Tyramine action on motoneuron excitability andadaptable tyramine/octopamine ratios adjustDrosophila locomotion to nutritional stateNatalie Schützlera, Chantal Girwerta, Isabell Hüglia, Giriram Mohanab, Jean-Yves Roignantb, Stefanie Ryglewskia,1,and Carsten Ducha,1

aInstitute of Developmental Biology and Neurobiology, Johannes Gutenberg University Mainz, 55099 Mainz, Germany; and bInstitute of Molecular Biology,Johannes Gutenberg University Mainz, 55099 Mainz, Germany

Edited by John G. Hildebrand, University of Arizona, Tucson, AZ, and approved January 9, 2019 (received for review August 7, 2018)

Adrenergic signaling profoundly modulates animal behavior. Forexample, the invertebrate counterpart of norepinephrine, octopamine,and its biological precursor and functional antagonist, tyramine,adjust motor behavior to different nutritional states. In Drosophilalarvae, food deprivation increases locomotor speed via octopamine-mediated structural plasticity of neuromuscular synapses, whereastyramine reduces locomotor speed, but the underlying cellular andmolecular mechanisms remain unknown. We show that tyramine isreleased into the CNS to reduce motoneuron intrinsic excitabilityand responses to excitatory cholinergic input, both by tyraminehonoka

receptor activation and by downstream decrease of L-type calciumcurrent. This central effect of tyramine on motoneurons is requiredfor the adaptive reduction of locomotor activity after feeding. Sim-ilarly, peripheral octopamine action on motoneurons has beenreported to be required for increasing locomotion upon starva-tion. We further show that the level of tyramine-β-hydroxylase(TBH), the enzyme that converts tyramine into octopamine inaminergic neurons, is increased by food deprivation, thus selectingbetween antagonistic amine actions on motoneurons. Therefore,octopamine and tyramine provide global but distinctly differentmechanisms to regulate motoneuron excitability and behavioralplasticity, and their antagonistic actions are balanced within adynamic range by nutritional effects on TBH.

biogenic amine | calcium channel | insect | Dmca1D | neuromodulation

Biogenic amines adjust the properties of single neurons, syn-apses, and neural circuits in multiple parts of the CNS (1) to

regulate arousal, mood, and other global brain states that pro-foundly influence behavior (2, 3). Given the multitude of aminetargets and effects, it is often difficult to predict the behavioralconsequences of altered amine signaling and vice versa to pin-point the sites and mechanisms of amine action which underliebehavioral adaptations. For example, norepinephrine (NE) andits invertebrate counterpart, the monoamine octopamine (OA)(4, 5), are potent modulators of arousal (4, 6), but they alsotarget specific neurons or microcircuits to affect cognitive andemotional behaviors (7–10), sensory processing (11), and motorbehavior (12).Vertebrate locomotion is enhanced by the release of NE and

5HT into spinal cord circuitry from descending brainstem neu-rons (12, 13). In invertebrates OA and its biological precursor,tyramine (TA), control locomotor activities such as crawling (14)and flight (15) in an antagonistic manner. OA enhances loco-motion during fight-or-flight reactions or states of hunger,whereas TA reduces locomotion during rest and digest. Para-doxically, OA and its precursor TA are both released from thesame tyrosine decarboxylase 2 (TDC2)-expressing neurons (16).In Drosophila larvae OA is released at the neuromuscular junc-tion (NMJ) during states of hunger and causes expansion ofneuromuscular terminals, which, in turn, is required for increasedlocomotion (17, 18). By contrast, feeding or increased TA signalingreduce locomotion (19). Therefore, OA and TA antagonistically

regulate locomotion in the context of nutritional state. How-ever, the signals that select between TA and OA release andthe mechanisms by which TA reduces locomotor activity areunknown.We show that TA release from TDC2 neurons into the CNS

reduces Drosophila larval motoneuron (MN) excitability viahonoka receptor activation and downstream regulation ofDmca1D L-type Ca2+ channels. This mechanism is required fordecreases in locomotor activity following feeding. Starvationincreases the levels of tyramine-beta-hydroxylase (TBH), theenzyme that converts TA into OA. This likely balances OA-mediated increases versus TA-mediated decreases in locomo-tor speed. Therefore, adaptive changes of locomotor activity tonutritional state are mediated by regulation of the OA/TA syn-thesis pathway and subsequent amine actions on MNs.

ResultsOA and TA Adjust Drosophila Larval Locomotion to Nutritional State.We first confirmed that OA enhances locomotion during statesof hunger, whereas TA reduces locomotion during satiation (14,18, 19). We quantified crawling distance and speed as in Wanget al. (20) from crawling traces of 2-min duration in Canton S(CS) third-instar larvae that were starved for 2 h, fed normally, orfed with TA-containing food (Fig. 1 A–C). Starvation significantly

Significance

Biogenic amines, like norepinephrine, can act as neuromodulatorsto render animal behavior adaptive to changing external and in-ternal conditions. The Drosophila genetic model system has beenused extensively to identify aminergic neurons which modulatecognitive, emotional, andmotor behaviors, but the cellular targetsand physiological consequences of aminergic modulation remainpoorly understood. We show that nutritional state affects aminebiosynthesis to tune the relative levels of octopamine and tyra-mine in central Drosophila neurons. Starvation increases octop-amine and locomotion, whereas satiation increases tyramine,which acts directly on motoneurons to decrease L-type calciumcurrent and excitability. Both effects are required to adjust loco-motor behavior to nutritional state. Therefore, adaptive behaviorcan be controlled on the level of motoneuron modulation andamine biosynthesis.

Author contributions: C.D. designed research; N.S., C.G., I.H., G.M., and S.R. performedresearch; S.R. and C.D. contributed new reagents/analytic tools; N.S., C.G., G.M., J.-Y.R.,S.R., and C.D. analyzed data; and S.R. and C.D. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Published under the PNAS license.1To whom correspondence may be addressed. Email: ryglewsk@uni-mainz.de or cduch@uni-mainz.de.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1813554116/-/DCSupplemental.

Published online February 11, 2019.

www.pnas.org/cgi/doi/10.1073/pnas.1813554116 PNAS | February 26, 2019 | vol. 116 | no. 9 | 3805–3810

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increased total crawling distance (Fig. 1B) and mean crawlingspeed (measured every second and averaged over 120 s) (Fig. 1C),whereas feeding and TA significantly decreased speed and dis-tance (Fig. 1 A–C). Accordingly, null mutants for TBH (21), theenzyme that generates OA from its biological precursor TA, lackOA but have increased TA levels and show markedly reducedlocomotor activity (Fig. 1 A–C).

TDC2 Neurons Contact MNs in Central Ventral Nerve Cord Neuropils.OA and its precursor, TA, are both released from TDC2-positiveventral unpaired median neurons (VUM neurons) (22). At theNMJ, OA and TA are contained in synaptic vesicles at type IIterminals of TDC2 neurons (16). We tested on the light micro-scopic level whether the central arbors of TDC2 neurons alsopossessed potential output synapses and whether these synapsescontacted the dendrites of larval crawling MNs. Triple immu-nolabeling in the larval ventral nerve cord (VNC) for anti-TDC2(Fig. 1 D and Dii, magenta), for UAS-GFP expression under thecontrol of eve-GAL4 (see SI Appendix, Supplementary Methods)in RP2 and aCC MNs (Fig. 1 D and Di, green), and for thepresynaptic active zone protein bruchpilot (Brp) by NC82 anti-body staining (Fig. 1 D and Diii, cyan) indicated overlap betweenarbors of OA/TA-containing neurons and MN dendrites (Fig. 1E and F, white arrowheads). Moreover, many of these contact

sites also showed active zone label in TDC2-positive boutonlikethickenings (Fig. 1 E and F, white arrowheads). This indicatedthat presynaptic release sites of TDC2 neurons contacted MNdendrites. Although laser scanning light microscopy does notprovide sufficient spatial resolution to prove synaptic contacts bycolocalization analysis, these data hint at OA/TA action on MNsvia central release from TDC2 neurons.

TA Decreases MN Excitability.We next tested for the central effectsof TA and OA on MNs. The firing responses of larval RP2 MNsto somatic current injections were recorded in current clampmode. RP2 responded to square-pulse current injection with acharacteristic biphasic depolarization and a marked delay to thefirst action potential, both of which were previously described(23–25). A representative example trace for current injection justabove firing threshold is shown before any pharmacologicalmanipulation (Fig. 2A), 2 min after bath application of TA (Fig.2Ai), and following 2-min washout in saline (Fig. 2Aii). Bathapplication of TA (10−5 M) had two reversible effects: a re-duction in firing frequency and an increase in the delay to thefirst action potential (Fig. 2 A–Aii). Quantification (21 animals)revealed that both the increased delay to firing (Fig. 2B) and thedecreased firing rate (Fig. 2C) were statistically significant andfully reversible. This was unlikely the result of changes in passivemembrane properties because firing threshold (Fig. 2D) andinput resistance (Fig. 2E) were not altered by TA. Moreover, thesame effects were observed following thermogenetically inducedTA release from TDC2 neurons with targeted expression ofUAS-TRPA1 channels (Fig. 2F).Since ramp current injections better mimic the gradual de-

polarizations that occur in response to synaptic input from themotor network, we repeated the experiments with ramp proto-cols (see SI Appendix, Supplementary Methods). Again, bath ap-plication of TA significantly increased the delay to the first spike(Fig. 2 G and H) and reduced MN firing rates (Fig. 2 G and I).To test whether TA acted directly on MN membrane properties,we repeated similar experiments in synaptic isolation intemperature-sensitive shibirets mutants. At nonpermissive temper-ature with chemical synaptic transmission blocked, TA bath ap-plication still reduced MN firing rates significantly and reversibly(Fig. 2J), thus ruling out indirect effects via chemical synaptictransmission. Although the effective TA concentration at the MNmembrane remained unclear, our data indicated a bell-shapeddose–response relationship. Bath application of 10−6 and 10−5 MTA reduced MN excitability significantly, but 10−4 M TA had nosignificant effect (Fig. 2K). In sum, our data show that TA releasefrom TDC2 neurons into the central motor neuropil significantlyreduces MN excitability. Since TDC2 neurons also release OA(18), we next tested bath application of OA (10−6 and 10−5), butOA had no effect on MN responses to current injection (SI Ap-pendix, Fig. S1).

TA Acts on MN Honoka Receptors and Dmca1D Ca2+ Current.We nextaimed at identifying the TA receptor and downstream ionicmechanisms. Action potential shape seemed unaltered in TA(Fig. 2L), indicating that fast Na+ and K+ channels were notaffected. Given that TA increased the delay to the first spike(Fig. 2 A and B) and this delay is caused by the Drosophila Kv4channel homolog, Shal (23, 24), we tested whether Shal RNAiknockdown affected TA effects on MNs. Although Shal knock-down in RP2 MNs effectively reduced the delay to the first spike,TA still increased the remaining delay and decreased firingfrequency significantly and reversibly (SI Appendix, Fig. S2).Therefore, normal levels of Shal expression were not requiredfor TA-mediated modulation of MN excitability. Moreover, bathapplication of TA (10−5 M) had no obvious effect on MN totaltransient or total sustained outward current and no significanteffect on the maximum amplitude of transient outward current (SIAppendix, Fig. S2).Since Dmca1D L-type-like Ca2+ channels have multiple roles

in shaping Drosophila larval MN excitability (26), we next tested

Fig. 1. OA/TA-containing neurons adjust locomotor activity and contact MNdendrites. (A) Representative traces of 2 min of crawling filmed at 4 frames/s(*, starting position) from CS larvae after 2 h of starvation (first trace),continuous feeding (second trace), and feeding of TA (third trace) and froma fed TBH mutant (TβHnM18) animal (fourth trace). Starvation significantlyincreases (B) crawling distance and (C) speed (dark gray bars). TA signifi-cantly reduces locomotor activity (magenta bars). TBH mutants with no OAbut increased TA exhibited highly significantly reduced locomotor activity(light gray bars). **P < 0.01; ***P < 0.001; ANOVA with Newman–Keuls posthoc testing. (D–Diii) Maximum projection images of triple labeling of aCCand RP2 MNs with GFP (w;P{eve-GAL4.RN2}P,P{UAS-mcd8-GFP.L}LL5/+;act>>GAL4 UAS-FLP/+) (Di, green), VUM neurons with anti-TDC2 (Dii, ma-genta), and the presynaptic active zone with anti-NC82 (Brp, Diii, cyan).Dotted white boxes indicate enlargements shown in E–Eiii (total z distance of5 μm). White arrowheads demark spots with overlap of all three labels. Singleoptical sections (z = 0.5 μm) from areas in dotted white boxes are enlarged inF–Fiii. Arrows mark VUM neuron central arbor varicosities which are in directcontact with MN dendrites and contain the presynaptic marker Brp.

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for effects of TA on MN Ca2+ channels. Focal pressure appli-cation of the nAChR agonist nicotine (10−5 M) onto MN den-drites with targeted GCaMP6m expression reliably evokeddendritic Ca2+ signals (Fig. 3A and ref. 26). Bath application ofTA (10−5 M) reduced MN dendritic Ca2+ responses significantly(Fig. 3 Ai and B) and reversibly (Fig. 3 Aii and B). Please note thatintratrain variation of Ca2+ response amplitudes (Fig. 3 A, C, andE) was likely caused by dilution of nicotine in the puffing electrodeduring the intertrain intervals because, often, the first puff of atrain yielded the smallest response amplitude (SI Appendix, Sup-plementary Methods). However, intratrain amplitude variationswere much smaller than the reduction in response amplitude byTA. In null mutants for the TA receptor, honoka (hono) (refs. 27and 28), focal nicotine application reliably evoked MN dendriticCa2+ responses (Fig. 3C) with an amplitude similar to that incontrols (Fig. 3A), but bath application of TA had no significanteffect on these signals (Fig. 3 Ci and D). Therefore, TA action onthe hono receptor likely modulates MN dendritic Ca2+ responsesto nAChR activation. The downstream target of hono could eitherbe nAChRs or voltage-gated Ca2+ channels. RNAi knockdown forDmca1D in MNs reduced the amplitude of MN Ca2+ responses tonicotine puffs (26). However, imaging at the root of the dendritesof RP2 still yielded reliable Ca2+ responses (Fig. 3E), but theywere not affected by TA (Fig. 3 Ei and F). These data indicate thatTA modulation of MN responses to nAChR activation requiresDmca1D channels downstream of hono.We next tested whether TA affected MN excitability via the

same mechanisms by recording MN responses to somatic currentinjection with and without TA in hono mutants, following honoRNAi, or Dmca1D RNAi in MNs (Fig. 3 G–M). RP2 firing re-sponses indicated that TA had no effect on MN excitability inhono mutants (Fig. 3G). Quantification revealed that TA-mediated

decreases in firing rate (Figs. 2 and 3H, Left) were abolished by bathapplication of the TA receptor blocker yohimbine (Fig. 3H, secondfrom Left), in honomutants (Fig. 3H, third from Left), and by honoRNAi in MNs (Fig. 3H, Right). Similarly, the increase in the delayto the first spike that was caused by TA in controls (Fig. 3I, Left)was not observed in honomutants, in yohimbine, or following honoRNAi in MNs (Fig. 3I). We judge unspecific effects of geneticbackground unlikely because the effects of hono knockout wererecapitulated by RNAi knockdown and pharmacology.Finally, targeted expression of Dmca1D RNAi in MNs elimi-

nated the effects of TA on MN excitability (Fig. 3 J–L). AlthoughDmca1D mediates Drosophila larval MN somatodendritic Ca2+ cur-rent (26, 29), we also tested the second Drosophila high-voltage–activated Ca2+ channel (cacophony or Dmca1A). However, RNAiknockdown of Dmca1A in MNs (30) did not abolish the effectsof TA on MN excitability (SI Appendix, Fig. S3). Therefore, MNresponses to both synaptic input and somatic current injectionare significantly extenuated by TA action on hono receptors anddownstream reduction of Dmca1D Ca2+ current, but cacophonychannels are not required. A direct demonstration of TA effects onMN Ca2+ current amplitude is complicated by space clamp prob-lems due to the dendritic localization of Dmca1D channels (Fig. 3)and by Ca2+ current rundown over time in somatic patch clamprecordings. However, comparing normal rundown in the absenceand in the presence of TA (10−5 M) indicated a small but significantreduction of L-type-like Ca2+ current by TA (SI Appendix, Fig. S4).

TA Actions on MNs Are Required for Locomotor Adaptations. OAmodulation of MN axon terminals at the NMJ is required forstarvation-induced increases in Drosophila larval locomotion(18). We found that OA’s functional antagonist, TA, decreasesMN excitability via direct actions on MNs in the CNS. We nexttested whether the central effects of TA were required for de-creases in locomotion as observed following feeding. Feeding CScontrol animals with TA for 2 h significantly reduced crawlingspeed (Fig. 4 A and B, Left bars) and distance (Fig. 4 A and C, Leftbars). Please note that linear locomotion of Drosophila larvae isinterrupted by pausing and searching, characterized by turns ofmore than 20° per second (20). Therefore, decreased average speedand distance could result from either slower linear locomotion,more pauses, or both. Although TA increased the mean turningangle (from 22.5° ± 9.9° to 26.3° ± 11.9°), the number of pauses(from 40 ± 17 to 44 ± 19), and the relative time spent pausing(from 33 ± 14% to 36 ± 14%) by ∼10%, these differences werestatistically not significant (Student’s t test, P > 0.05). Therefore,increases in the pausing and searching mode alone cannot explainthe observed decreases in crawling speed and distance. Extenuatingeffects of TA on larval locomotion were not observed in honomutants (Fig. 4 A–C, Middle bars) and, importantly, following ex-pression of hono RNAi in MNs only (Fig. 4 A–C, Right bars).Therefore, TA-mediated decreases in MN excitability were re-quired for TA-mediated adaptations of locomotor behavior. Con-versely, potential additional effects of TA on higher brain centerswere not sufficient to mediate adaptive changes in locomotion.

Starvation Affects the Levels of TBH Enzyme in VUM Neurons. OArelease from TDC2-expressing VUM neuron neuromuscularaxon terminals is required for starvation-induced increases inlocomotion (18). We show that TA release from TDC2 neuroncentral arbors is required for decreases of locomotion that occurafter feeding. Therefore, TDC2 neurons may release two differentneurotransmitters which modulate locomotion antagonistically.This seems contradictory because locomotion-promoting (pe-ripheral OA release) and locomotion-inhibiting (central TA re-lease) effects should countervail, unless TDC2 neurons releasedmore OA during states of starvation but more TA during satia-tion. Given that TA is the biological precursor of OA and isconverted to OA by TBH, we tested whether starvation affectedTBH protein and mRNA levels. First, coimmunolabeling showeda punctuated TBH label in somata (Fig. 4 D and Di) and alongcentral arbors of TDC2 neurons (Fig. 4 Dii–Div). Visual inspection

Fig. 2. TA reduces MN electrical excitability. (A–Aii) Representative responseof a RP2 MN in CS to square-pulse current injection of 80-pA amplitude before(A), 2 min after TA application (10−5 M) (Ai), and following 2-min washout insaline (Aii). TA significantly and reversibly increases the delay to the first spike(pulse onset to first spike) (B) and decreases firing rate (mean firing rate of re-sponse spike train) (C) but has no effects on firing threshold (D) or input re-sistance (E ) (n = 21). Thermogenetic activation of TA-containing neurons(w;tdc2-GAL4,UAS-TRPA1;+ at 30 °C, control at 20°) reversibly decreases RP2firing rate (F) (n = 9). (G–Gii) Representative responses of RP2 to ramp currentinjection of 100-pA (black) and 180-pA (gray) amplitude before (G), 2 min afterbath application of TA (10−5 M) (Gi), and following 2 min of washout (Gii). TAsignificantly and reversibly increases the delay to the fist spike (ramp onset tofirst spike) (H) and decreases firing rate (mean from first to last spike duringramp; n = 21) (I). These effects remained in the absence of chemical synaptictransmission in shits (w1118 shi1) animals at nonpermissive temperature (30 °C)(J). Dose–response tests revealed significant effects of TA at 10−6 and 10−5 Mbut not at 10−4 M (K). Action potential (AP) shape was not affected by TA(black trace before TA, red trace with 10−5 M TA) (L). **P < 0.01; ***P < 0.001;Kruskal–Wallis ANOVA with Mann–Whitney U test pairwise comparisons. AHP,after hyperpolarization; n.s., not significant; sal, saline; syn isol, synaptic isolation.

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indicated increased anti-TBH labeling intensity upon starvation(2 h, Fig. 4E) compared with stage-matched, fed controls (Fig. 4F).Given that TBH puncta were consistently brighter in VUM neu-ron central processes than in somata, both could be separated bythresholding (see SI Appendix, Supplementary Methods). Quanti-fication revealed a significant increase of more than 50% in TBHimmunolabel intensity in TDC2 neuron central arbors upon star-vation (Fig. 4G, n = 10). Similarly, TBH labeling intensity in VUMneuron somata was significantly increased by starvation (Fig. 4 Hand I). Quantitative RT-PCR revealed a significant increase inTbh mRNA levels by about 50% upon starvation (Fig. 4J, n = 8data points with 10 animals each). By contrast, mRNA levels ofcontrol proteins were not affected (Fig. 4J, ribosomal protein 49).

DiscussionCentral TA Release from TDC2 Neurons Regulates MN Excitability.Although TA acts as an independent transmitter (31) to affect

numerous behaviors in nematodes (32), insects (15, 33), andmammals (34), the cellular sites and molecular mechanisms ofTA action remain incompletely understood. We provide evi-dence that TA is released from central arbors of TDC2 neuronsinto motor neuropils of the Drosophila larval CNS to directlymodulate MN excitability. First, bath application of TA de-creases MN firing responses to somatic current injection and theamplitude of MN dendritic Ca2+ responses to focal nAChR ac-tivation. Second, these effects remain after systemic blockade ofsynaptic transmission. Third, at the level of confocal laser scanningmicroscopy, contact sites of TDC2 neuron central arbors with MNdendrites exhibit boutonlike varicosities with local expression ofpresynaptic markers, which are structurally reminiscent of type IIaxon terminals at the NMJ (16). Fourth, the effects of TA bathapplication can be reproduced by thermogenetic stimulation ofTDC2 neurons and blocked by the TA receptor antagonist yo-himbine. Hence, MNs are likely the direct cellular targets of TA

Fig. 3. TA action on MNs requires the honoka receptor and Dmca1D Ca2+ channels. (A–Aii) Ca2+ signals in RP2 MN dendrites of a representative controlanimal (w;OK371-GAL4/20xUAS-IVS-GCaMP6m attP40;+) in response to focal pressure application of nicotine (10−5 M) through a glass electrode placed intothe motor neuropil within ∼10 μm of GCaMP6-labeled MN dendrites. Nicotine was puffed in two trains each before (A), 2 min in TA (gray shaded area) (Ai),and following 2 min washout (Aii). Each train consisted of four to five consecutive puffs (see black arrows) with a 15-s interpuff interval. Intertrain intervalwas 1 min, and TA wash in and washout durations were 2 min each. Upper row shows original images, and Lower row shows changes in relative GCaMPfluorescence (ΔF/F) over time. (B) TA significantly and reversibly reduces dendritic Ca2+ responses. (C and D) TA has no effect on MN dendritic Ca2+ responses inhonoka mutants (w;OK371-GAL4/20xUAS-IVS-GCaMP6m attP40;oct-tyrRhono/oct-tyrRhono). (E and F) TA has no effect on RP2 Ca2+ responses to nicotine fol-lowing RNAi knockdown of Dmca1D in MNs (w;OK371-GAL4/20xUAS-IVS-GCaMP6m attP40;UAS-Dmca1D-RNAi HMS00294/+). (G and I) Firing responses of RP2 tosomatic ramp current injections in honoka mutants (+;;P{lwB}Oct-TyrRhono). (G) Recordings before (Upper trace), 2 min in TA (10−5 M) (Middle trace), and after 2-minwashout (Lower trace) indicate no differences. (H) Quantification shows that TA induced reductions in firing frequency in controls but not following bath applicationof the TA receptor blocker yohimbine or in honoka mutants or following hono RNAi in MNs (w*;OK371-GAL4/+;UAS-hono-RNAiJF02967attP2/+). (I) Similarly, TA in-creased the delay to the first spike in controls but not in the presence of yohimbine, in honoka mutants, or following honoka RNAi in MNs. (J–M) Firing responses ofRP2 to ramp current injections in control (J) compared with Dmca1D RNAi in MNs (w*;OK371-GAL4/+; UAS-Dmca1D-RNAiHMS00294attP2/+) (K). Representative currentclamp traces before TA (Upper traces in J and K) during TA (Middle traces in J and K), and after washing (Lower traces in J and K) indicate that TA effects are reducedby Dmca1D RNAi in MNs. Quantification shows that Dmca1D RNAi in MNs abolishes the effects of TA on firing rate (L) and on the delay to the first spike(M ). ***P < 0.001. n.s., not significant.

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release from TDC2 neuron central arbors (Fig. 4K). Similarly, inCaenorhabditis elegans head oscillations are regulated by the ac-tivity of tyraminergic ring inter/motor neurons which presumablymodulate the activity of MNs to head muscles (31). In the spinalcord, MN excitability is regulated by biogenic amine release fromdescending brainstem neurons (12). Therefore, aminergic modu-lation of MN excitability seems a principle conserved from wormsand flies to mammals.

Honoka Receptor Activation Decreases MN L-Type Ca2+ Current.Drosophila expresses three G protein-coupled TA receptors,TAR (CG7431), TAR II (CG16766), and honoka (CG7485).Our data suggest that honoka mediates TA effects on MN ex-citability. First, honoka is abundantly expressed in the DrosophilaCNS. Second, TAR II can likely be excluded because it is in-sensitive to yohimbine (35), which effectively blocked TA effects.Third, effects of TA on MNs are abolished in honoka mutantsand by honoka RNAi. Honoka was first described as an OAreceptor (27), but later studies revealed a significantly higherbinding affinity to TA (36–38).Although intermediate steps remain to be investigated, we

identified MN L-type Ca2+ current through Dmca1D channels asan essential downstream target. Following targeted RNAiknockdown of Dmca1D in MNs TA has no effect on MN excit-ability. While this does not exclude the possibility of additionalion channels as downstream targets of honoka, it shows that areduction of L-type Ca2+ current is required for the observed TAeffects. TA-mediated decreases of MN L-type Ca2+ current mayeffectively decrease locomotor activity because Ca2+ influxthrough axonal and dendritic Dmca1D channels cooperativelyincreases MN intrinsic excitability and MN responses to excit-atory cholinergic synaptic input (26). In sum, we identify MNs asa cellular target and L-type Ca2+ channels as a molecular targetof behaviorally relevant TA action. Given the importance ofbiogenic amines in the control of motor, mood, social, and cognitive

behaviors, the identification of cellular and molecular amine targetsis of broad significance.

TA Action on MNs Is Sufficient for Adaptive Locomotor Changes. OAand TA mediate adaptive changes of locomotor behavior in re-sponse to nutritional state (see Fig. 4K for summary). Increasesin locomotor speed upon food deprivation require OA releasefrom VUM neuron neuromuscular terminals, which induces theexpansion of type II aminergic and type I glutamatergic MN axonterminals (18). Since each VUM neuron innervates numerousmuscles, this mechanism may globally enhance glutamatergicneuromuscular transmission during food seeking (18). By con-trast, feeding or increased TA signaling reduces locomotion (19).Although the cellular targets of OA and of TA action are MNs,the subcellular sites and mechanisms of action are strikinglydifferent. OA acts on Octβ2Rs at neuromuscular axon terminalsand induces structural plasticity (Fig. 4K, Right, and ref. 18),whereas TA acts in the CNS to induce electrical plasticity viahonoka receptor activation and Ca2+ channel modulation inMNs (Fig. 4K, Left). Given the broad arborizations of TDC2neurons through motor neuropils, central TA release mayglobally decrease MN excitability. TA actions on MNs are re-quired for adaptive changes of locomotor behavior becausehonoka RNAi knockdown in MNs abolished the effects of TAon locomotion. Therefore, adaptive changes in locomotor be-havior to nutritional state require global aminergic modulationof MNs, with antagonistic effects of OA and TA taking placethrough distinctly different mechanisms at different cellular com-partments of MNs (Fig. 4K).

TA/OA Conversion in TDC2 Neurons Depends on Nutritional State. TAand OA are both produced and released from TDC2 neurons.One possibility to select between TA-mediated decreases andOA-mediated increases in locomotion would be a regulatorymechanism for selective TA release from central arbors versusOA release from neuromuscular terminals, but we have no

Fig. 4. TA effects on crawling require the hono receptor and Dmca1D channels in MNs, and TBH levels are increased by starvation. (A) Representative crawlingtraces indicate that TA decreases crawling distance in CS controls (Upper traces) but not in honoka mutants (+;;oct-tyrRhono/oct-tyrRhono; Middle traces), or fol-lowing honoka RNAi in MNs (w*;OK371-GAL4/+; UAS-hono-RNAi JF02967attP2/+; Lower traces). TA significantly decreases crawling speed (B) and distance (C) incontrols but not in honoka mutants or following honoka RNAi in MNs. ***P < 0.001; Student’s t test. (D) Double immunolabeling for GFP (green) andTBH (magenta) in animals expressing GFP in TDC2-positive neurons (w;TDC2-GAL4/10xUAS-IVS-mcd8::GFP;+). (Di) TBH-positive puncta in TDC2 neuron somata.(Dii–Div) Z-projections of 2-μm thickness at different VNC depths (ventral, medial, and dorsal neuropils) reveal TBH positive puncta in central processes of TDC2neurons. (E and F) Representative anti-TBH labeling in starved (E) and nonstarved (F) CS larvae. Starvation significantly increases TBH label in varicositylikeprocesses (G) (Student’s t test; **P < 0.01) and in TDC2 neuron somata (H and I) (Student’s t test; **P < 0.01). (J) qRT-PCR reveals a significant increase in TBHmRNAbut not of the housekeeping gene Rp49 following 2 h of starvation (**P < 0.01). (K) Sketch summary of the proposed OA/TA mechanisms that adjust locomotionto nutritional state. ICa2+, Ca2+ current; int., integrated; n.s., not significant; PNS, peripheral nervous system; SV, synaptic vesicle.

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evidence for a mechanism that restricts synaptic vesicle re-lease to distinct subneuronal compartments. By contrast, weshow that starvation induced up-regulation of the TBH enzyme atboth the mRNA and the protein levels. Since TBH is likely the rate-limiting step in OA synthesis (39), this may augment the conversionof TA into OA and thus promote locomotion (Fig. 4K). IncreasedOA/TA ratios have also been documented upon stress in Dro-sophila (16), cockroaches (40), and locusts (41). In locusts,descending OA/TA projection neurons synthesize OA onlyupon stressful stimuli (42). We suggest that direct effects ofnutritional state on the transcriptional levels of the TBH en-zyme balance the antagonistic OA/TA system to adaptivelyadjust locomotor behavior (Fig. 4K). It will be important totest whether our findings can be generalized to other biogenicamines which exert distinctly different functions but are pro-duced in a shared biosynthesis pathway.

Experimental ProceduresDrosophila melanogaster were reared in plastic vials at 25 °C and a 12-hlight/dark cycle on a standard cornmeal, glucose, yeast, and agar diet. Allexperiments were carried out on third-instar larvae. CS was used as the wild-type control and the GAL4-UAS system to express transgenes in subsets of MNs(26). See SI Appendix, Supplementary Methods for fly strains and details on elec-trophysiology, imaging, behavioral analysis, immunohistochemistry, and qRT-PCR.

Larval preparation, electrophysiology, and Ca2+ imaging were conductedas previously described. MN excitability was determined in current clampmode by somatic current injections. Ramp current injections of 400-ms du-ration were applied from −40 to +100 or +200 pA final amplitude, andsquare pulses were applied from −40 to +100 pA in 10-pA increments (26).Ca2+ currents were recorded in a voltage clamp as previously described (30).Transient and sustained K+ outward currents were separated as in Kadaset al. (25). Ca2+ imaging was conducted as in Ryglewski et al. (43). ΔF/F wascalculated by the formula [F(stim) − F(rest)]/F(rest).

Immunohistochemistry for TBH was conducted with a rat-anti-DrosophilaTBH antibody (a gift from Dr. Maria Monastirioti, IMBB, Crete, Greece) at1:1,000 in PBS-TritonX (0.1%) at 4° overnight after fixation of a freshly dis-sected specimen in Bouin’s solution (5% acetic acid, 9% formaldehyde, 0.9%picric acid) and washing in PBS eight times for 20 min and in PBS-TritonX(0.5%) and 10% normal goat serum four times for 30 min. Primary anti-GFPand anti-NC82 and secondary antibodies, mounting, and confocal micros-copy were conducted as previously described (26).

Crawling of individual third-instar larvae on a 0.7% agarose gel arena(diameter of 14.5 cm) was filmed with a Basler acA 2000-165 camera at4 frames/s. Crawling traces of 2-min duration were analyzed as in Wang et al.(20) with Tracker software (Open Source Physics, Version 4.95).

ACKNOWLEDGMENTS. We thank Dr. M. Monastirioti (Crete, Greece) for theTBH antibody. Support by the German Research Foundation (DFG) (Grant DU331/6-2 to C.D. and Grant RY 117/3-1 to S.R.) is gratefully acknowledged.

1. Marder E (2012) Neuromodulation of neuronal circuits: Back to the future. Neuron 76:1–11.

2. Marder E, O’Leary T, Shruti S (2014) Neuromodulation of circuits with variable pa-rameters: Single neurons and small circuits reveal principles of state-dependent androbust neuromodulation. Annu Rev Neurosci 37:329–346.

3. Lee SH, Dan Y (2012) Neuromodulation of brain states. Neuron 76:209–222.4. Roeder T (2005) Tyramine and octopamine: Ruling behavior and metabolism. Annu

Rev Entomol 50:447–477.5. Verlinden H, et al. (2010) The role of octopamine in locusts and other arthropods.

J Insect Physiol 56:854–867.6. Berridge CW, Page ME, Valentino RJ, Foote SL (1993) Effects of locus coeruleus in-

activation on EEG activity in neocortex and hippocampus. Neuroscience 55:381–393.7. Arnsten AF, Wang MJ, Paspalas CD (2012) Neuromodulation of thought: Flexibilities

and vulnerabilities in prefrontal cortical network synapses. Neuron 76:223–239.8. Burke CJ, et al. (2012) Layered reward signalling through octopamine and dopamine

in Drosophila. Nature 492:433–437.9. Yanowitch R, Coccaro EF (2011) The neurochemistry of human aggression. Adv Genet

75:151–169.10. Watanabe K, et al. (2017) A circuit node that integrates convergent input from

neuromodulatory and social behavior-promoting neurons to control aggression inDrosophila. Neuron 95:1112–1128.e7.

11. Suver MP, Mamiya A, Dickinson MH (2012) Octopamine neurons mediate flight-induced modulation of visual processing in Drosophila. Curr Biol 22:2294–2302.

12. Heckman CJ, Lee RH, Brownstone RM (2003) Hyperexcitable dendrites in motoneu-rons and their neuromodulatory control during motor behavior. Trends Neurosci 26:688–695.

13. Jordan LM, Liu J, Hedlund PB, Akay T, Pearson KG (2008) Descending command sys-tems for the initiation of locomotion in mammals. Brain Res Brain Res Rev 57:183–191.

14. Fox LE, Soll DR, Wu CF (2006) Coordination and modulation of locomotion patterngenerators in Drosophila larvae: Effects of altered biogenic amine levels by the ty-ramine beta hydroxlyase mutation. J Neurosci 26:1486–1498.

15. Brembs B, Christiansen F, Pflüger HJ, Duch C (2007) Flight initiation and mainte-nance deficits in flies with genetically altered biogenic amine levels. J Neurosci 27:11122–11131.

16. Stocker B, et al. (2018) Structural and molecular properties of insect type II motoraxon terminals. Front Syst Neurosci 12:5.

17. Koon AC, Budnik V (2012) Inhibitory control of synaptic and behavioral plasticity byoctopaminergic signaling. J Neurosci 32:6312–6322.

18. Koon AC, et al. (2011) Autoregulatory and paracrine control of synaptic and behav-ioral plasticity by octopaminergic signaling. Nat Neurosci 14:190–199.

19. Saraswati S, Fox LE, Soll DR, Wu CF (2004) Tyramine and octopamine have oppositeeffects on the locomotion of Drosophila larvae. J Neurobiol 58:425–441.

20. Wang JW, et al. (1997) Morphometric description of the wandering behavior inDrosophila larvae: Aberrant locomotion in Na+ and K+ channel mutants revealed bycomputer-assisted motion analysis. J Neurogenet 11:231–254.

21. Monastirioti M, Linn CE, Jr, White K (1996) Characterization of Drosophila TBH geneand isolation of mutant flies lacking octopamine. J Neurosci 16:3900–3911.

22. Selcho M, Pauls D, El Jundi B, Stocker RF, Thum AS (2012) The role of octopamine andtyramine in Drosophila larval locomotion. J Comp Neurol 520:3764–3785.

23. Schaefer JE, Worrell JW, Levine RB (2010) Role of intrinsic properties in Drosophilamotoneuron recruitment during fictive crawling. J Neurophysiol 104:1257–1266.

24. Ping Y, et al. (2011) Shal/K(v)4 channels are required for maintaining excitability

during repetitive firing and normal locomotion in Drosophila. PLoS One 6:e16043.25. Kadas D, Ryglewski S, Duch C (2015) Transient BK outward current enhances moto-

neurone firing rates during Drosophila larval locomotion. J Physiol 593:4871–4888.26. Kadas D, et al. (2017) Dendritic and axonal L-type calcium channels cooperate to

enhance motoneuron firing output during Drosophila larval locomotion. J Neurosci

37:10971–10982.27. Arakawa S, et al. (1990) Cloning, localization, and permanent expression of a Dro-

sophila octopamine receptor. Neuron 4:343–354.28. Kutsukake M, Komatsu A, Yamamoto D, Ishiwa-Chigusa S (2000) A tyramine receptor

gene mutation causes a defective olfactory behavior in Drosophila melanogaster.

Gene 245:31–42.29. Worrell JW, Levine RB (2008) Characterization of voltage-dependent Ca2+ currents in

identified Drosophila motoneurons in situ. J Neurophysiol 100:868–878.30. Ryglewski S, Lance K, Levine RB, Duch C (2012) Ca(v)2 channels mediate low and high

voltage-activated calcium currents in Drosophila motoneurons. J Physiol 590:809–825.31. Alkema MJ, Hunter-Ensor M, Ringstad N, Horvitz HR (2005) Tyramine functions in-

dependently of octopamine in the Caenorhabditis elegans nervous system. Neuron46:247–260.

32. Pirri JK, McPherson AD, Donnelly JL, Francis MM, Alkema MJ (2009) A tyramine-gatedchloride channel coordinates distinct motor programs of a Caenorhabditis elegans

escape response. Neuron 62:526–538.33. Huang J, Liu W, Qi YX, Luo J, Montell C (2016) Neuromodulation of courtship drive

through tyramine-responsive neurons in the Drosophila brain. Curr Biol 26:

2246–2256.34. Rutigliano G, Accorroni A, Zucchi R (2018) The case for TAAR1 as a modulator of

central nervous system function. Front Pharmacol 8:987.35. Bayliss A, Roselli G, Evans PD (2013) A comparison of the signalling properties of two

tyramine receptors from Drosophila. J Neurochem 125:37–48.36. Reale V, Hannan F, Midgley JM, Evans PD (1997) The expression of a cloned Dro-

sophila octopamine/tyramine receptor in Xenopus oocytes. Brain Res 769:309–320.37. Robb S, et al. (1994) Agonist-specific coupling of a cloned Drosophila octopamine/

tyramine receptor to multiple second messenger systems. EMBO J 13:1325–1330.38. Saudou F, Amlaiky N, Plassat JL, Borrelli E, Hen R (1990) Cloning and characterization

of a Drosophila tyramine receptor. EMBO J 9:3611–3617.39. Homberg U, Seyfarth J, Binkle U, Monastirioti M, Alkema MJ (2013) Identification of

distinct tyraminergic and octopaminergic neurons innervating the central complex ofthe desert locust, Schistocerca gregaria. J Comp Neurol 521:2025–2041.

40. Châtel A, et al. (2013) Characterization of tyramine β-hydroxylase, an enzyme upre-gulated by stress in Periplaneta americana. J Mol Endocrinol 50:91–102.

41. Kononenko NL, Wolfenberg H, Pflüger HJ (2009) Tyramine as an independenttransmitter and a precursor of octopamine in the locust central nervous system: Animmunocytochemical study. J Comp Neurol 512:433–452.

42. Kononenko NL, et al. (November 16, 2018) A population of descending tyraminergic/octopaminergic projection neurons of the insect deutocerebrum. J Comp Neurol,10.1002/cne.24583.

43. Ryglewski S, Vonhoff F, Scheckel K, Duch C (2017) Intra-neuronal competition forsynaptic partners conserves the amount of dendritic building material. Neuron 93:632–645.e6.

3810 | www.pnas.org/cgi/doi/10.1073/pnas.1813554116 Schützler et al.

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