These data indicate that FLRT3 acts as a controlling factor of retinal vascular development and suggests that the action of FLRT3 depends on its interaction

with Unc5B. The structural data presented here indicate that distinct FLRT LRR surfaces mediate homophilic adhesion and Unc5-dependent repulsion. By using these surfaces, FLRTs can affect both adhesive and repulsive functions in the same receiving cell, e.g., neurons or vascular cells that coexpress FLRT and Unc5. We show that coexpressed FLRT and Unc5 act in parallel, and that cells must integrate these adhesive and repulsive effects. This separation Venetoclax datasheet of adhesive and repulsive functionalities allows FLRTs to regulate the behavior of migrating pyramidal neurons in distinct ways; FLRT2 repels Unc5D+ neurons and thereby controls their radial migration, while FLRT3-FLRT3 homophilic interactions regulate their tangential distribution. FLRT3 also controls retinal vascularization, possibly involving combinatorial signaling via FLRT and Unc5. To distinguish FLRTs from adhesion-only CAMs, we propose to define a new subgroup, here designated as repelling CAMs (reCAMs). reCAMs provide a guidance system that combines the finely tunable cell adhesion of classical INCB024360 molecular weight homophilic CAMs with repulsive functions through the addition of a heterophilic

receptor. We show here that FLRT-mediated adhesion involves the conserved concave surface on the LRR domain. This mode of homophilic binding resembles that of other LRR-type CAMs, for example, decorin (Scott et al., 2004). The FLRT-FLRT binding affinity is

weak (below the sensitivity of our SPR assay ∼100 μM), and FLRT oligomerization correlates with local concentration. Thus, FLRTs are ideal candidates for providing the finely tuned adhesive cell-cell traction required for cell migration. In contrast to the low-affinity adhesive binding, repulsive found FLRT-Unc5 interaction is of nanomolar affinity and mediated through a distinct binding surface on the FLRT LRR domain. The high degree of conservation within the binding surfaces of Unc5 and FLRT homologs suggests the interaction evolved before homolog diversification. The mode of interaction is atypical for LRR-type proteins, which mostly bind ligands via the concave surface of the domain, although some examples of ligand-binding surfaces other than the concave side exist (Bella et al., 2008). Our results with thalamic neurons and vascular cells indicate that coexpressed FLRTs act as attenuators of Unc5 repulsion. Stripe assays with FLRT3-positive, compared to FLRT3-negative, thalamic axons provide strong evidence that the attenuation results from FLRT-FLRT interaction in trans, rather than in cis, masking.

The molecular mechanism by which caspase effects are restricted to specific neuronal compartments (perhaps even specific synapses) is an important unanswered

question. Local breakdown of proteins—and selective MAPK inhibitor pruning of synapses—can additionally be achieved by spatiotemporal control of E3 ligase assembly. In Caenorhabditis elegans, localized inhibition of the assembly of an SCF complex through binding of core protein SKR-1 to a synaptic adhesion molecule, SYG-1, spares synapses from elimination ( Ding et al., 2007). It is unknown whether synapse elimination in mammals also relies on local regulation of E3 ubiquitin ligase. The morphological sculpting of certain synapses is regulated by an evolutionarily conserved RING domain E3 ligase Phr1 (also known as Highwire in Drosophila, and RPM-1 in C. elegans)

( Schaefer et al., 2000, Wan et al., 2000 and Zhen et al., 2000). In mammals, Phr1 functions to sculpt motor nerve terminals and is essential for formation of major CNS axon tracts ( Bloom et al., 2007). Interestingly, in mice, Phr1 is localized to the axonal shaft and excluded from growth cones, where the protein kinase DLK is restricted ( Lewcock et al., 2007). In the absence of Phr1, DLK aberrantly distributes to axons, leading to altered microtubule dynamics and axon-pathfinding deficits. Based on the reciprocal localization of DLK and Phr1, DLK was proposed as a Phr1 substrate, similar to the scenario in invertebrates ( Collins et al., 2006, Lewcock et al., 2007 and Nakata et al., 2005). However, no increase in DLK was detected find more in the central nervous system of Phr1 mutant mice and DLK is not required for Phr1 loss-of-function phenotypes ( Bloom et al., 2007). In fish, Phr1 localizes to growth cones and regulates pathfinding independent of DLK ( Hendricks and Jesuthasan, 2009). Collectively, these studies demonstrate that despite possible cell-type or species-specific differences

in the regulation of Phr1, this ubiquitin ligase regulates microtubule remodeling during development and is crucial for axon navigation. HECT domain Nedd4 is another ubiquitin ligase acting in axons; it promotes branching by targeting PTEN, a PIP3 phosphatase that negatively regulates axonal branching (Drinjakovic et al., 2010). Remarkably, Nedd4 also enhances Beta Amyloid the branching of dendrites by monoubiquitinating GTPase Rap2 and inhibiting its function (Kawabe et al., 2010). Thus an E3 ligase can target different substrates in different subcellular compartments to carry out similar cell biological functions. Neurons also utilize the lysosome system to degrade organelles and synaptic proteins. For example, following endocytosis, AMPARs either recycle back to the membrane or are sorted into lysosomes, depending on their subunit composition and whether AMPARs themselves or NMDARs were activated (Ehlers, 2000 and Lee et al., 2004).

This minimal stimulation protocol revealed that in slices from IO rats the minimally evoked EPSC amplitude (excluding failures; “potency”) was greater compared to that in slices from sham animals (Figure 8C). The potency for single fiber activation is dependent on quantal size and number of functional synaptic connections. There was a 37% increase in quantal size measured using Sr-evoked mEPSCs (Figure 7), whereas the potency increase was approximately 65%, demonstrating that IO nerve resection additionally causes an find more increase in the number of functional TC synapses in L4. Taken together,

the results show that IO nerve resection causes plasticity of the spared TC input by BMN 673 in vitro increasing both quantal amplitude and number of functional synapses. The present study investigates the mechanisms and sites of plasticity induced

by loss of whisker sensory input in 6-week-old rats using a combined MRI and slice electrophysiology approach. In contrast to the expectation that plasticity at this age is mediated by modification of cortico-cortical inputs, we found that a prominent plasticity of TC input to L4 underlies the robust increase in spared barrel cortex activation detectable by fMRI. This plasticity was due to a selective increase in quantal amplitude and number of functional synaptic contacts at the TC input to L4 stellate cells while maintaining excitatory/inhibitory balance. This combined MRI and slice electrophysiology approach therefore allows for an analysis of sites and mechanisms of plasticity, which could be broadly applied to many paradigms. The results show that TC inputs can mediate plasticity after the end of the previously defined critical period for this input. IO nerve resection was the sensory

manipulation used to induce experience-dependent plasticity in barrel cortex. The IO nerve carries all sensory information from the whiskers, but does not contain motor afferents; therefore, this manipulation results in a complete loss of whisker-dependent sensory input with no loss Amisulpride of motor innervation to the whiskers. The increase in cortical BOLD-fMRI responses after 2 weeks of IO nerve resection in response to electrical stimulation of the spared whisker pad is likely due to increased cortical neuronal activity. Although there are a few examples in which BOLD-fMRI has not been associated with corresponding changes in neuronal activity (Maier et al., 2008 and Sirotin and Das, 2009), a proportional increase of BOLD and neuronal signals has been observed in functional mapping studies across a variety of species including rodent, monkey, and human (Heeger et al., 2000, Logothetis et al., 2001, Ogawa et al., 2000 and Rees et al., 2000), including for somatosensory cortex (Goloshevsky et al., 2008 and Hyder et al., 2002).

1 mM EGTA, 5 mM MgCl2, 10 mM KCl, 5 mM NaF, 2 mM Na3VO4, 4 mM Na4P2O7, 1 mM PMSF, and 1% Triton X-100)

supplemented with a protease inhibitor cocktail (Nacalai Tesque). PIP5Kγ661 was immunoprecipitated with an anti-PIP5Kγ661 antibody conjugated with protein A sepharose (GE Healthcare). Proteins in the immunoprecipitate were blotted with anti-α adaptin, anti-β high throughput screening assay adaptin, and anti-PIP5Kγ antibodies. For time-lapse imaging, hippocampal neurons were plated on 35 mm PEI-coated glass-bottom dishes (thickness = 0.12–0.19 mm; Mattek or Asahi glass) and cultured in Neurobasal medium without phenol red (Invitrogen), with B-27 supplement (Invitrogen) and 0.5 mM L-glutamineas as described above for 16–20 DIV. The neurons were transiently transfected with plasmids for VN-β2 ear and VC-PIP5K-WT or VC-PIP5K-S645E and cultured for another 19–26 hr. During imaging, the glass-bottom dish was kept at 32°C by a microscope incubation

system (Tokai Hit). Time-lapse epifluorescent images were acquired with a Nikon Eclipse Ti inverted microscope DZNeP equipped with an Apochromatic 60× oil immersion objective (NA 1.49) and an ORCA-II-ER camera (Hamamatsu Photonics). Image capture and data acquisition were performed using NIS-Elements BR3.0 software (Nikon). Image sequences were subsequently processed with NIS-Elements and ImageJ software (1.42q, National Institutes of Health). For immunocytochemical analysis, hippocampal neurons cultured on PEI-coated glass coverslips were transfected as described above. After treatment with 50 μM NMDA for 5 min, the neurons were fixed, permeabilized, and blocked with a blocking solution containing 0.4% Triton X-100. PSD-95 and MAP2 were labeled with anti-PSD-95 (1:1,000) and anti-MAP2 antibodies (1:1,000), respectively, and visualized with Alexa 546 secondary antibodies (1:1,000). F-actin was labeled with rhodamine phalloidin. The numbers of the Venus punctate Pramipexole signals in the dendrites and the total length of the dendrites between 20 and 100 μm from the soma were measured. PIP5Kγ661 activity was determined as previously reported (Honda et al., 1999). For more detail, see Supplemental Experimental Procedures. The recombinant Sindbis virus for the expression

of GFP and wild-type or kinase-dead PIP5Kγ661 was constructed as described (Matsuda et al., 2003). Under the deep anesthesia with an intraperitoneal injection of ketamine/xylazine (80/20 mg/kg; Sigma), the recombinant Sindbis virus (2.5 μl; titer, 1.0 × 108−1.0 × 109 TU/ml) was stereotactically injected into the CA1 region of dorsal hippocampus of P14–21 ICR mice (2.0–2.3 mm posterior to the Bregma, 1.5–2.0 mm lateral to the midline, and 1.5–2.0 mm ventral from the pial surface). After 24–36 hr, infected cells were identified by the GFP expression, and hippocampal slices were used for electrophysiological analyses. Transverse hippocampal slices (300 μm thickness) were prepared from P14–21 ICR mice or virus-infected mice according to the institutional guidelines.

The discrepancy with earlier estimates is best explained

by an inability to draw AIS Na+ channels into the patch-clamp recording pipette due to tight coupling of these VEGFR inhibitor channels to the actin cytoskeleton (Kole et al., 2008). Consistent with this idea, much larger Na+ currents are observed in patch-clamp recordings from the AIS after chemical disruption of the actin cytoskeleton (Kole et al., 2008) and in recordings from axon blebs (Hu et al., 2009 and Schmidt-Hieber and Bischofberger, 2010). Axonal blebs are swellings where the axon has been cut at the surface of the brain slice and then sealed over and, therefore, presumably do not have an intact cytoskeleton (Hu et al., 2009). As they are larger than the Doxorubicin price axon they provide a more accessible location for making axonal recordings (Shu et al., 2006). In addition, disruption of myelination at the cut end allows one to record from myelinated axons at locations that would otherwise not

be possible. While recording from axon blebs has technical advantages, it should be recognized that they are damaged regions of the axon. As such, channel expression at these axon structures may not be representative of that in the intact axon. Despite this caveat, these recent functional estimates suggest the AIS Na+ channel density is indeed high (∼110 to 300 channels/μm, assuming a 17 pS single-channel conductance), giving a conductance density of 2,000 to 5,000 pS/μm2. For comparison, the Na+ channel density in the squid giant axon is around 1,200 pS/μm2 (Hodgkin and Huxley, 1952). While there

is now general Phosphoprotein phosphatase consensus that the density of Na+ channels is high in the AIS, whereas it is low in dendritic regions (Magee and Johnston, 1995 and Stuart and Sakmann, 1994), how the density of Na+ channels at the soma compares to that in the AIS is still debated. As mentioned above, recent electrophysiological estimates provide evidence that the density of Na+ channels at the AIS is much higher than at the soma (see Figure 2B). Consistent with this idea, Lorincz and Nusser (2010) using quantitative freeze-fracture immunogold labeling found that the number of Nav1.6 channels in the AIS of hippocampal pyramidal neurons was ∼40-fold higher than that found at the soma (Figure 2A2). In sharp contrast, a recent study using Na+ dye imaging together with modeling predicted that the difference in Na+ channel density between the AIS and the soma in cortical pyramidal neurons is only 3-fold (Fleidervish et al., 2010). Presumably, methodological differences underlie this apparent discrepancy. Immunocytochemical studies suffer from the fact that they do not provide information on functional channels, whereas channel density estimates based on Na+ imaging rely on accurate modeling of Na+ diffusion.

, 1997, Jones et al., 1997 and Kralic et al., 2002a). By contrast, deletion of the γ2 subunit results in only a modest reduction of GABA binding sites (−22%), and the γ2 subunit is therefore largely dispensable for assembly of α and β subunits (Günther et al., 1995). Intriguingly, a recent

study analyzing the expression of GABAARs in transfected human embryo kidney (HEK) cells suggests that GABA might act as an intracellular chaperone important for GABAAR biogenesis in the early secretory pathway (Eshaq et al., 2010). Consistent with such a function, the above-mentioned N-terminal assembly signals are located proximal learn more to the GABA- and benzodiazepine-binding sites of GABAARs (Boileau et al., 1999 and Teissére and Czajkowski, 2001). The importance of subunit N-terminal domains for receptor assembly in vivo is exemplified by a naturally occurring point mutation (R43Q) in the γ2 subunit that is associated with childhood absence epilepsy and febrile seizures (Wallace et al., 2001, Kang and Macdonald, 2004, Hales et al., 2005, Frugier et al., 2007 and Tan et al., 2007). Moreover, a small naturally occurring N-terminal deletion mutant of the rat α6 subunit abolishes assembly of corresponding

receptors (Korpi et al., 1994). The rules that govern differential assembly in cells that coexpress multiple GABAAR subtypes remain little explored, although some evidence indicates that assembly may be mass-driven by the rate of cotranslation of compatible subunits. Transgenic mice that express Selleckchem Dolutegravir ectopic α6 subunits in hippocampal pyramidal cells exhibit a gain of extrasynaptic α6βγ2 receptors at a cost of postsynaptic receptors (Wisden et al., 2002). Deletion of the α1 subunit Hydroxychloroquine in mice leads to compensatory

upregulation of receptors containing other α subunits (Sur et al., 2001, Kralic et al., 2002a, Kralic et al., 2002b and Kralic et al., 2006). Furthermore, a residue (R66) in the N-terminal domain of the α1 subunit is essential for assembly of α1β2 receptors but dispensable for formation of α1β1 and α1β3 complexes (Bollan et al., 2003b). Recent evidence further suggests that entry of transport competent GABAAR assemblies into the secretory pathway depends on subunit glycosylation (Tanaka et al., 2008 and Lo et al., 2010). The exit of GABAARs from the ER is limited by constitutive ER-associated degradation (ERAD) of α and β subunits (Gallagher et al., 2007, Saliba et al., 2007 and Bradley et al., 2008), suggesting that receptor assembly is relatively inefficient (Figure 2). ERAD of GABAARs is further enhanced by chronic blockade of neural activity (Saliba et al., 2009). Neural activity blockade-induced ubiquitination and degradation of GABAAR subunits involves reduced Ca2+ entry through voltage-gated Ca2+ channels (VGCCs).

The difference in exon numbering for SHANK2 in different organisms in other reports is likely due to the pattern of uncharacterized exons or alternative splicing ( Leblond et al., 2012; Lim et al., 1999; McWilliams et al., 2004). The deletion of exon 7 and exon 6-7 of Shank2a (exons 17 and exons 16–17 of predicted full-length Shank2) resulted in a frame shift of the open reading frame immediately after exon 7. Therefore, the molecular nature of these two targeted mutations is predicted to be very similar at protein level. Analyses of protein composition, synaptic development and function, and Everolimus mouse behaviors have revealed similarity but also significant differences

between these two lines of Shank2 mutant mice ( Table 3). Below, we utilize the exon 6–7 nomenclature based on numbering from promoter 2 of Shank2a/ProSAP1a. Full-length exon numbering is depicted in Figure 3B. Biochemically, protein composition at synapses was altered in both Shank2 Δex7 and Δex6–7 mice but with slight differences. In Shank2 Δex7−/− mice, GluN1 and GluN2B NMDA-type

glutamate receptors in hippocampus and GluN1, GluN2A, and GluA1 in striatum are increased ( Schmeisser et al., 2012). Interestingly, Shank3 was upregulated in striatum of Shank2 Δex7−/− mice. In Shank2 Δex 6-7−/− mice, reduction of phosphorylated CaMKIIα/β (T286), ERK1/2, p38, and GluA1 (S831/S845) CX-5461 in vitro was observed in hippocampus ( Won et al., Quetiapine 2012). Similar to Shank2 Δex7−/− mice ( Schmeisser et al., 2012), GluN1 is increased in the hippocampus of Shank2 Δex6–7−/− mice ( Won et al., 2012). Whereas baseline synaptic transmission was reduced in Shank2 Δex7−/− mice ( Schmeisser et al., 2012), normal synaptic transmission was observed in Shank2 Δex6–7−/− mice ( Won et al., 2012). mEPSCs recorded from CA1 hippocampal neurons were unaltered in Shank2 Δex6–7−/− mice, but reduced in Shank2 Δex7−/− mice. Interestingly, the ratio of NMDA/AMPA currents was reduced at CA1 synapses in Shank2 Δex6–7−/− mice

but increased at the same synapses of Shank2 Δex7−/− mice. NMDA receptor-dependent LTP in hippocampal CA1 synapses was increased and LTD was unaffected in Shank2 Δex7−/− mice. In contrast, both NMDA receptor-dependent LTP and LTD at CA1 synapses were reduced in Shank2 Δex6–7−/− mice. Behaviorally, hyperactivity, impaired social interaction, altered ultrasonic vocalizations, and increased self-grooming were observed in both Shank2 Δex6–7−/− and Shank2 Δex7−/− mice. Spatial learning and memory was impaired in Shank2 Δex6–7−/− but normal in Shank2 Δex7−/− mice. The basis for apparent discrepancies in synaptic physiology but similar behavioral profiles between Shank2 Δex6–7−/− and Shank2 Δex7−/− mice is not immediately clear and further investigation is warranted.

As one might imagine, this could be a serious challenge to calibrating voltage signals in small dendrites or dendritic spines, although researchers can use, and have used, the neuron’s own electrical signals, such as back-propagating action potentials, as internal standards for calibration (Nuriya et al., 2006). Finally, the relatively high speed of the electrical responses of mammalian neurons also generates a serious challenge for voltage measurements. While infinite temporal resolution would be welcome, in

practice most questions can be addressed with one CH5424802 mouse millisecond resolution. As we will discuss in the next section, there are a variety of chromophores with different response times; but unfortunately, the fastest ones normally provide the smallest signals,

which has been a long-standing problem in voltage imaging (Waggoner, 1979). The reader can appreciate from the previous list of problems that for effective voltage imaging one needs to solve some nontrivial challenges. At the same time, as mentioned, the electric field at the plasma membrane is very strong and can easily alter the physical, chemical, environmental, and spectral properties of any molecule located within it. This creates the potential to tap into a rich toolbox of different physicochemical principles

and harness them to measure changes in the electric field. As we will see, there is a great diversity of approaches this website that have achieved meaningful optical voltage measurements, a tribute to the determination and ingenuity of the scientists involved ( Cohen, 1989 and Cohen and Lesher, 1986). Most of the successful experiments with voltage imaging so far have been accomplished using single photon excitation with visible light, where the absorption cross-sections of the indicators are large. Also, some light sources (arc lamps, or now LEDs) can have very low noise, making it relatively easy to detect minute changes in signal, with ratiometric measurements at multiple absorption or emission wavelengths providing additional noise immunity and sensitivity ( Yuste et al., 1997 and Zhang Galactokinase et al., 1998). With typical light sources, wide field excitation is possible, and many photons can be collected from spatially extended areas, such as a section of dendrite, the entire soma, or many cells and their processes, increasing the integrated signal. But all of the typical problems of single-photon excitation apply—there is low penetration into scattering media like intact vertebrate brain tissue, and no native sectioning capability, requiring the use of confocal microscopes to afford cellular resolution.

Currents generated around AP threshold in CA3 neurons at −40 mV also increased following NO treatment or conditioning (Ctrl: −25 ± 80 pA, n = 9; NO: 377 ± 88 pA, n = 5; PC: 282 ± 145 pA, n = 5; p < 0.05), and this was suppressed by r-stromatoxin-1 (NO+Strtx: 82 ± 93 pA, n = 4) and by 7-NI treatment during conditioning (PC+7-NI: 107 ± 59 pA, n = 3), confirming a NO-dependent Kv2 current activation at potentials around AP threshold. Further evidence of the conductance

change was obtained by tail current measurements from the MNTB (Figures 3I and 3J) and CA3 (Figures 3K and 3L). Fit of a Boltzmann function showed that NO signaling (NO donor or PC) caused a marked leftward shift of the activation curve (V1/2) in neurons from both brain regions that was blocked by 7-NI or by glutamate receptor antagonism during the conditioning paradigm (Figures Gefitinib cost 3J and 3L). It is not possible to precisely equate half-activation voltages between recombinant and native K+ channels (because there are many unknowns in terms of heteromeric assembly, accessory proteins, and phosphorylation FG-4592 supplier states), but such a leftward shift is consistent with a reduced contribution from Kv3 channels that have

a more positive half-activation voltage (Hernández-Pineda et al., 1999, Kanemasa et al., 1995 and Rudy and McBain, 2001) than Kv2 channels (Guan et al., 2007, Johnston et al., 2008 and Kramer et al., 1998). Additional evidence Batroxobin for expression of Kv3 and Kv2.1 channels came from immunohistochemistry and qPCR experiments, showing Kv3.1b, Kv3.3, and Kv2.1 protein (Figures 4A, 4C, and 4D) and Kv3.1a, Kv3.1b, Kv3.2, and Kv3.3 mRNA (Figure 4B) in CA3 pyramidal cell bodies. We could not detect immunostaining (not shown) or substantial mRNA for Kv3.4. Together, these data confirmed that Kv3 channels are present in hippocampal CA3 pyramidal neurons as reported previously (Perney et al., 1992 and Weiser et al., 1994). We excluded significant contributions from Kv1, Kv4, and BK K+ channel families: Kv1 was routinely blocked with dendrotoxin-I (100 nM; data

not shown); Kv4 was inactivated by the conditioning voltage of the I/V protocol (Figure S2); and the NO-potentiated current was not a BK because this was TEA insensitive. We conclude that NO signaling mediates an activity-dependent adaptation in postsynaptic excitability by suppressing Kv3 and potentiating Kv2 currents in both the brain stem and hippocampus. These results suggest that neuronal delayed rectifiers are malleable; under low-activity conditions, Kv3 contributes to outward rectification, but during more active periods, Kv2 channels become dominant. This idea was tested in both MNTB and CA3 by examining the effect of TEA (1 mM) on AP waveforms under control conditions (before conditioning), on exposure to NO donors, or after synaptic conditioning (Figure 5).

This work was supported by the NSF (IOS 0542372, P.S.; DMR-0820492, D.K. [MRSEC program]), the HFSP (RGY0042- P.S.), the NIH (core grant P30 NS45713

to the Brandeis Biology Department; F31 DC011467, D.M.Z.; R00 GM87533, R.A.B.), the DGIST MIREBrain and Convergence Science Center (12-BD-0403) and Basic Science Research Program (2012009385) of the Ministry of Education, Science and Technology, Korea (K.K.), the Natural Sciences and Engineering Research Council of Canada (PGS-D3), and the Brandeis National Committee (S.J.N.), a gift from the Jensam Foundation (C.I.B.), and Olaparib datasheet the Howard Hughes Medical Institute (C.I.B.). C.I.B. is an Investigator of the Howard Hughes Medical Institute. Author contributions: H.J., K.K., S.J.N., and D.M.Z. performed the experiments; E.M., D.K. and R.B. provided reagents; Volasertib supplier H.J., K.K., C.I.B., and P.S. analyzed and interpreted data; C.I.B. and P.S. wrote

the manuscript. “
“In most species, males and females display sex-specific behavioral repertoires. Courtship and mating behaviors elicited by pheromones are among the most obvious sexually dimorphic repertoires because they are innate and stereotyped (Stowers and Logan, 2010). What are the neural differences that give rise to different behaviors in each sex? Behavioral differences could be due to differences in the ability of each sex to detect pheromone or to differences in the processing of pheromone sensory information. For example, female mice with an impaired vomeronasal organ exhibit male mating behaviors, suggesting that the underlying neural circuitry is the same in both sexes but only active in males (Kimchi et al., 2007). It may be that females Protein kinase N1 are capable of smelling pheromones that males cannot and that smelling these compounds represses male mating. In this case, the difference is at the level of detection. Alternatively, male flies detect

pheromone identically to females (Kurtovic et al., 2007) but possess male-specific ganglia that initiate male courtship behavior (Clyne and Miesenböck, 2008; Kohatsu et al., 2011), even in an animal that is otherwise female (Kimura et al., 2008). Here, both sexes smell the same compound, cis-vaccenyl acetate, but male and female higher brain centers generate different responses ( Kurtovic et al., 2007). Thus, in this case, the difference is at the level of processing. The two mechanisms are not mutually exclusive. In Manduca sexta, transplanting the nascent male sensory apparatus (his antennae) to a female larva induces male development in the female brain, and the adult animal has male behaviors ( Schneiderman et al., 1986). The reciprocal switch generates an animal that has female behaviors ( Kalberer et al., 2010). In this case, a difference in detection induces sexually dimorphic wiring, resulting in a difference in processing. Behavior that depends only on differences in detection could be easily modulated, for example, by regulating chemoreceptor expression.