (Chalcidoidea: Pteromalidae: Pteromalinae), a cosmopolitan synanthropic synovigenic autogenous solitary idiobiont larval and pupal ectoparasitoid of several granivorous coleopteran species [50, 51]. This species exhibits extensive terebra movements during the assessment of a potential host and eventual subsequent egg placement [45, 47]. We aimed (1) to analyse the oviposition process in vivo, (2) to describe the ovipositor of L. distinguendus, including all inherent cuticular elements and muscles, (3) to examine the mechanics and mode of function of the musculoskeletal system, including the actuation of the various ovipositor movements, (4) to investigate the underlying working mechanisms of the terebra steering movements and (5) to discuss their eco-evolutionary significance.

Results and discussion

We combined behavioural analyses involving high-resolution video recordings with morphological investigations based on microscopical and microtomographical techniques. These studies have enabled us to present a thorough morphological and mechanical analysis of the musculoskeletal ovipositor system that steers the various movements executed by the female L. distinguendus (Fig. 1) during oviposition. In particular, we focused on the employment of the terebra and on its form, structure and material properties.

Morphological terms are applied according to the Hymenoptera Anatomy Ontology (HAO; [52,53,54]; a table of all 210 terms relevant to the hymenopteran ovipositor system, their definitions and 513 synonyms commonly found in literature is given in Table 2 in the Appendix 1).

In cases in which our findings have been confirmed by other studies, these are indicated below with ‘cf.’.

Oviposition process and employment of the terebra

Previous studies describing the behavioural sequences of the attempts of L. distinguendus [45, 50, 55, 56] and other pteromalids [40, 46, 57] to oviposit have been unable to provide an analysis of the events that take place within the cavity of the substrate. Therefore, we mainly focus on the employment of the terebra and its movements in the following (Fig. 2; Additional file 1).

Search for the host's habitat: L. distinguendus parasitizes concealed granivorous host larvae (Fig. 2a). The parasitoids mainly use volatile chemicals to locate the habitat of their hosts: faecal cues from the host itself and herbivory-associated chemicals in the seed induced by the mechanical damage caused by the host larvae [51, 55, 58].

Search for an infested substrate: Once L. distinguendus finds the host’s location (infested grains; blotting paper with the host faeces in our experimental setup), the wasp starts to walk on the substrate followed by antennal drumming with the flagellum directed towards the ground (Additional file 1, min. 0:05–0:07; cf. [45, 47, 50, 55, 56]). The female parasitoid is able to discriminate between healthy and infested grains [59].

Penetration of the substrate: Once the female wasp has selected a small spot with its antennae, it brings its terebra into the drilling position by a downward bending of the metasoma so that its tip taps the surface. The terebra is guided and stabilized by the 3rd valvulae in order to prevent buckling despite axial compressive forces occurring during the initial puncturing of the substrate, i.e. the pericarp of the grain. Once the apex of the terebra is engaged in the substrate, the metasoma with the 2nd valvifer and the attached 3rd valvulae are lifted upwards out of the way (Fig. 2b–d; Additional file 1, min. 0:06–0:11; cf. [40, 45, 47, 50, 55, 56]). The initial puncturing (i.e. pericarp surface penetration) is necessary for the 1st and 2nd valvulae to be anchored in the substrate so that the subsequent ‘push-pull’ mechanism can be initiated. Thereby, the wasp exhibits alternate reciprocal movements of the paired 1st valvulae, which can be seen as trembling movements of the posteroventral part of the metasoma (i.e. the 2nd valvifers and the female T9). Only one of the 1st valvulae is pushed into the substrate at a time, while the other 1st valvula and the 2nd valvula, which are anchored in the substrate, are simultaneously pulled [60,61,62]. The apical sawteeth thereby increase the friction with the surrounding substrate. The tension in the two anchored ‘stationaryʼ elements increases their bending stiffness and, hence, they can serve as guides for the particular 1st valvula being pushed into the substrate [44, 60]. Small pushing movements of the 2nd valvula caused by the relative movements of the 2nd valvifers cannot be excluded (cf. [32]). The simultaneous pushing and pulling of the various terebral elements minimizes the net compressive force on the substrate and thus the chance of buckling of the terebra [32, 44, 60]. The ‘push-pull’ mechanism enables drilling without torque and with very low axial load, although these cannot be completely avoided [44, 62]. During the drilling process (Additional file 1, min. 0:12–0:20; cf. [47]), the wasp combines the ‘push-pull’ mechanism with slight rotations of the terebra [44, 60]. Moreover, a fluid is constantly secreted at the apex and also along the shaft of the terebra. This secretion putatively prevents particles from entering the terebra but might also act as a (cooling) lubricant (cf. [63, 64]).

Search for a potential host within the substrate: As soon as the wasp has penetrated the grain in which a potential host larva is living, it attempts to locate the host larva in its concealed cavity with its terebra (Additional file 1, min. 4:02–4:32). Thereby, the metasoma is frequently rotated by up to 35° from the longitudinal body axis of the wasp (cf. [45]); this influences the orientation of the terebra. However, the wasp also expresses steering movements of the terebra in several directions that are independent of the orientation of the metasoma (see subchapter ‘Mechanisms of terebra bending and rotation’ below).

Penetration of the targeted host's skin: Once the wasp has succeeded in reaching its host, it pushes its terebra straight down to its fullest extent and penetrates the skin of the beetle larva several times with rapid stabbing movements of the terebra (Additional file 1, min. 0:21–0:27, 2:33–2:47; cf. [47, 55]) achieved by fast alternate movements of the 1st valvulae.

Injection of venom: The host larva is usually pierced several times (cf. [45, 47]), with the 1st valvulae performing fast alternate movements. Venom is injected into the host’s body and permanently paralyses the host (Additional file 1, min. 0:21–0:27, 2:33–2:47; cf. [47]) thereby preventing its further development. This is crucial for ectoparasitoids, since movements of the host larva within a small cavity might damage the externally attached parasitoid [65].

Assessment of the host: The permanent paralysis of the host larva presumably allows an easier and more accurate assessment to be carried out by the female wasp, which can now actively steer its terebra (Fig. 2e–l; Additional file 1, min. 0:30–1:10, 2:48–3:17; cf. [47]; see subchapter ‘Mechanisms of terebra bending and rotation’ below). However, some passive bending of the terebra might also occur because of its deflection on the host surface. A small actively actuated bending of the apex of the terebra would therefore be sufficient to indicate the direction of the bending movement. The assessment of the host is not primarily carried out by the terebra tapping of the host surface, but by the puncture and the assessment of the host’s haemolymph (cf. [66, 67]).

Formation of a feeding tube for host feeding: In most parasitization attempts, the female wasps create a feeding tube. Thereby, a secretion, which is produced by the large colleterial glands [68], oozes from the entire terebra [66, 69]. The terebra is moved up and down and is also putatively rotated to a certain degree to ensure an even distribution of the secretion, which hardens in the air and remains for a couple of minutes, forming a feeding tube (cf. [47]). As a result of capillary forces, the haemolymph of the host flows upwards within the tube. The wasp now appears to lick the end of the feeding tube. The absorbed haemolymph serves both as protein-rich nutrition that is needed for egg maturation [70] and allows an assessment of the quality of the potential host [66, 67].

Ovicide/larvacide of the competitors' eggs/larvae: In our artificial setup, we have not tested whether the female wasps attempt to kill their conspecifics’ eggs or larvae. Ovicidal and larvicidal behaviour has not as yet been observed in L. distinguendus.

Search for a suitable place for oviposition: If the female wasp deems the host larva to be of good quality, it searches for a suitable oviposition site on the host surface. It appears to estimate the available space within the cavity to ensure that the growing larva has enough room for development (Fig. 2e–l; Additional file 1, min. 3:19–3:34; cf. [47]).

Oviposition: Rapid longitudinal alternate movements of the paired 1st valvulae serve to pass the egg along the terebra (Fig. 2m–p; Additional file 1, min. 1:56–2:26, 3:36–4:00; cf. [47, 71]). The diameter of the egg is significantly larger compared with that of the egg canal. The egg is thus strongly deformed during ovipositing. It does not emerge at the very apex of the terebra but is pushed out ventrally between the two paired 1st valvulae in a region about 100–200 µm proximal to the apex (Fig. 2p). Finally, the egg is attached to the surface of the host. In a few cases, it was also observed to be attached to the surface of the cavity near the host larva. Finally, the wasp withdraws its terebra. Female L. distinguendus only lay one egg per host [45, 55].

Morphological structure of the musculoskeletal ovipositor system

The musculoskeletal ovipositor system of L. distinguendus consists of three pairs of valvulae, two pairs of valvifers, the female T9, three paired articulations and a set of nine paired muscles.

Because of its bilateral bauplan, all the ovipositor elements and muscles are paired apart from the distal region of the 2nd valvula and the female T9. Paired morphological structures are only described in the singular form in the following, i.e. the elements of the left side only, although they have a mirror image on the right side.

The anatomy of the venom system and of the female internal reproductive system is not discussed thoroughly in the following (for chalcidoids, see [19,20,21,22,23,24,25, 35,36,37,38, 72,73,74,75,76,77,78,79]; for parasitoid hymenopterans in general, see [26, 27]).

Cuticular elements of the ovipositor

1st valvula (1vv; Figs. 2m–p, 3a, b, f, 4a–d, g–k, and 5a, c): Basally, the thin 1st valvula is continuous with the 1st valvifer via its dorsal ramus (dr1; Figs. 3d, e, g, 5a, c, d, and 6c, j). The 1st valvula has a crescent-shaped cross-section over most of its length (1vv; Fig. 4c). The aulax (au; Figs. 3a and 4g, i, k) of L. distinguendus does not reach the apex of the 1st valvula but tapers off around 50 µm before it. The distal end of the aulax features a coeloconic sensillum (cs; Figs. 3a and 4i, j; sensu [80]), presumably monitoring the position of the 1st valvula relative to the 2nd valvula (cf. [81]). Further sensilla can be seen at regular intervals on the lateral sides (blue ‘notches’ in Fig. 3f), which might have a mechano- and/or chemosensory function. However, the sensillar equipment of the terebra was not further investigated in this study (but see [82,83,84]). Dorsomedially to the aulax, the medial wall of each 1st valvula is thickened (Fig. 4c). The ventral part of the medial wall is thin and formed into a large membranous fold (when at rest) that projects inwards and overlaps ventrally (Fig. 4a–c; cf. [39]). These thin chitinous folds are considered effectively to seal the crack between the paired 1st valvulae in order to prevent the loss of venom and/or oviposition fluids [39]. The 1st valvula laterally bears two small sawteeth (st1; Fig. 3a) that are of decreasing size at its apex and that are most probably used to penetrate the substrate and the host’s skin. On the dorsomedial side of their apices, the 1st valvulae are connected by the olistheter-like interlock of the 1st valvulae (il1; Fig. 4h, i, k), presumably preventing them from being torn apart during the initial puncturing of the substrate and during drilling. The egg exits the egg canal proximad to these structures and ventrally between the paired 1st valvulae (Fig. 2p; Additional file 1, min. 1:56–2:26, 3:36–4:00). Such interlocking structures are also found in other pteromalids and some species of Aphelinidae, Chalcididae, Eulophidae, Eurytomidae, Ormyridae, Tanaostigmatidae and Trichogrammatidae [39]. In all chalcidoids, the ventral ramus of the 1st valvula is completely reduced [29] and the valvilli inside the egg canal are absent [85].

2nd valvula (2vv; Figs. 2m–p, 3a, c, f–h, 4a–g, i, k, and 5a–c): Proximally, the bulbs of the 2nd valvula (blb; Figs. 3g, h, 4a, and 6e–h) are basally articulated with the 2nd valvifer via the basal articulation (ba; Figs. 3h and 6f). At its basal part, the 2nd valvula bears the processus articuaris laterally on the bulbs, and the processus musculares dorsally on the anteriorly directed horn-like processes of the bulbs. On its ventral side, the 2nd valvula bears the rhachises (rh; Figs. 3c and 4g). The 2nd valvula of L. distinguendus consists of two longitudinally split, asymmetrically overlapping and more-heavily sclerotized halves (2vv; Figs. 3a, f–h and 4a–c; Additional file 2) that are thickened medially (2vv; Fig. 4c). The two halves are dorsally connected for most of their length by a conjunctiva called the notal membrane (nm; Fig. 4f) [17, 19,20,21,22,23,24,25, 40] but are fused at the apex (2vv; Figs. 3a and 4d, i, k). Proximally, the notal membrane is modified into a transversely striate band called the laminated bridge (lb; Figs. 3g, h, 4a, b, e, 5a–c, and 6c, f, h) [19, 20, 22, 40]. The modified 2nd valvula with its longitudinally split and overlapping halves presumably permit a greater distortion of the valvula and appear to be a synapomorphy for all Chalcidoidea, except for Mymaridae [39]. The ventral side is formed by the ventral wall of the 2nd valvula (vw2; Fig. 4a–c, f; sensu [86]), which extends from the base almost to the apex. This creates a lumen (lu2; Fig. 4a–d, f, i, k). The rhachises are attached to this lamella-like process over most of their length, except for the apex. Ventrolaterally to the rhachises lie lateral extensions of the 2nd valvula (le; Fig. 4c; sensu [86]). The apex of the 2nd valvulae of L. distinguendus features seven sawteeth that are placed laterally and staggered relative to one another (st2; Figs. 3a, f and 4i) with sensilla being found in between them. The laterally positioned sawteeth are postulated to act like a screw during the alternate rotational movements of the terebra during substrate penetration [22] and seem to be present in all chalcidoid species that undertake drilling actions [25] (Additional file 3).

Terebra (trb; Figs. 1, 2d–p, 3a and 6c–h): The acicular terebra consists of the paired 1st valvulae and the 2nd valvula and has a smooth surface. The terebra of L. distinguendus (and other chalcidoid wasps) features a single opening at the basal end, where the common oviduct (co; Fig. 5) seamlessly merges with the base of the egg canal (cf. [19,20,21,22,23, 25]). In chalcidoid wasps (such as L. distinguendus and Microterys flavus (Howard, 1881) (Encyrtidae) (data not yet published)), both the orifice of the venom gland reservoir (ovr; Fig. 5b–d; Additional file 4, min. 0:30–0:31) and the dorsolaterally situated Dufour’s gland duct (Dgd; Fig. 5) empty into the common oviduct (cf. [73, 77]) before the latter fuses with the egg canal (unlike in ichneumonoid wasps; cf. [16, 87]). The junction lies directly anterior to the basal articulation (ba; Figs. 3h and 6f) and is indicated by the furcula (Fig. 3g; Additional file 2, min. 0:21–0:36). The complete length of the egg canal thus functions as a conduit not only for the egg itself, but also for the expulsion of venom or other fluids during oviposition. The diameter of the terebra is even along its length (Fig. 4c; Additional file 3) between the broad basal bulbs (Figs. 3g, h and 4a,b) and the distally tapering apex (Figs. 3a and 4d, i, k). The rhachises (rh; Figs. 3c and 4g) on the ventral side of the 2nd valvula are interlocked with the aulaces (au; Figs. 3a, b and 4g, i, k) on the dorsal side of the opposing 1st valvulae via the olistheter system (oth; Fig. 4c); this enables the 1st valvulae to move along the 2nd valvulae while they are still connected to each other. The olistheters of L. distinguendus does not extend along the entire length of the terebra but end around 50 µm before its apex (Fig. 3a). The distally directed scale-like structures on the contact surfaces of both the rhachises and the aulaces (sc; Fig. 3b, c) presumably reduce frictional forces by minimizing the contact area of the olistheter elements [88]. However, these scale-likes structures potentially also forward a liquid lubricant from the colleterial glands (= accessory glands) to the apex of the olistheter system further to reduce friction in between the moving valvulae (cf. [89]). This arrangement might also enable particles to be continuously flushed out the olistheter system during drilling or venom injection. The scale-like structures might additionally create anisotropic conditions in the olistheter and thus prevent the 1st valvulae from randomly sliding back during drilling and piercing (cf. [31]). The longitudinally split and asymmetrically overlapping halves of the 2nd valvula presumably allow lateral sliding to occur towards or away from each other. Moreover, the rachises of L. distinguendus are suspended from lamellar structures of the ventral wall of the 2nd valvula (vw2; Fig. 4b, c) over their entire length, except for the apex (Fig. 4d). Thus, both the 1st and 2nd valvulae, which are connected via the olistheter system, are presumably movable in their position and may diverge tangentially. Moreover, the dorsally thickened walls of the 1st valvulae can be bent away from the midline and, in doing so, can take up the ventral membranous slack, further increasing the volume inside. This is thought to be an adaptation in several chalcidoid taxa to facilitate deformation of the terebra and temporarily to enlargement of the egg canal (ec; Fig. 4c), which is mainly formed by the two paired 1st valvulae, in order to accommodate the passing egg [39]. The olistheter system thereby must sustain the forces exerted by the egg [62, 71]. However, the maximal diameter of the apical half of the terebra is limited by the diameter of the puncture site in the substrate during oviposition. The areas of the rhachises at the basal bulbous part of the 2nd valvula presumably are also flexible (purple areas of the cuticle in Fig. 4a, b presumably indicating a higher resilin content). The internal microsculpture of the medial wall of the egg canal consists of distally orientated leaf-like ctenidia (ct; Fig. 3c, h) that contain large amounts of resilin (ct; Fig. 3h; Additional file 2, min. 0:05–0:20) and are found from the proximal basis to the region before the apex. The ctenidia help to push the deformable egg along the egg canal by alternate movements of the 1st valvulae, prevent regression [71, 88] and are also hypothesized to forward a liquid lubricant for the moving valvulae and thus to reduce friction [88, 90] and/or to produce a feeding tube. Both the 1st and 2nd valvulae have tapered apices. The terebra apex in many hymenopteran taxa is heavily sclerotized and hardened with metal atoms, such as calcium (Ca), manganese (Mn) and zinc (Zn). This enables the piercing of hard substrates, reduces wear and tear and prevents buckling [15, 62, 81, 91,92,93,94].

3rd valvula (3vv; Figs. 3a, 4d and 6a–d): The relatively short semi-tubular 3rd valvula of L. distinguendus emerges at the posterior end of the 2nd valvifer (Fig. 6a–d) and ensheaths and protects the distal part of the terebra when at rest (Fig. 4d). The distally directed microsetae on the medial surface of the 3rd valvula (Fig. 3a) are thought to be involved in the cleaning of the terebra between oviposition episodes [16, 83]. The 3rd valvula might also have a sensory function [27].

1st valvifer (1vf; Figs. 3d and 6a–d, i, j): The 1st valvifer of L. distinguendus and other chalcidoids is bow-shaped [17, 19,20,21,22,23,24,25, 35,36,37]. The anteroventral angle of the 1st valvifer features a horizontal ridge, which has a medial–lateral orientation (Fig. 6i) and which is part of the tergo-valvifer articulation (tva; Figs. 3d and 6b, i, j). The posteroventral corner of the 1st valvifer is bifurcated (Fig. 6i) and is part of the intervalvifer articulation (iva; Figs. 3d, g and 6b, i, j). The interarticular ridge (iar; Fig. 6i) lies between the two articulations and might serve mechanically to stabilize the 1st valvifer. The anterodorsal angle of the 1st valvifer is continuous with the dorsal ramus of the 1st valvula (dr1; Figs. 3d, e, g, 5a, c, d, and 6c, j), which is interlocked with the dorsal projection of the 2nd valvifer (dp2; Fig. 5c, d; cf. [31]) by a system analogous to the olistheter. This tight interlocking guides the dorsal ramus and prevents it from buckling when pushing forces are applied during the protraction of the 1st valvula. Since the dorsal ramus constantly slides around the proximal bulbous end of the 2nd valvula during pro- and retraction, the ramus needs to be flexible in the sagittal plane and thus presumably contains high proportions of the elastic rubber-like protein resilin in its cuticle (cf. [95,96,97,98]).

2nd valvifer (2vf; Figs. 3d, 4a–c, 5a, c, d and 6a–d, f): The 2nd valvifer is elongated and its posterior part is placed medially of the female T9 (Fig. 6b). A conjunctive, called the genital membrane (not shown), connects the ventral margins of the paired 2nd valvifers arching above the 2nd valvula. The anterior part of the 2nd valvifer of L. distinguendus extends dorsally in a semi-circular shape and dorsally bears the dorsal projection of the 2nd valvifer (dp2; Fig. 5c, d), which is interlocked with the dorsal ramus of the 1st valvula via an interlocking system similar to the olistheter (cf. [31]). At its posterodorsal end and posterior to its medial ridge (mr2; Fig. 6f), the anterior part of the 2nd valvifer features the post-ramus flap (prf; Figs. 3d and 6b; sensu [22]), on which the dorsal projection continues, thus allowing a greater arc of movement of the 1st valvifer and therefore a greater retraction of the 1st valvula. The 2nd valvifer features two sensillar patches: (1) the sensillar patch (sp; Fig. 3d) located anteroventrally to the intervalvifer articulation (iva; Figs. 3d, g and 6b, i, j) and (2) the row of sensilla (sr; Fig. 3e) on the dorsal margin of the 2nd valvifer. These two sensillar patches are in contact with the ventromedial side of the 1st valvifer and the dorsal ramus of the 1st valvula, respectively, and probably monitor the movements of the 1st valvula indirectly. The dorsal margins and the dorsal flanges are strengthened by cuticular ridges that putatively have a stabilizing function and prevent deformation (i.a. at the intervalvifer articulation). The posterodorsal ends of the 2nd valvifers are connected by the median bridge (mb2; Fig. 6c). The venom gland reservoir (vr; Fig. 5a, b; Additional file 2, min. 0:37–0:52; Additional file 4, min. 0:24–0:31; = acid gland reservoir sensu [19,20,21,22,23,24,25, 73]) is situated in between the 2nd valvifers with its proximal end lying near the base of the terebra. The Dufour’s gland (Dg; Fig. 5a; Additional file 4, min. 0:21–0:31; = alkaline gland sensu [19,20,21,22,23,24,25, 73]) is situated dorsolaterally to the venom gland reservoir (cf. [77, 79]).

Female T9 (T9; Figs. 3d and 6a–d): The female T9 of L. distinguendus is U-shaped and situated lateral to the posterior part of the 2nd valvifers (Fig. 6b). Its elongated anteriorly projecting arms articulate with the 1st valvifers via the tergo-valvifer articulations (tva; Figs. 3d and 6b, i, j). The cordate apodeme (not shown) on the anterior margin of the female T9 is located posterior to the articulation. The dorsal margins are strengthened by the anterior flange of T9, which presumably mechanically stabilizes the female T9 during oviposition. Medially, the anterior flange of T9 bifurcates and forms a dorsomedial crest-like ridge that runs almost the entire length of the female T9. This ridge serves as a muscle attachment area both medially and laterally and presumably increases the mechanical stability of the female T9.

Articulations of the musculoskeletal ovipositor system

Basal articulation (ba; Figs. 3h and 6f): The two articular surfaces of this ball-and-socket-like articulation are located on the socket-like pars articularis of the anteroventral part of the 2nd valvifer and the ball-like processus articulated laterally on the bulb of the 2nd valvula. This rotational joint presumably also allows some pivotal and rotational movements of the 2nd valvula and thus of the whole terebra.

Intervalvifer articulation (iva; Figs. 3d, g and 6b, i, j): The 1st and 2nd valvifer are connected via the intervalvifer articulation, a rotational joint that allows a rotation of the 1st valvifer in the sagittal plane only [32]. This articulation consists of the bifurcated posteroventral corner of the 1st valvifer (iva; Fig. 6i), which encloses the articulation site at the 2nd valvifer. Thereby, one furcal structure of the 1st valvifer is placed medially and one laterally to the 2nd valvifer.

Tergo-valvifer articulation (tva; Figs. 3d and 6b, i, j): The 1st valvifer lies adjacent to the female T9 via the tergo-valvifer articulation, which is situated dorsally to the intervalvifer articulation. It is a rotational joint that allows the 1st valvifer to rotate in the sagittal plane only [32]. This articulation consists of a horizontal ridge at the 1st valvifer (tva; Fig. 6i) and a corresponding counterpart at the female T9 situated near the cordate apodeme.

Ovipositor muscles

In total, nine paired ovipositor muscles have been identified that drive and actuate the associated skeletal apparatus (Table 1). Three of these muscles (i.e. the 1st valvifer-genital membrane muscle, the ventral 2nd valvifer-venom gland reservoir muscle and the T9-genital membrane muscle) are described here for the first time in chalcidoids.

Table 1 Ovipositor muscles of Lariophagus distinguendus (abbreviations in brackets) and their origin, insertion (cf. Additional file 4) and presumed function as verified in the present contribution

Full size table

1st valvifer-genital membrane muscle (m-1vf-gm; Figs. 4c and 6d, e, f): This muscle is the only muscle of the 1st valvifer. It originates at the medial surface of the posteroventral part of the 1st valvifer, i.e. at the centre between the tergo-valvifer and the intervalvifer articulation (Fig. 6c, d, f), and inserts anteriorly on the genital membrane (Fig. 4c). We here describe the m-1vf-gm for the first time in Chalcidoidea. Previous authors (e.g. [17, 19,20,21,22,23,24,25, 36]) might have overlooked its presence because of to its minute size.

Dorsal 2nd valvifer-venom gland reservoir muscle (m-d-2vf-vr; Figs. 5a–c and 6d, e, f): This muscle originates at the medial surface of the most anterior part of the 2nd valvifer (Fig. 6c–f) and inserts dorsally at the anterior part of the venom gland reservoir (Fig. 5a, b), which is located ventrally of the common oviduct. Most previous authors (e.g. [17, 19,20,21,22,23,24,25]) have overlooked the presence of this muscle; it was only mentioned by [73].

Ventral 2nd valvifer-venom gland reservoir muscle (m-v-2vf-vr-a/b; Figs. 5a, c, d and 6d–f): This muscle forms two distinct bundles. Its anterodorsal part (m-v-2vf-vr-a) originates at the medial surface of the most anterior part of the 2nd valvifer, ventrally to the origin region of the dorsal 2nd valvifer-venom gland reservoir muscle (Fig. 6d–f), and inserts laterally at the orifice the venom gland reservoir (Fig. 5c, d). The other part (m-v-2vf-vr-b) originates at the medial surface of the anterior part of the 2nd valvifer, posteroventrally to the origin region of part a (Fig. 6d–f), and inserts laterally at the orifice of the venom gland reservoir, ventrally to the insertion of part a and shortly before the orifice of the venom gland reservoir enters the common oviduct (Fig. 5c, d). To our knowledge, this muscle has also not yet been described in chalcidoids (but see [99,100,101] for the description of a similar set of muscles in ants).

Anterior 2nd valvifer-2nd valvula muscle (m-a-2vf-2vv; Fig. 6d, f, g, h): This muscle originates at the medial region along the anterodorsal arch of the 2nd valvifer (Fig. 6c, d) and inserts at the processus articularis, located laterally on the bulbs of the 2nd valvula (Fig. 6f–h).

Posterior 2nd valvifer-2nd valvula muscle (m-p-2vf-2vv; Fig. 6d, f, g, h): This muscle originates at the medial region along the ventral part of the 2nd valvifer (Fig. 6c, d) and inserts at the processus musculares, located dorsally on the anteriorly directed horn-like processes of the bulbs of the 2nd valvula (Fig. 6f–h).

Dorsal T9-2nd valvifer muscle (m-d-T9-2vf-a/b; Fig. 6d): This muscle is modified in its insertion and forms two distinct muscle bundles. One part of this muscle (m-d-T9-2vf-a) originates at the lateral region along the posterodorsal part of the female T9, i.e. laterally along its dorsomedial ridge (Fig. 6a–d), and inserts at the anterior section of the dorsal flange of the 2nd valvifer, posterior to its medial ridge (Fig. 6c, d, f). The other part (m-d-T9-2vf-b) originates at the medial region along the posterodorsal part of the female T9, i.e. ventromedially to its dorsomedial ridge (Fig. 6c, d), and inserts at the anterior section of the dorsal flange of the 2nd valvifer via a tendon (t-m-d-T9-2vf-a; Fig. 3g), located ventrally to the insertion region of m-d-T9-2vf-a (Fig. 6c, d).

Ventral T9-2nd valvifer muscle (m-v-T9-2vf; Fig. 6d): This muscle originates at the cordate apodeme, which is located at the anterior margin of the female T9, posteriorly to the tergo-valvifer articulation (Fig. 6c, d), and inserts at the medial surface along the posterior section of the dorsal flange of the 2nd valvifer (Fig. 6c, d).

Posterior T9-2nd valvifer muscle (m-p-T9-2vf; Fig. 6d): This muscle originates at the medial surface of the posterodorsal part of the female T9 (Fig. 6c, d) and inserts at the median bridge of the 2nd valvifers. Previous studies on the chalcidoid ovipositor [17, 19,20,21,22,23,24,25] report only one muscle originating in the posterior region of the female T9. The authors presumably were unable to distinguish this muscle from the T9-genital membrane muscle described below.

T9-genital membrane muscle (m-T9-gm; Fig. 6d): This muscle originates at the medial surface of the posterodorsal part of the female T9, dorsally of the origin region of the posterior T9-2nd valvifer muscle (Fig. 6c, d), and inserts posteriorly at the genital membrane. We here describe the m-T9-gm for the first time in Chalcidoidea.

Mechanics and mode of function of the ovipositor system

The set of nine paired ovipositor muscles in L. distinguendus comprises two pairs of two antagonistically working muscles that are mainly responsible for the various ovipositor movements, three muscles stabilizing the musculoskeletal system, and two muscles related to the function of the venom gland reservoir (Table 1).

Depression and elevation of the terebra: The 2nd valvula is connected with the 2nd valvifer by a rotational joint called the basal articulation (ba; Figs. 3h, 6f and 7a). Two muscles (m-a-2vf-2vv, m-p-2vf-2vv) insert at the bulbous region around this articulation. The insertion region of the posterior 2nd valvifer-2nd valvula muscle (m-p-2vf-2vv; Fig. 6f–h) at the 2nd valvula is located dorsal of the basal articulation, whereas its region of origin at the 2nd valvifer is located posteroventral to it (Fig. 6c, d). Taxa from other superfamilies use the m-p-2vf-2vv to depress their terebra towards an active probing position (e.g. Ichneumonoidea [31, 32]). However, female L. distinguendus have never been observed to depress their terebra in such a manner. Instead, these wasps bend their whole metasoma downwards to bring their terebra into the drilling position. Once the apex of the terebra is engaged in the substrate, the metasoma is lifted upwards again, while the terebra remains in its depressed position (Fig. 2b–d; Additional file 1, min. 0:06–0:11; cf. [45, 47]). This behaviour has also been reported for other pteromalids [40, 46] and species of Torymidae [23], Eurytomidae [21], Encyrtidae (data not yet published) and Eulophidae [19]. Therefore, in pteromalids (and possibly also in other chalcidoid taxa), we assume that the m-p-2vf-2vv is adapted in its function (see paragraph ‘Rotation of the terebra’ of the subchapter ‘Mechanisms of terebra bending and rotation’ below). During this indirect depression of the terebra, the bulbs of the 2nd valvula might be pulled out of the socket-like anterior ends of the 2nd valvifer ventrally by pushing them slightly apart, resulting in a slight translation of the pivot point (= joint axis or fulcrum) of the basal articulation (cf. [32]). The insertion region of the anterior 2nd valvifer-2nd valvula muscle (m-a-2vf-2vv; Fig. 6f–h) at the 2nd valvula is situated posteroventrally of both the basal articulation and the insertion region of m-p-2vf-2vv, whereas its origin at the 2nd valvifer is located posterodorsally of this articulation (Fig. 6c, d). After an oviposition attempt, the terebra is withdrawn from the substrate. Since slender structures such as the terebra can support much higher tensile than compressive stresses, the withdrawal does not damage it [62]. A contraction of the anterior 2nd valvifer-2nd valvula muscle (F_m-a-2vf-2vv; Fig. 7a) presumably initiates the elevation of the terebra (arrow 9; Fig. 7a; Table 1). The passive rebound of the bulbs of the 2nd valvula into the socket-like anterior ends of the 2nd valvifer presumably further supports the elevation of the terebra passively and helps to stabilize it in its resting position (cf. elevation of the terebra in ceraphronoids, which completely lack the m-a-2vf-2vv [102]). The anatomical cluster comprising the 2nd valvifer, the 2nd valvula and the two muscles connecting them is a simple mechanical system in which the 2nd valvula is a two-armed class 1 lever, whereby the effective (= mechanical) inlever arm and the joint angle (attachment angle) of m-a-2vf-2vv change over the range of motion (cf. [31]).

Pro- and retraction of the 1st valvulae: Three muscles (m-d-T9-2vf, m-v-T9-2vf, m-p-T9-2vf) connect the 2nd valvifer with the female T9. Both of these cuticular structures are connected with the 1st valvifer via the intervalvifer articulation and the tergo-valvifer articulation (iva/tva; Figs. 3d, 6b, i, j and 7a), respectively. The insertion region of both parts of the dorsal T9-2nd valvifer muscle (m-d-T9-2vf-a/b; Fig. 6c, d) at the 2nd valvifer are situated anterodorsally, whereas their regions of origin at the female T9 are located posterodorsally of both articulations (Fig. 6c, d). A simultaneous contraction of m-d-T9-2vf-a and m-d-T9-2vf-b (summarized as F_m-d-T9-2vf; Fig. 7a) slides the 2nd valvifer posteriorly with respect to the female T9 (arrow 1; Fig. 7a). This causes the 1st valvifer to tilt anteriorly (arrow 2; Fig. 7a), because it is articulated with both the 2nd valvifer and the female T9 via rotational joints. The 1st valvifer acts as a lever that transforms its tilting movement to the dorsal ramus of the 1st valvula (arrow 3; Fig. 7a). Its tight interlocking with the dorsal projection of the 2nd valvifer prevents it from buckling and transmits the movements to the apex of the 1st valvula, causing it to slide distally relative to the 2nd valvula, i.e. to protract (arrow 4; Fig. 7a; Table 1). In the active probing position, the dorsal ramus is less curved, which presumably reduces friction [32]. The region of origin of the antagonistically acting ventral T9-2nd valvifer muscle at the female T9 (m-v-T9-2vf; Fig. 6c, d) is situated posterodorsally near the intervalvifer articulation and posterior to the tergo-valvifer articulation, whereas its insertion region at the 2nd valvifer is located posteroventrally of both these articulations (Fig. 6c, d). Its contraction (F_m-v-T9-2vf; Fig. 7a) slides the 2nd valvifer anteriorly with respect to the female T9 (arrow 5; Fig. 7a), thus indirectly causing the 1st valvifer to tilt posteriorly (arrow 6; Fig. 7a) and the 1st valvula to slide proximally relative to the 2nd valvula, i.e. to retract (arrows 7, 8; Fig. 7a; Table 1). The vibration-like rapid reciprocal alternate pro- and retracting movements of the 1st valvulae are crucial for drilling and precise egg laying (Fig. 2m–p; Additional file 1, min. 1:11–1:35, 1:56–2:22, 3:36–4:00; cf. [32, 44, 47]). The following assumptions have been made for a simplified estimation of the torques (M) exerted by the forces of the dorsal and ventral T9-2nd valvifer muscles (F_m-d-T9-2vf/F_m-v-T9-2vf; Fig. 7a): (1) The 2nd valvifer acts as the frame of reference; therefore, the intervalvifer articulation (iva; Figs. 6i, j and 7a) acts as a pivot point around which the 1st valvifer tilts; (2) the movements of 2nd valvifer and the female T9 are constrained by the musculoskeletal system and cannot twist around the articulations but only slide telescopically towards or against each other along the anterior–posterior axis; and (3) frictional forces in the system can be neglected. In reality, all cuticular elements can move relatively to each other. However, under these assumptions, the horizontal force vector components acting in the anterior–posterior axis (F_{m-d-T9-2vf(x)-in}/F_{m-v-T9-2vf(x)-in}; Fig. 6j) act at the 1st valvifer at the tergo-valvifer articulation (tva; Figs. 6i, j and 7a). Therefore, the torques (M) of F_m-d-T9-2vf and F_m-v-T9-2vf that act at the intervalvifer articulation in the resting position can be estimated by using the horizontal vector components (F_{m-d-T9-2vf(x)-in}/F_{m-v-T9-2vf(x)-in}; Fig. 6j) of the maximum force of a muscle, the length of the anatomical inlever arm (a; Fig. 6j), i.e. the distance between the intervalvifer and the tergo-valvifer articulation, and the joint angle (α; Fig. 6j) according to the equations:

$$M_{{\text{m-d-T9-2vf}}} = F_{{{\text{m-d-T9-2vf}}\left( {\text{x}} \right){\text{-in}}}} \cdot {\text{a}} \cdot \sin(\alpha)$$

(1)

$$M_{{\text{m-v-T9-2vf}}} = F_{{{\text{m-v-T9-2vf}}\left( {\text{x}} \right){\text{-in}}}} \cdot {\text{a}} \cdot \sin(\alpha)$$

(2)

The 1st valvifer acts as a one-armed class 3 lever (force arm smaller than load arm) with the anatomical inlever (a; Fig. 6j) and the anatomical outlever (b; Fig. 6j), the latter being the distance between the intervalvifer articulation and the point at which the 1st valvifer continues as dorsal ramus of the 1st valvula (arrowhead; Fig. 6j). The resulting pro- and retracting forces at the dorsal ramus of the 1st valvula (F_{m-d-T9-2vf-out}/F_{m-v-T9-2vf-out}; Fig. 6j) can be estimated using the horizontal vector components (F_{m-d-T9-2vf(x)-in}/F_{m-v-T9-2vf(x)-in}; Fig. 6j) of the forces acting on the 1st valvifer at the tergo-valvifer articulation, the length of the effective inlever arm (a' = a · sin(α); Fig. 6j) and the effective outlever arm (b' = b · sin(β); Fig. 6j) according to the equations:

$$F_{{\text{m-d-T9-2vf-out}}} = \left( {F_{{{\text{m-d-T9-2vf}}\left( {\text{x}} \right){\text{-in}}}} \cdot {\text{a'}} } \right)/ \,\, {\text{b'}}$$

(3)

$$F_{{\text{m-v-T9-2vf-out}}} = \left( {F_{{{\text{m-v-T9-2vf}}\left( {\text{x}} \right){\text{-in}}}} \cdot {\text{a'}} } \right)/ \, \,{\text{b'}}$$

(4)

The shape of the 1st valvifer and the positions of the intervalvifer and the tergo-valvifer articulations influence the way that the 1st valvula is moved. A comparatively high quotient of the effective outlever to the effective inlever (b'/a' ratio), as observed in L. distinguendus (and other chalcidoid taxa [17, 21,22,23,24]), results in a smaller force output but an increase in the potential maximum velocity and mechanical deflection, i.e. an increase in the speed and the movement distance of the 1st valvula [31, 32, 87, 102].

Stabilization of the ovipositor: The small 1st valvifer genital membrane muscle (m-1vf-gm; Figs. 4c and 6d–f) presumably acts as a tensor muscle and stabilizes the 1st valvifers when performing the rapid pivoting movements during substrate drilling, host envenomation and oviposition (Table 1). Additionally, it might also contribute to bringing the 1st valvula into its aligned configuration [32]. The tension of both the T9-genital membrane muscle (m-T9-gm; Fig. 6d) and the posterior T9-2nd valvifer muscle (m-p-T9-2vfv; Fig. 6d) might predominantly serve the stabilization of the ovipositor during oviposition by holding the 2nd valvifers and the female T9 in position and preventing them from rotating around the articulations (Table 1). M-p-T9-2vf is also hypothesized to provide the 3rd valvulae with a certain degree of mobility [20, 22]. However, given its insertion on the median bridge of the 2nd valvifer, this is only possible if a contraction of this muscle is able to cause an elastic deformation of the median bridge, which is connected with the base of the 3rd valvula.

Support of the venom and reproductive system: The dorsal 2nd valvifer-venom gland reservoir muscle (m-d-2vf-vr; Figs. 5a–c and 6d–f) inserts dorsally at the venom gland reservoir. Its contraction presumably supports the discharge of the secretion from both the venom gland reservoir and the Dufour’s gland (Table 1; cf. [73, 99, 100, 103]). However, given its medial insertion, it might also act as a tensor muscle stabilizing the 2nd valvifer during the ovipositor movements. The two parts of the ventral 2nd valvifer-venom gland reservoir muscle (m-v-2vf-vr-a/b; Figs. 5a, c, d and 6d–f) insert laterally at the orifice the venom gland reservoir shortly before the latter enters the common oviduct. A contraction of this muscle presumably increases the diameter of the orifice, thereby controlling the venom discharge (Table 1). [35] described a muscle originating at the medial walls of the abdominal sternum 7 and inserting at the vagina; this muscle is postulated to assist in the expulsion of eggs.

Mechanisms of terebra bending and rotation

Various joint-free movement mechanisms have been described in animals (reviewed in [104]), and a variety of steering mechanisms, summarized in the following, have been proposed for the terebra of parasitoid wasps alone (cf. [62]).

The passive bending of the terebra originates from mechanical interactions of the inserted terebra with the surrounding substrate, e.g. the movements of the terebra of the fruit-fly parasitoid Diachasmimorpha longicaudata (Ashmead, 1905) (Braconidae) originate from the interplay between the surrounding substrate and relative movements of the valvulae. The relative position of the individual valvulae featuring geometrically asymmetric bevelled apices create various degrees of geometric asymmetry of the terebra apex. Consequently, the asymmetric substrate reaction forces acting on the apex push it away from a straight path [44], leading to a passive bending of the terebra, which is further facilitated by stiffness gradients in the cuticle of the apical part of the valvulae [105]. The structure and spacing of the ovipositor teeth are also thought to be involved in the passive bending movements of the terebra within plant substrates [106]. Passive bending mechanisms of the terebra are also likely to occur in species of Cynipidae (‘ovipositor searching’ sensu [107]) and Figitidae [108, 109] while they search for potential host larvae that live in plant substrates, and in species of Torymidae [110] and Agaonidae (fig wasps) [81, 106, 111] during the navigation of the terebra through the plant substrate.

The active bending of the terebra occurs when the bending moments originate from the relative movements of the valvulae, actuated by muscles inside the metasoma, e.g. (1) in species of the Aulacidae and Gasteruptiidae, abrupt terminal stops of the aulaces or protuberances in the ventrolateral side of the 2nd valvula interact with the rhachises or corresponding bosses of the 1st valvulae when the 1st valvulae are protracted and, thus, allow some dorsal bending of the terebra [42]; (2) in several species of the Braconidae, pre-apical ‘stop regions’ of the rhachis (e.g. swollen regions with scale-like sculptures located centrally within a corresponding widened part of the aulax at rest) increase friction if the 1st valvulae are retracted or extended thereby building up tension and compression and, thus, cause the terebra to curve because of the bending moment distribution [43] (cf. slide-lock working principle according to [104]); (3) in the braconid subfamily Doryctinae, a retraction of the 1st valvula causes the thinned outer walls of the aulaces to restrain the rhachis that features ancillary teeth, consequently resulting in a ventrad bending movement of the terebra [43]; (4) in the braconid genus Zaglyptogastra, the distal part of the terebra is formed into multi-arched and unevenly sclerotized regions, with the intermodal arched sections being more heavily sclerotized than the thinner nodes, and thus the protrusion of the 1st valvulae causes a flattening out of the nodal regions and the ventral flexing of the terebra [41]; (5) in several species of the Ichneumonidae, a largely longitudinally divided 2nd valvula, which is fused only at the apex, might allow the terebra to bend left or right when one part of the 2nd valvula is retracted [27]. In all these active bending mechanisms, the extent of bending movement can be controlled by adjustment of the amplitude of pro-/retraction of the individual valvulae [62]. Most of these parasitoid wasps are able to bend their terebra both dorso–ventrally and laterally, since multilateral steering can be achieved by the interplay of at least three elements [62], or by a rotational movement occurring simultaneously with the bending movement.

Passive and active bending mechanisms can technically act simultaneously or sequentially within the same structure.

During the oviposition process, female L. distinguendus were observed actively to bend their terebra in the air (i.e. in a cavity within a substrate) in which passive bending mechanisms can be excluded. The wasps were also observed to be able to pro- and retract the 1st valvula simultaneously with the bending movements (Fig. 2i–l; Additional file 1, min. 0:30–1:10, 1:36–1:44, 3:18–3:34) and independently of the bending state of the 2nd valvula and thus the whole terebra. The 1st valvulae can be protracted far forward and be retracted to a certain degree without significantly changing the bending of the whole terebra (arrowheads; Fig. 2i, k, m–o). This implies that the friction forces between the valvulae, i.e. in the olistheter system, are low. It further implies that no ‘stop regions’ or similar significant mechanical interactions occur between the 1st and 2nd valvulae in L. distinguendus. Despite rigorous searches (with scanning electron (SEM) and confocal laser scanning microscope (CLSM)), neither apical ‘stop structures’ in the olistheter nor evidence of a cluster-like occurrence of resilin in the terebra of L. distinguendus have been found (Additional file 2). Therefore, we conclude that the active bending mechanisms (1)–(4) mentioned above are not relevant for the terebra bending movements of L. distinguendus. The bending mechanisms for lateral and dorso–ventral bending and the rotation of the terebra of L. distinguendus are discussed in the following.

Lateral bending of the terebra: During the oviposition process of L. distinguendus, the terebra is anchored in the substrate. In this active probing position (Fig. 2a, c–l), a contraction of the anterior 2nd valvifer-2nd valvula muscles cannot elevate the terebra back into the resting position. In this case, however, a contraction of one of the m-a-2vf-2vv (F_m-a-2vf-2vv; Fig. 7b) presumably pulls the corresponding bulb and thus the longitudinally split and asymmetrically overlapping 2nd valvula dorsad along its longitudinal axis (arrow 10; Fig. 7b, c) because of the orientation of the muscle and the resulting direction of the force vector. Since the two halves of the 2nd valvula are fused at the apex (Figs. 3a and 4d, i, k), this movement causes the distal part of the terebra (the part inside the cavity within the substrate) to bend to the left or right (Fig. 2i–l; Additional file 1, min. 1:02–1:10, 1:36–1:44): a contraction of the left m-a-2vf-2vv causes the 2nd valvula and thus the whole terebra to bend to the left (arrow 11; Fig. 7b, c; Table 1), whereas a contraction of the right m-a-2vf-2vv causes a bend to the right. The m-a-2vf-2vv in L. distinguendus is thus adapted to its lateral bending function of the terebra and no longer serves mainly as its elevator. Furthermore, in the active probing position, a simultaneous contraction of the paired posterior 2nd valvifer 2nd valvula muscles (F_m-p-2vf-2vv; Fig. 7b) could also move the two overlapping halves of the 2nd valvula tangentially towards each other. However, the extent to which the resulting partial deformation of the 2nd valvula potentially allows local bending needs to be further investigated.

Dorso–ventral bending of the terebra: Female L. distinguendus can protract their 1st valvulae far beyond the apex of the 2nd valvula. However, these movements do not cause a dorsad bending movement of the terebra, indicating that no structures in the olistheter impede the movements of the 1st and 2nd valvulae relative to each other. However, a simultaneous retraction of both the 1st valvulae has been postulated to place the terebra under unilateral tension causing the apex to bend ventrad, and a retraction of a single 1st valvula causing the terebra to bend ventrad right or ventrad left [22, 25].

Rotation of the terebra: In the active probing position with the terebra being anchored in the substrate, a contraction of one of the posterior 2nd valvifer-2nd valvula muscles (F_m-p-2vf-2vv; Fig. 7b) presumably causes the base of the 2nd valvula and thus the whole terebra to rotate at the basal articulation along its longitudinal axis to a certain degree. Even terebra rotations of up to 90° have been observed (Fig. 2n, o; Additional file 1, min. 1:45–1:54), although such extreme rotations are in part attributable to movements of the whole metasoma. Because of the orientation of the muscle in the active probing position and the resulting direction of the force vector, a contraction of the left m-p-2vf-2vv causes the 2nd valvula and thus the terebra to rotate anti-clockwise when viewed from the dorsal side (arrow 12; Fig. 7b, c; Table 1), whereas a contraction of the right m-p-2vf-2vv causes a rotation in a clockwise direction. Contraction of the m-a-2vf-2vv might further support these rotational movements. Alternating contractions of the left and right m-a-2vf-2vv and m-p-2vf-2vv cause a reciprocal rotation of the terebra, as can be observed during substrate penetration and drilling (cf. [25, 36, 60]). A rotation occurring simultaneously with lateral bending movements of the terebra allows the bending to become effective in various directions. The morphological structure of the basal articulations is well adapted for rotational movements (cf. [20]). Since the tendon of m-p-2vf-2vv runs over the curved dorsal side of the bulbous proximal end of the 2nd valvula, the effective inlever will presumably only change little over the range of motion. However, angular changes have a large impact on the torque that can be generated (cf. [32]) and on the resulting rotation. This mechanism of terebra rotation has also been postulated for other pteromalid [22], chalcidid [36] and aphelinid species [25].

Terebra bending movements in L. distinguendus do not result from mechanical interactions between the 1st and 2nd valvulae (as postulated for some species of Aulacidae and Gasteruptiidae [42] and Braconidae [41, 43]⁠). However, the slide-lock working principle (cf. [104]) is attained in a different way. The mechanism relevant for terebra bending in L. distinguendus shows similarities with that postulated for species of ophioniform Ichneumonidae, which feature a largely split 2nd valvulae that, like the one of L. distinguendus, is fused at the apex only [27, 39]. In these ichneumonid wasps, the pteromalid L. distinguendus and possibly also other chalcidoid taxa (see subchapter ‘Eco-evolutionary significance of terebra movements’ below), the terebra bending is presumably initiated by a bending of the 2nd valvula solely. The 1st valvulae, which are connected to it via the olistheter, can thus be pro- and retracted to a certain degree without significantly changing the bending state of the 2nd valvula and thus the whole terebra. This can be advantageous, e.g. for the penetration of the host larva’s skin for precise oviposition, whereby, in a bent state of the terebra, often alternating pro- and retraction movements of the 1st valvulae are required.

Whenever the 2nd valvula is bent in the lateral plane, one side (and thus also the lateral side of the corresponding 1st valvula) is under compression, with the opposite side being under tension. Both the bending and torsional stiffness of the terebra depend on its geometry, i.e. its cross-sectional shape (cf. [62]), and its material composition, i.e. chitin embedded in a protein matrix of variable mechanical properties (depending on its contents of resilin, arthropodin and sclerotin). The material stiffness of insect cuticle, expressed as Young’s modulus, has previously been estimated to range between 0.5–20 GPa [60, 81, 112].

Eco-evolutionary significance of terebra movements

The structure of the terebra of Chalcidoidea, featuring a longitudinally split 2nd valvula with overlapping, asymmetric halves, is strongly consistent in structure within families and basically similar across families (with the exception of the primitive Mymaridae [18, 39], which recently have been identified as a sister group to all remaining Chalcidoidea [10, 49]), but is unique among other superfamilies of parasitoid Hymenoptera [39]. The similar structure of the terebra of chalcidoid taxa might indicate similar underlying working mechanisms, since form and function are strongly connected [113, 114]. Other chalcidoids are therefore also likely to be able to steer their terebra in a similar manner to that of L. distinguendus, as such terebra steering movements have also been observed in other species of Pteromalidae during the assessment of a potential host and egg placement [40, 46], in Eurytomidae during egg placement [21], in Eupelmidae during the assessment of a potential host [115] and the ovicide or larvacide of the competitors’ eggs and larvae, respectively [116], in species of the Aphelinidae during the ovicide of the competitors’ eggs [117,118,119] and in species of Torymidae [110] and Agaonidae for accurate egg deposition in the plant substrate [81, 106, 111] (although the latter two groups probably use passive bending mechanisms, unlike L. distinguendus).

Oviposition is crucial for the reproductive success of insects; thus, oviposition behaviour and ovipositor structure have a central adaptive role [83, 90, 93, 106, 120, 121] that should directly affect fitness. The improved manoeuvrability of the metasoma of the Apocrita, which is essential in the female wasp’s probing behaviour when searching or assessing a potential host, is attributed to the evolution of the wasp waist (a construction between the 1st and 2nd abdominal segment). The presence of a waist was a major innovation in the evolution of Hymenoptera and presumably contributed to the rapid diversification of Apocrita, since it allowed the successful attack of a variety of new hosts [3,4,5]. However, some chalcidoid wasps, e.g. species of Trichogrammatidae [122], have secondarily lost their wasp waist, presumably during miniaturization. Moreover, the vast majority of Chalcidoidea, although targeting the largest diversity of host taxa among parasitoid wasps [12], are idiobiont ectoparasitoids that develop on enclosed host stages with reduced mobility. Depositing eggs within a substrate provides them and the hatched larvae with the protection of a concealed environment (but without being exposed to the host’s immune system, as are endoparasitoids). Thus, in most chalcidoid wasps, as in L. distinguendus, a manoeuvrable metasoma does not improve the ability to reach hosts hiding in concealed cavities in hard substrates, since the position of the terebra is anchored at the puncture site. Moreover, drilling is extremely energy- and time-consuming (drilling a hole through a seed grain accounts for approximately 15% of the daily energy budget in a female eupelmid Eupelmus vuilleti (Crawford, 1913) [123]) and risky, as the wasps are exposed to predators. In L. distinguendus and presumably most chalcidoids that parasitize hosts hidden in hard substrates, the ability actively to bend and rotate the terebra in various directions is crucial during the search for a potential host within the substrate (or the cavity within), a targeted venom injection (e.g. directly into the ganglia [124, 125] or fat bodies of large hosts [126]), the assessment of the potential host, the ovicide and larvicide of the competitors’ eggs and larvae, respectively, the search for a suitable place for oviposition and controlled egg placement (Fig. 7d).

In Chalcidoidea, multiple morpho-physiological and behavioural traits have evolved in correlation with the use of hosts concealed deep within hard substrates, apparently related to several functional demands including host localization, substrate penetration, oviposition and emergence from the substrate (cf. [14]). Modifications or specializations of these traits, such as the ability actively to steer the terebra, may have interacted synergistically to open up new evolutionary pathways (cf. [5]). Adaptations in oviposition behaviour combined with morphological modifications of the terebra and adaptations in the function of certain muscles (i.e. the anterior and posterior 2nd valvifer-2nd valvula muscles) have potentially facilitated the evolution of terebra steering mechanisms, which in turn have facilitated the acquisition of new hosts to a parasitoid’s host range. These shifts in host exploitation allow niche partitioning among co-occurring species (cf. [127]) and presumably have led to rapid adaptive radiations in Chalcidoidea [10] (for speciation process in L. distinguendus see [128,129,130]). Thus, the ability actively to steer the terebra potentially has been a central factor in the evolution of the parasitoid life history strategy and the diversification of chalcidoid wasps, resulting in the evolutionary success of this group with its tremendous extant species richness [10, 49]. The Chalcidoidea are the most diverse group of parasitoid hymenopterans, with estimations of more than 500,000 chalcidoid species, the vast majority of them being parasitoids, out of a total of 680,000 parasitoid hymenopteran species [6, 7]. Even larger species numbers might exist because of the possibility of a vast underlying biodiversity of cryptic species (cf. [7, 8, 11]).

Conclusions

Adaptations in oviposition behaviour combined with morphological modifications of the terebra and adaptations in the function of certain muscles allow L. distinguendus and presumably also other chalcidoid wasps to steer their terebra in various directions, a crucial skill for the successful oviposition of hosts that are concealed in substrates. Therefore, the evolution of the ability actively to steer the terebra can be considered as a putative key innovation that has largely contributed to the acquisition of new hosts to a parasitoid’s host range. Here, we have identified the structural adaptations, i.e. the longitudinally split and asymmetrically overlapping halves of the 2nd valvula that are fused at the apex and the functional adaptions of its associated muscles, and the mechanisms behind these innovations. Further comparative studies are needed to reveal the way in which morpho-physiological, behavioural, ecological and life history traits have interacted during the evolution and resulted in the enormous radiation of Chalcidoidea.

The terebra of hymenopterans in general and chalcidoids in particular can also act as a suitable biological concept generator, with further investigations into this matter possibly being helpful in the development and the design of slender miniaturized man-made probing tools (cf. [131,132,133,134]) for curved steering and drilling.

Methods

The L. distinguendus used in this study originate from the laboratory colonies of FuturA GmbH (Borchen, Germany) and Biologische Beratung GmbH (Berlin, Germany), where they were bred on the larvae of Sitophilus oryzae (Linnaeus, 1763) (Coleoptera: Curculionidae) that develop endophytically in grain of the common wheat Triticum aestivum L.. This host indicates that the L. distinguendus used in this study probably belong to the karyotype with 5 chromosomes (‘Sitophilus Clade 1’ sensu [129] or ‘GW-lineage’ sensu [130], respectively) of the L. distinguendus species complex that comprises at least two morphologically indistinguishable cryptic species [135]. The clade/lineage presumably evolved by a host shift from drugstore beetles (Stegobium paniceum (Linnaeus, 1758)) to weevils of the genus Sitophilus [129]. This shift was probably related to the ability to learn from host-related cues [128].

For lateral overview images, female wasps were imaged with a digital microscope of the type Keyence Digital Microscope VHX-7000 (Keyence Corporation, Osaka, Japan) by using focus stacking.

Images were processed (white/black balancing, cropping) with GIMP version 2.10.30 (https://www.gimp.org; RRID:SCR_003182). The schematic drawings were created with Inkscape version 1.1 (https://www.inkscape.org; RRID:SCR_014479).

High-resolution videography

The oviposition process of L. distinguendus was recorded in an artificial two-part Plexiglas chamber. Each lower chamber element featured a notch (ca. 4 · 1 · 1 mm) at the upper side of the front. Each upper element also featured a notch (ca. 4 · 2 · 4 mm) positioned at the front and lying exactly on that of the lower element. A piece of blotting paper was clamped in between the two elements to divide the space created by the two notches into two compartments. This paper was placed in the breeding substrate of S. oryzae for several days before the recording trials for it to take on the hosts’ scent (faecal cues) and thus to trigger the wasps to attempt to oviposit. The two chamber elements were then fixed with screws. A female L. distinguendus was placed into the upper compartment and a S. oryzae larva in the lower. The front and upper openings of the chamber were subsequently each closed with a clean glass coverslip. The process of oviposition was filmed in a horizonal position by using a Nikon DSC D90 camera (Nikon Corporation, Tokyo, Japan) mounted on a Leica MZ 12.5 stereomicroscope (Leica Microystems GmbH, Wetzlar, Germany) and with two LED cold-light sources KL 300 LED (Schott AG, Jena, Germany) for sufficient illumination. The focus was adjusted manually.

Scanning electron microscopy (SEM)

For scanning electron microscopy (SEM), we dissected the ovipositor from the genital chamber of ethanol-fixed animals by using fine forceps. Specimens were dehydrated in a graded ethanol (C₂H₆O) series (30, 50, 70, 80, 90, 95 and twice 100% for 30 min each concentration) and air-dried for at least one week in a desiccator with silica gel (Carl Roth GmbH & Co. KG, Karlsruhe, Germany). We mounted the samples with double-sided adhesive carbon tape onto stubs and sputter coated them with 19 nm pure gold (Au) using an Emitech K550X (Quorum Technologies Ltd, West Sussex, UK). Investigation and imaging were performed with a scanning electron microscope of the type Zeiss EVO LS 10 (Carl Zeiss Microscopy GmbH, Jena, Germany) and the software SmartSEM version V05.04.05.00 (Carl Zeiss Microscopy GmbH, Jena, Germany).

Confocal laser scanning microscopy (CLSM) and wide-field epifluorescence microscopy (WFM)

For confocal laser scanning microscopy (CLSM), specimens preserved in 70% ethanol were transferred to glycerol, dissected and further stored in a glycerol (ChemWorld, Kennesaw, GA, USA) droplet on concave microscope slides. Specimens were imaged between two #1.5 coverslips with a confocal laser scanning microscope of the type Nikon A1R-HD (Nikon Corporation, Tokyo, Japan). We used three excitation wavelengths, namely 409, 487 and 560 nm, and three emission ranges, namely 435–470, 500–540 and 570–645 nm. The resulting image sets were assigned pseudo-colours that reflected the fluorescence spectra (blue, green and red, respectively). Volume-rendered micrographs and media files were created using Fiji [136] (https://imagej.net/Fiji; RRID:SCR_002285).

For wide-field epifluorescence microscopy (WFM), we dissected the ovipositor from freshly killed individuals and washed them in distilled water. We mounted them carefully onto cleaned microscope slides (76 mm · 26 mm, VWR International, Radnor, PA, USA), embedded them in glycerol (Croma-Pharma GmbH, Loebendorf, Austria) without staining for observation with an epifluorescence microscope of the type Zeiss Axio Imager M2 (Carl Zeiss Microscopy GmbH, Jena, Germany) equipped with an ORCA-Flash4.0 V2 Digital CMOS camera C11440-22CU (Hamamatsu Photonics K.K., Hamamatsu, Japan) and the software ZEN 2 pro (blue edition) (Carl Zeiss Microscopy GmbH, Jena, Germany). We used Plan-Apochromat objectives and the following wavelength filters: DAPI (blue, excitation 353 nm, emission 465 nm), ATTO488 (green, excitation 500 nm, emission 525 nm) and Cy5 (red, excitation 650 nm, emission 673 nm).

We superimposed both the CLSM and WFM images in order to show the autofluorescence of the cuticular structures in order to analyse their material composition. Cuticular structures that predominantly show blue autofluorescence are composed of high proportions of the soft and highly elastic rubber-like amorphous protein resilin [98], which has an autofluorescence at a narrow band around 415 nm wavelength [95], whereas cuticular structures that autofluoresce in green are chitinuous and non- or weakly sclerotized and those that exhibit red autofluorescence are heavily sclerotized [98, 137].

Sample preparation, light microscopy (LM), transmission electron microscopy (TEM) and image processing

Each female L. distinguendus was anaesthetized with carbon dioxide (CO₂) before its metasoma was removed and submersed in fixative solution containing 2.5% glutaraldehyde (C₅H₈O₂) and 5% sucrose (C₁₂H₂₂O₁₁) buffered with 0.1 M cacodylate (C₂H₇AsO₂) buffer (pH 7.4). During this process, the samples were stored in the fixative in small glass vials held in an ice bath at approximately 4 °C for 12 h, following which they were rinsed three times in pre-chilled 0.1 M cacodylate buffer (pH 7.4) for 10 min. After a 4 h period of post-fixation in 1% osmium tetroxide (OsO₄) solution buffered with 0.1 M cacodylate buffer (pH 7.4) in an ice bath, the samples were again rinsed three times in the same buffer. The subsequent steps were performed at room temperature. The samples were dehydrated through a graded ethanol (C₂H₆O) series (30, 50, three times 70, 75, 80, 85, 90, 95 and 100% for three times, 10 min each concentration), containing en-bloc staining by using a saturated solution of 70% ethanolic uranyl acetate (C₄H₆O₆U) for 12 h. The fully dehydrated samples were then passed through two changes of 100% propylene oxide (C₃H₆O) for 1 h per change and then through increasing concentrations of Spurr low-viscosity embedding resin (Polysciences Inc., Warrington, PA, USA) in propylene oxide with C₃H₆O:Spurr ratios of 3:1, 1:1, 1:3 and 1:7 for 1 h per change and 100% Spurr for 17 h on a rotatory shaker. As a last incubation step, the samples were placed in fresh pure resin for embedment in silicon moulds and polymerized at 70°C for 8 h.

Semithin (600 nm) and ultrathin (60 nm) sections were cut perpendicularly to the terebra of L. distinguendus by using an ultramicrotome of the type Leica Ultracut UTC (Leica Microsystems GmbH, Wetzlar, Germany) equipped with a DiATOME histo Jumbo diamond knife (45° knife angle; DiATOME Ltd, Nidau, Switzerland) with a large boat for continuous serial semithin sectioning or a DiATOME ultra diamond knife (35° knife angle; DiATOME Ltd, Nidau, Switzerland) for ultrathin sectioning. We conducted two complete section series through the whole metasoma; one continuous series of semithin sections and one with consecutive alternating series of 20 semithin and 10 ultrathin sections. Microscope slides (76 mm · 26 mm, VWR International, Radnor, PA, USA) for the mounting of semithin serial sections were preliminary stored in a bath containing 96% ethanol and 25% ammonia (NH₃) at a C₂H₆O:NH₃ ratio of 9:1 for at least one week and finally cleaned and stored in distilled water shortly before use. Semithin serial section bands were directly mounted onto these slides and stained with toluidine blue (C₁₅H₁₆ClN₃S) for 60 s on a hot plate at 80°C. After being rinsed with distilled water and dried, the stained sections were embedded in Euparal (Waldeck GmbH & Co. KG, Münster, Germany). Ultrathin sections were placed on Formvar-coated copper slot grids and post-stained with 2% ethanolic uranyl acetate and lead citrate according to Venable and Coggeshall [138] for 20 and 10 min, respectively.

To image the semithin sections, we used a light microscope of the type Zeiss Axioplan (Carl Zeiss Microscopy GmbH, Jena, Germany) equipped with a Nikon D7100 single-lens reflex digital camera (Nikon Corporation, Tokyo, Japan) and the software Helicon Remote version 3.6.2.w (Helicon Soft Ltd, Kharkiv, Ukraine) (for focus stacking Helicon Focus version 6.3.7 Pro; RRID:SCR_014462). After initial image processing (white balancing, colour contrasting, black and white converting, cropping) in Adobe Lightroom version 6.0 (Adobe Systems, San José, CA, USA), the image stack was calibrated with Fiji [136] (https://imagej.net/Fiji; RRID:SCR_002285), a distribution of the software ImageJ2 version 2.3.0/1.53f [139, 140] (https://imagej.net; RRID:SCR_003070), and imported to the plugin TrakEM2 [141] (RRID:SCR_008954). A preliminary least square rigid alignment followed by an elastic alignment of the image stack was performed using the ‘Elastic Stack alignment’ plugin [142] in order to create an aligned image stack.

To investigate and image the ultrathin sections, we used a transmission electron microscope of the type Philips/FEI Tecnai 10 (FEI Company, Hillsboro, OR, USA) operated at 80 kV equipped with a side-mounted Gatan Rio9 CMOS camera (Gatan Inc., Pleasanton, CA, USA) and the software Tecnai G² User Interface version 2.1.5 (FEI Company, Hillsboro, OR, USA) and DigitalMicrograph version 3.32.2403.0 (Gatan Inc., Pleasanton, CA, USA), respectively.

Synchrotron X-ray phase-contrast microtomography (SR-µCT) and image processing

Two metasomas of ethanol-fixed female L. distinguendus were dehydrated stepwise in ethanol and critical-point-dried by using a Polaron 3100 (Quorum Technologies Ltd, West Sussex, UK) to avoid shrinking artefacts by water loss during the tomography procedure. The anterior ends of the metasomas were glued onto plastic pins and mounted onto the goniometer head of the sample stage for tomography. Synchrotron X-ray phase-contrast microtomography (SR-µCT) [143] was performed at the beamline ID19 at the European Synchrotron Radiation Facility (ESRF; Grenoble, France) at 19.5 keV (wavelength 8 · 10⁻¹¹ m). 6000 projections were recorded over a 180° rotation with an effective detector pixel size of 0.3 µm, and a corresponding field of view of 0.63 · 0.63 mm. The detector consisted of a 4.5 μm thick LSO:Tb (Tb-doped Lu₂SiO₅) single-crystal scintillator lens (magnification 20×, numerical aperture (NA) 0.4) coupled to a sCMOS-based camera type pco.edge 5.5 (Excelitas PCO GmbH, Kelheim, Germany) [144, 145]. The detector-to-sample distance was set to 10 mm. Two separate overlapping image stacks were acquired since the structures of interest were larger than the field of view. The sample was therefore repositioned in between the imaging procedure, resulting in a certain overlap of the two consecutive images. The 3D voxels datasets were reconstructed from 2D radiographs by using the filtered back-projection algorithm [146, 147] developed for parallel-beam tomography.

The two resulting tomograms were registered and calibrated with Fiji [136] (https://imagej.net/Fiji; RRID:SCR_002285) and further imported to the plugin TrakEM2 [141] (RRID:SCR_008954) for stitching and cropping. Export of the aligned image stack was performed using a custom script, allowing the export of 16bit image stacks from TrackEM. Subsequently, the resulting image stack was imported to Amira version 6.0 (FEI Company, Hillsboro, OR, USA; RRID:SCR_014305) to pre-segment the various elements of the ovipositor and the whole metasoma in the software’s segmentation editor by manually labelling every 25th virtual slice and assigning them to different ‘materials’. These labels served as the input for automated segmentation by using the Biomedical Image Segmentation App ‘Biomedisa’ [148] (https://biomedisa.org). After some minor manual corrections to the segmentation results of the ‘Biomedisa’ output by using Amira, we converted them into polygon meshes. We thereby applied some minor smoothing (unconstrained smoothing, smoothing extent: 2) and polygon reduction to create the final 3D model (surface mesh).

Availability of data and materials

All data supporting the conclusions of this article are included within the article and its additional files. The full resolution videos (Additional file 1, Additional file 2, Additional file 3, and Additional file 4) and the analysed raw datasets are available from the corresponding author on reasonable request.

Abbreviations

1vf:: 1st valvifer
1vv:: 1st valvula
2vf:: 2nd valvifer
2vv:: 2nd valvula
3vv:: 3rd valvula
au:: Aulax
ba:: Basal articulation
blb:: Bulb
CLSM:: Confocal laser scanning microscopy
co:: Common oviduct
cs:: Coeloconic sensillum
ct:: Ctenidium
den:: Dendrite
Dg:: Dufour’s gland
Dgd:: Dufour’s gland duct
dp2:: Dorsal projection of the 2nd valvifer
dr1:: Dorsal ramus of the 1st valvula
ec:: Egg canal
F :: Force
F _(x) :: Horizontal vector component of a force
fu:: Furcula
iar:: Interarticular ridge of the 1st valvifer
il1:: Interlock of the 1st valvulae
iva:: Intervalvifer articulation
lb:: Laminated bridge
le:: Lateral extensions of the 2nd valvula
LM:: Light microscopy
lu2:: Lumen of the 2nd valvula
M :: Torque
m-1vf-gm:: 1st valvifer-genital membrane muscle
m-a-2vf-2vv:: Anterior 2nd valvifer-2nd valvula muscle
m-d-2vf-vr:: Dorsal 2nd valvifer-venom gland reservoir muscle
m-d-T9-2vf-a:: Dorsal T9-2nd valvifer muscle part a
m-d-T9-2vf-b:: Dorsal T9-2nd valvifer muscle part b
m-p-2vf-2vv:: Posterior 2nd valvifer-2nd valvula muscle
m-p-T9-2vf:: Posterior T9-2nd valvifer muscle
m-T9-gm:: T9-genital membrane muscle
m-v-2vf-vr:: Ventral 2nd valvifer-venom gland reservoir muscle
m-v-2vf-vr-a:: Ventral 2nd valvifer-venom gland reservoir muscle part a
m-v-2vf-vr-b:: Ventral 2nd valvifer-venom gland reservoir muscle part b
m-v-T9-2vf:: Ventral T9-2nd valvifer muscle
mb2:: Median bridge of the 2nd valvifers
me:: Metasoma
mr2:: Medial ridge of the 2nd valvifer
oth:: Olistheter
ovr:: Orifice of the venom gland reservoir
prf:: Post-ramus flap
rh:: Rhachis
sc:: Scale-like structure
SEM:: Scanning electron microscopy
sp:: Sensillar patch of the 2nd valvifer
sr:: Sensillar row of the 2nd valvifer
SR-µCT:: Synchrotron X-ray phase-contrast microtomography
st1:: Sawtooth of the 1st valvula
st2:: Sawtooth of the 2nd valvula
t-m-d-T9-2vf-a:: Tendon of the dorsal 2nd valvifer-T9 muscle part a
T9:: Female T9 (9th abdominal tergum)
TEM:: Transmission electron microscope
trb:: Terebra
tva:: Tergo-valvifer articulation
vr:: Venom gland reservoir of the 2nd valvifer
vw2:: Ventral wall of the 2nd valvula
WFM:: Widefield epifluorescence microscopy

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Acknowledgements

The authors thank the following colleagues for their help: Matthias Schöller (Biologische Beratung GmbH) for providing wasps and their hosts; Jana Collatz, Johannes LM Steidle and Urs Wyss for providing inspiring film sequences (Entofilm) and valuable tips; Paavo Bergmann for assistance with histological techniques, Julia Straube with ultramicrotomy, Monika Meinert with SEM, Lorenz Henneberg and York-Dieter Stierhof with WFM, Verena Pietzsch and James H. Nebelsick with the Keyence digital microscope; Robin Kraft, Benjamin Sampalla, Lea von Berg, Erich Lara Spiessberger, Margarita Yavorskaya and Manfred Drack for inspiring discussions; Theresa Jones for improvements and linguistic corrections of the manuscript, and three referees for their helpful comments. The authors gratefully acknowledge the Tübingen Structural Microscopy Core Facility (funded by the Excellence Strategy of the German Federal and State Governments) and the University of New Hampshire Instrumentation Center for their support and assistance in this work.

Funding

This work was funded by the German Research Foundation (DFG) as part of the Transregional Collaborative Research Centre (SFB-TRR) 141 ‘Biological Design and Integrative Structures’ (project A03 ‘Inspired by plants and animals: actively actuated rod-shaped structures exhibiting adaptive stiffness and joint-free kinematics’). The experiment (LS-2342) at the ESRF was funded by the European Union (EU). The authors acknowledge support from the Open Access Publishing Fund of the University of Tübingen. Open Access funding enabled and organized by Projekt DEAL.

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Authors and Affiliations

Evolutionary Biology of Invertebrates, Institute of Evolution and Ecology, University of Tübingen, Auf der Morgenstelle 28, 72076, Tübingen, Germany
Benjamin Eggs, Stefan Fischer, Michael Csader & Oliver Betz
Tübingen Structural Microscopy Core Facility (TSM), University of Tübingen, Schnarrenbergstraße 94–96, 72076, Tübingen, Germany
Stefan Fischer
State Museum of Natural History Karlsruhe, Erbprinzenstraße 13, 76133, Karlsruhe, Germany
Michael Csader
Department of Biological Sciences, University of New Hampshire Collection of Insects and Other Arthropods, University of New Hampshire, Spaulding Hall, Durham, NH, 03824, USA
István Mikó
ESRF – The European Synchrotron, Structure of Materials Group – ID19, CS 40220, 38043, Grenoble Cedex 9, France
Alexander Rack

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Benjamin Eggs
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Stefan Fischer
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Michael Csader
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István Mikó
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Alexander Rack
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Oliver Betz
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Contributions

BE and OB prepared the study design; AR and OB obtained the SR-µCT data; AR deployed and adjusted the beamline settings for SR-µCT; BE and SF prepared the samples (fixation, embedding) and processed the SR-µCT and light microscopical images; BE carried out the ultramicrotomy, LM, WFM, TEM and high-resolution videography, analysed the data and wrote the manuscript; BE and MC carried out the SEM; IM carried out the CLSM; SF, MC, IM, AR and OB discussed the results and revised the manuscript. All authors read and approved the final manuscript.

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Correspondence to Benjamin Eggs.

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No approval of research ethics committees was required to achieve the goals of this study, as it involved experimental work with an unregulated invertebrate species.

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Supplementary Information

Additional file 1. Video sequences of female Lariophagus distinguendus parasitizing larvae of Sitophilus granarius in an artificial film chamber (cf. Fig. 2b–p). The terebra bending and rotating movements during host assessment and the alternate movements of the paired 1st valvulae can be observed.

Additional file 2. Superimposed CLSM images of the ovipositor of Lariophagus distinguendus (dorsal view, cf. Fig. 3g, h).

Additional file 3. Animation of the aligned semithin sections through the terebra of Lariophagus distinguendus (from distal to proximal; cf. Fig. 4a–d). The jittering movements of the two halves of the 2nd valvula in the middle of the image stack result from section and compression artefacts.

Additional file 4. Animation of the rotating segmented 3D model of the musculoskeletal ovipositor system of Lariophagus distinguendus (cf. Figs. 5 and 6).

Appendix 1

See Table 2.

Table 2 Morphological terms relevant to the hymenopteran ovipositor system

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Eggs, B., Fischer, S., Csader, M. et al. Terebra steering in chalcidoid wasps. Front Zool 20, 26 (2023). https://doi.org/10.1186/s12983-023-00503-1

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Received: 05 May 2023
Accepted: 13 July 2023
Published: 08 August 2023
DOI: https://doi.org/10.1186/s12983-023-00503-1

Terebra steering in chalcidoid wasps

Abstract

Background

Results and discussion

Oviposition process and employment of the terebra

Morphological structure of the musculoskeletal ovipositor system

Cuticular elements of the ovipositor

Articulations of the musculoskeletal ovipositor system

Ovipositor muscles

Mechanics and mode of function of the ovipositor system

Mechanisms of terebra bending and rotation

Eco-evolutionary significance of terebra movements

Conclusions

Methods

High-resolution videography

Scanning electron microscopy (SEM)

Confocal laser scanning microscopy (CLSM) and wide-field epifluorescence microscopy (WFM)

Sample preparation, light microscopy (LM), transmission electron microscopy (TEM) and image processing

Synchrotron X-ray phase-contrast microtomography (SR-µCT) and image processing

Availability of data and materials

Abbreviations

References

Acknowledgements

Funding

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Authors and Affiliations

Contributions

Corresponding author

Ethics declarations

Ethics approval and consent to participate

Consent for publication

Competing interests

Additional information

Publisher's Note

Supplementary Information

Appendix 1

Appendix 1

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Keywords

Frontiers in Zoology

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