Since the dawn of agriculture, humanity has been in an arms race with insect pests. Traditionally, a set of integrated cultivation strategies tailored to local settings helped keeping pest insects at bay, including associational resistance through varietal mixtures and intercropping (Abate et al., 2000; Snyder et al., 2020; Wuest et al., 2021). With the advent of agrochemicals, monocultures superseded traditional strategies. However, their profound externalities on ecosystem resilience and global climate (Altieri, 2009; Shukla et al., 2019) have resuscitated interest in more sustainable alternatives, frequently grafted on traditional strategies. Trending terms such as agroecology, and climate-smart, regenerative, or organic agriculture highlight the search for solutions that harmonize food production and pest control with ecological sustainability. Some innovative practices have been important sources of inspiration. Among these, the push–pull strategy in which maize is intercropped with the legume, Desmodium, is arguably the most well known (Cook et al., 2007).

The push–pull strategies aim to reduce the abundance of insect pests in crops through repelling the ovipositing herbivores from the crop, while simultaneously attracting the pest outside the field (Miller and Cowles, 1990). Using this ‘stimulo-deterrent diversion’ principle, a push–pull strategy was devised to combat lepidopteran pests in sub-Saharan smallholder maize farming (Khan et al., 1997b; Khan et al., 2010). Embroidering on the common practice of smallholder farmers to intercrop maize with e.g. edible pulses, the strategy uses the perennial fodder legume Desmodium as intercrop in maize plots. Desmodium reportedly constitutively releases large amounts of volatile monoterpenes and sesquiterpenes, such as (E)-4,8-dimethyl-1,3,7-nonatriene ((E)-DMNT), (E)-β-ocimene and cedrene, that repel (push) lepidopteran pests and attract natural enemies (pull) (Hassanali et al., 2008; Khan et al., 1997a; Khan et al., 2000). A ‘trap crop’ sown as border crop (another ‘pull’ component), typically Napier grass, complements the strategy, as it induces oviposition in Lepidoptera, but reduces larval survival compared to maize (Khan et al., 1997a; Khan et al., 2000; Khan et al., 2016). This cropping strategy thus suppresses infestations with various lepidopteran pests, including Chilo partellus and Busseola fusca, as well as Spodoptera frugiperda, a polyphagous invasive pest that is ravaging maize and vegetable production and threatens food security in sub-Saharan Africa (Midega et al., 2018; Feldmann et al., 2019). This intercropping strategy has found widespread adoption in East Africa (Khan, 2011; Niassy et al., 2022; Nkurunziza, 2021; ICIPE, 2019; Government of Ruanda, 2011; Kenya National Assembly, 2019). As a hallmark of sustainable pest control, it also serves as a tremendous source of inspiration for intervention strategies in other cropping systems.

The ‘push’ volatile terpenoids reported in previous studies (Khan et al., 1997a; Khan et al., 2000) are usually released in detectable amounts by plants after induction by herbivory. Although several plants do release them constitutively, such as Melinis minutiflora (Kimani et al., 2000), constitutive release of volatile terpenoids is not known from legumes. We sought to understand the role of soil-borne interactions, particularly the soil microbiome, in the constitutive release of these volatiles. This is of particular interest given that push–pull intercropping of maize and Desmodium causes substantial shifts in below-ground ecosystems, including increased soil microbe diversification, increased soil nitrogen and carbon, increased plant defense through plant–soil feedback, and suppression of parasitic weeds and pathogenic microbes (Mutyambai et al., 2019; Mwakilili et al., 2021). Indeed, soil and root–microbe interactions have been found to induce pathways that lead to release of volatile terpenoids (Mutyambai et al., 2019; Malone et al., 2020). We therefore verified if the ‘constitutive’ release of volatile terpenoids was, in fact, induced or enhanced by soil-borne interactions. The root–microbe interactions are of particular interest, given the intimate association of legumes with specific microbial groups e.g. rhizobia and mycorrhizae.

Different from the expectations, in the headspace of intact Desmodium intortum, which is by far the most commonly used intercrop in push–pull technology, the presence of the earlier described (Hassanali et al., 2008; Khan et al., 1997a; Khan et al., 2000) volatile terpenoids was not detectable (Figure 1A, B, Figure 1—figure supplements 1 and 3). This was independent of the soil in which D. intortum was grown, whether live soil (organic potting soil, organic clay Swedish soil, or African clay loam soil from D. intortum plots), autoclaved soil, or autoclaved soils inoculated with mycorrhiza or rhizobacteria (Figure 2—figure supplements 1–4) was used. Similar results were obtained with D. uncinatum, a species that has also frequently been used in push–pull cropping systems (Figure 1—figure supplement 2). In contrast, we did confirm that M. minutiflora, a poacean plant used previously as a push intercrop, releases a diverse blend of terpenoids regardless of herbivory ( Figure 1—figure supplements 1–3). Intact Desmodium plants thus did not release the earlier described repellent compounds in detectable quantities, independent of soil interactions.

Figure 1 with 3 supplements see all

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Although the constitutive release of repellent volatile terpenoids is an important precondition for push–pull, inadvertent herbivory of Desmodium could result in a volatile emission similar to those reported in earlier studies for intact plants (Khan et al., 2000). In our experiments, only small amounts of volatile terpenoids were detected in the headspace of D. intortum plants when fed upon by S. frugiperda larvae (Figure 1A-C; Figure 2). This is in contrast with maize, which, in line with previous studies (Turlings et al., 1990; Degen et al., 2004), was detected to release large amounts of volatile terpenoids in response to herbivory, with emission peaking between 24 and 48 hr following infestation, and declining over the course of 7 days (Figure 1C). Herbivory of M. minutiflora did not significantly boost the release of volatile terpenoids above the already high constitutive release (Figure 1B, Figure 1—figure supplements 1 and 3).

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Arguably, greenhouse conditions are not representative of field conditions and additional, unknown factors in the field may cause the release of volatile terpenoids by Desmodium. We therefore analyzed 50 headspace samples from D. intortum from seven locations in Tanzania and Uganda. Also under field conditions, very few D. intortum plants released detectable amounts of terpenoids in the headspace. The few plants that did release terpenoids, did so in comparatively low amounts (Figure 2, Figure 2—figure supplement 1). This was most likely induced by herbivory, which upon further inspection was visible on these sampled plants. This confirms the greenhouse experiments and underlines that Desmodium does not constitutively release detectable amounts of volatile terpenoids, whereas following induction, the release is comparatively low. Although it cannot be excluded that other conditions or more substantial herbivore attack may induce higher release of volatile terpenoids, our experiments conducted under different growth conditions, and in different geographic regions show that this is likely rare, which would make it tenuous to be at the core of a generic strategy. In contrast, the headspace of field-sampled maize, most of which displayed some herbivore damage, did contain typical herbivore-induced volatile terpenoids (Turlings et al., 1990; Degen et al., 2004; von Mérey et al., 2013; Figure 2, Figure 2—figure supplement 1), with variations in volatile release likely reflecting differing levels of, and age since herbivore infestations, which could not be controlled in the field. The findings reported here are results of greenhouse experiments and field experiments from three geographical areas (Tanzanian highlands and lowlands, and Uganda) involving a variety of abiotic and biotic factors (including genetic background of D. intortum and potential herbivory). However, how these, and other environmental factors may have influenced volatile emission have not yet been investigated specifically and warrant further study.

Although volatile terpenoids were sparsely observed in the headspace of Desmodium, and seemed to be an unlikely cause of oviposition repellence, we tested the oviposition repellency in bioassays. In a modified wind tunnel, gravid S. frugiperda were given a choice between maize plants with either D. intortum or artificial plants in the background (Figure 1D, Figure 3—figure supplement 1). Adult females landed on either maize plant and the number of egg batches were not significantly different, underlining that odor from D. intortum that was placed upwind from the maize plants, did not elicit significant oviposition repellence in gravid S. frugiperda (Figure 3B). Contrary to our findings, D. intortum volatile emission appeared to be repellent for S. frugiperda in a recent study (Sobhy et al., 2022). However, the volatile profile of the D. intortum plants in the choice tests (Sobhy et al., 2022) was very different from intact plants we have studied and was reminiscent of the Desmodium plants under active herbivory infestation by larvae (Figure 1—figure supplement 1).

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In order to explain the suppression of lepidopteran pests using Desmodium as intercrop, one needs to invoke a different mechanism than odor-based ‘stimulo-deterrent diversion’ or ‘push–pull’. To investigate possible alternatives we scored S. frugiperda oviposition preference, larval feeding preference, and larval survival on maize and Desmodium. First, in two-choice tests S. frugiperda preferred to lay eggs on maize over Desmodium. Yet, the preference was not strong, as females also oviposited on Desmodium. In the field, one could perhaps expect a further shift toward Desmodium, particularly when maize is small and Desmodium, a perennial, well developed.

Though, irrespective of female oviposition choice, larvae of many lepidopteran species are known to disperse from the plant on which they hatched. Neonate, first instar larvae rapidly disperse to avoid sibling competition. Besides locomotion, they also passively disperse with wind through spinning silk threads allowing them to ‘parachute’ between plants (Njuguna et al., 2021; Rojas et al., 2018; Sokame et al., 2020). Later larval stages, which no longer disperse with wind, have been observed to actively disperse across the soil in search for new host plants (Sokame et al., 2020; Berger, 1994; van Rensburg et al., 1988). Given the dense, contiguous ground cover provided by Desmodium in the interrows, stochastically a large majority of dispersing larvae would end up on Desmodium, particularly when maize plants are small and Desmodium, a perennial, large. We asked whether these larvae would feed and survive on Desmodium. In feeding choice assays, the number of first instar S. frugiperda larvae on leaf discs and the area consumed was significantly higher for Desmodium compared to maize (Figure 4). However, in survival analyses, the development of those fed on Desmodium stagnated, with hardly any larva molting to the second instar, and none reaching pupation (Figure 5). Several sensory modalities including vision (Han et al., 2024), olfaction (Castrejon et al., 2006), taste (Castrejon et al., 2006; Sun et al., 2022), and tactile stimuli can potentially influence the consumption patterns observed in the larval choice assays. While further studies are needed to elucidate the mechanism of preference, the data demonstrate that Desmodium is a palatable plant for dispersing larvae, yet does not support larval development.

Figure 4

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Figure 5

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In addition to stagnating development, we found that larvae moved slowly on Desmodium leaves and stems, and many were entirely immobilized, particularly visible at later larval instars. Closer scrutiny of D. intortum surfaces revealed a dense network of non-glandular, uniseriate, and uncinate trichomes, with densities and a distribution depending on the surface type (Figure 6A, D, F; Figure 6—figure supplement 1). The stems and main veins of the leaves were particularly densely populated with large uncinate trichomes. First instar larvae were somewhat freely moving and grazing between trichomes of the stem (Figure 6—figure supplement 1C), but older larvae were seen impaled and immobilized by them (Figure 6C, D; Figure 6—figure supplement 1D–F). Occasionally, even ovipositing S. frugiperda were immobilized at their ovipositor on D. intortum (Figure 6—figure supplement 1G). Whereas trichomes were flexible at the base, they were fortified with silica toward the tip (Figure 6F), equipping the plant with an effective mechanism to obstruct, damage and immobilize herbivores. Also beneficial insects (Figure 6—figure supplement 1I) and even vertebrates can be trapped by Desmodium (Coleman, 2016). Incidentally, the presence of small uncinate trichomes on leaves could also have affected the movement of first instar larvae and thereby support the observed preference pattern observed earlier (Figure 4) and survival rate (Figure 5). Stellar non-glandular trichomes were shown to decrease feeding of Manduca secta larvae on several Solanaceae species (Kariyat et al., 2018), and uniserate non-glandular trichomes of bottle gourd (Lagenaria siceraria) affect the feeding and survival of Trichoplusia ni (Kaur and Kariyat, 2023). How the size, shape and density of these surface structures affect lepidopteran behavior in a species- and stage-specific manner needs to be addressed in future studies. Uncinate non-glandular trichomes are used by many other plant species (Ballhorn et al., 2013; Gilbert, 1971), and may serve multiple purposes including seed or fruit dispersal (Xing et al., 2017; Sorensen, 1986; Freitas et al., 2014).

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We thus infer that in the field Desmodium trichomes affect fitness of lepidopteran larvae, both directly and indirectly. First, Desmodium entices larval feeding, but truncates development. Second, trichomes on Desmodium hinder movement, damage the cuticle and even entirely immobilize larvae on the plant, increasing developmental time, exposure to natural enemies and overall mortality (Kaur and Kariyat, 2023; Kariyat et al., 2017; Kaur and Kariyat, 2020). Third, ingestion of trichomes damages the intestinal lining and affects digestion, development and survival in closely related species (Kaur and Kariyat, 2020; Acevedo et al., 2021). Indeed, while first instar larvae fed around the large uncinate trichomes, larger larvae did ingest trichomes as evidenced by trichomes found in larval frass. Effectively, rather than functioning as a repellent intercrop, Desmodium appears to be a trap crop for larvae.

We hypothesize that ‘push’ does not describe the mode of action of Desmodium. Instead, the plant exhibits properties reminiscent of a ‘pull’ crop, a ‘trap crop’. Although superficially similar in mode of action to the ‘pull’ border crop Napier grass, Desmodium is distinctly different, as it is preferred by larvae, not by adults (Khan et al., 1997b; Hassanali et al., 2008). In addition, Desmodium forms a mechanical barrier to dispersing larvae. Further field studies need to detail how oviposition preference of different stemborer species, larval dispersal, development and survival on Desmodium, mechanical obstruction by Desmodium, and additional mechanisms such as parasitization and predation rates, interplay with crop phenology in suppressing various lepidopteran species across the cropping season. Knowing the exact mode of action is critical if we, for instance, wish to substitute the fodder crop Desmodium with a food crop to enhance food security, or design push–pull inspired, pest-suppressive conditions for other crops.

The observation that Desmodium does not emit detectable amounts of volatile terpenoids and does not repel S. frugiperda, contrasts with the large number of publications and the global attention that maize–Desmodium push–pull technology has garnered over more than two decades. Indeed, the idea of the ‘push’ crop Desmodium repelling moths is found in numerous papers since its first mention around the year 2000. However, close scrutiny of the literature revealed a limited amount of primary data, except for a recent paper discussed below (Sobhy et al., 2022). The data presented here suggest that the mechanism of push–pull requires detailed studies. This is important as, although push–pull clearly suppresses lepidopteran pests (Cook et al., 2007; Khan et al., 2010; Hassanali et al., 2008; Khan et al., 1997a; Khan et al., 2000; Khan et al., 2016; Midega et al., 2018; Feldmann et al., 2019; Mutyambai et al., 2019), knowing the precise mechanism is essential to optimize the strategy to and troubleshoot it when it underperforms. How the mechanical defense described here impacts herbivore population, growth rate, the rate of parasitization and predation, depends on other biotic and abiotic factors and needs to be further studied as well. With the mechanism at hand, the strategy can also be further tailored to the needs of local smallholder farmers e.g. replacing Desmodium with food crops with similar properties (Kariyat et al., 2018; Kaur and Kariyat, 2023; Ballhorn et al., 2013; Gilbert, 1971; Xing et al., 2017; Acevedo et al., 2021; Johnson, 1953; QulRing et al., 1992), as well as rationally translating the concept to other cropping systems.

Ideas and speculations

In our experiments, we exclusively detected volatile terpenoids when D. intortum was damaged by herbivores. In a recent study, aimed at testing the repellence of Desmodium to S. frugiperda under laboratory conditions, D. intortum plants may appear to emit volatile terpenoids constitutively (Sobhy et al., 2022). However, besides difference in odor collection methodologies (see below), the objective and experimental design of that study differed substantially from the current study, which makes comparison with the current study tenuous. Sohby et al. primarily aimed to assess the preference of S. frugiperda for maize alone or in combination with Desmodium without focusing on herbivore induction. At a first glance the volatile profile of intact Desmodium and maize in Sobhy et al., 2022 would appear herbivore-induced, as these profiles compare well with our herbivore-induced Desmodium and maize. However, the absence of both positive and negative control plants (induced and non-induced Desmodium and other push-plants such as M. minutiflora) in Sobhy et al., 2022, makes a direct comparison difficult. The current study provided contrasts through the inclusion of these controls. Desmodium, under a large range of conditions in the laboratory and the field, did not release detectable amounts of induced volatile terpenoids, and comparatively little when induced. While the antenna of S. frugiperda females (Sobhy et al., 2022) can detect volatile terpenoids, in our study these appeared only to be released in detectable, small amounts upon herbivory of Desmodium plants.

Besides differences in the hypotheses and design of the studies, a methodological factor, the volatile collection methods, is also worth considering. Static (this study) versus dynamic headspace sampling (Sobhy et al., 2022) in combination with other factors such as the adsorbent used, impacts volatiles collected in many ways, and in turn its interpretation ( Tholl, 2020; Tholl et al., 2006; Raguso and Pellmyr, 1998; Ouyang and Pawliszyn, 2008; Prosen and Zupančič-Kralj, 1999). Solid-phase microextraction (SPME) used here allows for time and cost-efficient collection of large numbers of samples, but this is non-exhaustive and with limited possibilities of absolute quantification (Figure 7). Compared to SPME, using adsorbent filled volatile traps in a dynamic headspace is an exhaustive volatile collection method ( Figure 7) and its sensitivity and quantifiability is better compared to SPME ( Tholl, 2020; Ouyang and Pawliszyn, 2008). In addition, the vastly more tedious collection procedures of dynamic headspace sampling substantially reduce the number of biological replicates that can be handled ( Tholl, 2020; Prosen and Zupančič-Kralj, 1999).

Figure 7

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The limitations of these methods point out the importance of designing the extraction protocols carefully including relevant blank samples and using internal reference compounds. Furthermore, the choice of method needs to be anchored in the research questions and hypotheses, which influences the design of the experiments and the choice of relevant positive and negative controls to include.

Further studies should quantitatively evaluate emission of behaviorally active volatile terpenoids from maize, Desmodium and e.g. Napier grass under various realistic field (intercrop) conditions. These will be helpful to understand odor release against a background of volatile terpenoids in the cropping system, and how insects may navigate in these odor spaces. Other factors that deserve further study include for instance the root–root and root–microbiome mediated interactions, as well as possible plant–plant volatile communication. Recent studies have indicated their importance in shaping the above-ground chemical defenses (Chen et al., 2019; Tao et al., 2017; Erb et al., 2015). Since Desmodium and maize are planted in close proximity to each other, these root–root connections may further shape the plant–herbivore interactions under field conditions.

Data associated with volatile analysis and behavioral bioassays are available on figshare at https://doi.org/10.6084/m9.figshare.19297730 and GC–MS raw data is available at https://doi.org/10.6084/m9.figshare.25592544.

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Author details

1. Anna L Erdei

   1. Department of Plant Protection Biology, Swedish University of Agricultural Sciences, Alnarp, Sweden
   2. Department of Chemical Ecology, HUN-REN Centre for Agricultural Research Plant Protection Institute, Budapest, Hungary

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing

   Contributed equally with

   Aneth B David

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7819-1287

2. Aneth B David

   1. Department of Plant Protection Biology, Swedish University of Agricultural Sciences, Alnarp, Sweden
   2. Department of Molecular Biology and Biotechnology, University of Dar-es-Salaam (UDSM), Salaam, United Republic of Tanzania

   Contribution

   Conceptualization, Investigation, Visualization, Methodology, Writing - review and editing

   Contributed equally with

   Anna L Erdei

   Competing interests

   No competing interests declared

3. Eleni C Savvidou

   1. Department of Plant Protection Biology, Swedish University of Agricultural Sciences, Alnarp, Sweden
   2. Department of Agriculture Crop Production and Rural Environment, University of Thessaly, Volos, Greece

   Contribution

   Investigation, Methodology

   Competing interests

   No competing interests declared

4. Vaida Džemedžionaitė

   Department of Plant Protection Biology, Swedish University of Agricultural Sciences, Alnarp, Sweden

   Contribution

   Validation, Investigation, Methodology

   Competing interests

   No competing interests declared

5. Advaith Chakravarthy

   Department of Plant Protection Biology, Swedish University of Agricultural Sciences, Alnarp, Sweden

   Contribution

   Investigation, Methodology

   Competing interests

   No competing interests declared

6. Béla P Molnár

   Department of Chemical Ecology, HUN-REN Centre for Agricultural Research Plant Protection Institute, Budapest, Hungary

   Contribution

   Conceptualization, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3192-0868

7. Teun Dekker

   Department of Plant Protection Biology, Swedish University of Agricultural Sciences, Alnarp, Sweden

   Contribution

   Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing - original draft, Writing - review and editing

   For correspondence

   teun.dekker@slu.se

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5395-6602

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