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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Nature. Author manuscript; available in PMC 2013 July 17.
Published in final edited form as:
PMCID: PMC3590906

Asymmetric neurotransmitter release enables rapid odor lateralization in Drosophila


In Drosophila, most individual olfactory receptor neurons (ORNs) project bilaterally to both sides of the brain1,2. Having bilateral rather than unilateral projections may represent a useful redundancy. However, bilateral ORN projections to the brain should also compromise the ability to lateralize odors. Nevertheless, walking or flying Drosophila reportedly turn toward their more strongly stimulated antenna3-5. Here we show that each ORN spike releases ~40% more neurotransmitter from the axon branch ipsilateral to the soma, as compared to the contralateral branch. As a result, when an odor activates the antennae asymmetrically, ipsilateral central neurons begin to spike a few milliseconds before contralateral neurons, and ipsilateral central neurons also fire at a 30-50% higher rate. We show that a walking fly can detect a 5% asymmetry in total ORN input to its left and right antennal lobes, and can turn toward the odor in less time than it requires the fly to complete a stride. These results demonstrate that neurotransmitter release properties can be tuned independently at output synapses formed by a single axon onto two target cells with identical functions and morphologies. Our data also show that small differences in spike timing and spike rate can produce reliable differences in olfactory behavior.

Insects navigating toward an odor can compare signals from their two antennae, and generally turn in the direction of the stronger signal. Confounding this strategy (by crossing and fixing the antennae, thereby spatially reversing them) impairs olfactory navigation in bees, ants, and locusts6-8. Drosophila resemble other insects in using this strategy3-5, but they are unlike other insects in having mainly bilateral ORN projections. In the Drosophila brain, both ipsi- and contralateral ORN axons synapse onto antennal lobe projection neurons (PNs)9, and ipsi- and contralateral synapses have approximately the same strength10. A minority of Drosophila olfactory glomeruli receive unilateral ORN projections1,11, raising the question of whether these unilateral glomeruli are what enable lateralization.

To investigate lateralization behavior, we built a spherical treadmill to measure olfactory behavior in walking Drosophila (Fig. 1a and Supplementary Fig. 1). To prevent the head from moving, it was glued to the body, and the two antennae were independently stimulated with odor. When we delivered fermented peach volatiles to one antenna and clean air to the other, flies on the treadmill made a fictive turn toward the odor (Fig. 1b and c). If odor or clean air was delivered symmetrically to the two antennae, the fly continued to walk straight (Fig. 1d).

Figure 1
Drosophila can lateralize odors based on bilateral receptor neuron input to a single pair of glomeruli. a. Schematic diagram of a fly on a spherical treadmill, with odor tubes directed at each antenna. b. Representative running trajectories from a single ...

Fruit volatiles typically activate multiple ORN types12. Therefore, we next used a monomolecular odor (pentanoic acid) to more specifically target a glomerulus that receives bilateral ORN input. This odor has just one known high-affinity receptor in the antenna11,12. This receptor corresponds to glomerulus DM6, which receives bilateral ORN innervation2,9. Pentanoic acid elicited turning behavior as robust as that elicited by peach volatiles (Fig. 1d). Turning was significantly reduced by a mutation (Orco2) which silences bilateral ORNs, including the DM6 ORNs (Fig. 1d). The ORN types that project unilaterally do not express the Orco gene11,13, and so this result implies that asymmetric input to strictly bilateral glomeruli can produce turning.

To target a single bilateral ORN type more selectively, we expressed channelrhodopsin-2 (ChR2) in the ORNs that express the olfactory receptor Or42b and project to glomerulus DM1 (Fig. 1e). These ORNs fired a burst of 3 or 4 spikes during a 50-ms light pulse directed at the antenna with a fine fiber optic filament (Fig. 1f). In the same fly, light had no effect on adjacent ORNs that did not express ChR2 (Fig. 1f). Illuminating one antenna produced a turn toward the stimulus (Fig. 1g; Supplementary Figure 2). Conversely, light offset produced a compensatory turn in the other direction (Fig. 1g-i, Supplementary Information Footnote 1). The turns evoked by ChR2 in one ORN type were as large as when ChR2 was expressed in most ORN types (under the control of the Orco-Gal4 line; Fig. 1h). Thus, bilateral ORNs can support lateralization behavior, and turning can be elicited by just a few spikes per neuron in one ORN type.

The temporal control enabled by the optogenetic approach allowed us to precisely determine the latency of the behavioral response. Surprisingly, turning began within ~ 70 ms of the ORN response onset (Fig. 1j). This is faster than the fly's stride period14 (about 100 ms, Supplementary Fig. 1). Thus, there must be a rapid mechanism in the brain for extracting lateralized information from sensory neurons.

We next asked how asymmetric odor stimuli are encoded at the level of PN spikes. We removed one antenna in order to lateralize the odor stimulus, and we made simultaneous cell-attached recordings from PNs ipsi- and contralateral to the intact antenna (Fig. 2a, b). We recorded from pairs of PNs in the same glomerulus on different sides of the brain (“sister PNs”), using GFP to target our electrodes to PNs in glomerulus DM6 or DM1. We used odors that preferentially activate either DM6 ORNs or DM1 ORNs (pentanoic acid or dilute ethyl acetate12). In these experiments, we found a small but consistent asymmetry in PN odor responses. This was apparent in the latency to the first odor-evoked spike: the first ipsilateral spike occurred 2.47±0.70 ms earlier than the first contralateral spike in DM6 PNs, and 1.01±0.41 ms earlier in DM1 PNs (n=16 and 6, Fig. 2c). The latency difference was statistically significant for DM6 PN pairs, although it fell short of significance for DM1 PN pairs (p < 0.005 and p = 0.06, Wilcoxon signed-rank tests). In addition, we found an asymmetry in odour-evoked firing rates: ipsilateral firing rates were on average about 50% higher than contralateral firing rates for DM6 PNs, with a larger asymmetry for DM1 PNs (Fig. 2d–g). The asymmetry was observed even during the spontaneous firing of these cells, and it was proportionately similar for all odour concentrations (Fig. 2d, g). The asymmetry in spikes must be due to an asymmetry in synaptic currents. Indeed, asymmetric stimulation of the antennae produced systematically larger inward currents in ipsi- versus contralateral PNs (Supplementary Fig. 3).

Figure 2
Lateralized odors produce an asymmetry in spike latency and spike rate in antennal lobe projection neurons. a. Schematic of simultaneous cell-attached recording configuration. An ORN axon innervates a glomerulus on each side of the midline, where it synapses ...

Synaptic currents in PNs reflect the combined effects of ORNs and local neurons (LNs), most of which are GABAergic15-17. We first asked if the asymmetry in PN activity requires GABAergic inhibition, by bath-applying GABAA and GABAB receptor antagonists. These antagonists elevated PN firing rates, but the difference between ipsi- and contralateral PN firing rates was not significantly altered (Fig. 3a,b).

Figure 3
The asymmetry arises at the level of ORN-to-PN synapses. a. Mean DM6 PN responses to pentanoic acid (10-2) in normal saline. b. Same as (a) after adding the GABAA receptor antagonist picrotoxin (5 μM) and the GABAB receptor antagonist CGP54626 ...

Our results suggest that the asymmetry originates at the level of ORN input to PNs. To compare ORN-to-PN synapses in ipsi- versus contralateral PNs, we made simultaneous whole-cell recordings of spontaneous excitatory postsynaptic currents (sEPSCs) in pairs of sister PNs. These sEPSCs are known to arise from ORN-to-PN synapses9,10. Each ORN-to-PN synapse consists of many release sites with high release probability, and so spike-evoked synaptic events are large and reliable10. In recordings from sister PNs in glomerulus DM6, virtually all sEPSCs occurred in a paired fashion (Fig. 3c). Almost 100% of the sEPSCs in one cell had a corresponding paired event in the other cell, and this percentage was not significantly different in ipsi- versus contralateral PNs (98.3 ± 0.6% versus 96.4 ± 1.3%, n = 15 pairs, p = 0.11, Wilcoxon signed-rank test). This result indicates that the number of unitary ORN-to-PN synaptic connections is essentially identical on the ipsi- and contralateral sides. This result also argues that action potential conduction failures do not occur to any appreciable degree in the contralateral axon branches of ORNs, although we cannot completely exclude the idea that this occurs at a low rate.

These dual whole-cell recordings allowed us to measure the mean difference in ipsi- versus contralateral sEPSC arrival times (0.80 ± 0.51 ms), which represents the delay imposed by axonal conduction between sister glomeruli. This delay is less than the contralateral lag in first spike latency in glomerulus DM6 (2.47 ms, see above). This makes sense, because the lag in first spike latency is the result of two delays: the delay imposed by axonal conduction, and the delay that results from less net synaptic excitation in the contralateral PN (Supplementary Fig. 3) which means that the contralateral PN requires more integration time to reach spike threshold.

Comparing the amplitude of paired sEPSCs on an event-by-event basis, we found considerable variation in the relative amplitude of ipsi- versus contralateral EPSCs (Fig. 3d). This is expected, given that synaptic vesicle release is a stochastic process occurring independently at ipsi- and contralateral synapses. When many sEPSCs were considered together, there was a consistent asymmetry in the amplitude of sEPSCs (Fig. 3d). Averaged across all experiments, ipsilateral sEPSCs were 39% larger than their contralateral counterparts (Fig. 3e). Thus, although ORN spikes reliably invade both ipsi- and contralateral axon branches, each ORN spike typically has a larger effect on ipsilateral PNs.

In principle, the asymmetry in sEPSC amplitudes could have either a pre- or postsynaptic locus. To determine if we can observe this asymmetry at the level of presynaptic release sites, we expressed synaptobrevin:GFP (a marker of presynaptic release sites) in DM6 ORNs. We removed one antenna and allowed three days for the cut ORN axons to degenerate, leaving only the axons from the intact antenna. We found that total synaptobrevin fluorescence was on average 41 ± 16% higher on the ipsilateral side, and the ipsi/contra ratio of synaptobrevin fluorescence was significantly greater than 1 (p < 0.05, n = 5, t-test, Fig. 4a). This suggests an asymmetry in the number or size of neurotransmitter release sites. This is consistent with a previous study reporting that a plasma membrane marker was more abundant on the ipsilateral side18. That study also severed one antenna and waited three days before imaging. Functional remodeling can occur over that time period18,19, so both findings should be interpreted cautiously.

Figure 4
The asymmetry in ORN-to-PN synapses has a presynaptic origin. a. Synaptobrevin:GFP (green) was expressed in DM6 ORNs to label neurotransmitter release sites, and one antenna was removed 3 days prior to allow the cut axons to degenerate. The brain is viewed ...

Next, to visualize presynaptic calcium, we expressed the genetically-encoded calcium indicator GCaMP3.0 in ORNs (using the pebbled-Gal4 line) and used two-photon microscopy to visualize calcium signals in ORN axon terminals. To lateralize the odor, we removed one antenna immediately prior to the experiment (Fig. 4b). We used pentanoic acid to evoke a fluorescence increase preferentially in glomerulus DM6 (Fig. 4c). We found that the size of the calcium response was significantly larger on the side of the brain ipsilateral to the intact antenna (Fig. 4c and d). This asymmetry did not require feedback from central circuits, because it persisted after washing in mecamylamine (to block nicotinic acetylcholine receptors, which mediate ORN-to-PN synaptic transmission10) along with picrotoxin and CGP54626 (Fig. 4e). In separate experiments, we saw similarly asymmetric calcium signals in glomerulus DL5, using an odor stimulus which is relatively selective for DL5 ORNs20 (trans-2-hexenal, 10-5 dilution, data not shown). We also saw a similar asymmetry in presynaptic currents, using simultaneous bilateral field potential recordings (Supplementary Figure 4). Together, these results demonstrate that the asymmetry in EPSC amplitudes has a presynaptic origin.

Interestingly, we found a roughly equal asymmetry (~40%) in the amplitude of sEPSCs, the level of synaptobrevin fluorescence, and the odour-evoked GCaMP3 fluorescence change. The simplest explanation for these results is that the ipsilateral arbor is 40% larger than the contralateral arbor. If everything else is equal, then this mechanism should produce a proportional change in all these measurements (Supplementary Information Note 2).

Our results are consistent with the finding that, in flies with one antenna removed, 2-deoxyglucose uptake is higher on the side ipsilateral to the intact antenna21. A previous study using calcium imaging has reported that asymmetric ORN stimulation can produce stronger signals in ipsilateral PNs than in contralateral PNs22, which is also broadly consistent with our results. In contrast to our results, however, this asymmetry was restricted to just a few glomeruli. This finding is puzzling, given that a variety of stimuli can produce turning behavior (Fig. 1 and refs 3-5). The same study also concluded that GABAergic inhibition mediates the ipsi/contra asymmetry in PNs22. These discrepancies may reflect limitations of the imaging techniques used in that study, as well as differences between stimuli or glomeruli.

Another previous study failed to find any significant ipsi-contra differences in the strength of ORN-PN synapses10. Those measurements involved sequential recordings from sister PNs, rather than simultaneous recordings. Also, that study measured EPSCs arising from a single ORN per PN. Here, we have better statistical power because we made simultaneous recordings from sister PNs, and because we sampled sEPSCs arising from many ORNs. Our finding that ipsi- and contralateral synapses differ only modestly explains why this difference has been difficult to resolve previously.

It is well-known that a single axon can form neurotransmitter release sites with different properties onto different target cell types (e.g., principal neurons versus interneurons)23. Here, we have shown that ORN axons discriminate between two classes of target cells (left and right) that are morphologically and functionally identical, and that share the same lineage and birth dates24. This is reminiscent of the circuitry of the leech midbody ganglion, where individual mechanoreceptor axons make stronger synapses onto ipsi- versus contralateral sister cells that are otherwise functionally identical25. How are these target cells distinguished? Right and left sister neurons might express different molecular tags26, thereby allowing a given axon to recognize them as ipsi- versus contralateral to itself. Alternatively, an axon might form more release sites at proximal locations as compared to distal locations; this cell-intrinsic mechanism would suffice because contralateral sites are always more distal than ipsilateral sites.

Our results reveal that even small signals in the Drosophila nervous system can be behaviorally relevant. The stimuli we used in our optogenetic experiments produced only a slight fractional difference in input to the right and left sides of the brain. This difference amounted to 5% over the 50-ms stimulus period (Supplementary Information Footnote 3). The finding that this incremental difference is relevant for behavior should motivate the continued development of sensitive methods for monitoring neural activity in the fly brain.

Methods Summary

The odor delivery device used for the olfactory behavioral experiments was specially designed to deliver no lateralized mechanical cues, and we performed control experiments to confirm that no turning was observed when no odor was present (Supplementary Fig. 5). The spherical treadmill apparatus was constructed by floating a small plastic sphere on a jet of compressed air. We measured the forward and lateral velocities of the sphere in the apparatus by placing the sensor from an optical mouse underneath the sphere. Photostimulation of Drosophila ORNs was achieved by butt-coupling a blue LED to a fiber optic filament (50 μm diameter) and positioning the tip of the fiber ~150 μm away from the fly's antenna. In all behavioral experiments using light to stimulate ORNs, the eyes and ocelli of the fly were shielded from light by painting them with ink. In vivo extracellular recordings from ORNs and patch-clamp recordings from PNs were performed as previously described9,27. In all electrophysiology or calcium imaging experiments (except ORN recordings), one antenna was removed just prior to the experiment by a person who was not the experimenter, and the experimenter remained blind to which side of the brain was ipsilateral to the intact antenna. All analysis was also performed blind to which side was ipsilateral. Calcium imaging experiments were performed on a custom-built two-photon microscope. All measurements represent mean ± s.e.m. computed across experiments. See Supplementary Methods for details on the spherical treadmill, odor delivery, optogenetic stimuli, and analysis of sEPSCs.

Supplementary Material


We are grateful to Michael Dickinson, Vivek Jayaraman, Liqun Luo, Dan Tracey, and Leslie Vosshall for gifts of fly stocks. Allison Baker helped construct and improve the spherical treadmill apparatus. Members of the Wilson lab provided feedback on the manuscript. This work was supported by a research project grant from the National Institutes of Health (R01DC008174). R.I.W. is an HHMI Early Career Scientist. B.D. and J.K. were supported by the Rowland Junior Fellows Program.


Author Contributions: Q.G. and R.I.W. designed the experiments. Q.G. performed all the experiments, except for the calcium imaging, which was performed by Q.G. and E.J.H, and the synaptobrevin imaging, which was performed by R.I.W. Q.G. analyzed the data. J.K. and B.L.D. helped design and build the spherical treadmill apparatus. Q.G. and R.I.W. wrote the manuscript.


1. Stocker RF, Lienhard MC, Borst A, Fischbach KF. Neuronal architecture of the antennal lobe in Drosophila melanogaster. Cell Tissue Res. 1990;262:9–34. [PubMed]
2. Couto A, Alenius M, Dickson BJ. Molecular, anatomical, and functional organization of the Drosophila olfactory system. Curr. Biol. 2005;15:1535–1547. [PubMed]
3. Borst A, Heisenberg M. Osmotropotaxis in Drosophila melanogaster. J. Comp. Physiol. [A] 1982;147:479–484.
4. Duistermars BJ, Chow DM, Frye MA. Flies require bilateral sensory input to track odor gradients in flight. Curr. Biol. 2009;19:1301–1307. [PMC free article] [PubMed]
5. Flugge C. Geruchliche Raumorientierung von Drosophila melanogaster. Z. Vergl. Phys. 1934;20:463–500.
6. Kennedy JS, Moorehouse JE. Laboratory observations on locust responses to wind-borne grass odour Entomol. Exp. Appl. 1969;12:487–503.
7. Martin H. Osmotropotaxis in the honey-bee. Nature. 1965;208:59–63.
8. Hangartner W. Spezifität und Inaktivierung des Spurpheromons von Lasius fuliginosus (Latr.) und Orientierung der Arbeiterinnen im Duftfeld. Z. vergl. Physiol. 1967;57:103–136.
9. Kazama H, Wilson RI. Origins of correlated activity in an olfactory circuit. Nat. Neurosci. 2009;12:1136–1144. [PMC free article] [PubMed]
10. Kazama H, Wilson RI. Homeostatic matching and nonlinear amplification at genetically-identified central synapses. Neuron. 2008;58:401–413. [PMC free article] [PubMed]
11. Silbering AF, et al. Complementary function and integrated wiring of the evolutionarily distinct Drosophila olfactory subsystems. J Neurosci. 2011;31:13357–13375. [PubMed]
12. Hallem EA, Carlson JR. Coding of odors by a receptor repertoire. Cell. 2006;125:143–160. [PubMed]
13. Benton R, Vannice KS, Gomez-Diaz C, Vosshall LB. Variant ionotropic glutamate receptors as chemosensory receptors in Drosophila. Cell. 2009;136:149–162. [PMC free article] [PubMed]
14. Strauss R, Heisenberg M. Coordination of legs during straight walking and turning in Drosophila melanogaster. J Comp Physiol A. 1990;167:403–412. [PubMed]
15. Chou YH, et al. Diversity and wiring variability of olfactory local interneurons in the Drosophila antennal lobe. Nat. Neurosci. 2010;13:439–449. [PMC free article] [PubMed]
16. Okada R, Awasaki T, Ito K. Gamma-aminobuyric acid (GABA)-mediated neural connections in the Drosophila antennal lobe. J. Comp. Neurol. 2009;514:74–91. [PubMed]
17. Das A, et al. Drosophila olfactory local interneurons and projection neurons derive from a common neuroblast lineage specified by the empty spiracles gene. Neural Dev. 2008;3:33. [PMC free article] [PubMed]
18. Berdnik D, Chihara T, Couto A, Luo L. Wiring stability of the adult Drosophila olfactory circuit after lesion. J. Neurosci. 2006;26:3367–3376. [PubMed]
19. Kazama H, Yaksi E, Wilson RI. Cell death triggers olfactory circuit plasticity via glial signaling in Drosophila. J. Neurosci. 2011;31:7619–7630. [PMC free article] [PubMed]
20. Olsen SR, Bhandawat V, Wilson RI. Divisive normalization in olfactory population codes. Neuron. 2010;66:287–299. [PMC free article] [PubMed]
21. Rodrigues V. Spatial coding of olfactory information in the antennal lobe of Drosophila melanogaster. Brain Res. 1988;453:299–307. [PubMed]
22. Agarwal G, Isacoff E. Specializations of a pheromonal glomerulus in the Drosophila olfactory system. J. Neurophysiol. 2011;105:1711–1721. [PubMed]
23. Pelkey KA, McBain CJ. Differential regulation at functionally divergent release sites along a common axon. Curr. Opin. Neurobiol. 2007;17:366–373. [PubMed]
24. Jefferis GS, Marin EC, Stocker RF, Luo L. Target neuron prespecification in the olfactory map of Drosophila. Nature. 2001;414:204–208. [PubMed]
25. Lockery SR, Kristan WB., Jr. Distributed processing of sensory information in the leech. II. Identification of interneurons contributing to the local bending reflex. J. Neurosci. 1190;10:1816–1829. [PubMed]
26. Chintapalli VR, et al. Functional correlates of positional and gender-specific renal asymmetry in Drosophila. PLoS One. 2012;7:e32577. [PMC free article] [PubMed]
27. Bhandawat V, Olsen SR, Schlief ML, Gouwens NW, Wilson RI. Sensory processing in the Drosophila antennal lobe increases the reliability and separability of ensemble odor representations. Nat. Neurosci. 2007;10:1474–1482. [PMC free article] [PubMed]
28. Bhandawat V, Maimon G, Dickinson MH, Wilson RI. Olfactory modulation of flight in Drosophila is sensitive, selective and rapid. J. Exp. Biol. 2010;213:3625–3635. [PubMed]
29. Fishilevich E, Vosshall LB. Genetic and functional subdivision of the Drosophila antennal lobe. Curr. Biol. 2005;15:1548–1553. [PubMed]
30. Larsson MC, et al. Or83b encodes a broadly expressed odorant receptor essential for Drosophila olfaction. Neuron. 2004;43:703–714. [PubMed]
31. Hwang RY, et al. Nociceptive neurons protect Drosophila larvae from parasitoid wasps. Curr. Biol. 2007;17:2105–2116. [PMC free article] [PubMed]
32. Tanaka NK, Awasaki T, Shimada T, Ito K. Integration of chemosensory pathways in the Drosophila second-order olfactory centers. Curr. Biol. 2004;14:449–457. [PubMed]
33. Lee T, Luo L. Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron. 1999;22:451–461. [PubMed]
34. Zhang YQ, Rodesch CK, Broadie K. Living synaptic vesicle marker: synaptotagmin-GFP. Genesis. 2002;34:142–145. [PubMed]
35. Sweeney LB, et al. Temporal target restriction of olfactory receptor neurons by Semaphorin-1a/PlexinA-mediated axon-axon interactions. Neuron. 2007;53:185–200. [PubMed]
36. Tian L, et al. Imaging neural activity in worms, flies and mice with improved GCaMP calcium indicators. Nat. Methods. 2009;6:875–881. [PMC free article] [PubMed]