https://orcid.org/0000-0002-6550-3851
Figures
Abstract
Chironomids are important laboratory model organisms used to assess toxicity in freshwater environments. Cell and tissue features are not commonly used as chironomid markers to detect toxicity, but they could be extremely helpful in identifying acute and chronic effects of pollutants. The nervous system is an excellent cellular candidate since it is reactive to toxic substances. However, a detailed description of the chironomid nervous system is required prior to considering it as a candidate for a cellular toxicity marker. The present study describes the central ganglia, nerves, axons, and the neuromuscular system of Chironomus vitellinus (Freeman, 1961) to facilitate its use as a model organism in environmental studies. We find that the structure of the C. vitellinus central nervous system is identical to that observed in other Chironomus larvae. We then focused our study on the first abdominal segment and labeled the 31 hemi-segmental muscles according to a nomenclature based on their position and orientation. We also characterized their innervation and assigned the nerves a nomenclature based on their terminals’ location in the muscle tissue. Finally, we investigated the neuromuscular junctions (NMJs) throughout this segment and defined four types of NMJs illustrating their great variability in size and shape. We selected a model NMJ, VEL 2, and quantified its mean bouton number and muscle size. Together with documenting a neurobiological system that could be informative to insects’ comparative biology, these results could help establish the Chironomus NMJ as an aquatic toxicity marker.
Citation: Reyes-Maldonado R, Ramírez A, Marie B (2026) The neuromuscular system of Chironomus vitellinus (Diptera: Chironomidae). PLoS One 21(1): e0326394. https://doi.org/10.1371/journal.pone.0326394
Editor: Simon Belle, Swedish University of Agricultural Sciences: Sveriges lantbruksuniversitet, SWEDEN
Received: May 30, 2025; Accepted: December 9, 2025; Published: January 29, 2026
Copyright: © 2026 Reyes-Maldonado et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the manuscript and its Supporting information files.
Funding: This work was supported by the Puerto Rico Center for Environmental Neuroscience (PRCEN) under NSF grant-HRD 1736019 to RR-M, AR and BM and by the NIH-NIGMS grant GM103642 to BM. Infrastructure support and service were provided, in part, by the grant U54MD007600 from the National Institute on Minority Health and Health disparities (NIMHD). There was no additional external funding received for this study.
Competing interests: The authors have declared that no competing interests exist.
Introduction
The aquatic larvae of various species of chironomids (Diptera: Chironomidae) have become a model of choice for assessing freshwater contaminants. Indeed, their larvae are easy to maintain under laboratory conditions, resist environmental changes, feed and burrow in benthic sediments, respond to many pollutants, and have a short life cycle [1]. When assessing toxicity, a series of markers have been used, from biochemical and molecular markers (BMMs) [2,3] to morphological and fitness markers (MFMs) [1].
The main benefit of using BMMs to assess toxicity is that these kinds of markers show the effects of chemicals before adverse effects are detectable at higher levels of biological organization [4]. Also, BMMs can provide important insights regarding how organisms deal with toxic chemicals and the mechanisms of toxicity [5]. In the case of MFMs, the main benefit is that the effects observed on individuals can be used to anticipate ecological effects. Despite the benefits offered by these markers, there are disadvantages. For example, BMMs are expensive to use in terms of the equipment needed and running costs [5] while MFMs are negatively influenced by the animal behavior and capacity for producing detoxification mechanisms [4].
Although BMMs and MFMs provide important information when assessing toxicity in freshwater environments, characterizing cellular and tissue markers (CTMs) can be a precious complementary approach [6]. Indeed, CTMs provide a complementary layer of evidence alongside BMMs and MFMs, helping connect and integrate changes resulting from chemical exposure at a molecular, biochemical, and physiological levels [7]. As a result, contaminant-induced cellular and tissue alterations can be related to the health and fitness of individuals allowing further extrapolation to population or community effects [7].
Acquiring CTMs often requires a specially trained investigator, specialized imaging and careful sample preparation and can be time consuming to standardize. However, once a technique is established, CTMs can offer endpoints with high ecological relevance and, in some contexts, a cost-effective way to verify toxicant effects [6].
Although a useful alternative, CTMs are rarely used as markers for assessing toxicity due to the lack of histological descriptions in this insect group. Having noticed this gap in knowledge, recent studies have provided new anatomical descriptions using modern histological techniques to identify new cellular and tissue markers (CTMs) for the assessment of freshwater contaminants [6,8]. Nevertheless, these studies do not focus on the use of the nervous tissue, a tissue that should be considered to assess toxicity given that contaminants can affect its function [9,10] and development [11,12].
The nervous system, as studied in the Drosophila melanogaster model, could be used as reference for the implementation of a quantifiable cellular marker for a Chironomus species. Because they are easily accessible, documenting neuromuscular junctions (NMJs, [13,14]) could provide a way to quantify and assess the toxicity of the animal milieu. Chironomus larval musculature and nervous system have been described separately [6,8,15]. Nevertheless, the general description of the larval musculature in its entirety has not been presented and no terminology allowing the identification of the different muscles has been defined [15]. Similarly, the central nervous system and its main projections to the periphery have only been described showing the different ganglia [6,8,15] and their corresponding main nerves [15]. Other studies have focused on describing the ultrastructure of the neural sheath, glial cells, and neurons, while others have identified the location of selected transmitters, transmitter related enzymes and neuropeptides through the ganglia and alimentary canal [16–18]. No study has provided a complete description of the neuromuscular system characterizing and naming the larval muscles and the nerve terminals innervating them.
NMJs, which are responsible for inducing muscle contraction and the animal’s locomotion [19], have been extensively studied in fruit flies [20], crustaceans [21], frogs [22], and mice [23]. In this study, we used immunohistochemical synaptic markers to describe in detail the neuromuscular anatomy (muscles, nerves and NMJs) of the first abdominal segment in the larvae of the broadly distributed chironomid Chironomus vitellinus. In addition, we selected a model NMJ to quantify features such as synaptic size. We assessed synaptic size by quantifying the number of synaptic boutons, the varicosities present at each terminal and containing the neurotransmitter release apparatus [24]. We hypothesize that our detailed descriptions of the neuromuscular system in chironomids will establish a framework for the study of the Chironomus NMJ as a CTM for future toxicological studies.
Materials and methods
Animal husbandry
This study used C. vitellinus as a model chironomid species. Previously, this species was referred to as Chironomus sp. ‘Florida’ in Reyes-Maldonado et al., 2021 [25] before being recently identified as C. vitellinus [26,27]. C. vitellinus has a widespread distribution, with reports from Africa, Asia, Oceania, and, more recently, the Americas [26]. Based on our research, these chironomids can be easily collected in both urban and rural areas of Puerto Rico, particularly in natural temporary pools and animal waterers.
The larvae used in this study were acquired from laboratory-reared colonies at the Institute of Neurobiology, University of Puerto Rico. These colonies were obtained from egg masses collected in the field using a previously described methodology [25]. The larvae studied were fed with TetraMin Tropical Flakes Fish Food provided as a slurry at 2 mg/ individual/ day and maintained at 27°C with a photoperiod of 12:12 light/dark. In order to reduce variability due to the differences in development we selected fourth Instar larvae. At this stage, larvae present large size musculature and synapses facilitating reliable dissection and imaging. They were identified by their head and body size (head capsule 0.39–0.58 mm; body length 6.38–14.19 mm, [25]) and bright red coloration (Fig 1A). Animals of both sexes were used in these procedures.
(A) The entire body of the 4th instar larvae is bright red with some green spots in the thorax in completely mature larvae. This stage is characterized by individuals having a length between 9-12 mm. (B-D) Schematic view of the dissection process designed for studying the neuromuscular tissue of C. vitellinus. (B) Initial position of larvae in the plate after pinning the larvae at its anterior and posterior extremities. The dashed lines and arrows represent the initial cuts of the dissection process. (C) Stretching the open larvae reveals extensive fat tissue, digestive tract, salivary glands, and tracheal system. (D) After removing the visceral tissue and fat, we can observe the segmental organization (T1 to A9) of both the CNS (chain of ganglia within the median plane) and the muscles (light grey rectangles). Scale: 1 mm.
Dissection
Tissue was dissected following an adapted methodology described by Sanhueza et al., 2016 [28]. Briefly, 4th instar larvae were attached, dorsal face up, to a Sylgard dish by pinning the space between the head and first abdominal segment and the last abdominal segment. Subsequently the larvae were covered with a modified Hemolymph-like 3 saline (HL3; 70mM NaCl2, 4mM KCl, 43mM MgCl2, 10 mM NaHCO3, 115mM Sucrose, 5mM HEPES; pH = 7.2) and dissected using microdissection scissors under the stereoscope. This saline solution was depleted of calcium to reduce synaptic activity and muscle contraction. The first incision was made transversally above the heart at abdominal segment 7/8. A second incision was made intersecting the first, proceeding anteriorly following the aorta until reaching the head of the animal. A third incision was made transversally at the end of the second incision to open the thoracic cavity (Fig 1B). Four pairs of pins were placed at each border of the 1st thoracic segment, 1st abdominal segment, 5th abdominal segment and the 7th abdominal segment in order to open the thoracic and abdominal cavities by stretching the tissue (Fig 1C and 1D).
Muscle characterization and imaging
Dissected tissue was stained using bromophenol blue (Millipore) diluted in Ringer’s saline solution (5 mg/ml) for 3 minutes. Muscle layers were successively removed starting with the most internal (visceral) muscles and finishing with the most external (cuticular) muscles. Within each layer, muscle arrangements were described by drawing under a stereomicroscope. Additional stained tissue was mounted on microscope slides, observed under an AmScope T490B-DK microscope, and photographed with a piece of metric paper (1 mm grid) using a 108MP phone camera (Motorola Edge, 2021) attached to the ocular. A scale was added to the obtained images in ImageJ (National Institutes of Health; http://imagej.nih.gov/ij/) by transforming pixel units to the known length of the metric paper grid. These images were used to produce more detailed drawings using the GNU Image Manipulation Program (GIMP 2.10.10, www.gimp.org). We defined a nomenclature based on position and orientation to label the muscles of the ventral and lateral region similar to previous work carried out on Drosophila larval anatomy [29,30].
Nervous tissue immunohistological characterization and imaging
After dissection, the stretched tissue was fixed with Bouin’s solution (Sigma) for 1 minute and washed three times with PBT solution [1x Phosphate-buffered Saline (PBS) and 0.1% Triton-X 100]. Guts and body fat were then carefully removed with tweezers and a bent pin head. After removing the pins, the tissue was washed and permeabilized in five PBT baths for an hour (12 min/bath).
Immunohistochemistry was used to visualize neuronal tissue. We assessed the cross reactivity of antibodies raised against Drosophila melanogaster proteins in the C. vitellinus tissue. The goat Cy3-conjugated anti-HRP antibody (1:300 in PBT; Jackson ImmunoResearch; [31] and the mouse anti-Syn antibody (1:20 in PBT; Developmental Studies Hybridoma Bank (DSHB) [32]) were tested. The immunohistochemistry process was initiated by incubating clean C. vitellinus tissue in the diluted primary antibody overnight at 4°C. This incubation was followed by five PBT washes during 1hr (12 min/wash) and a second 1hr incubation of the tissue was carried out in the dark with the diluted secondary antibody [Alexa Fluor 488-conjugated AffiniPure goat anti-mouseIgG (1:300; Jackson ImmunoResearch) and the Cy3 conjugated anti-HRP. A second wash of five changes of PBT in the dark was applied and the preparations were mounted on glass microscope slides using Vectashield (Vector Labs) as mounting media. The preparations were sealed with nail polish and stored in a cool dark place until analysis.
The specimens were scanned using a Nikon Eclipse Ti inverted A1R laser scanning confocal microscope. The NIS elements Advance Research 4.5 acquisition and analysis software was used for image acquisition. Complete body tissue recordings were obtained over the Z-axis (Z-stacks 0.5 µm) using a 20X objective with a numerical aperture (NA) of 0.75. Recording of specific features such as ganglia and nerves were obtained using a 40X objective (NA = 1.3; Z-stacks 0.2 µm) while the NMJs were obtained using a 40X objective (NA = 1.3; Z-stacks 0.2 µm) and a 100X objective (NA = 1.45; Z-stacks 0.1 µm). Large structures were recorded by scanning multiple areas in the X or Y axis with an overlap of 20% of the scanned area. Excitation was achieved using 489 nm and 561 nm lasers, corresponding to the excitation spectra of Alexa Fluor 488– and Alexa Fluor 568–equivalent fluorophores, respectively. Using Image J, images were produced as Maximum Intensity Projections over the Z‐axis of the stack of images. Figures were made using Adobe Photoshop 2023. In Fig 5, to better visualize some NMJ structures, the background of the image was attenuated using the GNU Image Manipulation Program (GIMP; https://www.gimp.org/). Briefly, the silhouette of the studied synaptic terminal was selected by hand on the color channel containing the HRP staining. The width of this selection was increased to 1 pixel size and later inverted to erase the area of no interest. GIMP was also used to create diagrams from raw images to show the location and appearance of ganglia, nerves and NMJs. A nomenclature to these structures was assigned based on previous work [15,29,30].
(A) General schematic view of the larval CNS with nomenclature of principal structures. Scale: 1 mm (B-D) Anti-HRP (red) and anti-Synapsin (green) immunofluorescence reveals the different CNS structures. Scale bar: 100 µm. (B) Representative confocal photograph of the brain (supraoesophageal ganglion), suboesophageal ganglion, prothoracic ganglion and mesothoracic ganglion (C) Representative confocal photograph of the metathoracic ganglion and the first abdominal ganglion (D) Representative confocal photograph of an abdominal ganglion from the 5th abdominal ganglion (E) Representative confocal photograph of terminal ganglia. Abbreviations: Tx: Thoracic segment x; Ax: Abdominal segment x; br.: Brain; s.oes.g. suboesophageal ganglion; pro.g.: prothoracic ganglion; mes.g.: mesothoracic ganglion; met.g.: metathoracic ganglion; xAb.g.: abdominal ganglion x; t.g.: terminal ganglion; VNC: ventral nerve cord.
(A) First abdominal segment as observed with the bromophenol blue staining. Scale is 200 µm. (B) Diagram showing the position and orientation of muscles in the first abdominal segment. Purple: dorsal muscles; Red: lateral muscles; Orange: ventral muscles. Light colors represent visceral/ interior muscles while dark colors represent cuticular/ exterior ones.
(A) Schematic representation of body wall muscles of one hemisegment with their respective peripheral nerve branches and neuromuscular junctions. Note that there are different layers of muscles and innervation going from the most cuticular/exterior to the more visceral/interior (B) Representation of the NMJs with their respective nerves in a view in which muscles were removed. Individual nerves are named and NMJs are drawn (C) View of the cuticular/ exterior layer of muscles and their innervation. Individual muscles are named (D) View of the visceral/ Interior layer of muscles, and their innervation shows the morphology of typical NMJs. (A-D) The midline, that separates the two bilaterally symmetrical hemisegments, is represented by a dotted line. Each nerve branch is represented with a single color to show target muscles. Muscle and nerve nomenclature are based on position, orientation, and number [29,30]. Abbreviation: Position V: ventral; VI: Ventral internal; VE: Ventral external L: lateral. Orientation: L: longitudinal O: oblique; T: transverse, SBM, segment border muscle.
Characterization of a model NMJ
The selected model NMJ was observed in 10 larval preparations (19 NMJs) immunolabeled with the anti-Syn and anti-HRP antibodies. The bouton numbers were quantified under a Nikon Eclipse 80i microscope at 60X. We quantified the mean number and the variability of the samples using “GraphPad Prism 6”. This software was also used to carry out the Shapiro-Wilk normality test to assess the distribution of the samples.
Results
Antibody reactivity in the nervous and muscle tissue of C. vitellinus
To investigate the different anatomical features of the C. vitellinus larvae, we decided to rely on two antibodies routinely used to characterize neurons and their synapses in another Diptera, Drosophila melanogaster. These antibodies presented immunoreactivity in C. vitellinus consistent with the described labeling of Drosophila tissue. Indeed, we were able to visualize neuronal membranes (anti-HRP; [31]) and synaptic vesicles (anti-Syn; [32]). In our study we used the neuronal membrane immunoreactivity revealed by the anti-HRP antibody to visualize the ganglia, nerves, and axons. The anti-Syn and anti-HRP antibodies were used to reveal the neuromuscular system. Like in Drosophila, synaptic boutons were visualized and quantified using the anti-Syn antibody.
Accessing the C. vitellinus tissue and observation of the basic larval structure
We dissected the 4th Instar larvae of C. vitellinus (Fig 1A). Upon dissection (described in methods; Fig 1B), the first structures exposed are the digestive tract and the salivary glands (Fig 1C). Once these structures are removed, an inner fatty layer can be observed covering the muscles and the central nervous system from the first to the last abdominal segment. This fatty layer is segmentally organized and reduces the visibility of the neuromuscular tissue especially around the central nervous system and the lateral muscles. Although this fatty layer cannot be eliminated, its careful partial removal in most of the areas of the preparation is required to visualize the neuromuscular system. The three thoracic and eight of the nine abdominal segments of the larva provide easily identifiable muscles, nerves and NMJs (schematic diagram; Fig 1D). While each thoracic segment (T1 to T3) presents a unique muscular organization, the arrangement of muscles is almost identical in the first seven abdominal segments (A1 to A7) and differs in the eighth and ninth abdominal segments. The muscles within different segments are innervated by a central nervous system (CNS) that is segmented and organized as a series of ganglia. We therefore decided to describe the CNS in more detail.
The central nervous system
Although the principal anatomical CNS features had already been described in other species within the Chironomus group [6,8,15], it was important to determine whether the anatomy of the C. vitellinus CNS was identical.
The C. vitellinus CNS is composed of a brain (also called supraoesophageal ganglion), a suboesophageal ganglion, three thoracic ganglia, and eight abdominal ganglia (Fig 2). The brain (br.) is located out of the head in the prothorax, and it is the largest and most prominent of the ganglia. Lying beneath the brain and in the same thoracic segment, the suboesophageal ganglion (s.oes.g.) and the prothoracic ganglion (pro.g.) stand out. The mesothoracic ganglion (mes.g.) and metathoracic ganglion (met.g.) are located in the mesothorax. The first abdominal ganglion (1 ab.g.) is in the metathorax, leaving the first abdominal cavity unoccupied. Abdominal ganglia 2–6 (2–6 ab.g.) occupy their respective abdominal segments. The 7th and 8th abdominal ganglia are both located in the 7th abdominal segment forming the terminal ganglion of the nerve cord (t.g.).
Nerves exit each ganglion to their respective region: nerves from the supra and suboesophageal ganglia reach only the head, the ones from the thoracic ganglia reach the thorax, and the ones from the abdominal ganglia reach their respective abdominal segments. Each ganglion is composed of two lobes and is interconnected to other ganglia through a series of nerve fibers, forming the ventral nerve cord (VNC).
Our results show that indeed the anatomical organization of the C. vitellinus CNS is identical to that observed in other Chironomus larvae such as C. dorsalis [15], C. sancticaroli [6], and C. columbiensis [8].
The musculature of the first abdominal segment
After describing the global larval architecture for both muscles and CNS, we decided to detail the organization of one segment. The first abdominal segment seemed the perfect candidate since it is easily accessible due to its central location and to the fact that the ganglion innervating its different muscles is located anterior to the segment itself (Fig 3). This last characteristic not only ensures unaltered visual access to the muscles, but it also facilitates potential future electrophysiological studies since the increased distance between muscle and ganglion also means longer and more accessible nerves.
Three orientations of muscles (longitudinal, transverse, and oblique) can be distinguished in the ventral, lateral and dorsal area of the thoracic and abdominal segments (Fig 3). The first abdominal hemisegment presents 31 muscles: 12 lateral, 10 ventral and 9 dorsal muscles. Regarding the orientation of the muscles in the ventral and lateral area, we can observe 8 oblique, 9 longitudinal and 5 transverse muscles per hemisegment. Muscle fibers are present in two layers, presenting 3 main muscles in the internal part and 7 muscles in the external part of the ventral region. The following five groups of muscles were defined: ventral external longitudinal (VEL), ventral internal longitudinal (VIL), ventral oblique (VO), lateral transverse (LT), and Lateral oblique (LO). The distribution of these muscles is described in detail in Fig 4. Fig 4C describes the external muscle layers and illustrates the nomenclature we used to identify the muscles namely SBM, VEL 1–4, VEL 6, VO1, LT 1–4, LO1, LO 4–9. Underneath this sheet of muscles, we can find the internal muscles (VIL 1–3, VEL 5; Fig 4D). As depicted in Fig 4, NMJs are visible and identifiable on every muscle. We therefore underwent the process of naming the nerves innervating the muscular architecture.
Nerves projecting to the first abdominal segment
We named “main nerves’‘ the two hemisegmental nerves originating from the first abdominal ganglia. There are six nerve tracts originating from the main nerve. We assigned these previously unnamed nerve tracts a nomenclature based on their nerve terminals’ location in the muscle tissue in a manner reminiscent of previously published work in Drosophila [13,30]. The six nerve tracts are: ventral nerve a, ventral nerve b, ventral nerve c, ventrolateral nerve, lateral nerve, and dorsal nerve (Fig 4B). The synaptic terminals of the ventral nerve a, b and c only reach ventral muscles; the synaptic terminals of the ventrolateral nerve reach a single ventral muscle but also few lateral muscles; the synaptic terminals of the lateral nerve reach only lateral muscles; and the synaptic terminals of the dorsal nerve reach all the dorsal muscles (Fig 4).
The first abdominal segment NMJs
We used the anti-Synapsin and the anti-HRP antibody labeling to observe the location of the NMJs in the C. vitellinus tissue. Because Synapsin is a synaptic vesicle protein [32], the anti-Synapsin fluorescence is more prominent in the synaptic boutons of the NMJ. Using this immunofluorescence, we noticed that some synapses showed clearly separated large-sized boutons with a round morphology, well distributed through the NMJ. Thanks to these characteristics, we were able to distinguish individual synaptic boutons, giving us the opportunity to quantify the number of boutons per synapse (for example Figs 5A and 6A). Any NMJ in which the anti-Synapsin immunoreactivity labeling lacked this description was cataloged as unquantifiable (for example Figs 5B and 6B). The anti-HRP antibody was used to visualize the neuronal membrane (axon, boutons, and inter-bouton synaptic area) and allowed the delineation of the NMJ and the identification of its origin. Integrating both antibodies, we determined the location of each NMJ and the axon from which it is originating.
NMJs are present throughout all the musculature and exhibit great variability in size and shape (Figs 5 and 6). We observed and defined four types of NMJs based on their sizes: large, medium, small, and very small (Fig 5A–5D). Large size NMJs are located in muscles VIL 1–3 and VEL 5 and show very small varicosities that do not produce quantifiable synaptic boutons (Figs 5A and 6A). Medium size NMJs are located in muscles VEL 2–4 and VEL 6 and show very well distributed varicosities that form quantifiable synaptic boutons (Figs 5B and 6B). Small size NMJs are located in muscles VIL 1 and LO 1 showing small varicosities that do not produce quantifiable synaptic boutons (Figs 5C and 6C). Very small synapses are located in muscles VO 1, LT 1–4, and LO 2–9 (Figs 5 and 6D–6F–6F). Synaptic terminals in LT 1–4 show a mix of tightly grouped and very distributed varicosities that produce quantifiable boutons of various sizes (Fig 6D). Varicosities in VO 1 and LO 2–9 are very condensed, producing mainly unquantifiable boutons, although quantifiable boutons can also be observed (Fig 6E and 6F).
Selection and description of the model NMJ VEL 2
We selected a model NMJ based on multiple criteria: 1) the NMJ is innervating a muscle that is easy to locate, 2) it is easy to differentiate from other synapses, and 3) it can be easily visualized. Furthermore, we considered the fact that its size is not an obstacle for its study. Indeed, a large synapse would not be adapted to a time- and energy- efficient study, while a small synapse might not allow the future characterization of undergrown synapses. In addition, its synaptic boutons must be easy to visualize and belong to a synapse with quantifiable boutons (see previous section).
The NMJ located on muscle VEL 2 fits all our criteria. This NMJ is a medium size synaptic terminal that originates from the “ventral nerve a” tract and is located in the external ventral muscle layer of the first abdominal segment. Its typical structure has two main branches covered with synaptic boutons, but secondary branches are also commonly observed. Its synaptic boutons are rounded, well distributed and cover a great part of the muscle (Fig 7A–7C). The number of synaptic boutons observed in different VEL 2 synapses is normally distributed (Shapiro-Wilk, p = 0.8) and ranges between 47–140 (X̅: 86.89; σ: 24.04, n = 19; Fig 7D and S1 Table). The muscle innervated by this terminal ranges from 22017 to 69982 μm2 (X̅: 44627; σ: 13452, n = 19; Fig 7E and S1 Table) and shows a normal distribution (Shapiro-Wilk, p = 0.82).
(A) Representative confocal photograph of a large ventral interior NMJ VIL1 revealed by anti-HRP (red) and anti-Synapsin (green) immunoreactivity. (A’) Diagram showing the location of the VIL1 muscle and its NMJ within the hemisegment (B) Representative confocal photograph and (B’) diagram of a medium ventral external NMJ on muscle VEL2 (C) Representative confocal photograph and (C’) diagram of a small NMJ on muscle LO1 (D) Representative confocal photograph and (D’) diagram of a very small NMJ on muscle VO1. Scale bar: 100 µm (A-D).
Anti-HRP and anti-Synapsin immunoreactivity reveal different synaptic bouton morphologies at different NMJs within the first abdominal segment. For example: (A) large size VLI1 NMJ, (B) medium size VEL2 NMJ, (C) Small size NMJ LO1, (D) Very small size LT2 NMJ, (E) Very small size LO 4-6 NMJ, (F) Very small size VO1 NMJ. Scale bar: 5 µm.
Anti-HRP (red) and anti-Synapsin (green) immunofluorescence reveals the structure of the VEL 2 synapse from 3 different animals. Scale is 50 µm. (D) Quantification of the number of synaptic boutons and (E) muscle size. Data is shown as a scatter plot and mean ± SEM. Data available in S1 Table.
In this work we have shown the basic architecture of the nervous system and muscles of C. vitellinus. We then focused our attention on a specific segment (Abdominal 1) and described in detail all the different muscles and their connecting nerves. Finally, we considered all the NMJs in this segment and selected a model NMJ for future study. While the inspiration for this work is the Drosophila NMJ, it remains to be seen whether we will be able to use the Chironomus NMJ to characterize changes in synaptic growth.
Discussion
The Chironomus larval NMJ shows great potential to be used as a cellular and tissue marker for future investigative work assessing rivers contamination. Indeed, Chironomus larvae inhabit river sediments (site of contamination) and work with Drosophila has shown that the NMJ responds to external/environmental factors (e.g., [33–35]). A description of the C. vitellinus neuromuscular system was therefore a necessary preliminary study.
Although partially known, the Chironomus neuromuscular anatomy has always been incomplete. Miall and Hammond’s work on Chironomus dorsalis presented a general view of the system, only describing the first layers of muscles without naming them and relating them to the nervous system [15]. Nerve tracts and the NMJs were not described by these authors nor by recent studies describing the nervous system in other Chironomus species [6,8,18]. The description presented in this work attempts to remediate these issues and presents a detailed anatomical description that could be used for toxicological studies.
The Chironomus larval abdominal muscular anatomy presented in this work is the most detailed available. It is unique when compared with the anatomy of other abdominal muscle arrangements described in other insect species such as D. melanogaster [29], Periplaneta americana [36], and Galleria mellonella [37]. Comparing the muscular anatomy of Chironomus with other insect anatomies is challenging since current insect descriptions use different body sections to characterize the muscle distribution. In this study, we focused on describing the first abdominal segment since the abdominal region of the larval stage is the most used for movement. Similar abdominal descriptions are common for the crawling larvae of insects (e.g., D. melanogaster [30]). Nevertheless, studies describing the anatomy of adult winged insects typically examine the thoracic section of the body given its relation to movement (e.g., Hemianax papuensis [38]). It is also true for the musculature descriptions of walking insects with well-developed appendages; in this case, the descriptions of the thoracic and appendage musculature are preferred (e.g., Locusta migratoria [39]).
When comparing the Chironomus anatomy to other crawling larvae such as D. melanogaster, we can notice that some muscle arrangements in the first abdominal segment are preserved while others are completely missing. This is the case for the D. melanogaster ventral acute and ventral transverse muscles, which are completely missing in Chironomus. In contrast, the lateral transverse and segment border muscles are preserved in both species. Another singular feature of the Chironomus musculature is that there are multiple lateral oblique (LO) muscles compared to the single LO muscle structure observed in D. melanogaster. The same is observed with the ventral oblique (VO) muscles, but this time the Chironomus anatomy only shows a single VO muscle structure while D. melanogaster shows multiple. The number of muscles per hemisegment in the first abdominal segment of the studied Chironomus species is similar to the number of muscles in D. melanogaster (31/Chironomus hemisegment vs 30/ D. melanogaster hemisegment [40] however, their distributions through the ventral, lateral and dorsal regions differ between species.
Interestingly we managed to transpose the knowledge acquired in Drosophila to Chironomus and produce relevant anti-HRP and anti-Synapsin immunostainings following identical protocols. Like in Drosophila, anti-HRP labeled the entire nervous tissue, including the brain, ventral nerve cord, and peripheral nerves in a way consistent with the specific labeling of the neuronal membrane [31]. For this reason, it is ideal to map the overall structure of the nervous system. The anti-Synapsin antibody also provides immunoreactivity resembling the one observed in Drosophila and specific to synaptic vesicles [32]. It is localized to the synaptic area and is missing from the axon and inter boutons segments. As such it is also an interesting tool to assess the number of boutons in Chironomus. The basic structure of the larval central nervous system was known in other Chironomus species [6,8,15] and C. vitellinus shows similar structure. We did, however, characterize a number of new features such as the different nerve tracts originating from the main nerve and their multiple NMJs. Nerve tracts trajectories in Chironomus bear similarities to the trajectories of the nerve tracts observed in D. melanogaster. The Chironomus dorsal nerve trajectory appears to be homologous with the Drosophila intersegmental nerve (ISN) trajectory, the Chironomus ventral a-c nerve trajectory to the Drosophila segmental nerves b-d (SNb-d) and the Chironomus lateral nerve trajectory to the Drosophila segmental nerve a (SNa). Nevertheless, tracing techniques such as retrograde labeling with different dyes (Inal et al., 2020) will be needed to corroborate the exact nerve trajectories in the abdominal segment of the Chironomus larvae. This method should also allow the identification of the motoneuron cell bodies innervating the muscles.
Synaptic terminals in the Chironomus tissue show a large variability of sizes paralleling the high variability in muscle size observed through the larval tissue. The great variability of synaptic varicosities made it difficult to identify suitable NMJs with quantifiable synaptic boutons. Similar variations in varicosities were previously observed in Chironomus tentans when trying to detect Bombesin immunoreactivity in nerve fibers of the larval alimentary canal [17]. The quantifiable NMJs in the ventral and lateral area can only be identified in the ventral external longitudinal (VEL) muscles. These muscles present an assortment of rounded and well-formed synaptic boutons making their quantification easy under the microscope. NMJs on lateral transverse (LT) muscles also present quantifiable synaptic boutons, but their quantification is difficult due to the presence of large amounts of fat tissue obstructing these terminals. It is important to stress the incredible diversity of synaptic morphologies that are present within the neuromuscular system of Chironomus vitellus. It will be interesting to ask whether, like in Drosophila, they are a combination of glutamatergic and peptidergic terminals [41]. Similarly, electrophysiological studies will be necessary to ask whether the motoneurons described here are tonic or phasic [42]. Validating antibodies with reactivity against active zones, postsynaptic glutamate receptors, and cytoskeletal markers in C. vitellinus should be a next step for the field to enable more detailed and quantitative NMJ analyses.
The model NMJ (VEL 2) was selected due to its visible location in the tissue and due to its quantifiable boutons. It is characterized by its large size, separated bouton arrangement and simple structure with elongated branches. The number of synaptic boutons of the Chironomus model NMJ shows similarities with the number of synaptic boutons observed in Drosophila at the model 6/7 NMJ. Authors have reported values averaging 75–100 boutons per NMJ in wild type Drosophila larvae [43–45]. For example, in our laboratory, a recent analysis of the Drosophila 6/7 NMJ at segment A3 shows a mean of 79 boutons when we examined 19 synapses. Our data ranged from synapses containing 48 boutons to synapses containing 116 boutons and the sample shows a standard deviation of 16 (S2 Table). The Chironomus model NMJ shows a mean average of 87 synaptic boutons for synapses ranging between 47 and 140 (standard deviation 24; n = 19; Fig 7D and S1 Table). The variability observed in Drosophila studies is lower than the ones observed in Chironomus. This is easily explained by the fact that the Drosophila strains are isogenic while the studied Chironomus animals are the progeny of genetically varied animals collected in the field. This observed variability in the number of boutons could also be due to factors such as larval size, sex, and age. Some of these factors are known to affect the structure and function of synaptic terminals in Drosophila [20,46] thus similar effects could be occurring in the Chironomus NMJ. Future experiments will have to be undertaken to assess whether these factors play a role. If they are, it should be possible to reduce the Chironomus synaptic size variability by only considering a well-defined subset of animals.
The present study of the Chironomus neuromuscular system provides the foundation upon which further research could develop. It identifies basic NMJ features that could be used as markers for assessing toxicity. In addition to quantifying the number of boutons and the muscle area, other features of the synapse (span, arborization, active zone numbers, postsynaptic differentiation) could be analyzed and quantified as new immunohistochemical tools become available. Given the outstanding neuromuscular anatomy of Chironomus, procedures such as electrophysiology could also be used to determine disruptions on neuronal signaling and synaptic transmission caused by toxic chemicals. Due to the versatility of this system, we are hopeful that it could be used to evaluate the responsiveness of the NMJ to biological, environmental, and toxicological variables.
Supporting information
S1 Table. Number of synaptic boutons and muscle sizes at the Chironomus VEL 2 NMJ segment A1.
Raw data collected from 19 synapses across 10 animals describing the number of synaptic boutons (Fig 7D) and the dimension of the muscle that is innervated (Fig 7E).
https://doi.org/10.1371/journal.pone.0326394.s001
(DOCX)
S2 Table. Number of synaptic boutons at the Drosophila m6/7 NMJ segment A3.
Raw data collected from 19 synapses across 10 animals describing the number of synaptic boutons. The mean, standard deviation and N number are shown.
https://doi.org/10.1371/journal.pone.0326394.s002
(DOCX)
Acknowledgments
The monoclonal anti-Synapsin antibody was obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at The University of Iowa, 45 Department of Biology, Iowa City, IA 52242. We thank Dr. Blagburn for his valuable comments on previous versions of this manuscript.
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