Glial-dependent clustering of voltage-gated ion channels in Drosophila precedes myelin formation
Abstract
Neuronal information conductance often involves the transmission of action potentials. The spreading of action potentials along the axonal process of a neuron is based on three physical parameters: the axial resistance of the axon, the axonal insulation by glial membranes, and the positioning of voltage-gated ion channels. In vertebrates, myelin and channel clustering allow fast saltatory conductance. Here, we show that in Drosophila melanogaster voltage-gated sodium and potassium channels, Para and Shal, co-localize and cluster in an area resembling the axon initial segment. The local enrichment of Para but not of Shal localization depends on the presence of peripheral wrapping glial cells. In larvae, relatively low levels of Para channels are needed to allow proper signal transduction and nerves are simply wrapped by glial cells. In adults, the concentration of Para increases and is prominently found at the axon initial segment of motor neurons. Concomitantly, these axon domains are covered by a mesh of glial processes forming a lacunar structure that possibly serves as an ion reservoir. Directly flanking this domain glial processes forming the lacunar area appear to collapse and closely apposed stacks of glial cell processes can be detected, resembling a myelin-like insulation. Thus, Drosophila development may reflect the evolution of myelin which forms in response to increased levels of clustered voltage-gated ion channels.
Editor's evaluation
Here, the authors characterize axon-initial segment-like structures in Drosophila using a variety of approaches spanning molecular genetic, confocal and super-resolution imaging, and ultrastructure. These important findings advance our understanding of how myelin evolved with potentially early roles in organizing ion channel clustering in axons. The convincing evidence supporting the conclusions includes advanced imaging combined with molecular genetic approaches, which will be of significant interest to neuroscientists and cell biologists.
https://doi.org/10.7554/eLife.85752.sa0Introduction
A functional nervous system requires the processing and transmission of information in the form of changing membrane potentials. To convey information along axons, neurons generate action potentials by opening of evolutionarily conserved voltage-gated sodium and potassium channels (Moran et al., 2015). Once an action potential is generated, it travels toward the synapse and the speed of information transfer is of obvious importance. It is long established that axonal conductance velocity depends on the resistance within the axon, which inversely correlates with its diameter. In addition, it depends on the resistance across the axonal membrane, which is increased by extensive glial wrapping. Furthermore, spacing of voltage-gated ion channels contributes to axonal conduction velocity (Eshed-Eisenbach and Peles, 2021; Freeman et al., 2016; Hodgkin and Huxley, 1952).
In vertebrates, unmyelinated axons generally have a small diameter with evenly distributed voltage-gated ion channels along their plasma membrane, and in consequence their conductance velocity is slow (Castelfranco and Hartline, 2015). To speed up conductance, axons grow to a larger diameter and show a clustering of voltage-gated ion channels at the axon initial segment (AIS) and the nodes of Ranvier. Together with the insulating glial-derived myelin sheet, this allows fast saltatory conductance (Arancibia-Cárcamo et al., 2017; Castelfranco and Hartline, 2015; Cohen et al., 2020; Dutta et al., 2018; Eshed-Eisenbach and Peles, 2021).
In invertebrates, mechanisms to increase conductance speed are thought to be limited by radial axonal growth, as seen in the giant fiber system of Drosophila or the giant axon of the squid (Allen et al., 2006; Hartline and Colman, 2007). No saltatory conductance has been described for invertebrates and it is assumed that voltage-gated ion channels distribute relatively evenly along axonal membranes. Nevertheless, myelin-like structures were found in several invertebrate species, including annelids, crustacean, and insects (Coggeshall and Fawcett, 1964; Davis et al., 1999; Günther, 1976; Hama, 1959; Hama, 1966; Hess, 1958; Heuser and Doggenweiler, 1966; Levi et al., 1966; Roots, 2008; Roots and Lane, 1983; Wigglesworth, 1959; Wilson and Hartline, 2011). However, it is unknown whether such myelin-like structures also impact the distribution of ion channels.
To address how glial cells affect axonal conductance velocity we turned to Drosophila. In the larvae, peripheral axons are engulfed by a single glial wrap resembling Remak fibers in the mammalian PNS (Matzat et al., 2015; Nave and Werner, 2014; Stork et al., 2008). In addition to insulating axons, we found that glial cells promote radial axonal growth. In the absence of wrapping glia axons are not only thin, but they are also characterized by a severe reduction in conductance velocity, which is stronger than predicted by the reduced axonal diameter (Hodgkin and Huxley, 1952; Kottmeier et al., 2020). Thus, wrapping glial cells might control localization of voltage-gated ion channels along the axonal plasma membrane.
Results
Distribution of the voltage-gated sodium channel Para
The Drosophila genome harbors only one voltage-gated sodium channel called Paralytic (Para), which is required for the generation of all action potentials (Kroll et al., 2015). To study the localization of Para and to test whether Drosophila glia affects its localization, we and others tagged the endogenous para locus with all predicted isoforms being modified (Ravenscroft et al., 2020; Venken et al., 2011; Figure 1A). In paramCherry flies, monomeric Cherry (mCherry) is inserted close to the Para N-terminus (Figure 1A). Homozygous or hemizygous paramCherry flies are viable with only mildly affected channel function (Figure 1B; Ravenscroft et al., 2020; Venken et al., 2011). ParamCherry localizes along many CNS and PNS axons of the larval nervous system (Figure 1—figure supplement 1A–C; Ravenscroft et al., 2020; Venken et al., 2011).
To independently assay Para localization, we generated antibodies against an N-terminal epitope shared by all predicted Para isoforms (Figure 1A). In western blots, anti-Para antibodies detect a band of the expected size (>250 kDa), which is shifted toward a higher molecular weight in protein extracts of homozygous paramCherry animals (Figure 1—figure supplement 1D). Immunohistochemistry detects Para localization in control first instar larvae but not in age-matched para null mutant animals, further validating the specificity of the antibodies (Figure 1D–E’). Whereas the pre-immune serum fails to detect any specific proteins (Figure 1F), anti-Para antibody staining of third instar larval filets revealed the localization of Para in the CNS and the PNS (Figure 1G) similar to what was noted for ParamCherry localization (Figure 1C). Thus, we anticipate that endogenously mCherry-tagged Para protein reflects the wild typic Para localization.
To test a possible differential distribution of Para in either sensory or motor axons, we utilized RNAi to remove mCherry expression in heterozygous paramCherry females. This leaves the wild type para allele intact and circumvents the early lethal phenotype associated with loss of para. Knockdown of mCherry expression in glutamatergic motor neurons (Mahr and Aberle, 2006) reveals paramCherry expression in cholinergic sensory neurons of third instar larvae. Here, Para appears to evenly localize along the abdominal nerves and is found at many processes within the CNS (Figure 2A). In contrast, silencing paramCherry in cholinergic neurons (Salvaterra and Kitamoto, 2001) reveals a predominant localization of Para in an axonal segment of motor axons at the PNS/CNS boundary of third instar larvae (Figure 2B), as suggested before (Ravenscroft et al., 2020).
Differential localization of voltage-gated potassium channels
Several genes encode voltage-gated potassium channels. Using endogenously tagged proteins, we find the Shaker potassium channel mostly in the synaptic neuropil regions (Figure 2C). The distribution of Shab resembles ParamCherry localization in sensory axons (Figure 2A and D), whereas Shal localizes in a pattern similar to Para localization on motor axons (Figure 2B and E). Thus, Drosophila larval motor axons appear to have an axonal segment resembling the vertebrate AIS harboring both voltage-gated sodium and potassium channels.
We then analyzed the localization of Para in the adult CNS. Here, too, global Para localization, as detected by anti-Para antibodies, matched the ParamCherry signal (Figure 2F and G). To differentiate between Para expression in motor and sensory neurons, we employed the recently developed FlpTag technique (Fendl et al., 2020). Here, cell type-specific expression of the Flp recombinase induces the inversion of a GFP-encoding exon located in the gene of interest. In paraFlpTag flies (Fendl et al., 2020), Flp expression in all motor neurons results in strong labeling of Para localization at a small part of the axon as it leaves the neuropil (Figure 2H and H’, arrowheads), indicating that an AIS is also found in adult motor axons. In contrast, expression of Flp in all sensory neurons using Chat-Gal4 UAS-flp reveals an even Para decoration of axons as they enter the CNS which fades out when axons reach into the neuropil (Figure 2I,I’, arrows).
High-resolution imaging reveals clustered localization of Para along motor axons
To obtain a higher spatial resolution of Para distribution, we used high-resolution Airyscan microscopy. In adult nerves, ParamCherry localization distal to the AIS is found in a clustered arrangement (Figure 3A–B’, arrowheads). To exclude that cluster formation is due to the fluorescence protein moiety, we performed anti-Para immunohistochemistry on primary Drosophila neural cells in culture, where axons form small fascicles with few accompanying glial cells (Figure 3C and C’). When such cultures are stained for Para distribution, we find Para channels localized in small clusters with a spacing of about 0.6–0.8 µm. However, in these neuronal cultures we cannot clearly define the number of Para expressing axons in a fascicle.
To further improve spatial resolution, we combined high-resolution imaging with the FlpTag labeling method. We restricted Flp expression to only one motor neuron in each larval hemineuromer using 94G06-Gal4 (Jenett et al., 2012; Pérez-Moreno and O’Kane, 2019), which results in the expression of GFP-tagged Para channels in only a single neuron. Whereas weak expression is noted around the nucleus, strong expression is seen in the AIS (Figure 3D, arrow, asterisks). Flanking the strong expression along the AIS, a clustered localization of ParaGFP could be noted (Figure 3E, arrowheads). Further super-resolution imaging of single motor axons decorated with GFP-tagged Para showed an average spacing of 0.620 µm (Figure 3F, F’, and G, n=91 clusters on 3 axons, quantification using Fiji). Interestingly, Para clusters appear to be organized along lines at the motor axon which was also found when analyzing the distribution of Para at the electron microscopic level (see below).
In sensory neurons increased Para localization is found at dendrites and the AIS
Having shown that in motor axons Para is concentrated in a clustered arrangement in an AIS of motor axons, we wondered whether similar distribution can be found in sensory axons. For this we expressed flp in multidendritic sensory neurons using the pickpocket Gal4 driver (ppk-Gal4). This allows labeling of the v’ada neurons (Figure 4A). Low levels of Para protein localize to the cell body and the distal shaft of the axon. Along the descending axon, Para localization increases only in some distance to the soma (Figure 4A, B, and C). However, para expression in sensory neurons is not as strong as in motor axons which may correspond to the notion that sensory axons are usually smaller axons. The relatively low expression levels did not allow super-resolution imaging and thus, we could not address whether Para is found in a clustered organization along axons of ppk positive sensory neurons. Interestingly, however, within some of the v’ada dendrites, Para accumulates in distinct clusters (Figure 4A’ and B’, arrows).
In conclusion, the above data show the presence of an AIS in Drosophila motor and sensory axons. In motor axons, where this domain likely serves as a spike initiation zone (Günay et al., 2015), Para channels are organized in a clustered arrangement.
Electron microscopic analysis of Para cluster formation
To determine the distribution of Para on the subcellular level, we integrated an Apex2 encoding exon in the para locus, which allows generating local osmiophilic diaminobenzidine (DAB) precipitates that are detectable in the electron microscope (Lam et al., 2015). The insertion of an Apex2 encoding exon in the N-terminus of Para affected para function less strongly than the insertion of an mCherry exon and resulted in a very weak hypomorphic para allele (Figure 1A and B). In adult flies, ParaApex2 is expressed in sufficient intensity to be detected along axons using the electron microscope. In small diameter peripheral axonal segments, weak ParaApex2 directed DAB precipitates are found (Figure 5A, white arrowheads). In contrast, in large diameter axons next to the CNS/PNS boundary intense DAB precipitates can be detected (Figure 5B). When we performed serial sectioning, the intensity of DAB labeling varied (Figure 5B–D), possibly reflecting the clustered localization of Para that we had found using the confocal microscope. We next determined the distribution of DAB precipitates along the circumference of an axon over 16 consecutive cross sections (Figure 5E and F). The resulting surface plot shows ParaApex2 localization of a small segment of the axon in a 3D space. This suggests that ParaApex2 clusters are organized in two lines along the ≈2.4 µm axonal circumference (Figure 5F and G), resembling the distribution of ParamCherry clusters along lines as detected using super-resolution light microscopy (Figure 3F). To further address the spacing of ParaApex2 clusters along the longitudinal axis of the axon, we performed longitudinal sections of Para-rich axon segments and determined the staining intensity along the plasma membrane by using Fiji (Figure 5H,I). Here, again a spatial modulation of the Para staining intensity is apparent, with a spacing of 0.706 µm (Figure 5I; see Figure 3G for quantification, n=54 cluster distances on 6 axons), which is similar to what we determined by confocal microscopy.
Para-rich axon segments are embedded in a lacunar system formed by tract glia
Interestingly, the large caliber axons decorated with highest levels of Para protein are embedded in a mesh-like glial organization, that resembles the lacunar system described earlier for the cockroach (Figure 5B–D, Figure 6A, B; Wigglesworth, 1960). The Drosophila glial lacunar system is characterized by intensive formation of glial processes around axons which are always larger than 0.5 µm in diameter (Figure 6A, B, asterisks). Glial cell processes have an average thickness of 35 nm (Figure 6A, B, n=189 processes, 4 nerves from 3 animals).
Next, we determined which glial cell type forms these lacunar structures. In the larva, the central ensheathing/wrapping glial cells express the 83E12-Gal4 driver (Peco et al., 2016; Pogodalla et al., 2021), whereas the peripheral wrapping glial cells can be addressed using the nrv2-Gal4 90C03-Gal80 driver (Kottmeier et al., 2020; Matzat et al., 2015; Stork et al., 2008; Figure 1—figure supplement 1A and B). In adults – but not in larvae – a specialized group of glial cells is found at the CNS/PNS boundary, called tract glia (Kremer et al., 2017) which overlaps with both the central ensheathing glia and the peripheral wrapping glia (Figure 6—figure supplement 1). Interestingly, a similarly distinct group of glial cells has been identified in the vertebrate nervous system (Fontenas and Kucenas, 2017; Kucenas et al., 2008; Kucenas et al., 2009). The position of the lacunae coincides with the location of the tract glial cells (Kremer et al., 2017) (compare Figures 2H, 6C). These glial cells express 75H03-Gal4, 83E12-Gal4 as well as the nrv2-Gal4 90C03-Gal80 driver (Figure 6—figure supplement 1). Multicolor flipout (MCFO2) labeling experiments (Nern et al., 2015) indicate tiling of these glial cells along the nerve with no overlap and no spaces in between individual glial cells (Figure 6D–F). To further determine which glial cell forms the lacunar structures, we generated flies harboring a UAS-Myr-Flag-Apex2-NES transgene (Apex2Myr, see Materials and methods) and expressed the myristoylated Apex2 with the different Gal4 drivers mentioned above. These experiments confirmed that most of the lacunar system is indeed generated by tract glial cell processes (Figure 6A, B).
Thus, in large caliber motor axons, most of the Para voltage-gated sodium channels is positioned close to the lacunar system, which had been previously speculated to serve as an extracellular ion reservoir needed for sustained generation of action potentials (Chandra and Singh, 1983; Leech and Swales, 1987; Maddrell and Treherne, 1967; Treherne and Schofield, 1981; Van Harreveld et al., 1969; Wigglesworth, 1960).
Myelin in the leg nerve is found close to the CNS
In vertebrates, clustering of voltage-gated ion channels occurs on the edges of myelinated axonal segments (internodes) (Arancibia-Cárcamo et al., 2017; Castelfranco and Hartline, 2015; Cohen et al., 2020; Dutta et al., 2018; Eshed-Eisenbach and Peles, 2021). Here, myelin not only participates in positioning of voltage-gated ion channels but also increases electric insulation and thus contributes to a faster conductance velocity. In Drosophila highest conductance velocity is likely to be required during fast and well-tuned locomotion in adults. Thus, we focused our search for myelin-like structures on adult leg nerves. Most of the 760 axons within an adult leg run in a single large nerve that exit the CNS at well-defined positions (Figure 7A-C, Figure 7—figure supplement 1A). Unlike the organization in larval nerves, axons running in the leg nerves are found in distinct zones depending on their diameter (Figure 7B and C). At the position of the femur, large axons are always covered by a single glial sheet. Small diameter axons are generally not individually wrapped but rather engulfed as a fascicle (Figure 7B and D). At the coxa, close to the CNS, we noted that large diameter axons were occasionally flanked by several glial membrane sheets (Figure 7E, asterisk). Up to 15 flat glial membrane sheets with a thickness of about 28 nm are found along larger axons (Figure 7E,F, Figure 7—figure supplement 1B). Axons with an intermediate diameter show individual glial wrapping with a single or very few glial sheets (Figure 7E and H). To quantify the occurrence of myelin-like structures we made semi-serial distal to proximal sections of six nerves every 5 µm across the entire lacunar area spanning 40–60 µm (Figure 7—figure supplement 2). The position where lacunar structures were first identified was set as zero. We then counted the occurrence of myelin-like structures in every section that we defined as ≥4 glial layers in close apposition. Here, we noted an increase in the number of myelin-like structures at the distal end of the lacunae (Figure 7—figure supplement 2B, Figure 7—figure supplement 3). No myelin-like structures were found at proximal positions close to the neuropil. The position of the up to four myelin-like structures found within a section plane was variable and could be either at the margin of the lacunae (Figure 7—figure supplement 3A) or could be found separating an area with small axons from an area with large axons (Figure 7—figure supplement 3B), or close to the blood-brain barrier (Figure 7—figure supplement 3C). In rare cases we noted formation of myelin-like membrane stacks without contact to axons in the lacunar region (Figure 7—figure supplement 3D). Myelin-like sheets contact serval axons (Figure 7—figure supplement 3A–C, E and F) but can also engulf single large axons with varying complexity of the membrane stacks (Figure 7—figure supplement 3G–H, Figure 7—figure supplement 4).
Myelin can be formed by central tract glia and peripheral wrapping glia
To determine which glial cell type is able to form myelin-like structures, we expressed Apex2Myr in specific glial cell types and analyzed whether DAB positive myelin-like stacks of glial cell processes can be detected in the electron microscope. Upon expression of Apex2Myr in CNS-derived tract glia 75H03-Gal4 DAB positive myelin stacks can be detected (Figure 7G (black arrowhead), Figure 7I, Figure 7—figure supplement 3, Figure 7—figure supplement 5). The finding that 75H03-Gal4 directed Apex2 labeling can be found next to unlabeled glial sheets (Figure 7G, white arrowhead) suggests that peripheral wrapping glial cells can also form myelin-like structures in the leg nerve. Drosophila myelin-like membrane stacks are generated by extensive membrane folding providing the disadvantage that axons are not entirely insulated (Figure 7I, Figure 7—figure supplements 4 and 5). However, we occasionally do find axons encircled by multiple glial wraps (Figure 7H, Figure 7—figure supplements 4 and 5). Interestingly, in some areas we noted almost compacted glial membrane sheets (Figure 7F,I, inlay boxed areas).
To further validate these findings we performed additional high-pressure freezing of prefixed samples to optimize tissue preservation (Möbius et al., 2016; Sosinsky et al., 2008). In such specimens compact stackings of thin glial membrane sheets can be detected, too (Figure 7J and K). In the compacted areas (Figure 7I–K), the interperiodic distance of the different glial layers is about 30 nm, which is considerably more than the interperiodic distance of 13 nm found in mouse peripheral myelin (Fledrich et al., 2018). The unique compact appearance of vertebrate myelin is mediated by the myelin basic protein (MBP) (Nave and Werner, 2021). In contrast to vertebrate myelin where extra- and intercellular space is removed, fly myelin-like structures only show an irregular compaction of the extracellular space.
Para localization depends on wrapping glial cells
Next, we wanted to test whether wrapping glial cells participate in the control of positioning voltage-gated ion channels. To address this, we ablated either peripheral wrapping glia or central ensheathing glia including the tract glia by directing the expression of the proapoptotic gene hid (Kottmeier et al., 2020; Pogodalla et al., 2021) and assayed the distribution of Shal and Para. Ablation of central or peripheral wrapping glial cells does not affect the distribution of the voltage-gated potassium channel Shal (Figure 8—figure supplement 1). Likewise, removal of the CNS-specific ensheathing glia does not affect Para localization in the larval nervous system (Figure 8—figure supplement 2A–E). In contrast, upon ablation of the peripheral wrapping glia a marked change in Para protein localization becomes obvious (Figure 8—figure supplement 2A–B’). In control larvae, anti-Para antibodies detect only a weak labeling of segmental nerves, but all nerves are intensely decorated with Para in wrapping glia ablated larvae. Whereas in wild type control larvae, 2.5 times more Para protein is found at the CNS/PNS transition zone compared to nerve segments on the muscle field, an almost even distribution is noted in glia ablated larvae (Figure 8C). In addition to the redistribution of Para protein along the axon, we also noted a twofold increase of para mRNA levels in further qRT-PCR experiments (Figure 8D).
Taken together, even in the small insect Drosophila melanogaster, myelin-like structures are formed (Figure 9). They are preferentially found distally to a lacunar region. The lacunae are formed by glial cell processes and comprise a large extracellular liquid filled space (Figure 9). Para voltage-gated sodium channels are differentially localized along sensory and motor neurons. In sensory neurons, Para expression is generally weaker and concentrates in an AIS but is also found in dendritic processes. In motor neurons Para localization is enriched in axonal segments that are running within the glial lacunar system. Interestingly, glia ablation experiments indicate that normal para mRNA expression as well as Para protein localization is dependent on the presence of wrapping glial cell processes. This suggests a signaling pathway from glia to the regulation of para transcription.
Discussion
In the vertebrate nervous system, saltatory conductance allows very fast spreading of information. This requires localized distribution of voltage-gated ion channels and concomitantly, the formation of the myelin sheath. The evolution of this complex structure is unclear. Here, we report glial-dependent localization of voltage-gated ion channels at an AIS-like domain of peripheral Drosophila larval motor axons. As more channels accumulate in adults, a lacunar system and adjacent myelin-like structures are formed by central tract glia and peripheral wrapping glia.
In myelinated axons of vertebrates, voltage-gated Na+ and K+ channels are clustered at the AIS and the nodes of Ranvier (Amor et al., 2014; Freeman et al., 2016; Nelson and Jenkins, 2017). In invertebrate neurons, the AIS corresponds to the spike initiation zone located distal to the soma and distal to the dendrite branching point. Such segments were found in Caenorhabditis elegans (Eichel et al., 2022) and have been previously postulated for Drosophila neurons due to the localization of a giant ankyrin, which in all systems appears to be an important scaffolding protein at the AIS, as well as the presence of voltage-gated ion channels (Dubessy et al., 2019; Freeman et al., 2015; Jegla et al., 2016; Ravenscroft et al., 2020; Trunova et al., 2011). Moreover, recent modeling approaches at the example of the pioneering aCC motor neuron predicted the localization of voltage-gated ion channels at the CNS/PNS boundary (Günay et al., 2015), which very well matches the localization of the voltage-gated ion channels Para and Shal, as reported here. Interestingly, in Drosophila para mRNA expression as well as Para protein localization depend on the presence of peripheral wrapping glia. In glia ablated nerves, Para expression is increased and decorates the entire axonal membrane. This loss of a clustered distribution may contribute to the pronounced reduction in axonal conductance velocity noted earlier in such glia ablated animals (Kottmeier et al., 2020). In addition, we found an increased para mRNA expression. How glial cells control Para localization and how this is then transduced to an increased expression of para remains to be further studied. Since alterations in glial differentiation caused by manipulation of FGF-receptor signaling specifically in peripheral wrapping glia does not cause a change in Para expression or localization (Figure 8—figure supplement 2F–H), proteins secreted by wrapping glia might be needed for the correct positioning of voltage-gated ion channels (Yuan and Ganetzky, 1999).
In the adult nervous system, the AIS-like domain is embedded in glial lacunar regions formed by wrapping glial cell processes. The increased expression of Para within the AIS-like segments of adult brains is expected to generate strong ephaptic coupling forces (Rey et al., 2022; Rey et al., 2021). These are caused by ion flux through open channels which generate an electric field that is able to influence the gating of ion channels in closely neighboring axons (Arvanitaki, 1942; Krnjevic, 1986; Rasminsky, 1980). Ephaptic coupling helps to synchronize firing axons (Anastassiou and Koch, 2015; Anastassiou et al., 2011; Han et al., 2018; Shneider and Pekker, 2015), but is also detrimental to the precision of neuronal signaling in closely apposed axons (Arvanitaki, 1942; Kottmeier et al., 2020).
Ephaptic coupling is counteracted by the glial lacunar system, that spatially separates axons and adds more levels of wrapping. Furthermore, it was postulated that the lacunar system provides a large extracellular ion reservoir (Wigglesworth, 1960). Given the tight apposition of axonal and glial membranes with most parts of the nerve, which is in the range of 20 nm, only a very small interstitial fluid volume is normally present. Thus, action potential generation would deplete sodium and potassium ions very fast, and would prevent sustained neuronal activity. The development of lacunar structures might therefore provide sufficient amount of ion and at the same time physically separates axons to reduce the likelihood of ephaptic coupling. It will be interesting to test this hypothesis in the future.
Close to the lacunar structures we detected myelin-like structures. It appears that the glial processes that form the lacunae collapse to form compact myelin-like membrane sheets. Interestingly, myelin-like structures are not formed at the lateral borders of the lacunae but rather form at its distal end. This indicates that insulation is likely not a key function of the myelin-like structures, but rather these structures originate as a consequence of the collapsed lacunar system. Concomitant with the occurrence of the myelin-like differentiations we note a decrease in the Para ion channel density. At the same time, the need for a large ion reservoir decreases, favoring the formation of myelin-like structures.
A hallmark of vertebrate myelin is the spiral growth of the insulating glial membrane. This is generally not observed in large fly nerves where glial membrane sheets rather fold back than spirally grow around a single axon. Compared to myelinated vertebrate axons, this provides the disadvantage that axons are not entirely insulated. However, spiral growth can be seen in small nerves where less extensive wrapping is noted. An additional unique feature of vertebrate myelin is its compact organization which is mediated by the MBP (Nave and Werner, 2021). In contrast to vertebrate myelin where extra- and intercellular space is removed, fly myelin-like structures only show a compaction of the extracellular space, which is expected to increase resistance as the number of freely moving ions is diminished. A fully compact myelin state would require MBP-like proteins which have not been identified in the fly genome.
In conclusion, the evolution of myelin appears reflected in the different developmental stages of Drosophila. First, voltage-gated ion channels are clustered at the AIS with the help of Drosophila glia. Second, upon increased expression of such ion channels in the adult nervous system, an ion reservoir might be formed by the lacunar system. The collapse of glial processes in the non-lacunar regions then provides the basis of myelin formation. In the future, it will be interesting to identify glial-derived signals that ensure channel positioning and determine how neuronal signaling adjusts channel expression and triggers formation of myelin.
Materials and methods
Drosophila genetics
Request a detailed protocolAll fly stocks were raised and kept at room temperature (RT) on standard Drosophila food. All crosses were raised at 25°C.
To determine temperature sensitivity, five 3-day-old male and female flies were transferred to an empty vial with a foam plug. The vials were incubated in a water bath at 42°C for 1 min and then placed at RT. Flies were monitored every 15 s for 5 min. At least 100 males and females for each genotype were tested.
For MCFO experiments early, white pupae were collected, put in a fresh vial and heat shocked at 37°C for 1 hr. Pupae were placed back to 25°C and dissected a few days after hatching.
To generate paramCherry flies we employed the MiMIC insertion strain paraMi8578 generated by the Bellen lab and we injected pBS-KS-attB1-2-PT-SA-SD-0-mCherry (DGRC Stock 1299; https://dgrc.bio.indiana.edu//stock/1299; RRID:DGRC_1299; Venken et al., 2011) into embryos with the following genotype: y w Φ31/paraMi08578. Following crosses to FM7c, y w flies were tested by PCR to identify successful insertion events.
To generate paraApex2 flies, we first removed mCherry encoding sequences from pBS-KS-attB1-2-PT-SA-SD-0-mCherry (DGRC#1299) using restriction enzymes and then inserted the apex2 coding sequence (Addgene #49386, using the primers AAGGATCCGGAAAGTCTTACCCAACTGT and AAGGATCCGGCATCAGCAAACCCAAG). pBS-KS-attB1-2-PT-SA-SD-0-Apex2 was used to establish a paraApex2 as described above. Flies were tested via single-fly PCR. To generate UAS-Myr-Flag-Apex2 flies, we cloned Apex2 using the primers CACCgactacaaggatgacgacgataa and cagggtcaggcgctcc into pUAST_Myr_rfA_attB, which was then inserted into the landing site 86Fb using established protocols (Bischof et al., 2007).
Western blot analysis
Request a detailed protocolTen adult fly heads were homogenized in 50 µl RIPA buffer on ice. They were centrifuged at 4°C for 20 min at 13,000 rpm. The supernatant was mixed with 5× reducing Lämmli buffer and incubated for 5 min at 65°C. Fifteen µl of the samples were separated to an 8% SDS-gel and subsequently blotted onto a PVDF membrane (Amersham Hybond-P PVDF Membrane, GE Healthcare). Anti-Para antibodies were generated against the following N-terminal sequence (CAEHEKQKELERKRAEGE), affinity purified, and were used in a 1/1000 dilution. Experiments were repeated three times.
Cell culture
Request a detailed protocolPrimary neural cell culture was preformed as described (Prokop et al., 2012). In brief, three to five stage 11 embryos were collected, chemically dechorionized, and homogenized in 100 µl dispersion medium. Following sedimentation for 5 min at 600 × g, cells were resuspended in 30 µl culture medium and applied to a glass bottom chamber (MatTek), sealed with a ConA-coated coverslip. Cultures were grown for 5–7 days. Experiments were repeated three times.
qPCR
Request a detailed protocolRNA was isolated from dissected larval brains using the RNeasy mini kit (QIAGEN) and cDNA was synthesized using Quantitect Reverse Transcription Kit (QIAGEN) according to the manufacturer’s instructions. qPCR for all samples was performed using a Taqman gene expression assay (Life Technologies) in a StepOne Real-Time PCR System (Thermo Fisher, para: Dm01813740_m1, RPL32: Dm02151827_g1). RPL32 was used as a housekeeping gene. Expression levels of Para were normalized to RPL32.
Immunohistochemistry
Request a detailed protocolLarval filets: L3 wandering larvae were collected in PBS on ice. Larvae were placed on a silicon pad and attached with two needles at both ends, with the dorsal side facing up. They were cut with a fine scissor at the posterior end. Following opening with a long cut from the posterior to the anterior end the tissue was stretched and attached to the silicon pad with additional four to six needles. Gut, fat body, and trachea were removed. Adult brains: Adult flies were anesthetized with CO2 and were dipped into 70% ethanol. The head capsule was cut open with fine scissors and the tissue surrounding the brain removed with forceps. Legs and wings were cut off and the thorax opened at the dorsal side. The ventral nerve cord was carefully freed from the tissue. For fixation, dissected samples were either covered for 3 min with Bouin’s solution or for 20 min with 4% PFA in PBS. Following washing with PBT samples were incubated for 1 hr in 10% goat serum in PBT. Primary antibody incubation was at 4°C followed. The following antibodies were used: anti-Para N-term, this study; anti-dsRed (Takara), anti-GFP (Abcam, Invitrogen), anti-Rumpel (Yildirim et al., 2022), anti-Repo (Hybridoma bank), rabbit α-V5 (1:500, Sigma-Aldrich), mouse α-HA (1:1000, Covance), rat α-Flag (1:200, Novus Biologicals). The appropriate secondary antibodies (Thermo Fisher) were incubated for 3 hr at RT. The tissues were covered with Vectashield mounting solution (Vector Laboratories) and stored at 4°C until imaging using an LSM880 Airyscan microscope, or a Elyra 7 microscope (Carl Zeiss AG Elyra 7 imaging, lateral resolution 80 nm with a voxel size of 30 nm × 30 nm × 100 nm). All stainings were repeated >5 times.
High-pressure freezing
Request a detailed protocolThree-week-old female flies were used with head, legs, and tip of abdomen removed. Following fixation in 4% formaldehyde (FA) in 0.1 M PHEM in a mild vacuum (–200 mbar), at RT for 45 min and three washes in 0.1 PHEM, the tissue was embedded in 3% low melting agarose for vibratome sectioning (Leica, VTS1200S). Samples were cut in PBS into 200 μm thick cross sections with 1 mm/s, 1.25 mm amplitude, and were placed into lecithin-coated 6 mm planchettes, filled with 20% PVP in 0.1 M PHEM and high-pressure frozen (Leica, HPM100). Seven specimens were sectioned. Freeze substitution was performed in 1 %OsO4, 0.2% glutaraldehyde, 3% water in acetone at –90°C and stepwise dehydrated over 3 days. Samples were embedded in mixtures of acetone and epon.
DAB staining and electron microscopy
Request a detailed protocolFlies were injected with 4% FA in 0.1 M HEPES buffer and fixed at RT for 45 min. Following washes and incubation in 20 mM glycine in 0.1 M HEPES, samples were incubated in 0.05% DAB in 0.1 M HEPES at RT for 40 min; 0.03% H2O2 was added and the reaction was stopped after 5–10 min. The tissue was then fixed in 4% FA and 0.2% glutaraldehyde in 0.1 M HEPES at RT for 3 hr. After three times rinsing the tissue was fixed in 4% FA at RT overnight. The FA was replaced by 2% OsO4 in 0.1 M HEPES for 1 hr on ice (dark). Uranyl acetate staining was performed en bloque using a 2% solution in H2O for 30 min (dark). Following an EtOH series (50%, 70%, 80%, 90%, and 96%) on ice for 3 min each step, final dehydration was done at RT with 2×100% EtOH for 15 min and two times propylene oxide for 30 min. Grids of high-pressure frozen samples were additionally counterstained with uranyl acetate and lead citrate. Following slow epon infiltration specimens were embedded in flat molds and polymerized at 60°C for 2 days.
Six specimens from three different fixation experiments were sectioned. Ultrathin sections were cut using a 35° ultra knife (Diatome) and collected in formvar-coated one slot copper grids. For imaging a Zeiss TEM 900 at 80 kV in combination with a Morada camera (EMSIS, Münster, Germany) operated by the software iTEM. Image processing was done using Adobe Photoshop and Fiji. Ultrathin sections of high-pressure frozen samples were examined at a Tecnai 12 biotwin (Thermo Fisher Scientific) and imaged with a 2K CCD veleta camera (EMSIS, Münster, Germany).
To plot the Para distribution across the axonal surface, an axon was serially sectioned (≈70 nm section thickness) and imaged. The images were cropped to the size of the axon in Fiji and aligned using Affinity Photo (software version 1.10.5.1342). The rotated/aligned images were loaded into Fiji and the segmented line tool was used to create ROIs on top of the axonal membrane in every section. The ROI was set to have the same starting point for the measurement. ParaApex2 staining intensity was measured along the circumference of an axon on 16 sequentially sectioned EM images. The relative gray values were binned by a factor of 100. This was then interpolated in 3D. We used biharmonic spline interpolation from the MATLAB Curve Fitting Toolbox (software version 9.13.0.2105380 (R2022b) Update 2) to generate a surface plot.
Script:
x=ParaE(:,1);
y=ParaE(:,3);
z=ParaE(:,2);
xlin = linspace(min(x), max(x), 100);
ylin = linspace(min(y), max(y), 100);
[X,Y]=meshgrid(xlin, ylin);
% Z=griddata(x,y,z,X,Y,'natural');
% Z=griddata(x,y,z,X,Y,'cubic');
Z=griddata(x,y,z,X,Y,'v4');
mesh(X,Y,Z)
axis tight; hold on
plot3(x,y,z,'.','MarkerSize',15)
Appendix 1
Data availability
All imaging and source data are available through: https://doi.org/10.57860/min_prj_000008. All Drosophila strains reported are available upon request to CK.
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Article and author information
Author details
Funding
Deutsche Forschungsgemeinschaft (SFB 1348 B5)
- Christian Klämbt
Deutsche Forschungsgemeinschaft (Kl 588 / 29)
- Christian Klämbt
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
Acknowledgements
We are grateful to all our colleagues for many discussions and P Deing, K Krukkert, K Mildner, and E Naffin for excellent technical assistance. T Zobel for help using the Elyra 7 microscope. B Zalc and KA. Nave for critical reading of the manuscript and many thoughtful suggestions. This work was supported by the Deutsche Forschungsgemeinschaft through funds to CK. (SFB 1348, B5, Kl 588/29).
Copyright
© 2023, Rey, Ohm 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.
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