For the control of locomotor behaviors such as flight or walking, the ability to stabilize gaze is of central importance for reducing motion blur (Land, 1999). In diverse animals, gaze stabilization is achieved through sensorimotor circuits that rapidly sense and correct any deviation from the intended course, typically by eliciting reflexive motor responses of their head and body (Hardcastle and Krapp, 2016; Land, 1999; Waespe and Henn, 1987). Of these, a class of responses called optomotor reflexes refer to those behaviors that occur in response to optic flow (Borst, 2014). Although gaze stabilization typically requires eye as well as neck movements, in most flying animals including insects (Hardcastle and Krapp, 2016; Kemppainen et al., 2022; Land, 1999), birds (Hardcastle and Krapp, 2016; Necker, 2007), and bats (Eitan et al., 2019), eye movements relative to the head are either reduced or absent. Such animals stabilize their gaze primarily using head and body movements guided by optomotor reflexes.

In insects, optomotor reflexes have mostly been studied in tethered flies (Diptera) using assays in which their heads were fixed to the thorax with glue while they were presented with a moving display (Borst, 2014). In such preparations, the entire image stabilization is achieved through compensatory wing movements which may be tracked by infrared sensors or through high-speed videography (e.g. Duistermars et al., 2007). The stroke amplitude difference between the wings can then be used to close the loop between display image and wing movement, allowing the fly to control the projection of the visual environment onto their eyes. This approach has been very successful in helping to uncover diverse visual capabilities of insects. However, because the head was glued to the thorax in these preparations, the role of head movements in image stabilization typically fell out of the purview of such studies. Recent work on the fruit flies Drosophila melanogaster suggests that head-fixation adversely affects mechanical power output for flight (Cellini and Mongeau, 2020). Moreover, head-fixation in tethered Drosophila impairs their ability to fixate on specific objects moving relative to ground; if an object is moving relative to a moving ground, wing movements depend on both object and ground motion, but head movements depend only on ground motion (Fox and Frye, 2014). Head-fixation has also been shown to affect steering maneuvers in locusts Locusta migratoria (Robert and Rowell, 1992). Head and wing movements are thus crucial in steering maneuvers during flight and contribute strongly to gaze stabilization.

In Diptera, head stabilization is mediated by feedback from multiple sensory systems. These include visual feedback from their compound eyes and the mechanosensory feedback and/or feedforward control from their halteres which are modified hindwings that detect aerial rotations (Hengstenberg, 1988; Hengstenberg, 1993; Schwyn et al., 2011). Additionally, mechanosensory proprioceptive structures in the ventral neck-prothorax region called prosternal organs also aid in head positioning (Paulk and Gilbert, 2006; Preuss and Hengstenberg, 1992). The relative contributions of these sensory systems depend on the angular speed of the maneuver because vision typically provides feedback at slower rates than mechanosensors (Hengstenberg, 1993). Importantly, during aerial maneuvers, haltere feedback combines non-linearly with visual feedback in some neck motor neurons to determine the compensatory head movements (Huston and Krapp, 2009). Thus, this system offers an interesting case study for how multisensory feedback from visual and mechanosensory modalities is integrated during locomotion. Because mechanosensory feedback from halteres is transduced at faster rates than visual feedback, it is hypothesized to play a key role in guiding wing movements during rapid maneuvers, whereas visual feedback mediates slower turns (Sherman and Dickinson, 2003).

Fewer studies have focused on head stabilization in freely flying insects as compared to tethered insects. Free-flight studies in diverse insects (honeybees Apis mellifera; Boeddeker and Hemmi, 2010; Wehner and Flatt, 1977, blowflies Calliphora vicina; Hateren and Schilstra, 1999) show that rapid head turns precede sharp saccadic turns, whereas gaze stabilization between saccades ensures that the visual field largely translates over the retina, thereby enabling depth perception. When flying through a patterned, oscillating drum surrounding their nest entrance, freely flying bees adjust their head angle based on wide-field image movement in their visual field (Boeddeker and Hemmi, 2010; Wehner and Flatt, 1977). Moreover, visually induced head roll response is limited beyond specific roll frequencies, suggesting constraints to visually mediated gaze stabilization (Boeddeker and Hemmi, 2010).

These studies demonstrate that optic flow plays a key role in head stabilization behavior in both tethered and freely flying insects. However, the role of mechanosensory feedback in gaze stabilization has been largely ignored in insects that lack halteres. Is head roll in these insects primarily visually driven or does it also require mechanosensory feedback from some haltere analogue? Head stabilization in wasps appears to be primarily visually mediated, although some degree of feedforward control has been suggested (Viollet and Zeil, 2013). Similarly, in day-flying insects such as dragonflies, head stabilization appears to be dominated by visual feedback, and these insects contain adaptations that ensure passive inertial head stabilization (Gorb, 1999; Hardcastle and Krapp, 2016; Mittelstaedt, 1950). Yet, being predatory insects, they achieve precise visual fixation of targets during prey capture in flight (Lin and Leonardo, 2017; Olberg et al., 2007). The role of mechanosensory feedback in head movements in dragonflies remains an open question, especially as their antennae are very small and they do not possess halteres (Gewecke et al., 1974). In contrast, nocturnal ants can stabilize their heads in darkness during walking, suggesting mechanosensory input for stabilization (Raderschall et al., 2016). However, the mechanosensors mediating head stabilization in non-Dipteran insects are yet to be identified.

One potential candidate for the mechanosensory mediation of head stabilization is the Johnston’s organ, which is a highly sensitive mechanosensory structure situated in the antennal pedicel-flagellar joint of all insects (e.g. Gewecke, 1974; Sant and Sane, 2018). In hawkmoths, the Johnston’s organ consists of hundreds of concentrically arranged scolopidial units that are range-fractionated, which means that the scolopidial units are tuned to different frequency ranges with high sensitivity (Dieudonné et al., 2014). Antennal mechanosensors contribute in sensing airflow (Roy Khurana and Sane, 2016; Natesan et al., 2019; Taylor and Krapp, 2007), and maintaining headwind orientation (Fuller et al., 2014). In moths, flight becomes unstable when the mechanosensory load on the Johnston’s organ is reduced by clipping off their flagella (Dahake et al., 2018; Sane et al., 2007). In walking crickets, there is evidence that antennae contribute to compensatory head roll (Horn and Bischof, 1983) and electrophysiological recordings from the neck motor system in flies further revealed that the ventral cervical nerve motor neuron in flies receive input from the antennae (Haag et al., 2010).

To determine the relative contributions of visual and mechanosensory inputs in gaze stabilization during flight, we studied head movements in the nocturnal hawkmoth, Daphnis nerii. In our experiments, moths were tethered to rotatable motor shafts, and we measured the compensatory counter-rotation of their head at two different angular rotations of their body under variable light conditions (Figure 1A–F; see Materials and methods). Additionally, we assessed their ability to use antennal mechanosensory feedback by ablating and reattaching their antennal flagellum. Our data show that at lower turning rates of the body, visual feedback is essential for head stabilization. However, at higher angular velocities, feedback from the Johnston’s organs plays a crucial role in the control of head rotations, consistent with the hypothesis that this feedback combines with vision to modulate compensatory head movements. We also show that head movements are essential for controlled flight. Thus, the observed reduction in flight performance due to the loss of mechanosensory feedback from the antenna may be related to impaired gaze stabilization.

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Head stabilization performance was characterized using three parameters: gain and phase of the response, and compensation error which is a metric that combines both gain and phase. The head-compensation error parameter was computed as the distance between the perfect head stabilization point and the experimental point from a trial (Figure 1E; for details, see Materials and methods, also see Figure 2—figure supplement 2). Note that all three parameters must be considered to fully interpret a compensatory head response. When the gain is zero, the system offers no response and hence the phase values at this gain are uninterpretable. For non-zero gain, the phase values indicate how closely the response matches the stimulus in phase. However, in many cases, heavily delayed responses may appear as leading the stimulus in phase. Also, because gain and phase are inherently linked, they cannot be interpreted or statistically compared separately. The compensation error is a metric that enables us to statistically analyze these results.

Visual cues affect head stabilization at low frequency

To assess the role of visual cues in mediating compensatory head rotations, we provided rotating stimuli to tethered moths at different light levels (Twilight condition: ~250 lux; dark condition: <0.01 lux; see Materials and methods). At each light level, the moths were rotated by ±30° about their longitudinal roll axis at oscillation frequencies of 2 Hz (Figure 2A) and 6 Hz (Figure 2B), while we measured their compensatory head movements.

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Under twilight condition at 2 Hz, the compensation error was low with a median ± standard error of mean of 0.4±0.02 (open circles, Figure 2A and C). However, when we rotated the moths at 2 Hz under dark condition (filled circles, Figure 2A and C, Figure 3—figure supplement 1A,C), a reduction in gain (twilight condition median: 0.6, dark condition median: 0.4) and a shift in phase (twilight condition circular median: 177.6°, dark condition circular median: 302.4°, Figure 3—figure supplement 3: Cases 1 and 2) increased the compensation error to 1.2±0.09, suggesting a severe reduction in their ability to generate compensatory head movements (p=0.0078, Supplementary file 1c, 1, Figure 3—figure supplement 1A,C). Additionally, the compensation error values of moths under dark conditions were more variable, which is also evident in the distribution of points in the polar scatter plots (compare open and filled circles, Figure 2A; see also Figure 2C, Supplementary file 1a). A different method of phase computation (see Materials and methods) yielded a nearly identical result (Figure 2—figure supplement 1, compensation error under twilight condition: 0.4±0.02, dark condition: 1.2±0.09, p=0.0078, Wilcoxon signed-rank test). Thus, visual input plays a crucial role in mediating head stabilization at 2 Hz (Video 1).

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We next increased the oscillation frequency to 6 Hz, and observed that under the twilight condition, the compensation error was 0.7±0.02 (open circles, Figure 2B–C). At 6 Hz, the phase-shift between the twilight and the dark condition was reduced in comparison to the 2 Hz case (twilight circular median: 173.7°, dark circular median: 214.2°) whereas the gain between the two conditions was similar (Figure 2B, Figure 3—figure supplement 1B,D, see also Figure 3—figure supplement 3 Cases 1 and 2). The compensation error (Figure 2C) remained largely unaltered with a median ± standard error of mean of 0.7±0.05 (p=0.5469, Supplementary file 1c, 2). However, similar to the 2 Hz case, compensation error was more variable under dark conditions (compare open and filled circles, Figure 2B, Supplementary file 1a). We got similar results using Fourier-based phase computation. (Figure 2—figure supplement 1B,C, compensation error under twilight condition: 0.6±0.02, dark condition: 0.7±0.06, p=0.4609, Wilcoxon signed-rank test). Thus, at 6 Hz, severe reduction of visual feedback did not greatly hamper head stabilization, suggesting that at roll rotation of higher frequency, an additional – perhaps mechanosensory – feedback played a role in head stabilization. However, the phase-shift and increased variability in the dark condition response suggests that some visual feedback may be required even at 6 Hz.

Antennal inputs affect head stabilization primarily at high frequency

Mechanosensory feedback from the antennae has been previously shown to mediate flight control in hawkmoths (Dahake et al., 2018; Sane et al., 2007). To test the hypothesis that it also plays a role in head stabilization, we clipped the antennal flagella, thus greatly reducing the inertial load of the antenna. We subjected these moths to roll stimuli (±30°) at oscillation frequencies of 2 Hz and 6 Hz in both twilight and dark conditions, and filmed the resulting head movements (Figure 3A–E).

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Under both twilight and dark conditions at 2 Hz (Figure 3E), the compensation error was significantly different between the flagella-intact (twilight: 0.4±0.02; dark: 1.2±0.09) and flagella-clipped groups (twilight: 0.5±0.01; dark: 1±0.01, p=0.0107, p=0.0133, Supplementary file 1c, 3, 4). Under the twilight condition, the gain values remained largely unaltered with a slight phase shift in the flagella-clipped moths in comparison to the flagella-intact ones from a circular median of 177.6° to 174.0° (open circles, Figures 2A and 3A, Figure 3—figure supplement 3 – Cases 1 and 3). Under the dark condition, the gain was very low in flagella-clipped moths with a median of 0.1 (filled circles, Figure 3A, Figure 3—figure supplement 1E,G, Figure 3—figure supplement 3 – Case 4). Because the gain values are very low, the phase values were uninterpretable in this context. Unlike in the flagella-intact group at 2 Hz under dark conditions, the variability of the compensation error was very low under flagella-clipped moths (Figure 3E, filled circles, Figures 2A and 3A, Supplementary file 1a). The response (open and filled circles, Figure 2A) of flagella-intact moths at 2 Hz in the dark could, therefore, potentially result from antennal mechanosensory input, in turn influencing compensatory head movements at the low frequency in the dark.

At 6 Hz (Figure 3E), the compensation error was significantly different between the control (twilight: 0.7±0.02, dark: 0.7±0.05) and flagella-clipped (twilight: 0.8±0.03, dark: 1.0±0.03) groups in both twilight (p=0.0064) and dark conditions (p=0.0011) (Supplementary file 1c, 5, 6). Under twilight conditions, we observed a shift in phase in the flagella-clipped moths compared to the flagella-intact group from a circular median of 173.7° to 127.8° although the gains were similar (open circles, Figures 2B and 3B, Figure 3—figure supplement 3 – Cases 1 and 3). Comparable to the responses observed at 2 Hz, the gain (median: 0.1, filled circles, Figure 3B, Figure 3—figure supplement 1H) and variability (Figure 3E, Supplementary file 1a) were lowest in flagella-clipped moths under dark conditions. These observations suggest that at 6 Hz, antennal mechanosensory input may be critical for head stabilization. Despite reduced antennal mechanosensory feedback, there are some indications of head stabilization under twilight conditions (compare open circles to filled circles, Figure 3B). This again suggests that visual feedback plays a role at this frequency. However, the compensation error of flagella-clipped moths under twilight condition at 6 Hz is significantly greater in comparison to 2 Hz (p=0.0156, Wilcoxon signed-rank test) because there is a shift in phase from a circular median of 174° to 127.8°. However, the gain values are similar (Figure 3—figure supplement 3, open circle: Figure 3A, B). Thus, the response due to visual feedback is reduced during faster body rolls.

Does restoring the mechanical load of flagella rescue head stabilization in the flagella-clipped moths? To determine this, we repeated the experiment in moths with reattached flagella. After reattachment of flagella, the head stabilization was similar to the performance of the moths with intact antennae, for all conditions. For both 2 and 6 Hz rotations, the compensation error in flagella-reattached moths (Figure 3C–E, Figure 3—figure supplement 1I-L) was not significantly different from the flagella-intact group under twilight and dark conditions (see Supplementary file 1c, 3–6 for statistical comparisons). Similar to the response of the flagella-intact group at 2 Hz, we observed a phase-shift between the twilight and dark conditions in the flagella-reattached moths from a circular median of 177° to 297.6° at 2 Hz and from 149.4° to 216° at 6 Hz (Figure 3—figure supplement 3 – Cases 5 and 6, Figures 2A and 3C). For 6 Hz rotations in the dark condition (Figure 3E), the compensation error of the flagella-reattached moths differed significantly from the flagella-clipped group (Supplementary file 1c. 6). The coherence estimates were also highly variable in flagella-clipped moths under the dark conditions in comparison to the corresponding flagella-intact and flagella-reattached groups (Figure 3—figure supplement 2A, Supplementary file 1b). Together, these experiments suggest that antennal mechanosensory input is required for head stabilization in hawkmoths primarily at higher frequency, besides some contribution at lower frequency as well (Video 2). Our results also indicate some contribution of visual feedback at higher frequency rotation.

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The role of Johnston’s organs in head stabilization

Johnston’s organs respond to extremely subtle deformations and vibrations of the flagellum relative to pedicel (Lin and Leonardo, 2017). Their role in flight stabilization has been highlighted in previous studies (Olberg et al., 2007; Paulk and Gilbert, 2006), and hence we specifically tested their contribution to head stabilization. In the experiments described above, the ability of the flagella-clipped moths to stabilize their heads was reduced although visual feedback may offset this effect to a certain extent at 2 Hz and to a lesser extent at 6 Hz. We therefore tested the hypothesis that this effect in moths with removed flagellar load was specifically due to the reduction of mechanosensory feedback from the Johnston’s organs. To reduce feedback from the Johnston’s organ, we glued the pedicel-flagellar joint of moths in which the Johnston’s organ is embedded. Data from these experiments were compared with sham-treated moths in which the ~2nd–3rd annuli in the flagella were glued, rather than the pedicel-flagellar joint (Figure 4A–E).

Figure 4

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Under twilight and dark conditions for 2 Hz roll stimulus, the compensation error between the sham (twilight: 0.4±0.02, dark: 1.2±0.08) and Johnston’s organ-glued (JO-glued) groups (twilight: 0.4±0.01, dark: 1±0.04) was not significantly different (Figure 4E, p=0.5358, p=0.0831, Supplementary file 1c, 7, 8). However, similar to the flagella-clipped group, the JO-glued moths exhibited very low gain (median: 0.11) under dark conditions and therefore, their phase values in this condition are not interpretable (filled circles, Figures 3A and 4C). Also, similar to the flagella-intact group, the sham group showed high variability in the dark conditions, and phase values shifted from 178.5° in twilight condition to 306.3° in dark condition (Supplementary file 1a, Figure 4E, filled circles, Figure 4A). At 6 Hz in both twilight and dark conditions, the compensation error differed significantly between the sham (twilight: 0.6±0.03, dark: 0.6±0.06) and JO-glued (twilight: 0.9±0.05, dark: 1±0.04) groups (Figure 4E, p=0.0097, p=0.0055, Supplementary file 1c, 9, 10). Thus, we observed similar effects in JO-glued and flagella-clipped moths (Figures 3A–B, E–4C–E), suggesting that feedback from the Johnston’s organ affects head stabilization. The coherence values were variable in the JO-glued group in comparison to those obtained in the sham group under dark conditions (Supplementary file 1b, Figure 3—figure supplement 2).

Restricted head movements cause loss of flight performance in freely flying moths

Together, the above experiments on tethered moths indicated that mechanosensory feedback from the antennal Johnston’s organ is involved in head stabilization, in addition to the visual feedback from its retina (Figure 5). We next hypothesized that flight impairment observed in previous studies in the flagella-clipped hawkmoths, Manduca sexta (Sane et al., 2007), may arise from inability of these hawkmoths to stabilize their head during flight. If that is the case, then we expect moths with restricted heads to show similar flight impairments as those observed in the flagella-clipped moths. We filmed the free-flight trajectories (see Materials and methods) of moths whose head movements were restricted. We compared these with control moths in which the head was free to move, and with sham moths which underwent the same procedure as experimental moths but without head restriction. In our experiments, out of 12 animals in each group, 11 control animals, 12 sham-treated, and 8 experimental head-restricted moths produced free-flight behavior (comparison with internal control: Figure 6—figure supplement 1A, B). The flight duration of control (73±17.0 s) and sham (34±9.3 s) groups were significantly longer compared to the moths with restricted head-motion (1.5±0.7 s) (Figure 6A, p=0.0001, Supplementary file 1c, 11). We also observed a difference in the flight behavior of the head-restricted moths in comparison to the control and sham moths. Whereas the control and sham moths took off smoothly and pitched up to a considerable height, the head-restricted moths were unable to pitch up, and consequently stayed close to the ground usually touching it, and became unstable as soon as they became airborne (Video 3). The frequency of these moths (1±0.1 per second) colliding with the walls of flight chamber was significantly greater than in control (0.4±0.05 per second) and sham (0.3±0.05 per second) moths, thus indicating impairment in their flight performance (Figure 6B, p=0.0002, Supplementary file 1c, 12).

Figure 5

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Figure 6 with 2 supplements see all

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Video 3

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In the free-flight assay described above, the moths were not presented with any specific stimuli to generate flight activity. Thus, the flight duration depended entirely on when the moth voluntarily chose to stop flapping and land on a surface, which caused substantial variability in the free-flight data on control moths. In the head-restricted moths however, flight duration is significantly lower compared to all other cases, implying that head-restricted moths cannot fly in a controlled manner and collide more frequently per unit time, when compared to moths in the control and sham group (Figure 6—figure supplement 1).

Compensatory head movements are crucial for stability in free-flight (Figure 6A–B). The data presented in this paper show that such compensatory head roll in tethered flight is mediated via a combination of visual and antennal mechanosensory feedback in the hawkmoth, D. nerii (Figure 5).

We have estimated power that is the probability of detecting a difference between the groups with a significance level of 0.05 across a range of sample sizes for all the comparisons which yielded significant difference. We have used the mean and standard deviation of each dataset for generating normal distributions. We have then used the relevant tests that is Wilcoxon signed-rank, Wilcoxon rank-sum, and Kruskal-Wallis tests for the power analysis. For each sample size, the loop was repeated 1000 times. In all the plots, the power of 0.8 is marked in dotted line. Our sample size in all cases is greater than or close to the power of 0.8.

Appendix 1—figure 1

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Appendix 1—figure 2

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Appendix 1—figure 3

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Appendix 1—figure 4

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All data related to this paper (both raw and processed) are available on the following link: https://data.mendeley.com/datasets/2trxj9gwsw/draft?a=89356c26-e581-40c6-935d-c1d4a0401074.

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Author details

1. Payel Chatterjee

   National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Validation, Visualization, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7019-6702

2. Agnish Dev Prusty

   National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India

   Contribution

   Data curation, Investigation, Methodology, Software, Validation, Visualization, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-4890-9163

3. Umesh Mohan

   National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Software, Validation, Visualization, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1992-0558

4. Sanjay P Sane

   National Centre for Biological Sciences, Tata Institute of Fundamental Research, Bangalore, India

   Contribution

   Conceptualization, Funding acquisition, Investigation, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing

   For correspondence

   sane@ncbs.res.in

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-8274-1181

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