Vibrations through substrates are an important source of information for diverse organisms, from nematodes to elephants. The fundamental challenge for small animals using vibrational communication is to move their limited mass fast enough to provide sufficient kinetic energy for effective information transfer through the substrate whilst optimising energy efficiency over repeated cycles. Here, we describe a vibratory organ found across a commercially important group of plant-feeding insects, the planthoppers (Hemiptera: Fulgoromorpha). This elastic recoil snapping organ generates substrate-borne broadband vibrations using fast, cyclical abdominal motion that transfers kinetic energy to the substrate through the legs. Elastic potential energy is stored and released twice using two different latched energy-storage mechanisms, each utilising a different form of elastic recoil to increase the speed of motion. Comparison to the acoustic tymbal organ of cicadas (Hemiptera: Cicadomorpha) reveals functional convergence in their use of elastic mechanisms to increase the efficacy of mechanical communication.
We begin by characterising the morphology of the newly-described snapping organ in our model species, Agalmatium bilobum (Fulgoromorpha: Issidae). The snapping organ can be found dorsally on each side of the body at the junction between the metathorax and the abdomen, spanning the first two abdominal segments (Fig 1A and 1B and S1 Movie). The organ has a W shape; a ridge (Fig 1B) articulates at its base with the thorax (first ‘V’) and fuses at its tip to the anterior arm of a Y-lobe (Fig 1B), which has resilin (Fig 1B) between its arms (second ‘V’; Figs 1B and S1). The posterior arm of the Y-lobe is fused with the second segment, tergum 2 (tg2; Fig 1B) of the abdomen. The Y-lobe is linked at its base to an internal spine (sp; Fig 2C) of the second segment via a membranous connector (Figs 1B, 2A and 2C). Eight muscle pairs are directly associated with the snapping organ (Fig 2 and S2 Table and S1 Text), comprising three pairs of dorsal longitudinal muscles (DLMs) and five pairs of dorsoventral muscles (DVMs). Four other muscle pairs are indirectly associated with the snapping organ (ventral longitudinal muscles [vlms] IIIvlm2, Ivlm1, and IIvlm2 and intersegmental dorsoventral muscle [IIisdvm]) (Fig 2A). The snapping organ is not sexually dimorphic. Homologous vibrational organs are present throughout the entire planthopper clade (Fig 3 and S1 Table). The defining features of the musculature (Fig 2 and S2 and S3 Tables), innervation (S2 Fig), and external morphology (the ridge, Y-lobe, and connector) of the snapping organ are consistent and identifiable, despite variation in its proportions and shape across the planthoppers (Fig 3B–3E and S1 Text). Two groups deviate from this general picture: part of the family Delphacidae, in which the exoskeleton and musculature have been drastically reorganized to form an entirely different type of vibrational organ (S3 Fig and S1 Text), and part of Derbidae, which have an externally obscure snapping organ and also possess tentative stridulatory structures (Fig 3 and S4 Table). Based on their phylogenetic position, the deviations observed in these two groups are likely to be derived (Fig 3). To determine the kinematics of the snapping organ, we used high-speed videography and laser vibrometry on our model species, A. bilobum (Fig 4 and S1 Movie and S1 Data). Each vibrational cycle began with the snapping organ in its relaxed position (Figs 1D and 4A). Subsequently, the thorax/midabdomen was raised over a 15 ms timescale (Fig 4B). The first mechanical impulse followed (loading vibrational peak), which resulted in closed Y-lobe arms, extended ridge, and the base of the Y-lobe pulled down and rotated clockwise (Fig 4C). The system resonated in response, giving a jagged waveform over a 15–20 ms timescale (Fig 4D). The cycle was completed by a second mechanical impulse (unloading vibrational peak), in which the Y-lobe arms reopened, the ridge retracted, and the base of the Y-lobe rose and rotated back (Fig 4E). This resulted in whole-system resonance ultimately returning the organ to the same relaxed position as at the beginning of the cycle. Each vibration generation cycle takes place within 120 ms, and the mechanism does not generate any audible acoustic noise [27]. We propose that each cycle of vibration generation consists of four main steps (Figs 1D and 4). Transition from the relaxed state to the cocked state was comparatively slow (on a timescale of 15 ms), and the movements of landmarks on the external exoskeleton suggest that this phase of the cycle was driven directly by DLM contraction (Figs 2 and 4B). Whilst we do not have direct recordings of muscle activity, the distance between the origin and insertion points of both DLMs shortens at this point in the cycle (Fig 4B), and there is no other muscle whose action could produce this strain. The distance between these points shortens even further at the transition from the cocked state to the loaded state (Fig 4C), but this change occurs too quickly to be explained by direct muscle action alone. Specifically, the rate of change in the kinetic energy of the abdomen during loading implies energy release at a much higher power density than the DLMs and DVMs combined (Idlm1, Idlm2, IIedvm1, IIedvm2) could possibly supply through contraction (7,080 W kg−1, which is nearly 15 times the typical 500 W kg−1 power density for a muscle [31]; see S1 Methods and S1 Data). It follows that some form of elastic recoil, which acts as a kind of mechanical power amplifier, must be involved in the transition between the cocked and loaded states. This fast phase (0 to peak velocity taking 0. 35 ms), which we term loading, is responsible for producing the first mechanical impulse transferring vibrational energy to the substrate. The distance between the origin and insertion points of the DVMs also shortens at this point in the cycle (Fig 4C), but contraction of these muscles alone cannot supply the mechanical energy at a high enough rate to explain the rapidity of the loading phase. Instead, the events at this transition are consistent with DVM contraction serving as an unlatching mechanism that triggers the rapid pulling down of the abdomen, followed by system resonance (Fig 4C). The next phase of the cycle, in which the system transitioned to its pretrigger state, was a slow phase, probably involving muscle relaxation, over a 15–20 ms timescale. The subtle shift of exoskeleton positions, and particularly the lengthening of the distance between the points of origin and insertion of the DLMs (Fig 4D), is consistent with the DLMs relaxing during this phase. In contrast, the distance between the points of origin and insertion of the DVMs remain constant through this phase of the cycle, suggesting that they remain in their contracted state. The final transition in the cycle was from the pretrigger state to the relaxed state. This second fast phase, which we term unloading, is responsible for producing the second mechanical impulse transferring vibrational energy to the substrate. The associated increase in distance between the points of origin and insertion of the DVMs (Fig 4E) suggests that unloading is triggered by DVM relaxation, which causes the rapid return of the snapping organ to its relaxed conformation through a second release of stored elastic potential energy. There is no evidence for muscle contraction at this phase of the cycle, and we therefore infer that this elastic potential energy is likely to be stored in the deformed exoskeletal elements of the snapping organ. To verify whether passive release of elastic potential energy could be responsible for the fast unloading phase, we built a simplified mathematical model of the snapping organ, in which we replaced the ridge and the anterior arm of the Y-lobe with a pair of rigid bars connected in series to the thorax by a pair of torsional springs (Figs 5A and S4). The stiffness constants of these torsional springs were determined experimentally in a static loading experiment (S1 Methods). The abdomen and posterior arm of the Y-lobe were modelled as a mass-spring–damper system attached to the free end of the second rigid bar (Figs 5A and S4), and the spring constants and damping coefficients of this system were fitted as free parameters (S1 Methods). Quantitative comparison of the measured and modelled motion supports our supposition that the unloading phase can be explained through passive recoil of the Y-lobe, in which mechanical energy is stored elastically (Fig 5B and S1 Methods). When released, the elastic potential energy of these stiff springs acts to move the mass of the abdomen back to its relaxed state, resulting in resonant motion of the abdominal mass. More harmonic content is apparent in the measured vibrations than the modelled ones, which is not surprising given the simplicity of the model, but importantly from the perspective of information transfer, both the measured and the modelled spectra involve a broad range of different frequencies (Fig 5C and 5D). The motion generated by the snapping organ during the two fast loading and unloading phases was on a timescale that would not have been possible through direct muscle action alone. The snapping organ instead uses two distinct elastic recoil mechanisms, each of which involves storing energy in springs, then releasing the stored energy quickly [8–11]. During the loading phase, the obvious candidate locations for elastic energy storage are the DLMs themselves, given that the exoskeleton itself deforms very little during loading (Fig 4B). This would mean that these muscles act both as engines, actively generating the force required for loading, and as springs, storing elastic energy within their deformed structure when subject to resistance against shortening from the exoskeleton. Muscles have previously been suggested to act as springs [10], and here the elastic energy storage is in the range achievable by the cross-bridges (energy density for paired Idlm1 and Idlm2 conservatively c. 2. 47 J kg−1) [32]. We therefore suggest that resistance to shortening of the contracted DLMs allows these muscles to act as an elastic spring during the loading phase [31], storing energy slowly, then releasing this quickly when triggered. A latch must be involved to prevent early release of energy, and a mechanical constraint at the base of the Y-lobe could act as a latch that is removed when the DVMs contract, acting to trigger the release of elastic potential energy stored in the DLMs. During the unloading phase, a more straightforward passive elastic recoil is the likely mechanism, as captured by our mathematical model (Fig 5). Energy is stored in stiff springs within the W-shaped exoskeleton linkage system that are deformed and therefore loaded during the loading phase (Fig 4C), but which return to their resting position and are therefore unloaded following the unloading phase (Fig 4E). The first elastic recoil event during the active loading phase thereby stores the energy that is released during the second elastic recoil event, which is the passive unloading phase. DVM relaxation is the likely trigger, with the membranous connector and acting as a possible cuticular latch preventing early release (Fig 2). Rapid recoil is made possible by DLM relaxation during the pretrigger step, and resilin between the Y-lobe arms (S1 Fig) will act to limit damage during recoil. Additional muscles may modulate the vibration during unloading (e. g. , IIIvlm2), but the muscles are far too small to account for the power density during unloading (c. 765,000 W kg−1 if normalising the mechanical power by IIIvlm2 mass; Fig 2A and S1 Data). In summary, the snapping organ uses two muscle contraction events per cycle and typically repeats its cycle every 0. 3–1 s [33], giving a muscle contraction frequency of under 5 Hz (S5A Fig). In contrast, the frequencies of the mechanical impulses resulting from this motion as measured on the midabdomen were broadband under 3 kHz (shown for recoil in Fig 5C and 5D). Crucially, from a communication perspective, the complete system also acts to transfer mechanical motion from the snapping organ to the substrate. This represents another form of mechanical power transformation, albeit one that is modulated by the substrate. For motion vertical to the plant stem for one individual, the velocity ratio of motion measured on the plant relative to motion measured on the insect midabdomen indicates that the velocity of motion is attenuated by 83% (average −15. 5 ± 6. 2 dB), with lower attenuation in velocity of motion between the prothorax and plant at 71% attenuation (average −10. 5 ± 5. 5 dB, S1 Data and S5 Fig).
The consistency of the snapping organ’s morphology, and its systematic distribution across planthoppers indicates that this most likely represents a conserved mechanism for generating abdominal vibrations across the Fulgoromorpha. Previous studies have only examined delphacid vibrational organs [24,26,34], but our analysis of their peculiar morphology indicates that the drumming organs of delphacids are the exception and not the rule. The consistency of snapping organ morphology across the rest of the planthoppers provides a clear mechanistic explanation for the observed uniformity of their vibrational signals [25,33]. These findings reflect the fundamental importance of vibrational signals in planthopper communication. The functional morphology of the snapping organ also reveals some remarkable functional convergences and some equally remarkable mechanistic differences between the mechanical communication mechanisms of planthoppers and their close relatives, the cicadas [12,24]. Both make use of paired elastic recoil mechanisms and low-frequency active muscle contractions to enhance the efficiency and efficacy of communication, using exoskeletal integration to transform mechanical impulses into substrate vibration [2,12]. Driven by a single muscle, the cicadas’ tymbal organs use buckling instability of multiple stiff ribs to store and release elastic energy, turning slow muscle contraction into fast motion as the ribs buckle [12]. Muscle relaxation and the release of energy stored in resilin pads causes the ribs to restraighten again, leading to a second step involving elastic energy release [15]. In contrast, the snapping organ uses two different energy-storage mechanisms for paired elastic recoil: elastic storage in contracted muscle for loading and elastic storage in the deformed exoskeleton for unloading. Instead of buckling like the ribs of a tymbal, the arms of the Y-lobe in the snapping organ use snapping motions similar to those used in fast raptorial strikes by jaws and claws [8,9]. Finally, whereas tymbal vibrations in most cicadas are often associated with resonant chambers that act to transform motion into loud acoustic signals [12], the snapping organ is specialised for substrate-borne vibration generation, with comparable muscle contraction rates that act to transfer mechanical energy into vibrations of the substrate [12]. In conclusion, the unique biomechanics of the snapping organ demonstrate the general importance of elastic recoil mechanisms in the fast motions of small arthropods, extending our knowledge of such mechanisms beyond the simpler one-off ballistic motions that characterise jumping, predatory strikes, and feeding. Elastic recoil is a very general mechanism allowing small animals to overcome the limitations of their size and enabling robust vibrational communication.
Individuals of A. bilobum, the model planthopper species used in this study, were collected in large numbers (n = 250) in late April 2017 as fourth/fifth-instar larvae or adults from Lycabettus Hill, Athens, Greece, and were imported to Oxford, UK under DEFRA Plant Health Licence no. 52972/198417/6. Larvae were reared into adulthood in mesh cages (47. 5 cm × 47. 5 cm × 47. 5 cm) kept at 22–29°C, 50% humidity, with a 16: 8 photoperiod (light/dark). In addition, the morphology of specimens from more than 130 taxa were examined, covering the entire phylogenetic spectrum of Fulgoromorpha. S1 Table details the techniques used to examine the morphology of the snapping organ for each species, along with its preservation method. Planthoppers belonging to 12 families (including three specimens of A. bilobum: adult male, female, and larva) were used for synchrotron radiation microcomputed tomography (SR-μCT) at the TOMCAT beamline, Swiss Light Source (SLS), Paul Scherrer Institute, Switzerland (S1 Table). All specimens were scanned at a beam energy of 15. 99 keV with a final pixel size of 1. 625 μm, allowing visualisation of even the smallest muscles and nerves of the snapping organ (Figs 1B and 2B and S2 and S3B–S3D), which were otherwise not detected by other techniques. Three-dimensional reconstruction was carried out using Amira 6. 1 software (Mercury Systems, Andover, MA, USA). All shown tomographic data (reconstructed TIFFs) for the two imaged species (A. bilobum and Stenocranus minutus) are freely available at CXIDB (http: //cxidb. org/id-93. html) [35]. Colouration and labelling of figures were performed in Adobe Illustrator CS6. In order to reveal the primary DVMs operating the snapping organ in A. bilobum, the ventral junction between the Y-lobe and tg2 were excised from an ethanol-preserved (70%) male (Fig 2C). The dissected sample was placed between two cover slips in 70% ethanol and was imaged with a laser confocal scanning microscope (Olympus FV1000; Olympus, Tokyo, Japan) at a laser wavelength of 488 nm. The morphologies of specimens belonging to all 21 planthopper families were examined under light microscopy. Images of the snapping organ of four species of planthoppers shown in Fig 3 were taken using a Leica M165c microscope equipped with a Leica DFC490 camera (Leica, Wetzlar, Germany). The final, stacked images were combined using Helicon Focus (Helicon Soft, Kharkiv, Ukraine). Image brightness adjustment was performed in Adobe Photoshop, and drawings were generated in Adobe Illustrator CS6. To record vibrational signals, planthoppers were placed on a dried grass (Schedonorus giganteus) stem (17 cm in height). The base of the stem was inserted inside an empty c. 1-cm–diameter tube and was held in place by aluminium foil. Vibrational signals were recorded by a laser Doppler vibrometer (Polytec PDV-100; Polytec, Waldbronn, Germany), focussed at different positions approximately orthogonal to the stem and bug. A sampling frequency of 9. 6 kHz was used for recordings at a gain of 100 mm/s/V. Recording started immediately once the planthoppers were placed on the stem. Each recording lasted 6 minutes and was repeated until the animal either ended its vibrational call or after four recordings if no songs were present. A total of 61 recordings were made, 31 on single planthoppers, 26 on male–female groups, and four on male–male groups, using a total of 19 individuals (12 males, 7 females). Recordings from two individuals are included in S1 Data, in which the laser was focussed on the plant stem (individual 1), bug prothorax (individual 1), bug genitalia (individual 2), or bug midabdomen (individual 1). All vibrometry recordings were similar in the type and pattern of motion observed, so the data presented in S1 Data and S5 Fig are assumed to be representative. Attenuation of motion during loading and unloading from the midabdomen to the plant stem and prothorax to the plant stem was calculated in decibels (S1 Data). Vibrometry figures were drawn using Raven Lite 2. 0 (Cornell Lab of Ornithology, Ithaca, NY, USA) and OriginPro 8. To stimulate vibration generation, we used playback tracks of recorded songs. The stem was vibrated 7. 3 cm from the base by a pin glued to a small piezo disc (RS Components, Corby, UK), which was glued on an inverted plastic cup. Playback songs consisted of prerecorded and amplified vibrational signals of both sexes. All males responded to the playback by emitting a series of pulses for several minutes. The motion of the snapping organ in A. bilobum was captured with a high-speed camera (Grasshopper3 2. 3 MP Colour USB3 Vision, Sony Pregius IMX174; Point Grey, Richmond, BC, Canada) mounted on a Leica S8 AP0 stereomicroscope, recording at a rate of 100 frames s−1. Videos were recorded directly to a computer using Spinnaker SDK-1. 3. 0. 21 software (Point Grey). A total of three males were video recorded, and a movie and still frames from one male are given in Fig 4 and S1 Movie. The males were filmed over multiple cycles, frames were classified into the different stages of the mechanism, and the clearest frames were chosen from these classified groups within Fig 4. Pixel coordinates of three points on the bug prothorax were quantified for each frame used in Fig 4 to check for alignment of the bug within the video frame over time. Standard deviation over the five frames for each of the three points was within the order of 0. 01 pixels, suggesting the bug has limited movement within the video frame over successive cycles (also supported by S1 Movie). Prior to recording, it was necessary to expose the snapping organ by removing the fore and hind wings with a scalpel. The males were then left on their host plant for one hour to recover after wing removal before playback recordings were started to stimulate vibration generation. Based on our observations, the motion captured in S1 Movie is representative of the vibration-generation mechanism across different individuals. The vibrometry recordings were analysed to calculate the peak energy and power of the loading and unloading motions (S1 Data). Maximum and minimum peak velocities and the timings of the peaks were extracted from the vibrometry data. The peak kinetic energy of the motion was calculated from the speed of the measured dorsoventral translation of the abdominal mass, and the corresponding mechanical power was determined by dividing this peak kinetic energy by the time taken to reach it from rest. The muscle power density that would be required to generate this motion through direct muscle contraction was calculated by dividing these values by the relevant muscle mass, as measured from SR-μCT measurements of A. bilobum, modelling muscles as cylinders with a density of 1,060 kg m−3 [36]. A mathematical model was developed to support the interpretation that unloading was due to elastic recoil of the system (Figs 5 and S4). The model included the abdomen as a mass attached to two rigid bars in series (anterior Y-lobe arm and ridge, respectively), each with a stiff rotational spring at their junctions. The anterior bar was fixed to a surface, representing the thorax. Springs and dampers acting on the mass of the abdomen modelled the combined action of the muscles, resilin, other exoskeletal components, and interior morphology on the motion of the mass in the dorso–ventral and anterior–posterior planes. Full details of the model are given in S1 Methods.
Animals use substrate-borne vibrations for eavesdropping and communication over an immense range of body size—from elephants to nematodes. Vibrational communication is especially challenging for small animals because of the high mechanical power that is needed to transmit information effectively over extended distances through a substrate. Here, we show that planthoppers, a commercially important group of insects, produce vibrations for communication using a reciprocal elastic recoil mechanism that proves remarkably effective at small body size. By combining morphological and biomechanical analyses of a previously overlooked vibratory organ on the abdomen, we show that planthoppers use fast, cyclical abdominal motions to generate substrate-borne vibrations. This novel, to our knowledge, mechanism, which we term the snapping organ, makes use of slow energy storage and fast elastic recoil twice during each cycle of motion, involving two distinct elastic elements. This cyclical mechanism allows planthoppers to transmit signal pulses containing a broad range of frequencies to the substrate. The mechanism is efficient, achieving fast cyclical motion without relying on high muscle power and mass, both of which are limited for animals of small size. The snapping organ is ubiquitous across planthoppers and presents an interesting example of how elastic mechanisms can be used to enable nonacoustic vibrational communication between animals.
medicine and health sciences
classical mechanics
abdomen
diagnostic radiology
vibration
social sciences
electromagnetic radiation
muscle contraction
mechanical energy
animal behavior
synchrotron radiation
zoology
research and analysis methods
muscle physiology
musculoskeletal system
animal communication
imaging techniques
behavior
abdominal muscles
muscles
tomography
physics
psychology
radiology and imaging
diagnostic medicine
anatomy
physiology
biology and life sciences
physical sciences
Planthopper bugs use a fast, cyclic elastic recoil mechanism for effective vibrational communication at small body size