Chemical Senses Advance Access originally published online on September 19, 2006
Chemical Senses 2006 31(9):821-834; doi:10.1093/chemse/bjl025
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Glomerular Targets of Heliothis subflexa Male Olfactory Receptor Neurons Housed within Long Trichoid Sensilla
1 Department of Entomology, Chemical Ecology Laboratory, Pennsylvania State University, University Park, PA 16802 USA 2 Department of Biology, University of Utah, Salt Lake City, UT 84112 USA
Correspondence to be sent to: Thomas C. Baker, Department of Entomology, Chemical Ecology Lab, Pennsylvania State University, University Park, PA 16802 USA. e-mail: tcb10{at}psu.edu
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Abstract |
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We used single-sensillum recordings to characterize male Heliothis subflexa antennal olfactory receptor neuron physiology in response to compounds related to their sex pheromone. The recordings were then followed by cobalt staining in order to trace the neurons' axons to their glomerular destinations in the antennal lobe. Receptor neurons responding to the major pheromone component, (Z)-11-hexadecenal, in the first type of sensillum, type-A, projected axons to the cumulus of the macroglomerular complex (MGC). In approximately 40% of the type-A sensilla, a colocalized receptor neuron was stained that projected consistently to the posterior complex 1 (PCx1), a specific glomerulus in an 8-glomerulus complex that we call the Posterior Complex (PCx). We found that receptor neurons residing in type-B sensilla and responding to a secondary pheromone component, (Z)-9-hexadecenal, send their axons to the dorsal medial glomerulus of the MGC. As in the type-A sensilla, we found a cocompartmentalized neuron within type-B sensilla that sends its axon to a different glomerulus of the PCx4. One neuron in type-C sensilla tuned to a third pheromone component, (Z)-11-hexadecenol, and a colocalized neuron responding to (Z)-11-hexadecenyl acetate projected their axons to the anteromedial and ventromedial glomeruli of the MGC, respectively.
Key words: cobalt staining, glomerulus, macroglomerular complex, olfactory receptor neuron, posterior complex, single-sensillum recording
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Introduction |
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Heliothis subflexa (Lepidoptera, Noctuidae) larvae are known to feed on plants in the genus Physalis, such as ground cherry. In Mexico, H. subflexa larval feeding on tomatillos can be so extensive that it is now considered to be a pest of this agricultural crop (Mitter et al. 1993
). Even though they are not yet serious agricultural pests in the United States, the successful hybridization of H. subflexa with the well-known pest species, Heliothis virescens, has interested researchers as a potential control technique for H. virescens because hybrid male offspring are sterile, as are backcross males (Laster 1972; Sheck and Gould 1995, 1996; Teal and Oostendorp 1995).
Morphological and molecular phylogenies of heliothine moths are in agreement that these 2 species are closely related (Mitter et al. 1993; Cho et al. 1995; Fang et al. 1997). They are sympatric in North America and share the same female-emitted major pheromone component, (Z)-11-hexadecenal (Z11-16:Ald) (Roelofs et al. 1974; Tumlinson et al. 1975; Klun et al. 1979, 1980, 1982). However, interspecific attraction of males and cross-mating do not occur in nature due to the presence of different minor pheromone components in the blends of the 2 species. Heliothis subflexa females release (Z)-9-hexadecenal (Z9-16:Ald) and (Z)-11-hexadecenol (Z11-16:OH) (Teal et al. 1981; Heath et al. 1991) as minor components that significantly increase attraction of H. subflexa males and are therefore part of this species' sex pheromone blend (Heath et al. 1990; Vickers 2002). Hexadecenal (16:Ald), (Z)-11-hexadecenyl acetate (Z11-16:Ac), (Z)-9-hexadecenyl acetate (Z9-16:Ac), (Z)-7-hexadecenyl acetate (Z7-16:Ac), and (Z)-9-hexadecenol (Z9-16:OH) have been identified as other constituents in the female pheromone gland (Teal et al. 1981; Klun et al. 1982; Heath et al. 1991), but as yet these compounds have not been shown to play a pheromonal role.
Another sympatric heliothine pest, Helicoverpa zea, like H. subflexa, emits Z11-16:Ald and Z9-16:Ald as essential pheromone components for male attraction (Pope et al. 1984; Vetter and Baker 1984). However, the H. subflexa blend will not attract H. zea males because the Z11-16:OH present in this blend has been shown to be antagonistic to H. zea attraction when added to its pheromone blend (Shaver et al. 1982; Quero and Baker 1999).
Much work has now been performed on the sex pheromone olfactory systems of these 3 North American heliothine moth species. These studies include examination of the electrophysiological response profiles of the olfactory receptor neurons (ORNs) (Almaas et al. 1991; Berg et al. 1995; Cossé et al. 1998; Baker et al. 2004; Lee et al. 2006) as well as neuroanatomical studies of the central nervous system pathways that integrate and relay the inputs from the ORNs (Christensen et al. 1991, 1995; Vickers et al. 1998; Vickers and Christensen 2003). Neuroanatomical studies of the ORN pathways into the antennal lobe, the first integrative center of the moth olfactory system, also now have begun to delineate exactly how ORN pheromone component–specific information converges onto specific glomeruli (knots of neuropil within which synaptic contacts are made between ORNs and central neurons) as well as how pheromone blend quality information is conveyed to higher centers by central olfactory projection neurons.
Such studies have been illuminating and have for instance shown how the pheromone component–tuned ORNs of H. virescens (Berg et al. 1998), Helicoverpa assulta (Berg et al. 2005), and H. zea (Lee et al. 2006) project their axons to component-specific homologous glomeruli in these species' macroglomerular complexes (MGCs). However, similar information about the ORNs of H. subflexa has been lacking. We therefore sought to clarify the primary ORN pathways to the MGC for H. subflexa by using the ORN cobalt-staining technique that has proven to be so informative for other noctuid moth species (Hansson et al. 1992; Hansson 1995; Ochieng et al. 1995; Todd et al. 1995; Berg et al. 1998, 2005; Lee et al. 2006).
We report here that the glomeruli in which male H. subflexa component-specific ORNs arborize correspond well to the glomerular arborization locations of similarly tuned projection interneurons exiting the H. subflexa antennal lobe to higher centers (Vickers and Christensen 2003). In addition, we found that the ORNs (Lee et al. 2006) that are cocompartmentalized with ORNs tuned to Z11-16:Ald or to Z9-16:Ald project their axons to a cluster of glomeruli located just posterior to the MGC, which we call the Posterior Complex (PCx) (Lee et al. 2006). The ORNs colocalized with Z11-16:Ald–sensitive ORNs always project to the same glomerulus, the PCx1, in the PCx, and the ORNs cocompartmentalized with Z9-16:Ald–sensitive ORNs always project to a different glomerulus in this complex, the PCx4.
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Materials and methods |
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Insects
A colony of H. subflexa was maintained at the University of Utah as described by Vickers (2002). Pupae were separated according to sex and shipped overnight to the Baker laboratory. Emerging adults were segregated into separate cages daily to obtain individuals of known age. Adults were maintained on a 16:8 light:dark photoperiod at 25 °C, 40–50% relative humidity.
Chemical stimuli
Five heliothine moth sex pheromone compounds were utilized to identify sensillar types according to Baker et al. (2004). The synthetic compounds, Z11-16:Ald, Z9-16:Ald, (Z)-9-tetradecenal (Z9-14:Ald), Z11-16:Ac, and Z11-16:OH were purchased from Bedoukian Research Inc. (Danbury, CT; purities >98% verified by gas chromatography). Serial dilutions were made in high performance liquid chromatography-grade hexane to 0.1 µg/µl and 1 µg/µl and stored at –20 °C. To prepare stimulus cartridges for each compound, 10 µl of each diluted solution was loaded onto a 0.7 x 2.5–cm filter paper strip, the solvent allowed to evaporate, and then the paper strip was placed in a glass Pasteur pipette (2.5 ml). For the mixture of Z11-Ald and (±)-linalool, 10 µg (±)-linalool (97%; Aldrich Chemical Company, Milwaukee, WI) in 10 µl hexane was added to a filter paper onto which 10 µg Z11-16:Ald had already been loaded, and the solvent was allowed to evaporate.
New cartridges were prepared every 5 days and stored at –20 °C when not in use after being sealed with aluminum foil.
Single-sensillum recording
The cut-sensillum technique (Kaissling 1974; Van der Pers and Den Otter 1978; Cossé et al. 1998) was used to record the activities of ORNs housed within individual antennal sensilla. A male H. subflexa was restrained inside a disposable pipette tip; the narrow end of the tip was cut to enlarge it to allow the male's head to pass through. The head and antennae were immobilized in the proper orientation using dental wax. A Ag/AgCl wire was inserted into the abdomen to serve as a ground and was then secured with an alligator clip attached in turn to the end of one of the micromanipulators. By maneuvering this micromanipulator, the tip of one of the many sexually dimorphic long trichoid sensilla was placed on the edge of a vertically positioned metal knife fashioned by sharpening an insect pin. The sensillum tip was cut off using a horizontally positioned glass knife held in place by a second micromanipulator. The cut end of the sensillum was then contacted first by a saline-filled glass micropipette and a connection established in order to determine the type of sensillum that was contacted, according to the differential responses of the ORNs to the array of pheromone odorants. The AC signals from the ORNs were amplified using a high-impedance amplifier (DAM50, World Precision Instruments, Sarasota, FL), and the action potentials were digitally recorded directly onto a PC by means of data processing software (Syntech Autospike 3.2; Hilversum, The Netherlands).
Volatile stimuli (50-ms pulses at 40 ml/s flow rate) were delivered into a continuous charcoal-filtered humidified airstream (
15 ml/s) by means of a stimulus flow-controller device (SFC-2; Syntech). The odor-laden air pulse passed through a 14-cm-long glass tube (8 mm inner diameter) whose outlet was positioned 2 cm from the antenna. Linear flow speed through this continuous airstream was 0.3 m/s.
Staining of ORNs with cobalt-lysine
We used a cobalt-lysine–staining technique, with a minor modification, following the protocol of Hansson et al. (1995), Ochieng et al. (1995), and Todd et al. (1995) in order to try to find the glomerular arborization destinations of the axons of each electrophysiologically identified ORN. A glass electrode filled with 0.5 M cobalt-lysine solution replaced the saline recording electrode after the ORNs residing in the sensillum were characterized using the saline electrode to discern their response specificity. The cobalt-lysine electrode then was used to contact the sensillum for 10 min, during which time one of the pheromone-related compounds to which one of the ORNs was tuned was puffed at a frequency of 0.7 Hz. Unlike in other noctuid moth species, the uptake of cobalt by heliothine moth ORNs is either not activity-dependent or occurs as a result of the ORNs' spontaneous activities alone (Lee et al. 2006). Nevertheless, we wanted to try to increase the chances for successful cobalt uptake by inducing the ORNs to fire in response to repetitive odorant stimulus puffs (Hansson et al. 1992; Todd et al. 1995). For these repetitive stimulations to type-A sensilla, 10 µg Z11-16:Ald or a mixture of 10 µg Z11-16:Ald plus 10 µg (±)-linalool was used. For type-B sensilla, only 10 µg Z9-16:Ald was used, and for type-C sensilla, 10 µg Z11-16:Ac, 10 µg Z11-16:OH, or blank cartridges were used.
The male was then removed from the electrophysiology rig and refrigerated for 2 days at 4 °C to allow sufficient time for the cobalt to migrate along the axons to the antennal lobe. The brain was then dissected from the head in saline (150 mM NaCl, 3 mM KCl, 3 mM CaCl2, 10 mM N-Tris (hydroxymethyl)methyl-2-aminoethanesulfonic acid, and pH 7.0) and treated with ammonium sulfide solution for the cobalt precipitation step. Brains were washed 4 times for 5 min each with saline that included 2% sucrose. After fixation, any cobalt-stained ORNs' axons were intensified with silver (Bacon and Altman 1977). Brains were dehydrated through a graded series of ethanol, cleared with methyl salicylate, and viewed as whole mounts under a light microscope. All brains were stored at 4 °C in 70% alcohol until being used for the next procedure.
Brains were preliminarily counterstained for differentially enhancing cell bodies and glomeruli using modified Lee's methylene blue-basic fuchsin solution (Bennett et al. 1976; Lee et al. 2006). Our modified solution used 0.5% methylene blue + 0.5% azure II in 1% borate: 0.5% basic fuchsin in 95% ethanol: 100% ethanol = 1:2:1—or 0.15% methylene blue, 0.15% azure II, and 0.3% basic fuchsin in 70% alcohol for 10 min. The stained whole brains were washed with 70% alcohol and dehydrated sequentially with 95%, pure ethanol (twice), and twice with propylene oxide. The infiltration procedure was then performed using a mixture of propylene oxide and Durcupan resin (Sigma-Aldrich, St Louis, MO). The propylene oxide was allowed to evaporate at room temperature (RT) overnight, and then the brains were embedded in pure Durcupan resin. Most brains were oriented for frontal sections, but a few were prepared specifically for horizontal sections. After curing at 60 °C for 2 days, brains were sectioned at a thickness of 10 µm.
Immunocytochemistry
For better interpretation of 3-dimensional (3D) glomerular topography, confocal microscopic images were collected after immunocytochemical stainings of glomeruli and then reconstructed to a 3D image (Figure 2D). We used a monoclonal anti-synaptotagmin of Drosophila from mouse as a primary antibody and goat anti-mouse immunoglobulin G antibody conjugated with Alexa Fluo555 (Molecular Probe, Eugene, OR) as secondary antibody. The anti-synaptotagmin antibody developed by Dr Kai Zinn was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the National Institute of Child Health and Human Development and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA. Brains were dissected in 4% paraformaldehyde in phosphate-buffered saline (PBS) with 0.2% Triton X-100 (PBST) and washed with PBST 4 times for 30 min at RT. After washing, brains were preincubated in 5% normal goat serum (NGS) in 0.5% PBST overnight at 4 °C and then incubated in primary antibody (1:100 in 2% NGS PBST) for 3 days at 4 °C. Brains were washed for 6 h at RT and then washed with PBST overnight at 4 °C and then incubated in secondary antibody (1:100 in 2% NGS PBST) for 3 days at 4 °C, always in darkness from this procedure. After washing with PBST for 6 h at RT and then washing overnight at 4 °C and with PBS 3 times for 1 h each at RT, brains were dehydrated through a graded series of ethanol and cleared with methyl salicylate. They were then observed using a 1 or 2 µm thickness of optical sections with the laser scanning confocal microscope (Olympus Fluoview 300, Olympus America Inc., Melville, NY) housed in the Huck Institutes for the Life Sciences at Pennsylvania State University. Alexa Fluo555 was excited with a green HeNe laser (543 nm) and images collected through a 565 long-pass filter.
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Image processing and figure preparation
Histological sections of cobalt-stained brains were captured with Spot RT II digital camera (Diagnostic Instrument, Sterling Heights, MI) attached to a compound microscope (Olympus BX50). The color, brightness, and contrast levels of digitally captured images were adjusted using Adobe Photoshop 7.0 (Adobe Systems, San Jose, CA) for better contrast. Schematic diagrams of Figure 2A,B,C were digitally drawn using Canvas 9 (ACD Systems, Miami, FL). Captured images of confocal microscopy were reconstructed (Figure 2D) using software developed by JC Fiala and KM Harris (downloaded from www.synapses.bu.edu).
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Results |
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Our histological examinations of the H. subflexa antennal lobe (Figure 1) revealed an MGC the same as described by Vickers and Christensen (2003)
, comprised of 4 glomeruli called the cumulus (Cu), the dorsomedial (DM), the anteromedial (AM), and the ventromedial (VM) (Figures 1 and 2). There were also numerous ordinary glomeruli clearly set apart from the 4 MGC glomeruli. In addition, we also found a distinct cluster of 8 glomeruli just posterior to the MGC in every male that we examined, and we named this the Posterior Complex (PCx) (Figures 1 and 2). Many of the PCx glomeruli lie in the most posterior area of this most dorsal portion of the antennal lobe (Figure 1). The PCx is also clearly separated from the large cluster of ordinary glomeruli (Figures 1 and 2). A large bundle of fibers projects out of the PCx, the main trunk of which is a confluence of the bundles from the various levels all oriented toward the center of the antennal lobe (Figures 1B and 2C). The fiber bundle from the PCx eventually merges with the bundle from the MGC (Figure 2C). The H. subflexa PCx appears similar to the PCx described for the first time in H. zea (Lee et al. 2006), except that the H. subflexa PCx has one more glomerulus than does H. zea. The numbers we assigned to the PCx glomeruli were determined as for H. zea, according to the order of their dorsal to ventral, anterior to posterior, and medial to lateral positions, respectively, after examining serial sections from 12 males (16 antennal lobes). The PCx morphology and topography were confirmed by the 3D reconstruction images (Figure 2D) obtained from serial 1- or 2-µm-thick optical sections from 8 antennal lobes (ALs) using laser scanning confocal microscopy. These reconstructions could be performed without the arbitrary reorientation of physical microtome sections as was required for reconstructing the antennal lobes in our cobalt-staining studies.
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Type-A sensilla
We attempted to stain type-A sensilla, as determined by their electrophysiological profiles in which ORN action potentials occurred only in response to the major pheromone component, Z11-16:Ald (Figure 3A) (Baker et al. 2004). Among the 72 such sensilla that were located, 28 produced successfully stained preparations showing arborizations in the antennal lobe. Out of these 28 preparations, 17 (60.7%) exhibited a single axonal stain projecting exclusively to the largest of the MGC glomeruli, the Cu (Figure 1, Table 1).
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The other 11 preparations (39.3%) exhibited 2 or more stained ORNs (Table 1). Without exception, one of these axons projected to the Cu and a second axon projected to the most dorsal of the PCx glomeruli, which we have labeled "PCx1" (Figures 3 and 4).
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An additional 2 sensilla out of those exhibiting projections to the Cu and PCx1 showed 2 or 3 extra axons projecting into ordinary glomeruli (Figure 4B,C). Also, an additional preparation exhibiting ORNs projecting to the Cu and PCx1 had its third axon projecting to the PCx4 of the PCx (Table 1).
Stimulation with repetitive puffs of either 10 µg Z11-16:Ald or a blend of 10 µg Z11-16:Ald and 10 µg linalool during contact with the cobalt-filled electrode did not significantly influence the selective uptake of cobalt stain by the ORNs in these type-A sensilla. Adding linalool to Z11-16:Ald was tried here to see if the heightened firing of the large-spiking ORN caused by the addition of this plant volatile (Ochieng and Baker 2002) would enhance cobalt uptake. However, the ORNs of heliothine moths appear to take up stain spontaneously, without the need for odor stimuli to be presented, perhaps due to spontaneous background firing (Berg et al. 1998; Lee et al. 2006). Therefore again, approximately 60% of the stained type-A sensillar preparations, regardless of stimulus, ended up with a single-stained ORN projecting to the Cu with the remaining 40% exhibiting 2 stained ORNs with the second ORN projecting to PCx1.
The second ORN in type-A sensilla that is colocalized with the ORN responding to Z11-16:Ald does not reliably respond to any odorant, either taken from a panel of over 60 general and plant odorants or from an array of heliothine moth pheromone-related compounds (Lee 2006).
Type-B sensilla
Type-B sensilla were identified by their characteristic ORN responsiveness to Z9-16:Ald, an important minor pheromone component of this species, plus a low degree of cross-responsiveness to Z9-14:Ald (Figure 5A) (Baker et al. 2004). Spontaneous activities in clean air showed spikes having 2 distinct waveforms. The larger spiking of these ORNs showed a more symmetrically bipolar waveform than the smaller spiking ORN (Figure 5A). Responses during stimulation showed that it is this larger spiking ORN that is sensitive to Z9-16:Ald and Z9-14:Ald stimulation (Figure 5A) (Baker et al. 2004). The second ORN, the one having the more asymmetric, unipolar action potential, showed no consistent activity in response to any of our test compounds (Figure 5A) (Baker et al. 2004; Lee 2006).
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We obtained 28 successful stains from ORNs in type-B sensilla in 58 preparations. The majority of these, 23 out of 28 (82.1%), resulted in 2 or more stained ORNs. In all cases, the 2 axons arborized in 2 specific glomeruli, the DM and the PCx4 (Table 1, Figures 2 and 5B). Eighteen of the 23 exhibited only 2 stained axons projecting to these 2 locations. The remaining 5 of these 23 exhibited an additional third or even a fourth stained ORN projecting either to an ordinary glomerulus (Figure 6A) or to the Cu (Table 1).
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The ORNs from the remaining 5 out of 28 stained type-B sensillar preparations (
18%) showed only a single stained ORN (Table 1). One such ORN had its single axon projecting only to the PCx4, and the ORNs from the other 4 sensilla stained only the DM. One of the DM-targeting sensilla had one or 2 extra axon(s) that were stained in the antennal nerve, but the stains faded out before reaching a target glomerulus (data not shown). Type-C sensilla
The physiology of H. subflexa type-C sensilla is characterized by differential activity profiles of 2 cocompartmentalized, pheromone odorant–sensitive neurons (Baker et al. 2004). Stimulation with the minor pheromone component Z11-16:OH elicits activity from an ORN exhibiting smaller action potentials, whereas larger action potentials are generated from the second ORN upon stimulation with either Z11-16:Ac or Z9-14:Ald (Figure 7A) (Baker et al. 2004). We obtained 19 successfully stained type-C sensillar preparations out of 23 attempts. Seventeen of these (89.5%) exhibited 2 stained axons projecting to the AM and VM glomeruli, the 2 smaller glomeruli of the MGC (Table 1; Figures 2, 7B,C, and 8A). The remaining 2 preparations resulted in a single stained axon projecting to the VM glomerulus. Three of these 17 exhibited additional stained axons targeting 1 or 2 ordinary glomeruli (Table 1; Figure 8B,C). There appeared to be no differential activity-dependent ORN cobalt uptake resulting from stimulation with either 10 µg Z11-16:Ac or 10 µg Z11-16:OH.
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Discussion |
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We have now described the pathways of axons from H. subflexa pheromone component–tuned ORNs residing in trichoid sensilla to their glomerular destinations in the antennal lobe.
When we couple our findings with those of Vickers and Christensen (2003), who described pheromone component–specific projection interneuron arborizations into each of the 4 H. subflexa MGC compartments, we conclude that there is a correspondence between pheromone odorant–specific input to, and output from, the different MGC compartments. These relationships for H. subflexa are depicted in Figure 9A.
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Although large numbers of our single-sensillar preparations exhibited 2 stained ORNs each projecting to a different glomerulus, the ORNs reached the same glomeruli with high fidelity from preparation to preparation. Pairs of glomeruli exhibiting stained axons were nearly perfectly correlated with their origin from type-A, type-B, or type-C sensilla. Preparations exhibiting only single stains were similarly highly correlated with their sensillar-type origin.
Sixty percent of the type-A sensillar preparations showed a single stained ORN projecting its axon without exception to the Cu. Another 40% of type-A sensillar preparations had 2 stained ORNs, with all these preparations exhibiting an ORN arborization in the Cu, plus a second stained axon targeting the PCx1 glomerulus of the PCx. No pheromone component–sensitive ORN in moths has ever been found to arborize anywhere but in the MGC, and we conclude that it is the Z11-16:Ald–tuned ORNs from these preparations that target the Cu.
This interpretation is consonant with what has been shown for the projections of Z11-16:Ald–tuned ORNs in H. virescens and H. zea (Berg et al. 1998; Lee et al. 2006). In addition, projection interneuron–staining patterns in the H. subflexa antennal lobe (Vickers and Christensen 2003) show definitively that the Cu is where Z11-16:Ald–tuned projection interneurons arborize. Given the linear pathways of ORN to glomerulus to projection interneuron that have been found in H. virescens and H. zea (Figure 9; Berg et al. 1998; Vickers et al. 1998; Lee et al. 2006), this again indicates that the Z11-16:Ald–tuned ORNs arborize in the Cu of the H. subflexa MGC.
The colocalized ORNs from type-A sensilla are therefore the ones in our cobalt stains that target the PCx1 glomerulus in the newly described PCx. The targeting of the PCx by the cocompartmentalized ORNs in type-A sensilla was definitively shown in H. zea (Lee et al. 2006). One of the most striking results from the current study is the finding that the ORNs in H. subflexa that are cocompartmentalized with pheromone component–sensitive ORNs in either type-A or type-B sensilla project reliably into their own specific glomeruli in a cluster of glomeruli having a morphology similar to the PCx of H. zea (Figure 9A,C; Lee et al. 2006). Moreover, as in H. zea, the colocalized, very small–spiking ORNs in type-A sensilla in H. subflexa (Figure 3A) always project into the most dorsal and medial of the 8 PCx glomeruli in H. subflexa, which we have labeled, as in H. zea, the PCx1 (Lee et al. 2006). Again as in H. zea, the PCx1-targeting neurons in H. subflexa were not found originating from any sensilla other than type-A; we conclude that these ORNs are not arbitrarily cocompartmentalized with the larger spiking ORNs in type-A sensilla.
Although the colocalized ORNs in type-A sensilla did not respond to any general odorants or pheromone-related analogs from a panel of more than 70 odorants (Lee 2006), we did see activity from this small-spiking ORN whenever there was firing from the large-spiking ORN in response to Z11-16:Ald or a structurally similar analog (Lee 2006).
We also tried stimulation with the C16 acids corresponding to the enzymatic degradation products of heliothine aldehyde pheromone components (Prestwich et al. 1989; Tasayco and Prestwich 1990a, 1990b) because in other noctuid species such as Trichoplusia ni and Agrotis segetum, the ORNs that are cocompartmentalized with major component-sensitive ORNs (Hansson et al. 1992; Todd et al. 1992, 1995) are highly responsive to the corresponding degradation products (alcohols) of the acetate major component (Prestwich et al. 1989). None of this large array of odorants evoked responses from the colocalized ORN; the correspondence of firing between the large-spiking and small-spiking ORNs was clear, however (Lee 2006). The anatomical conformity of the target glomeruli of the colocalized ORNs in type-A sensilla of these North American heliothine moths implies that if there is any odorant-specific processing performed within the PCx1, it may be similar across heliothine species. We have pointed out previously (Lee et al. 2006) the morphological similarities of the PCx we described in H. zea to glomeruli that have previously been labeled "ordinary" in H. virescens and H. assulta; all receiving input from cocompartmentalized ORNs having unknown tuning properties (Berg et al. 1998, 2005). In Figure 9, we have summarized the current status of ORN input projections to the MGC and PCx for these 4 species of heliothine moth.
Although there were 2 ORNs consistently stained in type-B sensilla with one projecting to the DM and the other to the PCx4, we conclude that the DM glomerulus receives input from Z9-16:Ald–tuned ORNs because in moths no pheromone component–sensitive ORNs, and only rarely projection interneurons, have been found to arborize outside of the MGC (Hansson and Christensen 1999; Vickers and Christensen 2003). This assignment is also consonant with the arborization of Z9-16:Ald–responsive projection interneurons that always target the DM glomerulus (Vickers and Christensen 2003). In addition, calcium images we have started to accumulate for H. subflexa show that the focal point of calcium activity in response to Z9-16:Ald antennal stimulation is in the DM glomerulus (MA Carlsson, BS Hansson, S-G Lee, TC Baker, unpublished data). There is thus a linear arrangement through the DM glomerulus of Z9-16:Ald–related input from ORNs and Z9-16:Ald–related output from projection interneurons. The PCx4 glomerulus of H. subflexa is therefore functionally and morphologically engaged in receiving information from the ORN that is colocalized in type-B sensilla with the ORN responsive to Z9-16:Ald/Z9-14:Ald. This companion ORN did not respond consistently to any of the approximately 70 test compounds with which it was challenged (Lee 2006). This type-B colocalized ORN produces spontaneous asymmetric action potentials, and the identical waveform of this type of ORN across all the type-B sensilla we recorded from suggests that it has a distinctive signal-generating system that is different from that of the Z9-16:Ald/Z9-14:Ald–responsive ORN in terms of its ion channel characteristics, especially during the outward current of hyperpolarization.
In H. virescens, one of 2 preparations of Berg et al. (1998) for the staining of type-B sensilla containing Z9-14:Ald–responding ORNs showed one axon projecting to an "ordinary glomerulus" located ventral to the MGC, in addition to an ORN projecting to the DM compartment of the MGC. The DM glomerulus was concluded to be the location receiving Z9-14:Ald–related input based, among other factors, on the DM arborization location of Z9-14:Ald–tuned projection interneurons (Vickers et al. 1998). Subsequently, a calcium imaging study confirmed the H. virescens DM glomerulus as the MGC glomerulus receiving ORN excitation related to Z9-14:Ald (Galizia et al. 2000). The ordinary glomerulus that is depicted in Berg et al. (1998) displays a similar, topographic, ventral proximity to the PCx4 glomerulus of H. subflexa (Figure 9A,B). No mention was made of the posterior depth of this glomerulus in relation to the DM glomerulus (Berg et al. 1998). The colocalized ORN that targets it has an undefined tuning profile because no compound was found that excited it (Berg et al. 1998).
The projections of the ORNs we found in type-C sensilla exhibit highly consistent patterns of labeling into the AM and VM, the 2 smaller glomeruli in the MGC. We were not able to directly identify which of these 2 ORNs target the AM or the VM glomerulus due to the nonselectivity of cobalt dye uptake during cobalt application. However, the arborization of a Z11-16:OH-responsive projection interneuron in the AM glomerulus (Vickers and Christensen 2003) and the increasingly well-established linearity of pheromone component olfactory pathways in heliothine moths (Berg et al. 1998; Vickers et al. 1998; Vickers and Christensen 2003) suggest that the AM glomerulus is the primary receiving center for action potentials from Z11-16:OH–tuned ORNs. By default, the VM glomerulus should be the target glomerulus for the ORN that responds to Z11-16:Ac and Z9-14:Ald. The first calcium images from H. subflexa now also support this assignment (MA Carlsson, BS Hansson, S-G Lee, TC Baker, unpublished data).
The frequency with which ordinary glomeruli received stains from third or fourth ORNs residing in H. subflexa types A, B, and C long trichoids was higher in H. subflexa than in other heliothines. This would imply that in this species, there may be more ORNs tuned to general odorants that are cocompartmentalized in these sensilla with ORNs tuned to pheromone components than there are in either H. zea (Lee et al. 2006) or H. virescens (Berg et al. 1998). The ordinary glomerulus staining ORNs arose with similar likelihood from type-A, type-B, and type-C sensilla, and so no one type of sensillum was associated with the presence of these ORNs. The stained arborizations were not in a wide variety of ordinary glomeruli but rather were restricted with high fidelity to but a few specific glomeruli. Arborizations of 7 of the 16 ORNs targeting ordinary glomeruli terminated in an ordinary glomerulus most closely adjacent and medial–posterior to the DM (Figures 4B and 8B). In addition, 2 ORNs from type-A, 2 from a type-B, and 3 from a type-C sensillum targeted a water drop–shaped glomerulus situated medially and several tens of micrometers posterior to the ordinary glomerulus mentioned above (Figures 4C, 6A, and 8C). We also found a type-A and a type-B sensillum each having an ORN projecting to the most anterior ordinary glomerulus in the antennal lobe. These results imply that these 16 ORNs terminating in ordinary glomeruli and originating variously from types A, B, and C trichoid sensilla might show 3 distinctive tuning spectra to odorant ligands.
It is unclear why, in addition to the MGC and ordinary glomeruli, that a third subcluster of glomeruli, the PCx, should exist in these heliothine moth species. Such a grouping implies a separate, broad function of this structure, just as the MGC cluster is related to pheromone compound inputs and the large cluster of ordinary glomeruli is related to all other odorant inputs. We have only performed staining experiments on physiologically identified ORNs in males of these species thus far. We hope to perform similar studies and find sensilla in females that have Z11-16:Ald–responsive ORNs in order to try staining any cocompartmentalized ORNs to determine their target glomeruli in the female antennal lobe. Our preliminary analysis of the female antennal lobes of H. zea and H. subflexa has thus far revealed a structure in both species similar to the male PCxs but appearing slightly different than those of the males. A more thorough morphological sex-specific comparison coupled with physiological and cobalt-staining studies using females should help provide more insight regarding any functional and morphological dimorphisms of the PCx in these species.
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This study was funded by grant #2003-01471 from the US Department of Agriculture NRI program to TCB. We thank to Drs J.C. Fiala and K.M. Harris for image analysis software.
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Accepted 27 July 2006
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