Defecation allows the body to eliminate waste, an essential step in food processing for animal survival. In contrast to the extensive studies of feeding, its obligate counterpart, defecation, has received much less attention until recently. In this study, we report our characterizations of the defecation behavior of Drosophila larvae and its neural basis. Drosophila larvae display defecation cycles of stereotypic frequency, involving sequential contraction of hindgut and anal sphincter. The defecation behavior requires two groups of motor neurons that innervate hindgut and anal sphincter, respectively, and can excite gut muscles directly. These two groups of motor neurons fire sequentially with the same periodicity as the defecation behavior, as revealed by in vivo Ca2+ imaging. Moreover, we identified a single mechanosensitive sensory neuron that innervates the anal slit and senses the opening of the intestine terminus. This anus sensory neuron relies on the TRP channel NOMPC but not on INACTIVE, NANCHUNG, or PIEZO for mechanotransduction.https://doi.org/10.7554/eLife.03293.001
In animals, the final stage in the digestion of food is the removal of waste from the body. Until recently, however, defecation has received less attention than other aspects of digestion such as feeding behavior and nutrition.
Fruit flies, also known as Drosophila, are commonly used in research as a model of animal biology. Food is moved through the digestive tract of fruit fly larvae when the muscles that circle the wall of the intestine contract. This process continues until the waste reaches the anus and is expelled from the body.
Now Zhang et al. have found that when fruit fly larvae defecate, the muscles at the end of the intestine contract just before the muscles in the anus contract. The nervous system controls these muscles via sequential firing of two sets of nerve cells that connect to the intestine and anus muscles, respectively.
Zhang et al. also identified a nerve cell that can sense when the anus is opened and relay this information back to the nervous system. The nerve cell is activated when stretched by the opening of the anus in a process that requires a protein called NOMPC.
Problems with defecation can lead to constipation and other diseases. For example, Hirschsprung's disease—a birth defect that affects one in 4000—is caused by abnormal development of the nerve cells that control muscles in the gut. Experiments on fruit flies could help us to understand how defecation works in humans and to develop new treatments for disease.https://doi.org/10.7554/eLife.03293.002
Defecation is important for food processing that provides nourishment to the animal. It eliminates waste (feces) from the digestive tract via the anus (Thomas, 1990; Heaton et al., 1992; Lembo and Camilleri, 2003), an unglamorous but essential body function. Compared to the extensively studied feeding behavior, defecation has received relatively little attention. Malfunction of defecation can lead to constipation and other diseases (Lembo and Camilleri, 2003), and abnormal development of neural circuits governing defecation may underlie birth defects such as Hirschsprung's disease due to elimination of intestinal ganglion cells required for bowel peristalsis (Romeo et al., 1994; Passarge, 2002), one of the major birth defects of the digestive system afflicting one in 4000 of the population.
Drosophila larvae provide a useful model system for the studies of feeding behavior and nutrition intake (Ikeya et al., 2002; Rulifson et al., 2002; Hwangbo et al., 2004; Bader et al., 2007). With an array of feeding assays and powerful genetic tools, these animals have yielded valuable information regarding the basis of the feeding behavior (Shen, 2012; Zhao and Campos, 2012; Bhatt and Neckameyer, 2013). However, modulation of defecation behaviors has received much less attention until recently (Edgecomb et al., 1994; Cognigni et al., 2011). Harnessing the experimental resources of this model system for the study of gut movements and the underlying neural basis should also help us understand the mechanisms of the defecation behavior.
In the larval intestines, peristaltic movements of the digestive tract push food from the anterior towards the posterior end. The rate of flow depends on various signals from gut cells and associated neurons (Benoit and Tracy, 2008; Schoofs et al., 2009). In Caenorhabditis elegans two groups of excitatory GABAergic motor neurons have been identified with partially redundant functions in activating enteric muscle cells (EMCs) (McIntire et al., 1993). Little is known about the motor control of gut movements in Drosophila larvae or any involvement of sensory neurons for defecation.
Mechanosensation is essential for many activities of Drosophila. Studies in adult flies have demonstrated that internal sensory neurons are important in regulating behaviors such as feeding, defecation, and egg laying (Yang et al., 2009). Whereas recent studies have identified mechanosensitive channels in specific sensory neurons in the larval body wall for harsh or gentle touch (Kim et al., 2012; Yan et al., 2013), whether and how a larva senses stretches of its internal organs is unknown nor have the neurons and channels mediating such mechanosensation been identified.
In this study we establish Drosophila larvae as a model system to study defecation behavior by performing studies of larvae 96 hr after egg laying (AEL). First, we show that Drosophila larvae exhibit rhythmic cycles of sequential contractions of the hindgut and the anal sphincter to expel feces. Second, we identify the motor neurons that innervate the hindgut and anal sphincter and show that these two groups of neurons fire sequentially with the same periodicity as the defecation cycle. Unexpectedly, we found that a single sensory neuron innervates the anal slit to sense its opening. Finally, we show that the TRP channel NOMPC but not other known mechanosensitive channels in Drosophila is required for the mechanosensation of this anus sensory neuron.
The Drosophila larval hindgut is the last part of the intestine, posterior to the Malpighian tubule, on the dorsal side under larval cuticle. At the posterior end of the hindgut is anal sphincter, which has a layer of thick sphincter muscles and a much narrower canal (Figure 1A, Figure1—figure supplement 1) (Murakami and Shiotsuki, 2001). Because the Drosophila larval body wall is transparent, contractions of the hindgut and anal sphincter can be monitored in vivo. Fluorescent markers, expressed with a hindgut-specific byn-Gal4 (Johansen et al., 2003), allowed visualization of contractions of the larval hindgut and anus sphincter in whole-mount of living larvae (Figure 1B). The defecation behavior consists of sequential contractions of the posterior hindgut and anal sphincter in a very stereotypical manner (Figure 1B,D), leading to opening of the anal slit to expel feces out of the lumen. This defecation process is repeated every 38 s at 25°C (Figure 1D). To demonstrate those gut movements triggered defecation, we fed the larvae with yeast laced with blue food dye and video taped their defecation cycle. As shown in Figure 1C and Video 1, each sequential contraction of hindgut and anus sphincter triggered a defecation cycle to expel feces out of the body.
To investigate the neural basis for the gut movements, we searched for the neuronal innervation of the hindgut and anal sphincter muscles. Since the axons that innervate the hindgut are from the most posterior pair of the axon bundles in the ventral nerve cord (VNC), the cell bodies of the neurons that innervate the hindgut and anal sphincter are most likely in the terminal segments of the VNC. We identified two groups of neurons, labeled by PDF-Gal4 and HGN1-Gal4 (Nassel et al., 1993; Edgecomb et al., 1994; Renn et al., 1999; Landgraf et al., 2003; Cognigni et al., 2011), which innervate the posterior hindgut and anal sphincter, respectively (Figure 1E–G). These neurons have their cell bodies in the terminal segments of VNC (Figure 1E) and send their axons along the midline of the ventral body wall to the posterior end of the larva, where they enter the hindgut. Within the hindgut, the HGN1 axons extend posteriorly to the anus sphincter surface to form dense arborizations over the muscles, while the PDF axons arborize over the posterior two-third of the hindgut with refined branches (Figure 1F–H). The PDF and HGN1 neurons are glutamatergic, as they could be labeled with antibody staining against Drosophila vGlut (Daniels et al., 2008) (Figure 1K,L). The axonal branches of PDF neurons on the hindgut can also be labeled with vGlut-Gal4 (Mahr and Aberle, 2006) (Figure 1—figure supplement 2), indicating that they are likely glutamatergic motor neurons. The labeled neurons in both cases have their axon terminals in close proximity of the gut muscles and form abundant bouton structures (Figure 1I,J). These results suggest that PDF and HGN1 neurons, which are likely motor neurons, might play a role in regulating hindgut contractions (Figure 1M).
In order to explore the functional connection between HGN1 neurons and anal sphincter muscles, we expressed Channlerhodopsin-2 (ChR2), a light activated cation channel, in the HGN1 neurons and recorded the excitatory junction potentials (EJPs) in the gut muscles before and after activating ChR2 by light. The gut muscles received tonic excitatory inputs (Figure 2A). Due to the fillet recording methods we used to gain access of anus sphincter muscles, this firing pattern might differ from those in intact animals. Illumination of the larval VNC with blue light caused a dramatic increase of EJPs in the anus sphincter muscles in the larva with ChR2 expression in the HGN1 neurons but not in the control animals (Figure 2B,C), providing evidence for HGN1 innervation of sphincter muscles. The PDF neurons have been previously shown to promote visceral muscle contractions (Talsma et al., 2012). Activation of PDF neurons expressing ChR2 with blue light also triggered a dramatic increase of EJPs in the anus sphincter muscles (Figure 2B,C), indicating that PDF neurons also play a role in regulating anal sphincter contractions, although PDF neurons do not directly form synapse with these muscles. This light-induced activation was absent in UAS or Gal4 control larvae and dependent on retinal, which is the chromosphere for ChR2 channels (Figure 2B,C).
To test whether direct activation of PDF or HGN1 neurons could trigger gut muscle contraction, we first expressed ChR2 in the PDF neurons and use a Minos insertion line to label the hindgut. Because the blue light used to excite GFP in the hindgut can also activate ChR2, we could activate the neurons expressing ChR2 and monitor the gut movements at the same time. Both the hindgut and anal sphincter contracted strongly (Figure 2D) upon stimulation. In contrast, with ChR2 expression in the HGN1 neurons, the anal sphincter but not hindgut contracted upon blue light stimulation of HGN1 neurons (Figure 2D and Video 2). The contractions were absent in larvae with only UAS-ChR2 or larvae fed with regular food without retinal.
To test whether PDF and HGN1 neurons are important for the normal defecation behaviors, we expressed Kir2.1 in these neurons to inhibit their activities. Silencing PDF neurons caused the interval of the defecation to increase from 38 s to 94 s (Figure 3A). Silencing the HGN1 neurons did not significantly alter the interval of anus sphincter opening (Figure 3A). Conceivably the peristaltic hindgut movements driven by PDF neuronal activity could have generated sufficient pressure to force open the anus sphincter. Indeed, silencing both the PDF neurons and HGN1 neurons caused the larva to display barely any hindgut movement over 5 min thus rendering it difficult to estimate the defecation interval, in contrast to the nearly eight cycles of contraction over 5 min—corresponding to a defecation interval of 38 s—in control animals (Figure 3B). These results suggest that the PDF neurons and HGN1 neurons are required for the normal defecation behavior.
To test whether the periodic defecation cycle of Drosophila larvae is associated with rhythmic firing of the PDF and HGN1 neurons innervating the hindgut and anal sphincter, we performed in vivo whole-mount Ca2+ imaging by monitoring the neuronal activity with a genetically encoded Ca2+ indicator GCaMP5 (Tian et al., 2009) driven by neuronal-specific Gal4s. Indeed, the PDF neurons displayed a stereotypical periodic firing pattern as revealed by Ca2+ elevation in both soma and dendrite area (Figure 4A,C). The average interval between peak Ca2+ signals is 38 s, which is highly consistent with the temporal pattern of contraction of the hindgut (Figure 1D and Figure 4M). We also monitored the Ca2+ signals of the HGN1 neurons and found that the HGN1 neurons also exhibited oscillation of Ca2+ levels with a periodicity of 38 s (Figure 4B,D,M, Video 3). Furthermore, Ca2+ signals of HGN1 neurons spread from dendrites to soma with a fixed latency (Video 3), indicating that the HGN1 neurons may receive excitatory inputs with a periodicity of 38 s.
To further investigate the properties of HGN1 neurons, we performed Ca2+ imaging and whole-cell patch clamp recording of these neurons in dissected VNC. HGN1 neurons exhibited periodic Ca2+ activities though with less regularity in this isolated preparation (Figure 4—figure supplement 1A,B). They also displayed clusters of EPSCs (Figure 4—figure supplement 1C), indicating that they received excitatory input from upstream neurons. Whole-cell patch recording of HGN1 neurons revealed that they fired bursts of action potentials (Figure 4—figure supplement 1D), similar to what was seen in other Drosophila larval motor neurons (Cattaert and Birman, 2001; Fox et al., 2006; Imlach et al., 2012).
The arborizations of PDF neurons and HGN1 neurons overlap extensively along the midline of the posterior VNC (Figure 4E,F and Video 4), raising the question whether they interact with each other. Green fluorescent protein reconstitution across synaptic partners (GRASP) has been developed as a technique to indicate synaptic connection of two neurons, each expressing one component of the split GFP (Feinberg et al., 2008; Gordon and Scott, 2009; Gong et al., 2010; Han et al., 2012). We expressed the two GFP components separately in PDF neurons and HGN1 neurons by using two different binary expression systems, PDF-LexA and HGN1-Gal4. An intense GFP signal was observed in the area where the processes of these two groups of neurons overlap (Figure 4G), while neither PDF neuron nor HGN1 neuron expressing one part of the split GFP of GRASP generated any fluorescent signals by itself (Figure 4—figure supplement 2), suggesting that a functional connection might exist between PDF and HGN1 neurons. By labeling the PDF neurons simultaneously with the dendritic RFP marker DenMark and the axonal GFP marker sytGFP, we found that the PDF neurons send their axons to the area where their processes overlap with HGN1 neuron dendrites (Figure 4—figure supplement 3).
To test whether the sequential contractions of the hindgut and the anal sphincter are associated with sequential firings of the PDF neurons and HGN1 neurons, we employed two Gal4 drivers to express GCaMP5 in both PDF neurons and HGN1 neurons at the same time. The cell bodies of PDF neurons are near the ventral surface of the VNC, while the cell bodies of HGN1 neurons are more dorsal and posterior (Figure 4E), making it possible to distinguish the two groups of cell bodies when monitored laterally (Figure 4E). By monitoring the Ca2+ signals in PDF neurons and HGN1 neurons simultaneously, we found that the Ca2+ level began to rise in PDF neuron cell bodies and then spread to the area occupied by arborizations of both groups of neurons, followed by Ca2+ elevation in the cell bodies of HGN1 neurons with a very short delay, indicating that these two groups of neurons have coordinated firing patterns (Figure 4H–L and Video 5).
Next, we asked whether there are sensory neurons for sensing movements of the gut or anus. The dendritic arborization (da) neurons are primary sensory neurons, which cover the entire body wall of a Drosophila larva. They are important for sensing chemical, thermal, light, and mechanical stimulations (Tracey et al., 2003; Xiang et al., 2010; Kim et al., 2012; Yan et al., 2013). In the vicinity of the anal slit, we found a specialized PPK-Gal4-labeled neuron. The cell body of this neuron resides on the anterior side of the anal slit, and its dendritic arbors surround the entire anal slit (Figure 5A). The majority of its dendrites forms a thin layer of arbors and covers the body wall around the anal slit (Figure 5A). There are also some arborizations extending along the anus sphincter (Figure 5A). The axon of this sensory neuron joins the nerve bundle that includes the axons of other da neurons and projects to the terminal segment of the VNC.
This single anus sensory neuron (ASN) has its cell body stochastically located on one side of the midline but its dendrites are highly symmetric. When the anal slit opens, the dendrites are dramatically stretched (Figure 5B). To investigate whether this stretch could activate the ASN, we carried out in vivo imaging of the ASN Ca2+ level with GCaMP5. We found that opening of the anal slit is accompanied with Ca2+ elevation in both dendrites and soma of this neuron (Figure 5C,D and Video 6). The periodicity of this Ca2+ response is similar to that of the movements of the anus and the oscillations of PDF neurons and HGN1 neurons in the VNC (Figure 5E). This result indicates that the ASN is mechanosensitive and participates in sensing the radial stretch caused by opening the anus.
We next tested whether the activation of the ASN could affect the firing patterns of motor neurons in the VNC. This sensory neuron projects to the terminal segments of the VNC and its axon terminals overlap with the area occupied by the dendrites of the HGN1 neurons and PDF neurons (Figure 5F). To investigate whether there might be direct synaptic contact between these neurons, we carried out GRASP analysis by expressing components of the split GFP in PDF neurons and the ASN. We found that there is very strong GRASP signal at the site where ASN axons and PDF dendrites overlap (Figure 5G), while neither PDF neurons nor ASN expressing one part of the split GFP of GRASP generated any fluorescent signals (Figure 5—figure supplement 1). Interestingly, the GRASP signal is asymmetric, in concordance with the localization of the ASN cell body to one side of the midline leading to a more intense axonal projection to the ipsilateral VNC, which is also evident with asymmetric Ca2+ elevation of the ASN axon terminals in the VNC (Video 7).
To further determine whether there is feedback from the ASN to activate the motor neurons in the VNC, we employed PPK-Gal4 to drive expression of ChR2 in the ASN so as to activate this neuron by blue light and monitored the EJPs of the gut muscles innervated by these motor neurons. Activation of ASN via blue light illumination induced large increases of EJPs in the majority of the anus sphincter muscles (Figure 5H,I).
To confirm that ASN rather than other PPK-Gal4-labelled neurons provides the feedback, we imaged PDF neurons and HGN1 neurons with GCaMP5 while inserting a tapered glass probe and advancing it to split open the anus sphincter, in a manipulation that mimicked the anus opening during defecation. We found that both PDF and HGN1 neurons responded to this local stimulation (Figure 5J). Stimulation of the anus with this glass probe induced an asymmetric Ca2+ increase in the VNC that is consistent with the asymmetric projection of ASN axons (Figure 5—figure supplement 2).
To study the functional importance of the ASN feedback to motor neurons, we used 2-photon laser to ablate the cell body of ASN at 48 hr after egg laying (AEL). The ASN was completely abolished 48 hr after laser ablation, while the other PPK neurons remained intact (Figure 5K). We then imaged the cell bodies of HGN1 neurons in the VNC and found abnormality of their rhythmic firing pattern, which displayed a much longer interval compared to control animals (Figure 5L).
These results suggest that the central neurons in the VNC receive excitatory feedback from ASN to increase motor neuron firing. Together with the GRASP analysis revealing the physical proximity of the processes of ASN and PDF neurons, our results indicate that the sensory neuron in the anus might sense the stretch when the anal slit is open and respond by provide feedback modulation of the PDF neurons and HGN1 neurons in the central nervous system.
To search for the putative mechanotransduction channel in the ASN, we examined the defecation behavior of several mechanotransduction channel mutants. We found that the defecation rhythm remains normal in the iav, nan, and piezo mutants. However, the defecation interval in the nompC mutants was significantly increased (Figure 6A). NOMPC is a TRPN channel essential for adult hearing and larval gentle touch (Gong et al., 2004; Yan et al., 2013). We found that NOMPC is highly expressed all over the dendrites and soma of the ASN, as revealed by staining with antibody against the NOMPC protein (Figure 6—figure supplement 1), raising the possibility that it might play a role in ASN sensing of radial stretch. Indeed, we found that the Ca2+ response is absent in the ASN of nompC null mutant larvae (Figure 6B,C, Video 8). This defect could be rescued by expressing wild-type NOMPC in the ASN with PPK-Gal4 (Figure 6B,C, Video 9). Our study thus identified a new role of the NOMPC channel, namely for sensing redial stretch of the intestinal terminus. NANCHUNG (NAN) and INACTIVE (IAV), two other TRP channels that often work in concert with NOMPC in other sensory neurons, are thought to form heterodimers and function in the mechanotransduction in Drosophila (Gong et al., 2004). We also tested the role of IAV in the ASN's stretch sensing. We found the ASN of iav1 mutant larvae exhibited Ca2+ response comparable to that in the wild-type larvae (Figure 6C and Video 10), suggesting that IAV is not required for the mechanotransduction of ASN.
This study establishes the Drosophila larva as a model system for studying the defecation behavior. We found that Drosophila larvae exhibit periodic defecation cycles, involving sequential contractions of the hindgut and the anal sphincter. We also found two groups of neurons, which innervate the hindgut and anal sphincter respectively, and can excite the hindgut and anal sphincter muscle in a sequential manner. In addition, we found a single sensory neuron that could sense the opening of the anal slit and send feedback to the motor neurons (Figure 7). Studies of C. elegans as a model system have investigated the defecation circuit (Thomas, 1990; Avery and Thomas, 1997; Branicky and Hekimi, 2006; Kwan et al., 2008). Studies of the adult fly have identified neurons regulating defecation behaviors subject to dietary and reproductive modulation (Cognigni et al., 2011). In this study of the defecation behavior in Drosophila larvae, we have identified not only the motor neurons innervating gut muscles but also a sensory neuron strategically located to sense radial stretch during defecation and provide feedback to the central nervous system.
Previous studies of the defecation behaviors of the adult fly (Cognigni et al., 2011) have revealed that its defecation rate is regulated by both the internal state and environment, rather than showing a robust rhythm. However, at the larval stage, the motor neurons and gut muscles as well as the sensory neuron responding to anus movement, all show very robust rhythmic activities. Given that feeding and defecation are dominant behaviors for third-instar larvae, conceivably robust rhythmic feeding and defecation behaviors may facilitate their nutrition intake and waste expulsion. In contrast, adult flies will likely encounter more complex environments and may need to conduct their defection behaviors in a more controllable manner.
Mechanosensation serves a number of important physiological functions in Drosophila larvae. The radial stretch sensation is a special type of mechanosensation essential for the function of many organs with luminal structures such as the digestive system and the blood vessels. However, how the organs sense radial stretch remains unclear.
We have identified a sensory neuron that can sense radial stretch with its highly specialized morphology in Drosophila larvae. In addition, we found that the TRP channel NOMPC but not other TRP channels tested, such as IAV that is often associated with NOMPC function, is required for normal ASN mechanotransduction. Interestingly, the ASN could be labeled by both class III da neuronal marker and class IV da neuronal maker, raising the question whether it might have the dual functions to sense different stimuli. The ASN may provide a neuronal model to study the distinct and cooperative roles of different channels in a single neuron in the sensing of different intensity of stimulation.
The two motor neurons and the sensory neuron ASN provide an entry point to elucidate defecation circuitry. The two motor neurons appear to be functionally connected, possibly involving synaptic connections between them, although we cannot exclude the possibility of multiple neurons being engaged in their functional connections. It remains to be determined as to how they are entrained with this rhythmic firing pattern, and whether it involves a central pattern generator upstream of PDF neurons. Interestingly, PDF is a peptide that has important roles in multiple neuropeptide signaling pathways in the fruit fly (Renn et al., 1999; Kim et al., 2013); it would be interesting to test whether this neuropeptide also plays a role in the regulation of defecation behaviors by PDF neurons in the VNC. It is also of interest to explore possible contributions of indirect effects of PDF over muscle contraction, such as an influence of tracheal branching in the hindgut that may affect muscle contractions (Linneweber et al., 2014). Recently, a study has suggested a novel role of HGN1 neurons in regulating the long-term food intake behaviors of adult flies (Olds and Xu, 2014). In our study we found that HGN1 neurons control the rhythmic pattern of larval defecation. These two studies suggest that Drosophila HGN1 neurons at different developmental stages might have multiple functions in regulating feeding and defecation behaviors.
Though separated in evolution millions years ago, the structures of Drosophila gut and human gut share striking similarity. There are circular and longitudinal muscles lining the gut ending with the anal sphincter that controls defecation (Netter, 1997; Murakami and Shiotsuki, 2001). It remains an open question as to the extent of similarity of the mechanisms that control the gut movements. Diseases such as Hirschsprung's disease and anorectal malformation with failure to pass meconium (Loening-Baucke and Kimura, 1999) are caused by developmental abnormality related to the gut and its innervation. Several genes and specific regions on the chromosomes have been shown or suggested to be associated with Hirschsprung's disease. Mutations in two human genes could lead to the absence of certain nerve cells in the colon (Puri and Shinkai, 2004). With the powerful genetic tools, further study of the Drosophila larval gut rhythmicity and its neural modulation will help us identify evolutionarily conserved features as well as strategies that may have been adopted by different organisms for their fitness.
All the larvae were raised in the normal fly medium (for the light activation assay, 100 µM all-trans retinal was added to the food). Flies are kept in 12 hr/12 hr dark/light circle at 25°C. PDF-Gal4, HGN1-Gal4, and UAS-ChR2 are from Bloomington stock center. GRASP was done using lines: PDF-loxA > loxAop-mCherry and HGN1-Gal4 > UAS-GFP or PPK-Gal4 > UAS-GFP. w; Gr28b[MB03888] is a Minos insertion which is from stock center (#24190), UAS-GCaMP5 fly line is from Loren L Looger lab in Janelia Farm. piezoKO is from A Patapoutian lab in Scripps.
GRASP between PDF neurons and HGN1 neurons: w; PDF-Lexa/UAS-CD4-sp1-10; HGN1-Gal4/LexAOp-CD4-sp11. GRASP between PPK neurons and HGN1 neurons: w; +/UAS-CD4-sp1-10; HGN1-Gal4/PPK-tdTomato-sp11. UAS-CD4-sp1-10; LexAOp-CD4-sp11 are from K Scott lab (UC Berkeley).
For the whole-mounting imaging, a freely moving larva was picked up and rinsed with distilled water. Then the larva was transferred into 4% PFA overnight at 4°C. The larva was put between cover glass and images were taken by Zeiss confocal microscopy. In some cases, the whole VNC or different part of the gut was dissected out and mounted on a cover glass in PBS for imaging.
For immunostaining of Drosophila larvae, third instar larvae were dissected in PBS. The whole hindgut and anus were isolated from their bodies. The tissues were then fixed in 4% PFA solution for 20 min at room temperature and treated with the primary antibody (NOMPC antibody from J Howard [Yale], vGlut antibody from G Davis [UCSF]) overnight at 4°C and secondary antibody for 2 hr at room temperature. Images were acquired with Leica SP5 confocal microscope.
Third instar larvae were picked up, rinsed, and transferred on to agar plate with yeast paste supplied with food dye (FD&C blue 1 and red 40, 1:1000). The larvae were fed with food dye for 2 hr and mounted between cover glasses for experiment.
A hindgut-specific byn-Gal4 was crossed with UAS-GFP to visualize the hindgut. A third instar larva was gently picked up from the food surface and rinsed with distilled water briefly. The larva was then transferred into a drop of PBS on the slide. A cover glass was put on the larva and pressed slightly to reduce larval movement. The larva was mounted ventral side up under a Leica stereoscope and video-taped for later analysis.
Mercury light filtered with a GFP filter was applied to the larva preparation with a certain duration of time. The movements of the hindgut and anus sphincter were video-taped for further analysis. The delay of contraction was calculated between the light onset and the anus sphincter contraction. Both hindgut and anus sphincter movements could be easily visualized with auto-fluorescence of food debris in the intestines.
Free moving third instar larva was pinned onto Sylgard coated chamber dorsal side up and filleted along the dorsal body wall. The larva was dissected in a saline containing: (in mM): 103 NaCl, 3 KCl, 5 TES, 10 trehalose, 10 glucose, 7 sucrose, 26 NaHCO3, 1 NaH2PO4, and 4 MgCl2, adjusted to pH 7.25 and 310 mOsm. 2 mM Ca2+ was added to the saline fresh before use. Fat bodies were gently removed from the gut surface. Additional pins were used to immobilize the gut. The preparation was visualized by Zeiss axioscope microscopy with 40× water lens. Sharp electrode with resistance around 80 MΩ was filled with 3 M KCl. The electrode tip was approached to the gut surface under the control of the MP-285 manipulator (Sutter, USA). The signal was acquired by the Axon 200B amplifier and filter at 2 kHz. The electrode was moved forward until the voltage suddenly dropped to around −40 mV.
Blue light was generated by mercury lamp with multiple filters. For ChR2 activation, a GFP filter was used to give out blue light with wavelength around 488 nm. The light application was controlled by a shutter equipped on the microscope. For pulse light activation, 2 s light pulse was repeated for three times. Recording data were analyzed with Clampfit and Matlab. The frequency before and during light application were calculated and compared as the index of activation.
The recordings were performed following the protocol described by Hu et al. (2010) with slight modifications. Briefly, the entire VNC of a third instar larva was dissected, and the peri-neural sheath was gently removed in recording saline containing 103 mM NaCl, 3 mM KCl, 5 mM TES, 10 mM trehalose, 10 mM glucose, 7 mM sucrose, 26 mM NaHCO3, 1 mM NaH2PO4, 1.5 mM CaCl2, and 4 mM MgCl2 (adjusted to 280 mOsm, pH 7.3). The dissected VNC were transferred to a glass-bottom recording chamber containing recording saline and immobilized with a platinum frame. The HGN1 neurons were identified by their GFP signals under a 40× water objective. Current-clamp and voltage-clamp recordings were performed using patch-clamp electrodes (9–10 MΩ) filled with internal solution (140 mM potassium D-gluconate, 10 mM HEPES, 4 mM MgATP, 0.5 mM Na3GTP, 1 mM EGTA, adjusted to 265 mOsm, pH 7.3). Cells were used for recording if the Rm value was greater than 500 MΩ and the membrane potential value was lower than −50 mV. A small constant hyperpolarizing current was injected during recording, immediately after break-in, to bring the membrane potential of neurons to approximately −60 mV. Cells were held at −60 mV in voltage-clamp mode for EPSC recordings. Signals were acquired with an Axon-700B multiclamp amplifier and were digitized at 10 kHz and filtered at 2 kHz using a 1322A D-A converter. Data were analyzed using Clampex 9.0 software (Molecular Devices).
A larva was mounted between two cover glasses with ventral side up. Its tail end was exposed for access of probe stimulation. A glass pipette was pulled and polished to form a taper-shaped probe with a diameter around 20 µm. The probe was spread on with grease to reduce friction. A piezo-controller was used to control the movement of the probe with fixed angle and increment. For imaging of ASN axons and PDF/HGN1 cell bodies, the probe was advanced 10 µm and the stimulation lasted for 1 s.
ASN ablation was carried out as previously described (Song et al., 2012). Briefly, a single second instar larvae 48 hr after egg laying (AEL) was mounted dorsal side up, and the cell body of the ASN was targeted using a focused 930-nm two-photon laser (∼350–700 mW) mounted on a custom-built Zeiss fluorescence microscope. Following lesion, animals were recovered on grape juice agar plates and imaged live at the appropriate stages.
A third instar larva was gently picked up from the food surface and rinsed with distilled water several times. The larva was transferred into a drop of PBS on the slide. A cover glass was put on the larva and pressed slightly to reduce larval movement. The preparation was then put under the Zeiss Pascal 510 confocal microscopy equipped with a 20× air objective. Time series images were acquired and used for analysis.
For HGN1/PDF dual imaging, the larva was mounted with the lateral side up, so that both groups of neurons could be visualized simultaneously.
For PPK neurons imaging, larvae were transferred to a piece of filter paper saturated with 100 mM sucrose for 4–6 hr to remove the food debris in the gut which could have potentially covered the neuron's images during experiments.
An automatic alignment was made to most image series since the larvae tend to move slightly during image acquisition. An ImageJ plugin ‘registration ROI’ was utilized to correct the movements of the images during recording.
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K VijayRaghavanReviewing Editor; National Centre for Biological Sciences, Tata Institute of Fundamental Research, India
eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.
Thank you for sending your work entitled “Identification of motor neurons and a mechanosensitive sensory neuron in the defecation circuitry of Drosophila larvae” for consideration at eLife. Your article has been evaluated by Vijay Raghavan (Senior editor) and 2 reviewers, one of whom, Howard Baylis, has agreed to reveal his identity.
The Senior editor and the two reviewers discussed their comments before we reached the decision that the manuscript needs revisions, and the Senior editor has assembled the following comments to help you prepare a revised submission.
This is an interesting paper that seeks to define the defecation programme in Drosophila larvae and to identify neuronal and molecular substrates for this behaviour. The paper has three main topics in the results; characterisation of the defecation motor programme, characterisation of three neuronal populations which control hind-gut muscle contractions and identification and characterisation of a neuronal TRP channel involved in mechanosensory feedback. Some of the data is of high quality (e.g. calcium imaging experiments) and the potential finding of a mechanosensitive neuron involved in the process is novel and potentially interesting. The work could have wider implications for understanding defecation and the generation of rhythmic behaviour, but there are substantial concerns with the paper as it stands.
Contrary to what the manuscript seems to imply, defecation behaviour has been previously studied in flies (see Edgecomb J Exp Biol 1994 and Cognigni et al Cell Metab 2011). In particular, Cognigni et al developed a method that allows quantification of diuresis, enteric function, and food intake, and showed complex dietary and reproductive modulation of some of these features. The characterisation of the neuronal circuitry underlying defecation per se is useful but linking to the above features is of broader interest and should be attempted. Furthermore, contrary to what the authors state, only one of the three neuronal populations that they “identify” is novel in this context, given that: 1) The PDF neurons have been previously shown not only to innervate the hindgut (Nassel et al J Comp Neurol 1993, Talsma et al 2012 PNAS), and also to promote visceral muscle contractions (Talsma et al PNAS 2012), and 2) The RN2 neurons have also been shown to innervate the hindgut and to regulate faecal output (Cognigni et al Cell Metab 2011, line referred to as HGN1-Gal4). [The authors should refer to the HGN1-Gal4 as this will allow other workers to connect the two studies.]
Both the Abstract and the body of the manuscript contain over-interpretations of the data and important omissions of previously published data. These need to be addressed in a revised manuscript. A revised manuscript should also show that what is being examined is indeed a circuit and that it controls actual defecation (as opposed to subtly modulating muscle contractions of the hind-gut). The experiments suggested below are achievable in a reasonable timeframe of about three months if the authors have the tools to manipulate those neurons with cellular resolution (all three drivers are much more broadly expressed so the observed behavioural effects, if any, could be secondary to interfering with all those other neurons). Substantial points are elaborated and listed below.
Characterization of “defecation behaviour” and its rhythmicity. The authors infer, but never show, effects of these neurons on defecation based on their effects on hindgut muscle contractions. Defecation rate could be subject to additional layers of neuronal control involving both visceral and somatic muscles, so the authors would need to show that these neurons do regulate defecation by, for example, using Fluoropoo (Cognigni et al Cell Metab 2011) if it works in larvae or by loading larval guts with dye-supplemented yeast and then recording defecation rate on unlabelled yeast. Related to this, the authors often make statements about defecation “rhythms/cycles” akin to those reported in C. elegans. These are based, as far as one can tell, on the rhythmic Ca2+ oscillations observed in resting PDF and RN2 neurons. However, the muscle recordings are not particularly rhythmic in resting conditions, and no evidence for rhythmic defecation events has been provided. For example, the pattern of EJPs shown in Figure 2A is described as rhythmic, however whilst it looks sporadic it does not look overtly rhythmic, analysis should be performed to support this statement and in support of a correlation between hind-gut contraction and defecation itself. Previously published data (Edgecomb J Exp Biol 1994 and Cognigni et al Cell Metab 2011) argues against a robust rhythm at least in adults, where defecation rate is regulated by both environmental (e.g. dietary content) and internal (e.g. reproductive) state. Thus, the conditions in which rhythmic defecation is observed should be precisely described.
Figure 3 which describes defecation data is not clear. What is being measured? The data in Figure 3b is described as hind-gut contraction, but 3A is simply described as “defecation” in the text. “Defecation” is too vague in this context, the exact step that is being measured needs to be defined. If the two measurements are the same thing, which one would hope, then the data in Figure 3A and B should be presented using the same measure, i.e. either both should be period in seconds or both should be defecation per period. This reflects a need as mentioned above to give precise details of the behavioural assays and data.
Gut peristalsis and defecation rate may vary depending on the larval stage. The authors seem to do all their experiments in L3 larva. Were these early, post-feeding, post critical weight or wandering L3s? Quantifications would need to be confined to one of these substages or data should be provided showing no differences between substages. Again there is a need for precise details of the behavioural assay and data.
Loss and gain-of-function phenotypes of PDF and RN2 neurons. Loss-of-function experiments were carried out constitutively throughout embryonic and larval development using Pdf-Gal4 and RN2-Gal4. Both drivers are expressed in other neurons that, when inactivated, could affect behaviours (e.g. feeding, circadian) that could indirectly impact gut movements (Thus, getting these experiments done at cellular resolution is important). Importantly, PDF has been shown to regulate tracheal branching in the hindgut, which could also affect muscle contractions (Linneweber et al Cell 2014). In the GoF muscle physiology experiments, no “raw” data has been provided for PDF neurons, and different experiments seem to have been conducted for the two neuronal populations. Only one control seems to have been used for the ChR2 experiments – several are normally required (+/- all-trans retinal, Gal4 and UAS parental controls). More details would also have to be provided about quantifications (how are the effects of tissue compression on fluorescence controlled? The hindgut diameter should become smaller. How would the ROI remain the same if the gut is actively contracting?).
The authors state that both PDF and RN2 neurons are motor neurons, based on comparisons with the gut innervation seen using VGLUT-Gal4. This is unconvincing. The authors should show the positions of the PDF, RN2 and OK371 axons on the hindgut muscles side by side, and use a broad neuronal marker like 22c10 to co-label the major nerves. What functional data shows a motor rather than neuromodulatory role for these neurons, especially for PDF?
The role of RN2 (see above too) is unclear. Activation of RN2 by ChR2 causes EJPs in the anal sphincter. This is the only direct evidence for the proposed role of RN2 as exciting the anal sphincter muscles. Other evidence suggests a more complex situation, ablation of RN2 does not prevent sphincter opening but does act additively with ablation of PDF on hindgut contraction period (Figure 3, also see comments re Figure 3 below). So does RN2 activation, by ChR2, alone have an effect on hind gut peristalsis? This data should be added to Figure 2D.
The sequential firing is very neat. There are several experimental predictions that come from this. One is that activation of PDF by ChR2 should induce anal muscle contraction (via RN2), data should therefore be included in Figure 2D on the anal sphincter muscle contraction when PDF is activated by ChR2. Similarly activation of PDF by ChR2 would be expected to give rise to anal sphincter EJPs. Another expectation is that inactivation of PDF may result in no Ca2+ signals in RN2, this could be tested by combining available reagents. Finally would it also be possible to test whether activation of PDF using ChR2 led to calcium signals in RN2?
The authors seem to have used HGN1-Gal4 to label the RN2 neurons. As described in Cognigni et al, HGN1-Gal4 is a particular line (the D insertion) resulting from mobilizing the initial RN2-Gal4 (Landgraf et al). This is an important distinction because other RN2-Gal4 lines are either expressed more broadly, or are not expressed in those hindgut-innervating neurons and HGN1-Gal4 should be used in the text as this will allow other workers to connect the two studies.
The data showing that PDF neurons are presynaptic to RN2 is not very convincing (especially the sytGFP/Denmark figure) - both neurons could be putting out dendrites in the same VNC region in order to respond to the same presynaptic partner.
The sensory role of the ASN is potentially interesting, but there is no data linking that particular ppk neuron (as opposed to other ppk+ neurons) with the hindgut muscle contraction phenotypes and/or defecation control. The authors could use the elegant approach previously used by this lab (Yang et al Neuron 2009, Fig 3) to strengthen this link.
ASN is identified as sensory neuron providing mechanosensory feedback from the anus. ASN shows periodic Ca2+ oscillations. These correlate to the opening of the anus and activity in PDF and RN2, however no functional test of the hypothesis that ASN activation lies downstream of PDF, RN2 and anal opening is shown. One experiment would be to show that inactivation of PDF results in a loss of Ca2+ signals in ASN. A more positive experiment would be to show that activation of RN2 (using ChR2 or dTRPA1) in turn activates ASN. The feedback experiments use a very indirect test of feedback onto PDF and/or RN2, EJPs in the anal sphincter muscles. This may be acting through ASN-PDF-RN2-anal sphincter muscle series. However there are other explanations, in particular it does not rule out other routes of ASN feedback onto the anal sphincter muscles. It would be much better to test the effect of ASN activity more directly on PDF. For example do NOMPC mutants have changes in PDF Ca2+ signals, which are alleviated by NOMPC rescue in ASN? Does ablation of ASN change PDF calcium signals? Even better would be activation of ASN and direct measurement of signals in PDF. This would add significant weight to the argument that ASN is a functional modify of PDF properties.https://doi.org/10.7554/eLife.03293.028
- Wei Zhang
- Zhiqiang Yan
- Lily Yeh Jan
- Yuh Nung Jan
- Wei Zhang
- Zhiqiang Yan
- Lily Yeh Jan
- Yuh Nung Jan
- Zhiqiang Yan
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank A Patapoutian and L Looger for fly lines. We thank J Howard and T Wang for antibodies. We thank S Younger, Y Song, M Klassen, and S Barbel for technical support. We thank members of the Jan lab for discussion. ZY was a recipient of the Long-Term Fellowship from the Human Frontier Science Program. ZY is supported by the Program for Professor of Special Appointment (Eastern Scholar) at Shanghai Institutions of Higher Learning and the Shanghai Rising-Star Program. This work was supported by NIH grants (R37NS040929 and 5R01MH084234) to YNJ. LYJ and YNJ are investigators of the Howard Hughes Medical Institute.
- K VijayRaghavan, Reviewing Editor, National Centre for Biological Sciences, Tata Institute of Fundamental Research, India
© 2014, Zhang et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.