of nomenclature of nuclear receptors.Pharmacol. Rev. 58, 685–704.

11. Yen, P.M. (2001). Physiological and molecularbasis of thyroid hormone action. Phys. Rev. 81,1097–1142.

12. Buchholz, D.R., Paul, B.D., Fu, L.Z., andShi, Y.B. (2006). Molecular and developmentalanalyses of thyroid hormone receptorfunction in Xenopus laevis, the African clawedfrog. General Comparative Endocrinol. 145,1–19.

13. Just, J.J., Kraus-Just, J., and Check, D.A.(1981). Survey of chordate metamorphosis.In Metamorphosis, Second Edition,L.I.G.a.E. Frieden, ed. (New York, NY: PlenumPress), pp. 265–326.

14. Delsuc, F., Brinkmann, H., Chourrout, D., andPhilippe, H. (2006). Tunicates and not

cephalochordates are the closest livingrelatives of vertebrates. Nature 439, 965–968.

15. Ogasawara, M. (2000). Overlapping expressionof amphioxus homologs of the thyroidtranscription factor-1 gene and thyroidperoxidase gene in the endostyle: insight intoevolution of the thyroid gland. Dev. Genes Evol.210, 231–242.

16. Wickstea, J.H. (1967). Branchiostomalanceolatum larvae - some experiments oneffect of thiouracil on metamorphosis. J. Mar.Biol. Assoc. UK 47, 49–61.

17. Yaoita, Y., and Brown, D.D. (1990). A correlationof thyroid hormone receptor gene expressionwith amphibian metamorphosis. Genes Dev. 4,1917–1924.

18. Tata, J.R. (2000). Autoinduction of nuclearhormone receptors during metamorphosis and

its significance. Insect Biochem. Mol. Biol. 30,645–651.

19. Truman, J.W., and Riddiford, L.M. (2002).Endocrine insights into the evolution ofmetamorphosis in insects. Annu. Rev. Entomol.47, 467–500.

Department of Molecular, Cellular andDevelopmental Biology and Department ofEcology and Evolutionary Biology,The University of Michigan, Ann Arbor,Michigan 48109-1048, USA.E-mail: rdenver@umich.edu

DOI: 10.1016/j.cub.2008.05.024

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Circadian Rhythms: Timing the Senseof Smell

It is well established that Drosophila olfaction is under circadian control, yet themechanisms underlying this temporal regulation have remained elusive. Thekinase GPRK2 has now been identified as a critical link between the circadianclock and olfactory responses.

Patrick Emery and Michael Francis

Most animals use specific behavioralstrategies to cope with daily changes intemperature, light intensities, and theirecological environment. For example,some species adopt a diurnal ora nocturnal behavior, while others aremostly active at dawn or dusk to avoidtemperature extremes. Thesebehavioral adaptations are largelyunder the control of cell-autonomouscircadian clocks found in brainpacemaker neurons. These molecularoscillators are composed ofa transcriptional feedback loop witha period adjusted to 24 hours bypost-translational regulation of theactivity and stability of its components[1]. Since most bodily functions, aswell as behavior, require propersynchronization to maximize resourceutilization while minimizing energyexpenditure, circadian clocks are alsofound in many peripheral organs andtissues, where they control localphysiology and metabolism [2].

Olfaction is essential for variousbehaviors influenced by circadianclocks (e.g., feeding and socialinteractions, such as mating andpredator avoidance) and is itself undercircadian regulation in mammals andinsects [3–5]. The Drosophila olfactorysystem provides a powerful modelsystem to explore circadian regulation

of sensory processing. Drosophilaolfactory sensory neurons (OSNs) arelocated in two pairs of specializedsensory organs, the antennae and themaxillary palps. Between one and fourOSNs are grouped into sensilla, whichare specialized hairs that dot thesensory organs, and can be classifiedinto three types — trichoid, basiconic,and coelomic — on the basis ofmorphological criteria [6]. Thestereotyped arrangement of theolfactory system has made it amenableto electrophysiological recordings. Theelectroantennogram (EAG) providesa measure of the compound responseof all antennal sensilla to a given odorstimulus, and extracellular-sharp-electrode recordings from individualsensilla have provided exquisitelydetailed information about theresponse properties (dendritic spikingactivity) of single OSNs. EAGrecordings demonstrated that odorresponses in Drosophila antennae areregulated by cell-autonomouscircadian clocks present within theOSNs [5,7]. The EAG amplitudechanges over the course of the day,with a marked increase during themiddle of the night. Interestingly, thiscorresponds to the time at which fliesare behaviorally most reactive toodorant stimuli [8].

So, how does circadian regulation ofodor sensation occur at the level of the

sensory neurons? Two papers fromHardin and colleagues [9,10], ina recent issue of Current Biology, shedlight on these issues. The olfactoryresponse is initiated by the binding ofvolatile compounds to olfactoryreceptors (ORs). In mammals andCaenorhabditis elegans, ORs belong tothe family of G-protein-coupledreceptors (GPCRs), characterized byhaving 7 transmembrane domains[11,12]. Since Drosophila ORs havea similar membrane topology [13,14],Tanoue and colleagues [10] decided todetermine whether genes known orpredicted to regulate GPCR function inDrosophila are under circadianregulation in antennae. G-proteinreceptor kinases (GPRKs) wereattractive candidates for these GPCRregulators. Indeed, in mammals,GPRKs phosphorylate activatedGPCRs. Arrestins then bind to thephosphorylated receptors, uncouplethem from their G-protein partners and,thus, terminate GPCR signaling [15].Consistent with this idea, Tanoue et al.[10] found that the expression ofa Drosophila GPRK, gprk2, iscontrolled by the circadian clock [10].Unexpectedly, however, the levels ofgprk2 mRNA and protein oscillatein-phase with the daily oscillations inEAG amplitude, suggesting a directcorrelation between GPRK2abundance and EAG amplitude, ratherthan an inhibitory role for GPRK2 in ORsignaling. This hypothesis was verifiedby measuring EAG amplitude in mutantflies with constantly high or low GPRK2expression. Surprisingly, the dailyvariations in GPRK2 levels are quitemodest, with night-time levels rising toconcentrations that are only 1.5–2 foldgreater than GPRK2 levels during theday. Increasing or decreasing GPRK2

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Figure 1. Circadian clocks within olfactory sensory neurons (OSNs) control GPRK2 levels, ORlocalization, and spike amplitude.

The Drosophila circadian pacemaker (symbolized by a clock dial) determines gprk2 mRNA lev-els by modulating gprk2 transcription or mRNA stability. The resulting increase in GPRK2 lev-els during the night (Zeitgeber Time (ZT) 17), compared with during the day (ZT5), promotesthe localization of odorant receptors (ORs) to the dendritic membrane of the OSNs via an un-known mechanism (here depicted as a positive regulation of OR vesicular transport). The ORsare non-specific cation channels composed of two subunits: an invariant subunit called OR83band an odor-specific subunit (ORX). Cyclic changes in dendritic OR levels result in circadianvariations in the amplitude of spontaneous dendritic spikes, and in the response of the wholeantennae to odors.

levels beyond this natural oscillationhad little effect on EAG amplitude(unless GPRK2 levels fell belowdetection threshold). Thus, thecircadian clock precisely tunes GPRK2levels to be in a dynamic range in whichrelatively small changes in proteinlevels have profound effects on EAGamplitude.

How might GPRK2 mediate changesin odor responsiveness? Krishnan et al.[9] found that redistribution of ORs tothe sensory dendrites accompaniesincreases in GPRK2 expression,suggesting that regulation might occurat the level of the OR (Figure 1). To testthis idea, the spontaneous activity fromindividual OSNs of several classes oftrichoid and basiconic sensilla wasmeasured [9]. In single-unit recordings,Krishnan et al. [9] observed rhythmic

changes in spike amplitude thatoccurred in-phase with the rhythmsof odor-evoked EAG responses,suggesting a common mechanism.Furthermore, two lines of evidencesuggested that circadian regulation ofspike amplitude involves modulationof OR activity or distribution. First,rhythms in spontaneous spikeamplitude were dependent on GPRK2;second, single-unit recordings fromDrosophila mutants lacking ORs didnot show rhythms in spike amplitude.Together, these data suggestthat circadian control of spikeamplitude arises from rhythmic,GPRK2-dependent localizationof ORs to dendrites. Interestingly,Krishnan et al. [9] did not observea corresponding circadian rhythm inthe spontaneous or odor-evoked spike

frequency of OSNs. Previous studiesmeasuring odor-evoked andspontaneous spike activity havedemonstrated the importance of spikefrequency rather than amplitude inneural coding of odor identity andconcentration, and suggested thatspike amplitude is an invariant propertyemerging from the unique membraneexcitability and morphologicalcharacteristics of each OSN class [16].The finding that spontaneous spikeamplitude is under circadian controlsuggests that spike amplitude may bea more dynamic property thanpreviously appreciated and leads to thespeculation that spike amplitude maybe an important factor in determiningsensitivity to olfactory cues.

Two recent studies havedemonstrated that insect ORs canthemselves act as ion channels [17,18].This finding draws a clear distinctionbetween the mechanism of GPRK2action in Drosophila OSNs and themechanism by which GPRKs terminateGPCR signals in mammalian systemsand raises some intriguing questionsfor future studies. How does GPRK2regulate OR function or distribution,and how do changes in OR propertiestranslate into rhythms in spikeamplitude? Perhaps GPRK2 regulatesthe coupling of OR ion channel activitywith activation of downstream ionchannels and, thereby, regulatesmembrane excitability properties andspontaneous spike amplitude. DoesGPRK2 contribute directly to theassembly of functional ORs or promotethe translocation of assembledchannels to the dendritic membrane? IsGPRK2 kinase activity required forcircadian rhythmicity in olfactoryresponses? And, importantly, how dochanges in OSN spike amplitudecontribute to responses to olfactorycues and modulate olfactory-drivenbehaviors? Interestingly, the phaseof spike-amplitude oscillationsdiffers between basiconic andtrichoid sensilla [9]. Since the latterplay an important role in the detectionof pheromones [19], whereas theformer appear to be tuned torecognize food odors [20], thisobservation suggests that circadianclocks within the OSNs might play animportant role in the temporalcompartmentalization of variousodor-guided behaviors.

In summary, Hardin and colleagues[9,10] have identified GPRK2 as a keyregulator of OR function and olfactory

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responses and demonstrated thatGPRK2 levels are under the control ofcircadian molecular pacemakerslocated within the OSNs. Whileunderstanding the precise mechanisticbasis of GPRK2 action will providea fertile area of inquiry for furtherstudies, this work has provided anexciting first glimpse into the circadiancontrol of olfaction. Future genetic,electrophysiological, and behavioralstudies are certain to reveal additionalinsights into olfactory rhythms and theirimplications for our rapidly advancingunderstanding of olfactory coding andfor the regulation of odor-guidedcircadian behaviors.

References1. Hardin, P.E. (2006). Essential and expendable

features of the circadian timekeepingmechanism. Curr. Opin. Neurobiol. 16, 686–692.

2. Stratmann, M., and Schibler, U. (2006).Properties, entrainment, and physiologicalfunctions of mammalian peripheral oscillators.J. Biol. Rhythms 21, 494–506.

3. Granados-Fuentes, D., Tseng, A., andHerzog, E.D. (2006). A circadian clock in theolfactory bulb controls olfactory responsivity.J. Neurosci. 26, 12219–12225.

4. Merlin, C., Lucas, P., Rochat, D.,Francois, M.C., Maibeche-Coisne, M., andJacquin-Joly, E. (2007). An antennal circadianclock and circadian rhythms in peripheral

pheromone reception in the moth Spodopteralittoralis. J. Biol. Rhythms 22, 502–514.

5. Krishnan, B., Dryer, S.E., and Hardin, P.E.(1999). Circadian rhythms in olfactoryresponses of Drosophila melanogaster. Nature400, 375–378.

6. Vosshall, L.B., and Stocker, R.F. (2007).Molecular architecture of smell and taste inDrosophila. Annu. Rev. Neurosci. 30, 505–533.

7. Tanoue, S., Krishnan, P., Krishnan, B.,Dryer, S.E., and Hardin, P.E. (2004). Circadianclocks in antennal neurons are necessary andsufficient for olfaction rhythms in Drosophila.Curr. Biol. 14, 638–649.

8. Zhou, X., Yuan, C., and Guo, A. (2005).Drosophila olfactory response rhythms requireclock genes but not pigment dispersing factoror lateral neurons. J. Biol. Rhythms 20, 237–244.

9. Krishnan, P., Chatterjee, A., Tanoue, S., andHardin, P.E. (2008). Spike amplitude of singleunit responses in antennal sensillae iscontrolled by the Drosophila circadian clock.Curr. Biol. 18, 803–807.

10. Tanoue, S., Krishnan, P., Chatterjee, A., andHardin, P.E. (2008). G-protein Receptor Kinase2 is required for rhythmic olfactory responses inDrosophila. Curr. Biol. 18, 787–794.

11. Buck, L., and Axel, R. (1991). A novel multigenefamily may encode odorant receptors:a molecular basis for odor recognition. Cell 65,175–187.

12. Troemel, E.R., Chou, J.H., Dwyer, N.D.,Colbert, H.A., and Bargmann, C.I. (1995).Divergent seven transmembrane receptorsare candidate chemosensory receptors inC. elegans. Cell 83, 207–218.

13. Clyne, P.J., Warr, C.G., Freeman, M.R.,Lessing, D., Kim, J., and Carlson, J.R. (1999). Anovel family of divergent seven-transmembraneproteins: candidate odorant receptors inDrosophila. Neuron 22, 327–338.

14. Vosshall, L.B., Amrein, H., Morozov, P.S.,Rzhetsky, A., and Axel, R. (1999). A spatial mapof olfactory receptor expression in theDrosophila antenna. Cell 96, 725–736.

15. Gainetdinov, R.R., Premont, R.T., Bohn, L.M.,Lefkowitz, R.J., and Caron, M.G. (2004).Desensitization of G protein-coupled receptorsand neuronal functions. Annu. Rev. Neurosci.27, 107–144.

16. Hallem, E.A., Ho, M.G., and Carlson, J.R. (2004).The molecular basis of odor coding in theDrosophila antenna. Cell 117, 965–979.

17. Sato, K., Pellegrino, M., Nakagawa, T.,Nakagawa, T., Vosshall, L.B., and Touhara, K.(2008). Insect olfactory receptors areheteromeric ligand-gated ion channels. Nature452, 1002–1006.

18. Wicher, D., Schafer, R., Bauernfeind, R.,Stensmyr, M.C., Heller, R., Heinemann, S.H.,and Hansson, B.S. (2008). Drosophila odorantreceptors are both ligand-gated and cyclic-nucleotide-activated cation channels. Nature452, 1007–1011.

19. Ha, T.S., and Smith, D.P. (2006). A pheromonereceptor mediates 11-cis-vaccenylacetate-induced responses in Drosophila.J. Neurosci. 26, 8727–8733.

20. Hallem, E.A., and Carlson, J.R. (2006). Codingof odors by a receptor repertoire. Cell 125,143–160.

University of Massachusetts Medical School,Department of Neurobiology, Worcester,Massachusetts 01605, USA.E-mail: Patrick.Emery@umassmed.edu,Michael.Francis@umassmed.edu

DOI: 10.1016/j.cub.2008.05.011

Gamete Fusion: Key Protein Identified

Is there a common mechanism of eukaryotic sex? Two recent reports highlightan ancient and widely distributed protein that is key to gamete fusion and isa potential target for malaria vaccines.

Christopher D. Goodmanand Geoffrey I. McFadden

Sex defines us — not as individualsor as a species but as eukaryotes.Reductive meiotic division to producehaploid gametes, and the subsequentfusion of these two cells allow for twovital processes: the reassortment ofalleles through recombination, and theregeneration of decrepit bodies byreversion to a single-celled statefollowed by new cell growth. Sex hasundoubtedly been a key to the successof eukaryotes, but it’s a tricky thing todo. The gametes must recognize eachother, then fuse; and both of theseprocesses must be managed carefullyas failures are disastrous. Despite thefundamental nature of this process,pulling back the veil to get a peek at themolecular mechanisms controllinggamete fusion hasn’t been easy, in

stark contrast to the exquisite detailin which the processes of intracellularmembrane fusion have beendescribed. But vesicle fusion withina cell has little in common with theunion of two intact cells, and only veryrecently have some of the proteinsinvolved in gamete fusion in plantsand animals been identified (reviewedin [1]). Still lacking is a basicunderstanding of the exact role theseproteins play in the complex series ofevents that leads to fertilization. Thisdeficiency in understanding stemsin part from the complex nature ofgamete development, which limitsthe availability of tractable in vitroexperimental systems, and alsofrom the difficulty in observingand manipulating the processesof fertilization in vivo [1].

Two new papers — one in a recentissue of Current Biology by Hirai et al.

[2] — have made significant stridestowards understanding thefundamental process of gamete fusion[2,3]. Both focus on a protein, knowneither as GCS1 (generative cell specific1) or HAP2 (hapless2), that was firstidentified in screens for male-fertilitymutants defective in pollen tube growthin the plants Arabidopsis [4] and Lilium[5], then again in a screen for mutantscompromised at later stages of gameteinteraction in the green algaeChlamydomonas reinhardtii [3]. GCS1thus has a clear role in plant and algalfertilization, but genome searches havealso revealed homologues in malariaparasites (Plasmodium), Africansleeping sickness parasites(Trypanosoma brucei), slime moulds,ciliates, choanoflagellates andcnidarians [5], suggesting that GCS1is an ancient and highly conservedcomponent of the gamete fusionapparatus. Hirai et al. [2] and Liu et al.[3] now provide elegant support forthis hypothesis.

The fact that both groups chose towork with malaria parasites toinvestigate GCS1 function markssomething of a coming of age for