Reversed functional topology in the antennal lobe of the male European corn borer

Moth pheromone communication is a schoolbook example of sexual selection and speciation. Many salient examples illustrate how evolutionary forces mold the female pheromone production as well as the male response to inter- and intraspecific signals. In addition, the moth pheromone system offers unique possibilities for studying the evolution of olfactory processing and its behavioral correlate, as the organization of the corresponding olfactory subcircuitry is relatively simple and behavioral responses to pheromones generally robust. Yet, the proximate mechanisms underlying shifts in pheromone preference are still elusive.

Sex pheromone components are detected by olfactory receptor neurons (ORNs),which, like other ORNs in insects, project into the first olfactory neuropil,the antennal lobe (AL), through the antennal nerve (AN)(Bretschneider, 1924). The AL comprises a number of glomeruli in which synaptic contacts between ORNs,projection neurons (PNs) and local interneurons are made. Male moths have a few enlarged glomeruli, which make up the macroglomerular complex (MGC),situated at the entrance of the AN. These glomeruli are dedicated to receiving information regarding female-produced sex pheromones(Bretschneider, 1924; Koontz and Schneider, 1987). In several moth species, the number of MGC glomeruli equals the number of behaviorally relevant pheromone components, with each ORN type projecting to one MGC glomerulus (Hansson et al.,1992; Ochieng et al.,1995; Todd et al.,1995; Berg et al.,1998). The ORNs project into the AL, where input is relayed onto PNs. Blend-specific PNs, which may be involved in blend recognition, have been found in several noctuid species(Christensen et al., 1989; Christensen et al., 1991; Hansson et al., 1994b; Anton and Hansson, 1994; Anton and Hansson, 1995; Wu et al., 1996).

The European corn borer, Ostrinia nubilalis Hübner(Lepidoptera: Pyralidae) has two sex pheromone components (Z)-11- and(E)-11-tetradecenyl acetate (Z11- and E11-14:OAc) and an interspecific behavioral antagonist (Z)-9-tetradecenyl acetate(Z9-14:OAc) (Glover et al.,1989). The species occurs in two strains that use opposite pheromone component ratios. Females of the Z-strain produce 97% Z11-14:OAc and 3% E11-14:OAc whereas E-strain females produce 99% E11-14:OAc and 1%Z11-14:OAc (Anglade et al.,1984; Klun and Robinson,1971; Klun et al.,1973). Consequently, the strains do not freely interbreed in sympatry (Cardé et al.,1978; Malausa et al.,2005).

Males of both strains have three types of sensilla trichodea on their antenna (type C, B and A) which each contain one, two or three neurons,respectively, responding to pheromone stimuli(Hansson et al., 1987; Hansson et al., 1994b). Sensillum type A contains three ORNs, the neuron characterized by a large spike amplitude responds to the major component, the neuron having a intermediate spike amplitude responds to the minor pheromone component; the third ORN produces small amplitude spikes in response to the behavioral antagonist. Sensillum type B houses two ORNs, one large spiking ORN responding to the major component, and a small-spiking ORN responding to the minor pheromone component. Sensillum type C contains one ORN responding either to the major pheromone component or to the behavioral antagonist(Hansson et al., 1987; Hallberg et al., 1994; Cossé et al., 1995). Genetic studies of the O. nubilalis pheromone communication system have indicated that female sex pheromone production and the male sensory setup are primarily controlled by a single autosomal factor(Hansson et al., 1987; Roelofs et al., 1987; Löfstedt et al., 1989; Roelofs and Glover, 1991).

Previous studies described the AL of O. nubilalis(Anton et al., 1997). However,the techniques at that time did not allow a clear resolution of the intricate structure of the MGC in this species. Here, we resolve in much more detail the neuroanatomy of the AL of the male and female European corn borer. Also, we morphologically and physiologically characterize pheromone sensitive ORNs and PNs. The results demonstrate that the single-gene-mediated shift is located upstream of the antennal lobes, i.e. at the level of the ORN.

The Z- and E-strains of European corn borer, Ostrinia nubilalisHübner (Lepidoptera: Pyralidae) were reared in the laboratory. The Z-strain culture originated from a 2004 adult collection from corn fields in Kéty, Tolna, Hungary. The E-strain was kindly provided by Dr Wendell Roelofs and originated from a collection of larvae, pupae and adults from corn stubble in New York state, USA. The cultures were maintained on a semi-artificial diet (Mani et al.,1978) at 25°C, RH 70% under a 18 h:6 h L:D photoperiod. The genetic purity of the cultures was monitored by gas chromatographic analysis(GC) of pheromone production in females.

Sexes were separated as pupae and kept in separate plastic boxes to avoid exposing adult males to female sex pheromone. Adults were fed 5% honey water solution throughout their adult lives. Moths of 1–4 days old were used.

Previous studies using antennal staining have demonstrated that under pulsed odor stimulation there is a preferential staining of the neuron sensitive to the stimulus (Hansson et al.,1992; Hansson,1997; Kirschner et al.,2006). Two techniques were used to obtain activity-dependent stains. A single sensillum from the second or third flagellar segment was characterized physiologically, after which a glass electrode stained with 1%neurobiotin (Molecular Probes, Carlsbad, CA, USA) in 0.25% KCl was placed over the sensillum. Alternatively, a small neurobiotin crystal was placed on the second or third flagellar segment between pheromone-sensitive sensilla. The animal was kept for 1 h under continuous pulsed (0.2 s stimulus, 4 s clean air, total flow 8 ml s^–1) stimulation with 100 ng of either Z11- or E11-14Ac. Moist filter paper surrounded the preparation to avoid dehydration and crystallization. Subsequently, the moths were decapitated and the heads were processed as described under `neuroanatomical techniques'. The success rate of specific stainings of single ORNs was low (∼14%). However,in preparations with multiple ORN stains, in every case where the axonal projections could be traced each ORN was found to arborize uniglomerularly.

A male moth was restrained in a plastic pipette tip. The moth was inserted from the wide end of the pipette tip with the head protruding from the tip. The head was immobilized with dental wax (Surgident, Heraeus Kulzer, Inc.,Armonk, NY, USA). The proboscis and one of the antennae were cut off and the scales on the head were removed. Incisions between the eyes were made creating a window through which the antennal lobes were visible. The muscles around the antenna were removed to allow for stable recordings. The moth was placed in an electrophysiological setup and the opened head was superfused with a ringer solution of pH 6.9 containing 8.55 g l^–1 sucrose(Christensen and Hildebrand,1987). The odorants were diluted in redistilled n-hexane and applied on a filter paper disc inside a Pasteur pipette. A 0.5 s stimulation was delivered at 4 ml s^–1 into a charcoal-filtered humidified air stream (0.5 ml min^–1) flowing over the ipsilateral antenna of the moth through an opening 20 cm from the antenna. The odor stimuli were presented at 10 s inter-stimulus intervals. Stimuli included Z11-14:OAc, E11-14:OAc and blends thereof, and Z9-14:OAc at a range of concentrations (1 ng–10 μg). A hexane blank served as control. The purity of the odorants was verified using GC. The recordings followed the procedures described by Christensen and Hildebrand(Christensen and Hildebrand,1987). Glass electrodes were stained with 1 mol l^–1 KCl, with the tip containing 1% neurobiotin. Using a micromanipulator, the recording electrode was inserted into the antennal lobe close to the point of entry of the antennal nerve, where many PN dendrites coalesce from the MGC. Usually, the most successful recordings were obtained with the electrode situated close to the surface. When intracellular contact was established, the ipsilateral antenna was stimulated and the activity of the neuron before, during and after stimulation was observed. The signal was amplified, digitally converted (IDAC-4 USB, Syntech, Kirchzarten, Germany) and visualized using a PC with AutoSpike 3.2 software (Syntech). Recordings of action potentials were stored on the PC and analyses were performed using AutoSpike software. The spikes were counted manually. The response of PNs was expressed as the number of spikes during an 0.25 s period after stimulus onset minus the number of spikes 0.25 s before stimulus onset (which represents the spontaneous activity of the neuron) and expressed as the number of spikes per second. Physiologically characterized neurons were stained with neurobiotin by passing 0.5–1.2 nA of constant depolarizing current through the recording electrode for 10–15 min. Brains were processed as described under `neuroanatomical techniques'.

Heads of decapitated moths were fixed in 4% formaldehyde containing 0.25%Triton X-100 in PBS overnight at 4°C and then dissected, washed 4×10min in 0.25% Triton X-100 in PBS and incubated in PBS (0.25% Triton X-100) with 5% α-synapsin (courtesy of Dr Buchner, University of Würzburg, Germany) antibody and 3% fluorescein Avidin (Invitrogen,Carlsbad, CA, USA) overnight on a rotator at room temperature. The next day,brains were washed 4×10min in PBS (0.25% Triton X-100); incubated with 1% α-mouse (goat) Alexa Fluor 546 (Invitrogen) in PBS (0.25% Triton X-100) for 4days at 4°C. Finally the brains were washed 4×10min in PBS (0.25% Triton X-100) and mounted in Vectashield Hard set (Vector Laboratories, Burlingame, CA, USA).

The mounted brains were examined under a Zeiss LSM 510 confocal microscope(Carl Zeiss, Jena, Germany) equipped with a 40×, 1.4 oil-immersion DIC objective lens. Structures indirectly labeled with fluorescein Avidin and Alexa Fluor 546 and were excited with an argon (488 nm) and a helium–neon laser (543 nm) and their fluorescence was detected after passing through a band pass (505–530 nm) and a long pass (560 nm)filter, respectively. Stacks of 50–200 confocal images were scanned and the images were stored at a size of 1024×1024 pixels. The three-dimensional reconstructions were done with AMIRA (Mercury Computer Systems SAS, Merignac Cedex, France) with 512×512 pixels image stacks. Every second section was reconstructed.

In each optical section, contours of glomeruli were demarcated by hand(i.e. image segmentation). The volumetric measurements were done using AMIRA software.

First, we elucidated the architecture of the primary olfactory neuropil,the antennal lobe (AL). α-Synapsin antibody staining penetrated the tissue well and enabled visualization of the entire O. nubilalis AL(Fig. 1A). Deeper, more posterior in the AL, where α-synapsin penetration and staining was weaker, phalloidin staining was helpful in resolving the glomerular boundaries. Based on our AMIRA reconstructions we estimated the number of glomeruli in Z-strain female and male O. nubilalis AL to be ∼64 and ∼66, respectively (Fig. 1). It should be noted that the precise number of identified glomeruli differed somewhat between preparations as a result of differences in the quality of staining, especially of more posterior glomeruli.

At the point where the AN enters the AL, Z-strain male O. nubilalis have a much larger set of glomeruli, the MGC known to be exclusively involved in sex pheromone processing. Three MGC glomeruli could be distinguished. Two large, highly convoluted and interdigitated glomeruli,variable in shape and dimension(297×10³±36×10³μm³,141×10³±17×10³μm³, N=3; Fig. 1B), one medial and one lateral. The accuracy of the demarcations of these two large glomeruli was corroborated using the stains of specific PNs (see below). A third large disc-shaped glomerulus was located posterior of the two large interdigitated glomeruli(89×10³±3.4×10³ μm³, N=6; Fig. 1C). Inexplicably, the medial glomerulus was always more strongly stained withα-synapsin than the lateral glomerulus(Fig. 1A,D). In females, two enlarged female glomeruli (LFGs) were found at the entrance of the AL. LFG1 had an estimated volume of 64×10³ μm³, which was on average 4.4 times larger than an ordinary glomerulus (average radius of 15 μm³; Fig. 1D). LFG2 had an estimated volume of 53×10³μm³(Fig. 1E,F).

Activity-dependent anterograde stainings of antennal sensilla using neurobiotin were used to establish the pattern of ORN arborizations in the MGC(Fig. 2A,B; Table 1). ORNs exclusively arborized in either the medial or lateral glomerulus. Single ORNs stained under stimulation with Z11-14:OAc arborized in the medial glomerulus, whereas stimulation with E11-14:OAc resulted in staining of the ORNs arborizing in the lateral glomerulus.

Extensive projection neuron (PN) recordings (538 contacts in total; Fig. 3F, Table 1, Fig. 4) were conducted and physiologically well-characterized PNs were stained with neurobiotin tracer. PN arborizations were checked against a background of α-synapsin staining. The stainings demonstrate that without exception Z11-14:OAc-specific PNs arborize in the medial glomerulus (Fig. 3A), whereas E11-14:OAc-specific PNs arborize in the somewhat smaller lateral glomerulus (Fig. 3B). The soma of specific neurons were located in the medial cell cluster. Specificity was evidenced in at least a 10¹–10³-fold difference in sensitivity to Z11- and E11-14:OAc. Neurons responding equally to both E11- and Z11-14:OAc were mostly local interneurons arborizing in most, if not all glomeruli. On rare occasions(two intracellular recordings, three successful stains) we encountered PNs that were more sensitive to a blend of Z11- and E11-14:OAc than either of the components separately (Fig. 3E). These PNs arborized in both the lateral and medial MGC glomeruli, sometimes apparently also in an ordinary glomerulus. The cell bodies of these PNs were located in the lateral cell cluster(Fig. 3E). Few recordings were obtained from PNs responding only to the antagonist, Z9-14:Ac. Only two stainings were obtained of an antagonist specific PN, with dendritic arborizations in the posterior disc-shaped glomerulus only (in yellow in Fig. 1C). However, the quality of the staining and dissection did not allow for reconstruction.

Overview staining with α-synapsin demonstrated that the architecture of the antennal lobes in the E-strain was highly similar to that of the Z-strain. The total volume of the MGC was similar in the two strains(Table 2). Activity-dependent neurobiotin stains of ORNs showed, similar to those from the Z-strain, that ORNs display uniglomerular arborizations, but with E11-14:OAc-sensitive ORNs projecting into the medial glomerulus, and Z11-14:OAc-sensitive neurons to the lateral glomerulus (Fig. 2C,D). The success rate of activity-dependent stains was low. However, the physiological characterization of peripheral input to the AL was totally corroborated by stains of physiologically characterized PNs. We recorded from a total of 1278 PNs. Twenty stainings of PNs responding specifically to one of the two pheromone components yielded a total of five single PN stainings(Table 1, Fig. 4). Specific neurons arborized again exclusively in one MGC glomerulus, with E11-14:OAc-responding PNs sending dendritic branches into the medial glomerulus and Z11-14:OAc-responding PNs arborizing into the lateral glomerulus(Fig. 3C,D). Both ORNs and PNs thus display innervation patterns opposite to those of the Z-strain. The two strains clearly have an identical MGC morphology, but a reversed functional topology. Table 2 summarizes the neuroanatomy and physiology of both strains of O. nubilalis.

Here we report on the neuroanatomy and neurophysiology of the AL of O. nubilalis. We were, for the first time, able to resolve the highly complex, intertwined MGC glomerular structures and to functionally classify the glomeruli this complex consists of.

The neuroanatomy of the MGC of both Z- and E-strain O. nubilalisis indistinguishable. However, our extensive physiological analyses of the MGC in- and output, revealed a reversed physiological specificity. In both strains ORNs responding to the major pheromone component arborized in the large,medial MGC glomerulus, whereas the minor pheromone component-specific ORNs arborized in the smaller, lateral one. A similar morphology but changed physiology was also reported for heliothine species. The MGC of Heliothis virescens and H. subflexa was indistinguishable, but the physiology has in part changed to accommodate for the shift in pheromone blend preference (Vickers and Christensen,2003).

We thus found a reversed functional topology in the MGC between the Z- and E-strain of the European corn borer. To account for the pheromone component representation in the two major MGC glomeruli, a similar change in the specificity and/or wiring of the sensory input must also have occurred. A possible explanation for our observations is a swap of olfactory receptors between ORNs within the same sensillum, while the ORN and PN arborization patterns in the MGC remain unaffected. Unlike the situation in mammals, insect ORs do not directly determine axonal targeting in the deutocerebrum(Dobritsa et al., 2003; Goldman et al., 2005; Endo et al., 2007; Ray et al., 2007). Previous genetic studies on the European corn borer revealed that a single,sex-chromosome linked factor is responsible for the reversed behavioral preference of the males for the two pheromone blends(Löfstedt et al., 1989;Dopman et al., 2004). In addition, an autosomal factor underlies a reversed action potential amplitude of Z11- and E11-14:OAc-responding ORNs between the Z- and E-strain (Hansson et al.,1987; Roelofs et al.,1987). Hybrids of Z- and E-strain O. nubilalis prefer an intermediate blend and show intermediate action potential amplitudes for both pheromone-responding cells. Yet, it is not clear how an OR swap would fit with the observation that the behavioral response is sex-linked, but the spike amplitude autosomal. Other possibilities include, for instance, that, instead of a single gene, a group of tightly-linked genes underlies the reversed antennal lobe physiology in O. nubilalis, which would allow for mechanisms such as rewiring of both ORNs to opposite MGC glomeruli. The matter is further complicated by the fact that Z11- and E11-14:OAc-sensitive ORNs are also found in other, much less frequent physiological subtypes of trichoid sensilla (Hallberg et al.,1994). O. nubilalis apparently expresses the same putative Z11- and E11-14:OAc-sensitive ORs in ORNs derived from different progenitor cells, while axons converge into the same glomerulus. Further research is needed on the cascade of events that determine the OR gene`choice' of ORNs, and ORN axonal targeting to elucidate the mechanisms underlying the reversed physiological specificity of MGC-innervating projection neurons in O. nubilalis.

Previous studies show that O. nubilalis has blend-specific AL neurons that are postulated to be crucial for the discrimination between blends (Anton et al., 1997). We also found PNs that displayed a stronger response to a blend of Z11 and E11 than the added responses to the components separately(Fig. 3E). The cell bodies of these PNs were located in the lateral cell cluster, which matches similar findings in B. mori (Kanzaki et al., 2003), A. segetum(Hansson et al., 1994a), H. zea and H. virescens(Christensen et al., 1989; Christensen et al., 1991)blend-specific PNs. It remains to be seen whether there is indeed a strict correlation between PN neuroanatomy, physiology and soma position. Of particular interest is whether such neurons are essential to the readout of the ratio of pheromone components and of behavioral importance. In Drosophila, such a relationship does not hold(Marin et al., 2002; Wong et al., 2002). How these differences may be relevant to olfactory behavior is thus still unclear.

Female O. nubilalis also possess enlarged glomeruli at the entrance of the antennal lobe, which are homologues to those found in other species and have been called large female glomeruli (LFG) in M. sexta(Roche King et al., 2000; Rossler et al., 1998). In females Heliothis virescens the ORNs tuned to one of the sex pheromone components arborize in the female-specific central large female glomerulus (cLFG) and other glomeruli in the AL(Hillier et al., 2006). By contrast, the LFGs of M. sexta seem to receive innervation from ORNs tuned to host odor volatiles. In electroantennogram (EAG) studies, antennae of females O. nubilalis Z-strain respond to Z11-14:OAc (Z.K.,unpublished observations). In further studies we will study the physiology of the female AL, including the two LFGs.

Our reconstruction of the ALs of O. nubilalis further shows that the total number of glomeruli (♀ ∼64 and ♂ ∼66) closely approximates that found in other Lepidoptera: Mamestra brassicae– 67 ♂, 68 ♀ (Rospars,1983), Manduca sexta – 63(Rospars and Hildebrand,2000), Heliothis virescens – 66 ♂, 62 ♀(Berg et al., 2002), Bombyx mor – ∼60 (Kanzaki et al.,2003), Agrotis ipsilon – 66(Greiner et al., 2004), which implies relative constancy in the number of different ORN types and the number of ORs expressed. The high numerical invariance of moth AL glomeruli, around∼64, in distantly related moth taxa is striking, especially considering the large niche diversification in Lepidoptera. By contrast, Hymenoptera species show a high variance in number of glomeruli even between closely related species or within castes of the same species [e.g. Apis mellifera worker: 166 glomeruli, drone: 103 glomeruli(Arnold et al., 1985); Vespa crabro ∼1000 glomeruli(Hanström, 1928)]. The invariance in the number of glomeruli raises the question of how the olfactory circuitry could accommodate the enormous niche differentiation observed in Lepidoptera. An alternative route for evolution of olfactory preference is evolution of the ORs themselves. Minor changes in amino acid sequences may affect the binding affinity (e.g. Dekker et al., 2006), although strikingly high conservation of physiological response characteristics of ORNs has been reported too(Stensmyr et al., 2003; Ray et al., 2007; McBride, 2007).

We thank Wen-Qi Wu for initial assistance with electrophysiology, and Sylvia Anton, Mikael Carlsson, Wiltrud Daniels, Medhat Sadek, Wendell Roelofs,Gábor Szöcs. We also would like to thank the two anonymous reviewers for their helpful suggestions. This project was supported by grants from the Swedish Research Council (VR) to B.S.H. and from Formas to T.D. It was also strongly supported by the Linnaeus Grant `Insect Chemical Ecology,Ethology and Evolution (ICE³)' and the Hungarian National Science Foundation (OTKA: K 71980).