1. Developmental Biology

Loss of dihydroceramide desaturase drives neurodegeneration by disrupting endoplasmic reticulum and lipid droplet homeostasis in glial cells

  1. Yuqing Zhu
  2. Kevin Cho
  3. Haluk Lacin
  4. Yi Zhu
  5. Jose T DiPaola
  6. Beth A Wilson
  7. Gary Patti
  8. James B Skeath  Is a corresponding author
  1. Department of Genetics, Washington University School of Medicine, United States
  2. Department of Chemistry, Washington University in St. Louis, United States
  3. Department of Medicine, Washington University School of Medicine, United States
  4. Center for Mass Spectrometry and Metabolic Tracing, Washington University in St. Louis, United States
  5. Division of Biological and Biomedical Systems, University of Missouri-Kansas City, United States
https://doi.org/10.7554/eLife.99344.3
Share this article
Cite this article
  1. Yuqing Zhu
  2. Kevin Cho
  3. Haluk Lacin
  4. Yi Zhu
  5. Jose T DiPaola
  6. Beth A Wilson
  7. Gary Patti
  8. James B Skeath
(2025)
Loss of dihydroceramide desaturase drives neurodegeneration by disrupting endoplasmic reticulum and lipid droplet homeostasis in glial cells
eLife 13:RP99344.
https://doi.org/10.7554/eLife.99344.3
  2. Download BibTeX
  3. Download .RIS

eLife Assessment

This study on the loss of DEGS1 in the developing larval brain convincingly shows the accumulation of dihydroceramide in the CNS which induces severe alterations in the morphology of glial subtypes as well as a reduction in glial number. The localization of DEGS1/ifc primarily to the ER is also compelling and interesting, and the loss of DEGS1/ifc clearly drives ER expansion and reduces the levels of TGs. This is an important contribution to the role of lipid metabolism in neural development and disease.

https://doi.org/10.7554/eLife.99344.3.sa0

Abstract

Dihydroceramide desaturases convert dihydroceramides to ceramides, the precursors of all complex sphingolipids. Reduction of DEGS1 dihydroceramide desaturase function causes pediatric neurodegenerative disorder hypomyelinating leukodystrophy-18 (HLD-18). We discovered that infertile crescent (ifc), the Drosophila DEGS1 homolog, is expressed primarily in glial cells to promote CNS development by guarding against neurodegeneration. Loss of ifc causes massive dihydroceramide accumulation and severe morphological defects in cortex glia, including endoplasmic reticulum (ER) expansion, failure of neuronal ensheathment, and lipid droplet depletion. RNAi knockdown of the upstream ceramide synthase schlank in glia of ifc mutants rescues ER expansion, suggesting dihydroceramide accumulation in the ER drives this phenotype. RNAi knockdown of ifc in glia but not neurons drives neuronal cell death, suggesting that ifc function in glia promotes neuronal survival. Our work identifies glia as the primary site of disease progression in HLD-18 and may inform on juvenile forms of ALS, which also feature elevated dihydroceramide levels.

eLife digest

Neurodegenerative diseases affect around 50 million people worldwide. They arise when neurons deteriorate and die. Neurodegeneration was thought to result from defects within neurons. But recent studies have shown that changes in brain cells known as glial cells – which surround, protect and nourish neurons – can also trigger this process.

The fat composition of the surrounding plasma membrane of glial cells differs from neurons and contains high levels of sphingolipids. These lipids regulate membrane fluidity – the movement of molecules within and through the membrane – and are also critical for cell signaling and the formation of nerve-insulating myelin sheaths.

All complex sphingolipids, such as sphingomyelins and gangliosides, are derived from ceramide. Enzymes called DEGS1 produce ceramide from dihydroceramide in the endoplasmic reticulum. Ceramides are then transported to the Golgi complex, where they are modified into complex sphingolipids.

In humans, mutations in the gene encoding DEGS1 cause a loss of the myelin sheath leading to a fatal neurodegenerative condition in children called hypomyelinating leukodystrophy-18. So far, it was unclear whether the accumulation of dihydroceramide or the depletion of ceramide might alter the function of neurons and glia enough to trigger neurodegeneration.

Zhu et al. addressed this question using genetically modified fruit fly larvae that lacked the DEGS1 gene. They discovered that in fruit flies, DEGS1 protects the nervous system from neurodegeneration by supporting the development and function of glial cells. In flies that lacked the gene, dihydroceramide accumulated in the central nervous system, which enlarged the endoplasmic reticulum in glial cells, causing them to swell. These morphological defects inhibited their ability to enwrap the cell bodies and axons of neurons with a supporting glial sheath. This suggests that a faulty DEGS1 gene may drive neurodegeneration as a secondary consequence of glial dysfunction.

By establishing a simple model system, Zhu et al. provide insight into how glial cells may contribute to neurodegeneration. Their results indicate that DEGS1 loss causes structural and functional defects in glial cells, preventing them from supporting neurons and ultimately leading to neurodegeneration. Because the ceramide synthesis pathway is conserved between fruit flies and humans, similar mechanisms likely contribute to neuronal degeneration in patients with DEGS1 mutations. A deeper understanding of these pathways could help identify strategies to slow the progression of hypomyelinating leukodystrophy-18 and open new avenues for therapy.

Introduction

Sphingolipids are key structural and functional components of the cell membrane in all eukaryotic cells and are enriched in glial cells, such as oligodendrocytes, where they comprise up to 30% of all membrane lipids (Jackman et al., 2009). All complex sphingolipids, like glycosylsphingolipids and sphingomyelin, derive from ceramide (Ghosh et al., 2013; Vacaru et al., 2013; Hannun and Obeid, 2018). In the de novo ceramide biosynthesis pathway, dihydroceramide desaturases, such as DEGS1, produce ceramide from dihydroceramide by catalyzing the formation of a trans double bond between carbons 4 and 5 of the sphingoid backbone, which enhances conformational plasticity (Li et al., 2002; Yasuda et al., 2016). In humans, bi-allelic mutations in DEGS1 cause hypomyelinating leukodystrophy-18 (HLD-18), a progressive, often fatal pediatric neurodegenerative disease marked by cerebral atrophy, white matter reduction, and hypomyelination (Dolgin et al., 2019; Karsai et al., 2019; Pant et al., 2019). The primary neural cell type impacted by loss of DEGS1 function and the cell biology of how disruption of ceramide synthesis leads to neurodegeneration remain unknown.

The de novo ceramide biosynthetic pathway is well conserved among higher metazoans (Hannun and Obeid, 2018; Fyrst et al., 2004; Acharya and Acharya, 2005; Dunn et al., 2019; Pan et al., 2023). De novo ceramide synthesis occurs in the endoplasmic reticulum (ER) and starts with the rate-limiting activity of the serine palmitoyltransferase (SPT) complex, which condenses serine and palmitoyl-CoA (lauoryl-CoA in flies) to form 3-Ketosphinganine, which is converted to sphinganine by 3-Ketosphinganine reductase. Ceramide synthases condense sphinganine with acyl-CoA to generate dihydroceramide, which is converted to ceramide by dihydroceramide desaturases. Ceramide is then efficiently transported by the specific ceramide transporter CERT from the ER to the Golgi (Kumagai et al., 2005), where it undergoes headgroup modifications to produce complex sphingolipids that eventually translocate to the plasma membrane (Kobayashi and Menon, 2018). Mutations in most members of the SPT complex, ceramide synthases, and DEGS1 lead to neurodegeneration (Hannun and Obeid, 2018; Dunn et al., 2019; Pan et al., 2023), identifying the de novo ceramide biosynthesis pathway as a hotspot for neurodegenerative disease mutations.

Consistent with its central role in ceramide biogenesis, reduction or loss of DEGS1 function in human patients or cell lines, mice, zebrafish, and flies drives dihydroceramide accumulation and ceramide depletion (Karsai et al., 2019; Pant et al., 2019; Holland et al., 2007; Jung et al., 2017). HLD-18 patients display reduced myelin sheath thickness in peripheral nerves, and knockdown of DEGS1 function in zebrafish reduces the number of myelin basic protein-positive oligodendrocytes (Karsai et al., 2019; Pant et al., 2019), suggesting DEGS1 regulates Schwann cell and oligodendrocyte development. In Drosophila, genetic ablation of infertile crescent (ifc), the fly DEGS1 ortholog, drives activity-dependent photoreceptor degeneration (Jung et al., 2017), suggesting that ifc function is crucial for neuronal homeostasis. In addition, forced expression of a wild-type ifc transgene in neurons, glia, or muscles was shown to rescue the ifc mutant phenotype (Jung et al., 2017). Whether ifc/DEGS1 acts primarily in glia, neurons, or other cells to regulate nervous system development then remains to be determined.

Research on DEGS1 points to dihydroceramide accumulation as the driver of nervous system defects caused by DEGS1 deficiency. Pharmacological or genetic inhibition of ceramide synthase function, which should reduce dihydroceramide levels, suppresses the observed reduction of MBP-positive oligodendrocytes in zebrafish and the activity-dependent photoreceptor degeneration in flies triggered by reduction of DEGS1/ifc function (Pant et al., 2019; Jung et al., 2017). Dihydroceramide accumulation may also contribute to other neurodegenerative diseases, as gain of function mutations in components of the SPT complex cause juvenile amyotrophic lateral sclerosis due to elevated sphingolipid biosynthesis, with dihydroceramides showing the greatest relative increase of all sphingolipids (Lone et al., 2022; Lone et al., 2023; Syeda et al., 2024; Dohrn et al., 2024; Srivastava et al., 2023). How dihydroceramide accumulation alters the cell biology of neurons, glia, or both to trigger myelination defects and neurodegeneration is unclear.

With its vast genetic toolkit, Drosophila is a powerful system in which to dissect the genes and pathways that regulate glial development and function (Freeman and Doherty, 2006; Coutinho-Budd and Freeman, 2013). In flies, six morphologically and functionally distinct glial subtypes regulate nervous system development and homeostasis (Yildirim et al., 2019; Corty and Coutinho-Budd, 2023). The surface perineurial and subperineurial glia act as physiochemical barriers to protect the nervous system and control metabolite exchange with the hemolymph, carrying out a similar function as the human blood–brain barrier (Stork et al., 2008; Volkenhoff et al., 2015). Residing between the surface glia and the neuropil, cortex glia ensheathe the cell bodies of neuroblasts and neurons in the CNS in protective, nutritive, honeycomb-like membrane sheaths. Ensheathing glia define the boundary of the neuropil and insulate axons, dendrites, and synapses from neuronal cell bodies in the CNS (Freeman, 2015). Astrocyte-like glia extend fine membrane protrusions that infiltrate the neuropil and form a meshwork of cellular processes that ensheathe synapses and regulate synaptic homeostasis in the CNS (Freeman, 2015). In the PNS, wrapping glia reside internally to the surface glia and insulate axons in the PNS to enhance neuronal signaling and provide energy support.

Using the Drosophila model, we found that ifc acts primarily in glia to regulate CNS development, with its loss disrupting glial morphology. Our work supports a model in which inappropriate accumulation and retention of dihydroceramide in the ER drives ER expansion, glial swelling, and the failure of glia to enwrap neurons, ultimately leading to neuronal degeneration as a secondary consequence of glial dysfunction. Given the conserved nature of de novo ceramide biosynthesis, our findings likely illuminate the exact mechanism through which elevated dihydroceramide levels drive neuronal degeneration and cell death in flies and humans.

Results

ifc contributes to the regulation of developmental timing and CNS structure

In an EMS-based genetic screen, we uncovered three non-complementing mutations that, when homozygous or trans-heterozygous to each other, resulted in identical phenotypes, including a 3-day or greater delay in reaching the late-third larval instar stage, reduced brain size, progressive ventral nerve cord elongation, axonal swelling, and lethality at the late larval or early pupal stage (Figure 1A; data not shown). Whole-genome sequencing revealed that ifc was independently mutated in each line: ifcjs1 and ifcjs2 encode V276D and G257S missense mutations, respectively, and ifcjs3 encodes a W162* nonsense mutation (Figure 1A, B). Sanger sequencing also uncovered a missense mutation, E241K, in the molecularly uncharacterized ifc (Jackman et al., 2009) allele (Figure 1B). All four mutations reside in the fatty acid desaturase domain, the hotspot for mutations in human DEGS1 that cause HLD-18 (Figure 1B, B').

Figure 1 with 7 supplements see all
Download asset Open asset
ifc regulates CNS and glial morphology.

(A) Ventral views of late-third instar larvae of indicated genotype showing 3xP3 RFP labeling of CNS and nerves. Arrowheads indicate nerve bulges; scale bar is 200 μm. Schematic of Ifc (B) and human DEGS1 (B’) proteins indicating location and nature of ifc mutations and 15 HLD-18-causing DEGS1 mutations (Dolgin et al., 2019; Karsai et al., 2019; Pant et al., 2019). (C) Schematic of de novo ceramide biosynthesis pathway indicating the subcellular location of ceramide synthesis and ceramide modifications. (D) Chemical structure of dihydroceramide and ceramide; arrow indicates trans carbon–carbon double bond between C4 and C5 in the sphingoid backbone created by the enzymatic action of Ifc/DEGS1. (E) Normalized quantification of the relative levels of dihydroceramide, ceramide, and six related sphingolipid species in the dissected CNS of wild-type and ifc−/− late-third instar larvae. (F) Ventral views of Drosophila CNS and peripheral nerves in wild-type and ifc−/− mutant late-third instar larvae labeled for NCAD to mark the neuropil, HRP to label axons, RFP to label glia, Dpn to label neuroblasts, ELAV to label neurons, and fatty acid binding protein (FABP) to label cortex glia. Anterior is up; scale bar is 100 μm for whole CNS images and 20 μm for peripheral nerve image. Statistics: **p < 0.01, ***p < 0.001, ****p < 0.0001.

As a prior study reported that a CRISPR-generated, gene-specific deletion of ifc, ifc-KO, resulted in early larval lethality (Jung et al., 2017), we first confirmed the genetic nature of our ifc alleles because they caused late larval-early pupal lethality. Complementation crosses of each ifc allele against a deficiency of the region (Df(2L)BSC184) and ifc-KO revealed that all combinations, including larvae trans-heterozygous for ifc-KO over Df(2L)BSC184, survived to the late larval-early pupal stage and yielded phenotypes identical to those detailed above for the newly uncovered ifc alleles. Flies homozygous for ifc-KO, however, died as early larvae. Further analysis uncovered second site mutation(s) in the 21E2 chromosomal region responsible for the early lethal phenotype of the ifc-KO chromosome (see Methods). When uncoupled from these mutation(s), larvae homozygous for the ‘clean’ ifc-KO chromosome developed to the late larval-early pupal stage and manifested phenotypes identical to the other ifc alleles. This analysis defined the correct lethal phase for ifc and identified our ifc alleles as strong loss of function mutations.

Loss of ifc function drives ceramide depletion and dihydroceramide accumulation

As ifc/DEGS1 converts dihydroceramide to ceramide, we used untargeted lipidomics on whole larvae and the isolated CNS from wild-type and ifc-KO/ifcJS3 larvae (hereafter termed ifc−/− larvae) to assess the effect of loss of ifc function on metabolites in the ceramide pathway (Figure 1C). Loss of ifc function resulted in a near complete loss of ceramides and a commensurate increase in dihydroceramides in the CNS and whole larvae (Figure 1E, Figure 1—figure supplement 1). Sphinganine, the metabolite directly upstream of dihydroceramide, also exhibited a significant increase in its levels in the absence of ifc function, while metabolites further upstream were unchanged in abundance or undetectable (Figure 1E, Figure 1—figure supplement 1). Ceramide derivatives like sphingosine, CPE, and glucosyl-ceramide (Gl-Cer) were reduced in levels and replaced by their cognate dihydroceramide forms (e.g., Glc-DiCer) (Figure 1E, Figure 1—figure supplement 1). Loss of the enzymatic function of Ifc then drives dihydroceramide accumulation and ceramide loss.

ifc governs glial morphology and survival

To connect this metabolic profile to a cellular phenotype, we assayed ifc function in the CNS. We leveraged the expression of fatty acid binding protein (FABP) as a marker of cortex glia (Kis et al., 2015) and that of the M{3xP3-RFP.attP} phi-C31 ‘landing pad’ transgene, which resided in the isogenic target chromosome of our screen, as a marker of most glia (Figure 1—figure supplements 2–3). In addition, we labeled neuroblasts with Deadpan (Dpn), neurons with ELAV, and axons with N-Cadherin (NCad). In ifc−/− larvae, we observed a clear reduction in Dpn-positive neuroblasts in the optic lobe, swelling of glia in peripheral nerves, enhanced RFP expression in the CNS, and the presence of large swollen, cortex glia identified by RFP labeling and FABP expression (Figure 1F, Figure 1—figure supplements 4 and 5). In wild-type larvae, cortex glia display compact cell bodies and fully enwrap individual neuronal cell bodies with their membrane sheaths (Figure 1F, Figure 1—figure supplement 4; Kis et al., 2015). In ifc−/− larvae, cortex glia display swollen cell bodies, fail to fully enwrap neuronal cell bodies, displace neurons from their regular arrangement, and appear to contain brightly fluorescent RFP-positive aggregates (Figure 1F, Figure 1—figure supplement 6). ifc is then necessary for glial development and function in the larval nervous system. We observed identical CNS phenotypes in larvae homozygous mutant for the ifcjs1 and ifcjs2 alleles (Figure 1—figure supplement 7).

To track the impact of ifc on glial morphology, we combined GAL4 lines specific for each glial subtype with a UAS-linked membrane-tagged GFP transgene (Myr-GFP) and the MultiColor FlpOut system (Kremer et al., 2017; Pfeiffer et al., 2010; Nern et al., 2015). Using this approach, we determined that loss of ifc function affects all CNS glial subtypes except perineurial glia (Figure 2E, E’, J, J’). Cortex glia appeared swollen, failed to enwrap neurons, and accumulated large amounts of Myr-GFP+ internal membranes (Figure 2A, A’, F, F’). Ensheathing glia (Figure 2B, B’, G, G’), and subperineurial glia (Figure 2D, D’, I, I’) also displayed swollen, disorganized cell bodies and accumulated Myr-GFP+ internal membranes. Astrocyte-like glia displayed smaller cell bodies, reduced membrane extensions, and disrupted organization along the dorsal–ventral nerve cord (Figure 2C, C’). We conclude that ifc regulates the morphology of most glial subtypes in the larval CNS.

Figure 2 with 1 supplement see all
Download asset Open asset
Loss of ifc disrupts glial morphology.

(A-E’) High magnification ventral views and X–Z and Y–Z projections of the nerve cord of wild-type and ifc−/− late-third instar larvae labeled for ELAV (magenta) for neurons and Myr-GFP (green) for cell membranes of indicated glial subtype. Anterior is up; scale bar is 40 μm. (F-J’) High magnification views of individual glial cells of indicated glial subtype in the nerve cord of wild-type and ifc−/− larvae created by the MultiColor-FlpOut method (Nern et al., 2015). Anterior is up; scale bar is 20 μm. (K–N) Quantification of total number of indicated glial subtype in the nerve cord of wild-type and ifc−/− late-third instar larvae (n = 7 for K, L, N; n = 6 for M). Statistics: *p < 0.05, ****p < 0.0001, and ns, not significant. The full genotype of flies shown in this figure can be found in Supplementary file 1.

Next, we asked if loss of ifc function alters the number of each glial subtype. Using the same glial subtype-specific GAL4 lines to drive a nuclear-localized GFP transgene, we counted the total number of all CNS glial subtypes, except perineurial glia, in wild-type and ifc−/− larvae. To remove the small size of the brain in ifc−/− larvae as a confounding factor, we focused our analysis on the ventral nerve cord. The number of subperineurial glia was unchanged between the two genotypes, but we observed a 12%, 40%, and 72% reduction in the number of astrocyte-like, ensheathing, and cortex glia, respectively, in ifc−/− larvae relative to wild-type (Figure 2K–N). Our data reveal a broad role for ifc in regulating glial cell morphology and number in the Drosophila larval CNS. Subsequent experiments revealed that a reduction in cell proliferation and an increase in apoptosis both contribute to the observed reduction in the number of cortex glia (Figure 2—figure supplement 1).

ifc acts in glia to regulate glial and CNS development

Prior work in zebrafish showed that DEGS1 knockdown reduced the number of myelin basic protein-positive oligodendrocytes (Pant et al., 2019); in flies, loss of ifc function in the eye drove photoreceptor degeneration (Jung et al., 2017). Neither study uncovered the cell type in which ifc/DEGS1 acts to regulate neural development. To address this question, we used the GAL4/UAS system, RNAi-mediated gene depletion, and gene rescue approaches to see if ifc acts in neurons or glia to control glial development and CNS morphology. First, we used a UAS-linked ifc-RNAi transgene to deplete ifc function in all neurons (elav-GAL4; repo-GAL80) or all glia (repo-GAL4). Focusing on FABP-positive cortex glia due to their easily scorable phenotype, we found that pan-glial, but not pan-neuronal, knockdown of ifc recapitulated the swollen cortex glia phenotype observed in ifc mutant larvae (Figure 3C, D, J).

Figure 3 with 1 supplement see all
Download asset Open asset
fc acts in glia to regulate CNS structure and glial morphology.

(A–I) Ventral views of photomontages of the CNS of late-third instar larvae labeled for fatty acid binding protein (FABP) (grayscale) to mark cortex glia in late-third instar larvae of indicated genotype. Neuronal-specific transgene expression was achieved by using elav-GAL4 combined with repo-GAL80; glial-specific transgene expression was achieved by using repo-GAL4. (J–K) Quantification of the number of swollen cortex glia in the abdominal segments of the CNS of late-third instar larvae of the indicated genotype for the RNAi (J) and gene rescue assays (K). Statistics: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant. The full genotype of flies shown in this figure can be found in Supplementary file 1.

To complement our RNAi approach, we asked if GAL4-driven expression of a wild-type Drosophila ifc or human DEGS1 transgene rescued the ifc−/− CNS phenotype. In the absence of a GAL4 driver, the ifc transgene drove weak rescue of the cortex glia phenotype (Figure 3E, K), consistent with modest GAL4-independent transgene expression reported for UAS-linked transgenes (Southall et al., 2013). Pan-neuronal expression of ifc drove modest rescue of the ifc CNS phenotype beyond that observed for the ifc transgene alone (Figure 3F, K), but pan-glial expression of ifc fully rescued the ifc mutant cortex glia phenotype and other CNS phenotypes (Figure 3H, K, Figure 3—figure supplement 1). Identical experiments using the human DEGS1 transgene revealed that only pan-glial DEGS1 expression provided rescuing activity, albeit at much weaker levels than the Drosophila ifc transgene (Figure 3G, I, K). Pan-glial expression of the Drosophila ifc transgene was, in fact, ~15-fold more potent than the human DEGS1 transgene at rescuing the ifc lethal phenotype to adulthood: When ifc was expressed in all glia, 57.9% of otherwise ifc mutant flies survived to adulthood (n = 2452), but when ifc was replaced by DEGS1 only 3.9% of otherwise ifc mutant flies reached adulthood (n = 1303). No ifc mutant larvae reached adulthood in the absence of either transgene (n = 1030). We infer that ifc acts primarily in glia to govern CNS development and that human DEGS1 can partially substitute for ifc function in flies despite a difference in the preferred length of the sphingoid backbone in flies versus mammals (Fyrst et al., 2004).

Results of our gene rescue experiments conflict with a prior study on ifc in which expression of ifc in neurons was found to rescue the ifc phenotype (Jung et al., 2017). In this context, we note that elav-GAL4 drives UAS-linked transgene expression not just in neurons, but also in glia at appreciable levels (Berger et al., 2007; Lacin et al., 2024) and thus needs to be paired with repo-GAL80 to restrict GAL4-mediated gene expression to neurons. Thus, ‘off-target’ expression in glial cells may account for the discrepant results. It is, however, more difficult to reconcile how neuronal or glial expression of ifc would rescue the observed lethality of the ifc-KO chromosome given the presence of additional lethal mutations in the 21E2 region of the second chromosome.

ifc is predominately expressed in glia and localizes to the ER

Next, we tracked ifc expression in the CNS via RNA in situ hybridization and an ifc-T2A-GAL4 transcriptional reporter. RNA in situ hybridization revealed that ifc is widely expressed in the CNS (Figure 4A), most obviously in the distinctive star-shaped astrocyte-like glia (Figure 4B), which are marked by Ebony expression (Ziegler et al., 2013). RNA in situ hybridization was not ideal for tracing ifc expression in other cells, likely due to signal diffusion. Thus, we paired the ifc-T2A-GAL4 transcriptional reporter with a nuclear RFP (nRFP) transgene (He et al., 2019) and confirmed authenticity of the ifc-T2A-GAL4 line by its strong expression in astrocyte-like glia (Figure 4C). Using this approach, we observed strong nRFP expression in all glial cells and modest nRFP expression in all neurons (Figure 4D, E, Figure 4—figure supplement 1), suggesting ifc is transcribed at higher levels in glial cells than neurons in the larval CNS.

Figure 4 with 1 supplement see all
Download asset Open asset
Loss of ifc drives ER expansion in cortex glia.

Dorsal (A, B–B’’, C–C’’) and ventral (D–D’’, E–E’’) views of the CNS of late-third instar wild-type larvae labeled for ifc RNA (gray in A; magenta in B’), ifc-GAL4>nRFP (magenta; C’–E’), EBONY to mark astrocytes (green; B, C), REPO to mark glia (green; D), and ELAV to mark neurons (green; E). Panels D–D’’ and E–E’’ show surface and interior views, respectively, along the Z-axis on the ventral side of the nerve cord. Arrowheads in E–E’’ identify neurons with low-level ifc-GAL4 expression. High magnification ventral views of thoracic segments in the CNS of wild-type late-third instar larvae labeled for GFP (green; F, G), CNX99A (magenta; F’), and ESYT (magenta; G’). (H–M) Late-third instar larvae of indicated genotype labeled for 3xP3-RFP (green; H’–M’), CNX99A (magenta; H, K), GOLGIN84 (magenta; I, L), and LAMP (magenta; J, M). Anterior is up; scale bar is 100 μm for panel A and 30 μm for panels B–M.

Using a fosmid transgene that harbors a GFP-tagged version of ifc flanked by ~36 kb of its endogenous genomic region and molecular markers for ER, cis-Golgi, and trans-Golgi (Sarov et al., 2016; Bergeron et al., 1994; Kikuma et al., 2017; Park et al., 2022), we found that the Ifc-GFP colocalized strongly with the ER markers Calnexin 99A (CNX99A) and ESYT (Figure 4F, F’, G, G’, Figure 1—figure supplement 7) and weakly with the cis-Golgi marker GOLGIN84 and the trans-Golgi marker GOLGIN245 (Figure 4—figure supplement 1). Our results indicate that Ifc localizes primarily to the ER, aligning with the presumed site of de novo ceramide biosynthesis and prior work on DEGS1 localization in cell lines (Karsai et al., 2019; Kawano et al., 2006).

Loss of ifc drives ER expansion and lipid droplet loss in cortex glia

We next asked if loss of ifc function altered ER, Golgi, or lysosome morphology. Focusing on cortex glia, we observed a clear expansion of the ER marker CNX99A (Figure 4H, H’, K, K’), a mild enrichment of the Golgi markers, GOLGIN84 and GOLGIN245, in diffuse ‘clouds’ (Figure 4I, I’, L, L’, Figure 4—figure supplement 1), and a reduction in expression of the lysosome marker LAMP (Figure 4J, J’, M, M’) in ifc−/− larvae. The expansion of ER markers in ifc mutant larvae compelled us to obtain high-resolution views of organelle structure in cortex glia via transmission electron microscopy (TEM). In wild-type, cortex glia display a compact cytoplasm that surrounds a large nucleus (Figure 5A, B) and extend glial sheaths that fully enwrap adjacent neuronal cell bodies (black arrows; Figure 5C, D). In ifc−/− larvae, cortex glia display enlarged cell bodies with a maze-like pattern of internal membranes (solid white arrows; Figure 5A’, B’, E, E’) and fail to enwrap neurons (hollow white arrows; Figure 5C’, D’). The internal membrane structures appear to assume an ER-like identity, as we observed significant overlap between the ER marker CNX99A and the membrane marker Myr-GFP when Myr-GFP was driven by a cortex glia-specific GAL4 line (Figure 5F). We observed similar yet milder effects on cell swelling and internal membrane accumulation in subperineurial and wrapping glia in abdominal nerves purple and pink shading, respectively (Figure 5G, G’, H, H’), indicating that loss of ifc drives internal membrane accumulation in and swelling of multiple glial subtypes.

Figure 5 with 1 supplement see all
Download asset Open asset
Loss of ifc leads to internal membrane accumulation and lipid droplet loss in cortex glia.

(A–E’) Transmission electron microscopy (TEM) images of cortex glia cell body (A, A’, B, B’) and neuronal cell bodies (C, C’, D, D’) at low (A, A’) and high (B, B’, C, C’, D, D’) magnification in the nerve cord of wild-type (A–D) and ifc−/− (A’, D’, E, E’) late-third instar larvae. (A, A’) Dotted lines demarcate cell boundary of cortex glia; yellow squares highlight regions magnified in B, B’, E’. Scale bar is 3 μm for A, A’ and 1 μm for B, B’. (B, B’) Cy denotes cytoplasm; Nu denotes nucleus. Solid white arrows highlight the layered internal membranes that occupy the cytoplasm of ifc−/− cortex glia. (C, C’, D, D’) Black arrows highlight cortex glia membrane extensions that enwrap neuronal cell bodies; hollow white arrows denote the absence of cortex glia membrane extensions; white asterisk denotes lipid droplets. Scale bar is 2 μm. (E, E’) An additional example of membrane-filled cortex glia cell body in ifc−/− larvae. Scale bar is 2 μm for E and 1 μm for E’. (F) Cortex glia in ifc mutant larvae labeled for Myr-GFP (green) to label membranes and CNX99A to label ER membranes. Scale bar is 30 μm. (G, H) Black and white and colored TEM cross-sections of peripheral nerves in wild-type and ifc−/− late-third instar larvae. Blue marks perineurial glia; purple marks subperineurial glia; pink marks wrapping glia. Scale bar: 2 μm. High magnification ventral views of abdominal segments in the ventral nerve cord of wild-type (I) and ifc mutant (I') third instar larvae labeled for BODIPY (green) to mark lipid droplets and fatty acid binding protein (FABP) (magenta) to label cortex glia. Anterior is up; scale bar is 30 μm. (J) Graph of log-fold change of transcription of five genes that promote membrane lipid synthesis in ifc−/− larvae relative to wild-type. A dotted line indicates a log2 fold change of 0.5 in the treatment group compared to the control group. (K–M) Quantification of the number (G) and area of lipid droplets (H, I) in the dissected CNS of wild-type and ifc−/− larvae. Statistics: ****p < 0.0001, and ns, not significant.

TEM analysis also revealed a near complete depletion of lipid droplets in the CNS of ifc−/− larvae (compare Figure 5A, C, D–A’, C’, D’; lipid droplets marked by asterisk; Figure 5M), which we confirmed using BODIPY to mark neutral lipids in FABP-positive cortex glia (Figure 5I–L). In the CNS, lipid droplets form primarily in cortex glia (Kis et al., 2015) and are thought to contribute to membrane lipid synthesis through their catabolism into free fatty acids versus acting as an energy source in the brain (Grabner et al., 2021). Consistent with the possibility that increased membrane lipid synthesis drives lipid droplet reduction, RNA-seq assays of dissected nerve cords revealed that loss of ifc drove transcriptional upregulation of genes that promote membrane lipid biogenesis, such as SREBP, the conserved master regulator of lipid biosynthesis, SCAP, an activator of SREBP, and Pcyt1/Pcyt2, which promote phosphatidylcholine (PC) and phosphatidylethanolamine (PE) synthesis (Eberlé et al., 2004; Osborne and Espenshade, 2009; Jacquemyn et al., 2017). The spliced form of Xbp-1 mRNA (Xbp-1s) (Figure 5J), which promotes membrane lipid synthesis required for ER biogenesis and is activated by the unfolded protein response (UPR) (Sriburi et al., 2007), is also upregulated in the CNS of ifc mutant larvae. Most ER chaperones, which are typically transcriptionally upregulated upon UPR activation (Bernales et al., 2006), were however downregulated (Figure 5—figure supplement 1), suggesting that in this case, misfolded protein is not the factor that triggers UPR activation and increased ER membrane biogenesis upon loss of ifc.

Loss of ifc increases the saturation levels of triacylglycerols and membrane phospholipids

Lipid droplets are composed largely of triacylglycerols (TGs), Jacquemyn et al., 2017 and we observed a five- and threefold drop in TG levels in the CNS and whole larvae of ifc mutant larvae relative to wild-type (Figure 6A). Our lipidomics analysis also revealed a shift of TGs toward higher saturation levels in the absence of ifc function (Figure 6A–C). Consistent with this, we observed transcriptional upregulation of most genes in the Lands cycle, which remodels phospholipids by replacing existing fatty acyl groups with new fatty acyl groups (Figure 5—figure supplement 1; Moessinger et al., 2014; Moessinger et al., 2011; Harayama et al., 2014; O’Donnell, 2022). As TG breakdown results in free fatty acids that can be used for membrane phospholipid synthesis, we asked if changes in TG levels and saturation were reflected in the levels or saturation of the membrane phospholipids PC, PE, and phosphatidylserine (PS). In the absence of the ifc function, PC and PE exhibited little change in quantity (Figure 6D, G), but the levels of the less abundant PS were increased threefold in the CNS of ifc mutant larvae relative to wild-type (compare Figure 6J to Figure 6D, G). All three phospholipids, however, displayed increased saturation levels: The relative levels of all PC, PE, and PS species were reduced, except for the most saturated form of each phospholipid – 18:1/18:1 – which is increased (Figure 6E, F, H–L). This increase was more pronounced in the CNS than whole larvae (Figure 6F, I, L), implying that loss of ifc function creates greater demand for lipid remodeling in the CNS.

Figure 6
Download asset Open asset
Phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), and triacylglycerol (TG) exhibit higher saturation levels in a CNS-specific manner in ifc mutant late-third instar larvae.

Quantification of total (A) and species-specific (B, C) TGs in whole larvae (A, C) and dissected CNS (A–C) of wild-type and ifc−/− larvae. Quantification of total (D) and species-specific (E, F) PCs in whole larvae (D, F) and dissected CNS (D–F) of wild-type and ifc−/− larvae. Quantification of total (G) and species-specific (H, I) PEs in whole larvae (G, H) and dissected CNS (G–I) of wild-type and ifc−/− larvae. Quantification of total (J) and species-specific (K, L) PSs in the whole larvae (J, L) and dissected CNS (J–L) of wild-type and ifc−/− larvae. Statistics: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant.

Reduction of dihydroceramide synthesis suppresses the ifc CNS phenotype

Following its synthesis, ceramide is transported by CERT from the ER to the Golgi, but CERT is less efficient at transporting dihydroceramide than ceramide (Pan et al., 2023). The expanded nature of the ER in ifc mutant larvae supports a model in which loss of ifc triggers excessive accumulation and retention of dihydroceramide in the ER due to CERT’s inefficiency of transporting dihydroceramide to the Golgi, driving ER expansion and glial swelling. If this model is correct, reduction of dihydroceramide levels should suppress the ifc CNS phenotype. In agreement with this model, we found that the schlankG0365 loss-of-function allele dominantly suppressed the enhanced RFP expression (Figure 7M) and CNS elongation phenotypes of ifc (Figure 7N). We also found that glial-specific depletion of schlank suppressed the internal membrane accumulation (Figure 7A, B), reduced lipid droplet size (Figure 7C, D, O), glial swelling (Figure 7E, F, Figure 7—figure supplement 1), enhanced RFP expression (Figure 7M), CNS elongation (Figure 7N), and reduced optic lobe (Figure 1—figure supplement 5) phenotypes observed in otherwise ifc mutant larvae. Our data support the model that inappropriate retention of dihydroceramide in the ER drives ER expansion and glial swelling and dysfunction.

Figure 7 with 3 supplements see all
Download asset Open asset
Glial-specific knockdown of ifc triggers neuronal cell death.

Transmission electron microscopy (TEM) images of cortex glia cell body (A, B) and neuronal cell bodies (C, D) at low (A) and high (B–D) magnifications in the nerve cord of ifc−/−; repo>schlank RNAi late-third instar larvae. Dotted lines demarcate cell boundary of cortex glia; yellow squares highlight regions magnified in A. Scale bar: 3 μm for A, 1 μm for B, and 2 μm for C, D. (B) Cy denotes cytoplasm; Nu denotes nucleus. (C, D) Black asterisk denotes lipid droplets. Ventral views of abdominal sections of CNS of ifc−/−; UAS-schlank RNAi/+ larvae (E) and ifc−/−; repoGAL4/UAS-schlank RNAi larvae (F) labeled for neurons (ELAV, green) and cortex glia (fatty acid binding protein [FABP], magenta/gray). Scale bar is 30 μm for E, F. Low (G–L) and high (G’–L’, G’’–L’’) magnification views of the brain (G–I) and nerve cord (J–L) of late-third instar larvae of the indicated genotypes labeled for ELAV (magenta or grayscale) and Caspase-3 (green). Arrows indicate regions of high Caspase-3 signal and/or apparent neuronal cell death identified by perforations in the neuronal cell layer. Scale bar is 50 μm for panels G–L and 10 μm for panels G’–L’’. Quantification of CNS elongation (M) and 3xP3 RFP intensity (N) in ifc mutants alone, ifc mutants with one copy of schlank[G0365] loss-of-function allele, or ifc mutants in which schlank function is reduced via RNAi in glial cells. (O) Quantification of the area of lipid droplets in dissected CNS of ifc mutants and ifc mutants in which schlank function is reduced via RNAi in glial cells. Anterior is up in all panels. (P, Q) Quantification of Cleaved Caspase-3 neurons for panels G–I (P) and J–L (Q). Statistics: **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant.

Glial-specific depletion of ifc function triggers neuronal cell death

Loss of DEGS1/ifc in human and flies has been shown to promote neurodegeneration and neuronal cell death (Karsai et al., 2019; Pant et al., 2019; Jung et al., 2017), but whether neuronal death results from an intrinsic defect in neurons or is induced by glial dysfunction remains unclear. By using Cleaved Caspase-3 as a marker of dying cells (Yu et al., 2002; Fan and Bergmann, 2010) and ELAV to track neurons, we tracked neuronal cell death in the brain and ventral nerve cord of wild-type and ifc−/− mutant late-third instar larvae. In wild-type, little to no cell death was evident in the brain or nerve cord, and neurons appeared smoothly packed next to each other; in contrast, in ifc−/− larvae, significant cell death was apparent in the brain and to a lesser degree in the nerve cord, with Caspase-3 staining often highlighting small perforations in the neuronal cell layer (Figure 7G, H, P, Figure 1—figure supplement 7). This perforated pattern was associated with and more expansive than Caspase-3 staining, indicating the perforations mark Caspase-positive dying neurons and Caspase-negative dead neurons. Glial-specific depletion of schlank function in otherwise ifc−/− larvae suppressed the neuronal cell death phenotype (Figure 7I, P), supporting the model that loss of ifc function specifically in glia, rather than a specific requirement for ifc function in neurons, drives neuronal cell death. To directly test whether the ifc function in glia is required to guard against neuronal cell death, we used the GAL4-UAS system and RNAi-mediated gene interference to deplete ifc function specifically in glia (repo-GAL4) or in neurons (elav-GAL4; repo-GAL80) and found that glial-specific, but not neuronal-specific, depletion of ifc function drove significant neuronal cell death in the brain and to a greater extent the nerve cord, a phenotype that was enhanced upon removal of one copy of ifc (Figure 7J, L, Q and Figure 7—figure supplement 2). Loss of ifc function then triggers glial dysfunction, which in turn drives neuronal cell death.

Discussion

Our work on ifc supports a model in which loss of ifc/DEGS1 function drives glial dysfunction through the accumulation and inappropriate retention of dihydroceramide in the ER of glia with this proximal defect driving ER expansion, glial swelling, and subsequent neuronal cell death and neurodegeneration. A large increase in dihydroceramide would also impact ER membrane structure: ER membranes are typically loosely packed, thin, semi-fluid structures, with sphingolipids comprising just 3% of ER phospholipids (Jacquemyn et al., 2017). Sphingolipids in general and dihydroceramide in specific are highly saturated lipids, promote lipid order, tighter lipid packing, membrane rigidity, and thicker membranes (O’Brien, 1965; Marcus and Popko, 2002). Our observation of thick ER membranes in cortex glia in ifc−/− larvae (Figure 5) aligns with increased dihydroceramide levels in the ER. In this context, we note that of many clinical observations made on an HLD-18 patient, one was widening of the ER in Schwann cells (Karsai et al., 2019), a finding that when combined with our work suggests that dihydroceramide accumulation in the ER is the proximal cause of HLD-18.

Although ER expansion represents the most proximal effect of loss of ifc/DEGS1 function and dihydroceramide accumulation on cortex glia, other organelles and their functions are likely also disrupted, minimally as a consequence of ER disruption. For example, the apparent reduction of the lysosome marker, LAMP, in cortex glia of ifc mutant larvae correlates with increased RFP levels and the presence of bright RFP puncta or aggregates in this cell type, suggesting impaired lysosome function contributes to increased RFP perdurance and aggregation. Defects in the activity of multiple organelles may collaborate to elicit the full phenotype manifested by loss of ifc/DEGS1 function, resulting in glial dysfunction and subsequent neuronal cell death.

Increased dihydroceramide levels may contribute to a broader spectrum of neurodegenerative diseases than simply HLD-18. Recent work reveals that gain of function mutations in SPTLC1 and SPTLC2, which encode components of the SPT complex that catalyzes the initial, rate-limiting step of de novo ceramide and sphingolipid biosynthesis (Fan and Bergmann, 2010), cause juvenile ALS via increased sphingolipid biosynthesis (Lone et al., 2022; Lone et al., 2023; Syeda et al., 2024; Dohrn et al., 2024; Srivastava et al., 2023). Of all sphingolipids, the relative levels of dihydroceramide were increased the most in patient plasma samples, suggesting that DEGS1 activity becomes limiting in the presence of enhanced SPT activity and that dihydroceramide accumulation contributes to juvenile ALS. Any mutations that increase metabolite flux through the ceramide pathway upstream of DEGS1 may then increase dihydroceramide levels, drive ER expansion and cell swelling, and lead to neurodegeneration, with disease severity predicated on the extent of excessive dihydroceramide accumulation. Model systems, like flies, can harness the power of genetic modifier screens to identify genes and pathways (potential therapeutic targets) that can be tweaked to ameliorate the effect of elevated dihydroceramide levels on neurodegeneration.

Our work appears to pinpoint glia as the cell type impacted by loss of ifc/DEGS1 function, specifically glia that exhibit great demand for membrane biogenesis like cortex glia in the fly and oligodendrocytes or Schwann cells in mammals. In larvae, ifc is expressed at higher levels in glial cells than in neurons, and its genetic function is required at a greater level in glia than neurons to govern CNS development. Our unpublished work on other genes in the ceramide metabolic pathway reveals similar glial-centric expression patterns in the larval CNS to that observed for ifc, suggesting they too function primarily in glia rather than neurons at this stage. In support of this model, a study on CPES, which converts ceramide into CPE, the fly analog of sphingomyelin, revealed that CPES is required in cortex glia to promote their morphology and homeostasis and to protect flies from photosensitive epilepsy (Kunduri et al., 2018). Similarly, ORMDL, a dedicated negative regulator of the SPT complex, is required in oligodendrocytes to maintain proper myelination in mice (Clarke et al., 2019). Given that many glial cell types are enriched in sphingolipids and exhibit a great demand for new membrane biogenesis during phases of rapid neurogenesis and axonogenesis, we believe that glial cells, such as cortex glia, oligodendrocytes, and Schwann cells, rather than neurons manifest a greater need for ifc/DEGS1 function and ceramide synthesis during developmental stages marked by significant nervous system growth.

Will this glial-centric model for ifc/DEGS1 function, and more generally ceramide synthesis, hold true in the adult when neurogenesis is largely complete and the demand for new membrane synthesis in glia dissipates? Recent work in the adult fly eye suggests it may not (Wang et al., 2022). argued that the GlcT enzyme, which converts ceramide to glucosylceramide, is expressed at much higher levels in neurons than glia, and that glucosylceramide is then transported from neurons to glial cells for its degradation, suggesting cell-type-specific compartmentalization of sphingolipid synthesis in neurons and its degradation in glia in the adult. In the future, it will be exciting to uncover whether genes of the sphingolipid metabolic pathway alter their cell-type-specific requirements as a function of developmental stage.

We note that cortex glia are the major phagocytic cell of the CNS and phagocytose neurons targeted for apoptosis as part of the normal developmental process (Freeman and Doherty, 2006; Coutinho-Budd and Freeman, 2013; Yildirim et al., 2019; Corty and Coutinho-Budd, 2023). Thus, while we favor the model that ifc triggers neuronal cell death due to glial dysfunction, it is also possible that increased detection of dying neurons arises due at least in part to a decreased ability of cortex glia to clear dying neurons from the CNS. At present, the large number of neurons that undergo developmentally programmed cell death combined with the significant disruption to brain and ventral nerve cord morphology caused by loss of ifc function renders this question difficult to address. Additional evidence does, however, support the idea that loss of ifc function drives excess neuronal cell death: Clonal analysis in the fly eye reveals that loss of ifc drives photoreceptor neuron degeneration (Jung et al., 2017), indicating that loss of ifc function drives neuronal cell death; cortex-glia-specific depletion of CPES, which acts downstream of ifc, disrupts neuronal function and induces photosensitive epilepsy in flies (Kunduri et al., 2018), indicating that genes in the ceramide pathway can act non-autonomously in glia to regulate neuronal function; recent genetic studies reveal that other glial cells can compensate for impaired cortex glial cell function by phagocytosing dying neurons (Beachum et al., 2024), and we observe that the cell membranes of subperineurial glia enwrap dying neurons in ifc mutant larvae (Figure 7—figure supplement 3), consistent with similar compensation occurring in this background, and in humans, loss-of-function mutations in DEGS1 cause neurodegeneration (Dolgin et al., 2019; Karsai et al., 2019; Pant et al., 2019). Clearly, future work is required to address this question for ifc/DEGS1 and perhaps other members of the ceramide biogenesis pathway.

Altered glial function may not only derive from dihydroceramide buildup in the ER, but also from altered cell membrane structure due to the replacement of ceramide and its derivatives, such as GlcCer and CPE, with the cognate forms of dihydroceramide. Relative to dihydroceramide species, the 4–5 trans carbon–carbon double bond in the sphingoid backbone of ceramide-containing sphingolipids enables them to form more stable hydrogen bonds with water molecules and facilitates their ability to associate with lipids of different saturation levels (Li et al., 2002). A high dihydroceramide to ceramide ratio has been shown to form rigid gel-like domains within model membranes and to destabilize biological membranes by promoting their permeabilization (Vieira et al., 2010; Hernández-Tiedra et al., 2016). As even minor alterations to membrane properties can disrupt glial morphology (Aggarwal et al., 2011), such alterations in membrane rigidity and stability may underlie the failure of cortex glia to enwrap adjacent neurons. The observed increase in saturation of PE, PC, and PS in the CNS of ifc mutant larvae may reflect a compensatory response employed by cells to stabilize cell membranes and guard cell integrity when challenged with elevated levels of dihydroceramide.

The expansion of ER membranes coupled with loss of lipid droplets in ifc mutant larvae suggests that the apparent demand for increased membrane phospholipid synthesis may drive lipid droplet depletion, as lipid droplet catabolism can release free fatty acids to serve as substrates for lipid synthesis. At some point, the depletion of lipid droplets, and perhaps free fatty acids as well, would be expected to exhaust the ability of cortex glia to produce additional membrane phospholipids required for fully enwrapping neuronal cell bodies. Under wild-type conditions, many lipid droplets are present in cortex glia during the rapid phase of neurogenesis that occurs in larvae. During this phase, lipid droplets likely support the ability of cortex glia to generate large quantities of membrane lipids to drive membrane growth needed to ensheath newly born neurons. Supporting this idea, lipid droplets disappear in the adult Drosophila CNS when neurogenesis is complete and cortex glia remodeling stops (Kis et al., 2015). We speculate that lipid droplet loss in ifc mutant larvae contributes to the inability of cortex glia to enwrap neuronal cell bodies. Prior work on lipid droplets in flies has focused on stress-induced lipid droplets generated in glia and their protective or deleterious roles in the nervous system (Bailey et al., 2015; Liu et al., 2015; Liu et al., 2017). Work in mice and humans has found that more lipid droplets are often associated with the pathogenesis of neurodegenerative diseases (Marshall et al., 2014; Farmer et al., 2020; Marschallinger et al., 2020), but our work correlates lipid droplet loss with CNS defects. In the future, it will be important to determine how lipid droplets impact nervous system development and disease.

Methods

Methods details

Fly husbandry

Flies were raised on standard molasses-based food at 25°C. Unless otherwise noted, wild-type is an otherwise wild-type stock harboring the M{3xP3-RFP.attP}ZH-51D insert in an isogenic second chromosome.

Mutagenesis

A standard autosomal recessive forward genetic screen was carried out using 25–30 μm EMS to mutagenize a M{3xP3-RFP.attP}ZH-51D isogenic second chromosome. Homozygous mutant third instar larvae were visually screened under a standard fluorescent microscope for defects in CNS morphology. A detailed description of the screen, all identified genes, and associated whole-genome sequences is described elsewhere (Lacin et al., 2024).

Creation of recombinant lines and identification of second site mutations in ifc-KO chromosome

Standard genetic methods were used to generate fly strains that contained specific combinations of GAL4 and UAS-linked transgenes in the ifcjs3 or ifc-KO background. During this process, we uncovered that the ifc-KO deletion could be unlinked from at least one second chromosomal mutation that caused early larval lethality, resulting in homozygous ifc-KO flies that survived to late L3 to early pupa. A subsequent EMS-based F2 lethal non-complementation screen using the M{3xP3-RFP.attP}ZH-51D isogenic second chromosome as target chromosome and screening against the ifc-KO chromosome identified multiple mutations in four complementation groups that led to an early larval lethal phenotype. Whole-genome sequencing identified these genes as Med15, lwr, Nle, and Sf3b1; all four genes reside within ~90 kb of each other in chromosomal bands 21D1–21E2 near the telomere in chromosome 2L, identifying the site of the associated lethal mutations and suggesting the actual lesion may be a small deletion that removes these genes. The following alleles of these genes are available at Bloomington Stock Center: Med15js1 (Q175*), Med15js2 (Q398*), Med15js3 (C655Y), lwrjs1 (I4N), lwrjs2 (E12K), Nlejs1 (W146R), Nlejs2 (L125P), Nlejs3 (Q242*), Sf3b1js1 (R1160*), Sf3b1js2 (Q1264*), Sf3b1js3 (570 bp deletion at the following coordinates chr2L571720–572290; this deletion removes amino acids 1031 through 1222 and introduces a frameshift into the reading frame).

Gene rescue and in vivo RNAi phenocopy assays

To restrict UAS-linked transgene expression specifically to glia, we used the repoGAL4 driver line. To restrict UAS-linked transgene expression specifically to neurons, we paired the elavGAL4 driver lines, which activate transgene expression strongly in all neurons and moderately in glia, with repoGAL80, which blocks GAL4-dependent activation in glia. We used P{VSH330794} (VDRC 330794) for the RNAi experiments and UAS-ifc (Bischof et al., 2013) and UAS-DEGS1 (BDSC 79200) for the gene rescue assays. All gene rescue experiments were performed in the ifcjs3/ifc-KO background with the UAS-transgene placed into the ifcjs3 background and the GAL4 drivers into the ifc-KO background.

The GAL4-UAS method was also used to assess the phenotype of each glial subtype. In combination with the UAS-Myr-GFP transgene, which labels cell membranes, we used the following GAL4 lines to trace the morphology of each glial subtype in wild-type and ifc−/− larvae: GMR85G01-GAL4 (perineurial glia; BDSC 40436), GMR54C07-GAL4 (subperineurial glia; BDSC 50472), GMR54H02-GAL4 (cortex glia; BDSC 45784), GMR56F03-GAL4 (ensheathing glia; BDSC 77469 and 39157), and GMR86E01-GAL4 (astrocyte-like glia; BDSC45914). The UAS-Myr-GFP transgene was placed into the ifcjs3 background; each glial GAL4 line was placed into the ifc-KO background.

The GAL4-UAS method was used to assess the ability of UAS-ifc and UAS-DEGS1 transgenes to rescue the lethality of otherwise ifc mutant larvae. ifc-KO/CyO Tb; repo-GAL4/TM6B Tb males were crossed to each of the following lines: ifcjs3/CyO Tb; UAS-ifc; ifcjs3/CyO Tb; UAS-DEGS1; ifcjs3/CyO Tb. All adult flies were sorted into Curly and non-Curly and then counted; all non-Curly flies lacked the TM6B Tb balancer indicating they carried repo-GAL4 and thus were likely rescued to viability by glial expression of ifc or DEGS1. Standard Mendelian ratios were then used to predict the expected number of ifc mutant flies if all survived to adulthood. The total number of observed adult ifc mutant flies, identified by lack of Curly wings, was then divided by this number to obtain the percentage of ifc mutant flies that survived to adulthood. 2452 total flies were assayed for the UAS-ifc cross, 1303 for the UAS-DEGS1 cross, and 1030 for the control cross.

DNA sequencing

Genomic DNA was obtained from wild-type larvae or larvae homozygous for each relevant mutant line and provided to GTAC (Washington University) or GENEWIZ for next-generation or Sanger sequencing.

RNA in situ hybridization ifc RNA probes for in situ hybridization chain reaction (HCR) were designed and made by Molecular Instruments (HCR RNA-FISH v3.0) (Choi et al., 2018). Wild-type CNS was harvested and fixed in 2% paraformaldehyde at late L3. The fixed CNS underwent gradual dehydration and rehydration, followed by standard hybridization and amplification steps of the HCR protocol (Choi et al., 2018). For double labeling with antibody, the post-HCR labeled CNS was briefly fixed for 30 min prior to standard antibody labeling protocol to identify specific CNS cell type(s) with highly localized ifc RNAs (Lacin et al., 2019; Duckhorn et al., 2022).

MultiColor FlpOut labeling of glial subtypes

For glial labeling in the control background, the MCFO1 line was crossed to GMR-GAL4 driver lines for each glial subtype (see Key Resources Table; Nern et al., 2015). For glial labeling in the ifc−/− background, the MCFO1 line was placed in the ifcJS3 background, each of the five glial-specific GMR-GAL4 lines was placed individually into the ifc-KO background, and then the MCFO, ifcJS3 line was crossed to each glial-specific GAL4, ifc-KO line. Flies were allowed to lay eggs for 24 hr at 25°C, and progeny were raised at 25°C for 4 days prior to heat-activated labeling. On day 4 after egg-laying, F1 larvae were incubated in a 37°C water bath for 5 min. When wild-type or ifc−/− mutant larvae reached late L3, which was days 5 and 6 for control and days 9 and 10 for ifc−/− mutants, the CNS was dissected, fixed, stained, and then analyzed under a Zeiss LSM 700 Confocal Microscope for the presence of clones, using Zen software.

Antibody generation

YenZym (CA, USA) was used as a commercial source to generate affinity purified antibodies against a synthetic peptide that corresponded to amino acids 85–100 (TLDGNKLTQEQKGDKP) of FABP isoform B. Briefly, the peptide was conjugated to KLH, used as an immunogen in rabbits to generate a peptide-specific antibody response, and antibodies specific to the peptide were affinity purified. The affinity-purified antibodies were confirmed to label cortex glia specifically based on comparison of antibody staining relative to Myr-GFP when a UAS-Myr-GFP was driven under control of the cortex-specific glial GAL4 driver GMR-54H02 (Figure 1—figure supplement 3). The FABP antibodies are used at 1:500–1:1000.

Immunofluorescence and lipid droplet staining

Gene expression analysis was performed essentially as described in Patel, 1994. Briefly, the larval CNS was dissected in PBS, fixed in 2.5% paraformaldehyde for 55 min, and washed in PTx 1× PBS, 0.1% Triton X-100. The fixed CNS was incubated in primary antibody with gentle rocking overnight at 4°C. Secondary antibody staining was conducted for at least 2 hr to overnight at room temperature. All samples were washed in PTx at least five times and rocked for an hour before and after secondary antibody staining. A detailed list of the primary and secondary antibodies is available in the Key Resources Table. Dissected CNS were mounted either in PTx or dehydrated through an ethanol series and cleared in xylenes prior to mounting in DPX mountant (Truman et al., 2004). All imaging was performed on a Zeiss LSM-700 Confocal Microscope, using Zen software.

For lipid droplet staining, the fixed CNS was incubated for 30 min at room temperature at 1:200 dilution of 1 mg/ml BODIPY 493/503 (Invitrogen: D3922). It was then rinsed thoroughly in PBS and immediately mounted for imaging on a Zeiss LSM-700 Confocal Microscope, using Zen software.

TEM imaging

For TEM, samples were immersion fixed overnight at 4°C in a solution containing 2% paraformaldehyde and 2.5% glutaraldehyde in 0.15 M cacodylate buffer with 2 mM CaCl2, pH 7.4. Samples were then rinsed in cacodylate buffer three times for 10 min each and subjected to a secondary fixation for 1 hr in 2% osmium tetroxide/1.5% potassium ferrocyanide in cacodylate buffer. Following this, samples were rinsed in ultrapure water three times for 10 min each and stained overnight in an aqueous solution of 1% uranyl acetate at 4°C. After staining was complete, samples were washed in ultrapure water 3 times for 10 min each, dehydrated in a graded acetone series (50%, 70%, 90%, 100% ×4) for 15 min in each step, and infiltrated with microwave assistance (Pelco BioWave Pro, Redding, CA) into Spurr’s resin. Samples were then cured in an oven at 60°C for 72 hr and post-curing, 70 nm thin sections were cut from the resin block, post-stained with uranyl acetate and Sato’s lead and imaged on a Transmission Electron Microscope (JEOL JEM-1400 Plus, Tokyo, Japan) operating at 120 keV.

Lipidomics

Untargeted lipidomics analysis was conducted on whole larva and dissected CNS of wild-type and ifc−/− mutants at the late-third instar stage. Five replicates were prepared for each set of experiments. For whole larvae, at least 15 larvae of each genotype were used for each replicate. For the dissected CNS, at least 50 wild-type and 60 ifc−/− CNS were used per replicate. Immediately following collection or dissection, larvae and the dissected CNS were flash frozen in liquid nitrogen and placed at –80°C.

Lipids were extracted from frozen whole larvae and dissected larval CNS samples by using an Omni Bead Ruptor Elite Homogenizer using acetonitrile:methanol:water (2:2:1; 40 μl/mg tissue). Two ultrahigh-performance LC (UHPLC)/MS systems were used in this work: a Thermo Vanquish Flex UHPLC system with a Thermo Scientific Orbitrap ID-X and an Agilent 1290 Infinity II UPLC system with an Agilent 6545 QTOF as described previously (Cho et al., 2021). Lipids were separated on a Waters Acquity HSS T3 column (150 × 2.1 mm, 1.8 mm). The mobile-phase solvents were composed of A: 0.1% formic acid, 10 mM ammonium formate, 2.5 μM medronic acid in 60:40 acetonitrile:water; and B = 0.1% formic acid, 10 mM ammonium formate in 90:10 2-propanol:acetonitrile. The column compartment was maintained at 60°C. The following linear gradient was applied at a flow rate of 0.25 ml min−1: 0–2 min, 30% B; 17 min, 75% B; 20 min, 85% B; 23–26 min, 100% B. The injection volume was 4 μl for all lipids analysis. Data were acquired in positive ion.

LC/MS data were processed and analyzed with the open-source Skyline software (Adams et al., 2020). Lipid MS/MS data were annotated with Agilent Lipid Annotator software.

RNA sequencing and analysis

To determine the CNS-specific transcriptional changes upon loss of ifc, RNA-seq was conducted on five replicates of dissected CNS tissue derived from wild-type and ifc−/− mutant late-third instar larvae. For each replicate, roughly 30–35 dissected CNS of wild-type or ifc−/− larvae were used. Invitrogen RNAqueous-Micro Total RNA Isolation Kit (AM1931) was used to extract RNA. Agilent 4200 TapeStation system was used for RNA quality control. All RNA samples were then provided to the Genome Technology Access Center in the McDonnell Genome Institute at Washington University for next-generation sequencing.

Samples were prepared according to library kit manufacturer’s protocol, indexed, pooled, and sequenced on an Illumina NovoSeq. Basecalls and demultiplexing were performed with Illumina’s bcl2fastq software with a maximum of one mismatch in the indexing read. RNA-seq reads were then aligned to the Ensembl release 76 primary assembly with STAR version 2.5.1a (Dobin et al., 2013) Gene counts were derived from the number of uniquely aligned unambiguous reads by Subread:featureCount version 1.4.6-p5 (Liao et al., 2014). Isoform expression of known Ensembl transcripts was estimated with Salmon version 0.8.2 (Patro et al., 2017). To find the most critical genes, the raw counts were variance stabilized with the R/Bioconductor package DESeq2 (Love et al., 2014) and were then analyzed via weighted gene correlation network analysis with the R/Bioconductor package WGCNA (Langfelder and Horvath, 2008).

Statistics

All data are presented as mean ± SEM. Statistical significance between groups was determined using Student’s t-test or one-way ANOVA with multiple comparisons, and with varying levels of significance assessed as *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, and ns, not significant. All experiments were done with a minimum of five biological replicates.

Material availability statement

All newly created reagents or strains are freely available by contacting the corresponding author and/or through public stock centers at which many of the strains were deposited.

Appendix 1

Appendix 1—key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Gene (Drosophila melanogaster)Infertile crescent (ifc)GenBankFLYB: FBgn-0001941
Genetic reagent (D. melanogaster)y[1] M{RFP [3xP3.PB] GFP [E.3xP3]=
vas-int.Dm}ZH-2A w[*];
M{3xP3RFP. attP}ZH-51D		Bloomington Drosophila Stock CenterRRID:BDSC24483Wild-type. Control for 3xP3 background.
Genetic reagent (D. melanogaster)Gpdh1[nSP4] ifc[1]
wg[spd-fg] pr[1]/CyO, cl[4]		Bloomington Drosophila Stock CenterRRID:BDSC4963
Genetic reagent (D. melanogaster)w[1118];
Df(2L)ED334/SM6a		Bloomington Drosophila Stock CenterRRID:BDSC9343Deficiency line that uncovers ifc
Genetic reagent (D. melanogaster)w[1118];
Df(2L)BSC184/CyO		Bloomington Drosophila Stock CenterRRID:BDSC9612Deficiency line that uncovers ifc
Genetic reagent (D. melanogaster)w[1118];
Df(2L)BSC353/CyO		Bloomington Drosophila Stock CenterRRID:BDSC24377Deficiency line that uncovers ifc
Genetic reagent (D. melanogaster)y[1] w[*];
TI{GFP[3xP3.cLa]=CRIMIC.TG4.2}
ifc[CR70115-TG4.2]/SM6a		Bloomington Drosophila Stock CenterRRID:BDSC92710
Genetic reagent (D. melanogaster)v[1]; Kr[If-1]/CyO;
P{y[+t7.7] v[+t1.8]=
UAS-Trans-Timer.v+}attP2		Bloomington Drosophila Stock CenterRRID:BDSC93411
Genetic reagent (D. melanogaster)y[1] w[*];
PBac {y[+mDint2]
w[+mC]=UAS-
hDEGS1.HA}VK00033		Bloomington Drosophila Stock CenterRRID:BDSC79200UAS line for human DEGS1
Genetic reagent (D. melanogaster)w[1118] P{y[+t7.7] w[+mC]=hs-FLP
G5.PEST}attP3;PBac{y[+mDint2]
w[+mC]=10xUAS(FRT.stop)
myr::smGdP-HA} VK0000 5 P{y[+t7.7]
w[+mC]=10xUAS(FRT.stop)
myr::smGdP-V5-THS 10x UAS
(FRT.stop)
myr::smGdP-FLAG} su(Hw)attP1		Bloomington Drosophila Stock CenterRRID:BDSC64085Referred to as “MCFO1”
Genetic reagent (D. melanogaster)w[1118] P{y[+t7.7] w[+mC]=hsFLP G5.PEST}attP3;
ifcJS3/CyOTb; PBac{y[+mDint2] w[+mC]=10xUAS
(FRT.stop)myr::smGd-PHA}VK0000 5 P{y[+t7.7]
w[+mC]=10xUAS(FRT.stop)
myr::smGdP-V5-THS 10x UAS(FRT.stop)
myr::smGdP FLAG} su(Hw)attP1		This studyFreely available from authorsMCFO1 line with ifcJS3/CyO Tb allele
Genetic reagent (D. melanogaster)w[*];
P{y[+t7.7]
w[+mC]=
10XUAS-IVS-myr::GFP}attP2		Bloomington Drosophila Stock CenterRRID:BDSC32197
Genetic reagent (D. melanogaster)w[1118];
P{y[+t7.7]
w[+mC]=GMR54H02-GAL4}attP2		Bloomington Drosophila Stock CenterRRID:BDSC45784
Genetic reagent (D. melanogaster)ifc-KO/CyO, P{2xTb[1]-RFP};
P{y[+t7.7]
w[+mC]=GMR54H02-GAL4}attP2		This studyFreely available from authorsCortex glia GMR GAL4 line with ifcK0 allele
Genetic reagent (D. melanogaster)w[1118];
P{y[+t7.7]
w[+mC]=GMR86E01-GAL4}attP2		Bloomington Drosophila Stock CenterRRID:BDSC45914Astrocyte-like glia GMR-GAL4
Genetic reagent (D. melanogaster)ifc-KO/CyO,
P{2xTb[1]-RFP};
P{y[+t7.7] w[+mC]
=GMR86E01-GAL4}attP2		This studyFreely available from authorsAstrocyte-like glia GMR-GAL4
with copy of ifc-KO
Genetic reagent (D. melanogaster)w[1118];
P{y[+t7.7]
w[+mC]=GMR56F03-GAL4}attP2		Bloomington Drosophila Stock CenterRRID:BDSC39157Ensheathing glia GMR-GAL4
Genetic reagent (D. melanogaster)ifc-KO/CyO,
P{2xTb[1]-RFP};
P{y[+t7.7]
w[+mC]
=GMR56F03-GAL4}attP2		This studyFreely available from authorsEnsheathing glia GMR-GAL4
with one copy of ifc-KO
Genetic reagent (D. melanogaster)w[1118];
P{y[+t7.7]
w[+mC]=GMR54C07-GAL4}attP2		Bloomington Drosophila Stock CenterRRID:BDSC50472Subperineurial glia GMR-GAL4
Genetic reagent (D. melanogaster)ifc-KO/CyO,
P{2xTb[1]-RFP};
P{y[+t7.7]
w[+mC]=GMR54C07-GAL4}attP2		This studyFreely available from authorsSubperineurial glia GMR-GAL4
with one copy of ifc-KO
Genetic reagent (D. melanogaster)w[1118];
P{y[+t7.7] w[+mC]=
GMR85G01-GAL4}attP2		Bloomington Drosophila Stock CenterRRID:BDSC40436Perineurial glia GMR-GAL4
Genetic reagent (D. melanogaster)ifc-KO/CyO,
P{2xTb[1]-RFP};
P{y[+t7.7] w[+mC]=
GMR85G01-GAL4}attP2		This studyFreely available from authorsPerineurial glia GMR-GAL4
with one copy of ifc-KO
Genetic reagent (D. melanogaster)P{w[+mW.hs]=GawB}elav[C155]Bloomington Drosophila Stock CenterRRID:BDSC458
Genetic reagent (D. melanogaster)FlyFos019206(pRedFlp-Hgr)
(ifc[27951]::2XTY1-SGFP-V5
-preTEV-BLRP-3XFLAG)dFRT		Vienna Dros. Research CenterRRID:VDRC318826
Genetic reagent (D. melanogaster)P{VSH330794}Vienna Dros. Research CenterRRID:VDRC330794RNAi line for ifc
Genetic reagent (D. melanogaster)M{UAS-ifc-ORF-3xHA.attP}86FbFlyORFFlyORF: F003887 (reference 72)
Genetic reagent (D. melanogaster)ifcJS1 M{3xP3-RFP.attP}ZH-51D/CyO TbThis studyFreely available from authorsV276D loss-of-function allele
Genetic reagent (D. melanogaster)ifcJS2 M{3xP3-RFP.attP}ZH-51D/CyO TbThis studyFreely available from authorsG257S loss-of-function allele
Genetic reagent (D. melanogaster)ifcJS3 M{3xP3-RFP.attP}ZH-51D/CyO TbThis studyFreely available from authorsW162* loss-of-function allele
Genetic reagent (D. melanogaster)elav-GAL4[C155];
Repo-GAL80/CyO Tb		This studyFreely available from authors
Genetic reagent (D. melanogaster)ifc-KO/CyO TbGift from Dr. Chih-Chiang ChanJung et al., 2017 (Ref: 17)
AntibodyRabbit anti-FABP polyclonalThis studyFreely available from authors1:500
AntibodyRabbit anti-EBONY polyclonalGift from Dr. Haluk LacinRRID:AB_23143541:500
AntibodyMouse anti-REPO monoclonalDSHBRRID:AB_5284481:100
AntibodyRat anti-ELAV monoclonalDSHBRRID:AB_5282181:100
AntibodyMouse anti-CNX 99A monoclonalDSHBRRID:AB_27220111:20
AntibodyMouse anti-GOLGIN84 monoclonalDSHBRRID:AB_27221131:20
AntibodyGoat anti-GOLGIN-245 polyclonalDSHBRRID:AB_26182601:500
AntibodyRabbit anti-ESYT polyclonalGift from Dr. Dion Dickman1:300
AntibodyGFP Antibody Dylight 488 Goat PolyclonalRockland (600-141-215)RRID:AB_19615161:1000
AntibodyAnti-LAMP1 antibodyAbcam (ab 30687)RRID:AB_7759731:500 (lyso-some marker)
AntibodyCleaved Caspase-3 (Asp175) polyclonalCell Signaling Technology (#9661)RRID:AB_23411881:400
AntibodyAnti-HA antibody produced in rabbitMillipore Sigma (H6908)RRID:AB_2600701:500
AntibodyANTI-FLAG antibody, Rat monoclonalMillipore Sigma (SAB4200071)RRID:AB_106033961:500
AntibodyChicken V5 Tag Polyclonal AntibodyBethyl laboratoriesRRID:AB_667411:500
AntibodyGoat anti-Chicken IgY Alexa 488Invitrogen (A-11039)RRID:AB_25340961:1000
AntibodyDonkey anti-Rat Alexa 555Invitrogen (A48270)RRID:AB_28963361:1000
AntibodyDonkey Anti-Rat Cy5Jackson Immuno-ResearchRRID:AB_23406721:1000
AntibodyDonkey Anti-Goat IgG Cy5Jackson Immuno-ResearchRRID:AB_23404151:1000
AntibodyDonkey Anti-Mouse IgG Cy5Jackson Immuno-ResearchRRID:AB_23408201:1000
AntibodyDonkey Anti-Rabbit IgG Cy5Jackson Immuno-ResearchRRID:AB_23406071:1000
Chemical compoundBODIPY 493/503InvitrogenD39221:200
SoftwareImageJ2 2.3.0/1.53qNIH (https://imagej.net/)RRID:SCR_003070
SoftwareZEN Microscopy SoftwareCarl Zeiss AG; Jena, DEURRID:SCR_013672
SoftwarePhotoshop 23.5.5Adobe; San Jose, CARRID:SCR_014199
SoftwareGraphPad Prism 10.2.2GraphPad; Boston, MARRID:SCR_002798

Data availability

Sequencing data have been deposited in Geo under accessions code: GSE263308. Untargeted lipidomics data have been deposited in the Metabolics Workbench under project ID - PR001967 and study ID - ST003162.

The following data sets were generated
    1. Skeath JB
    2. Zhu Y
    3. Zhu Y
    4. Lacin H
    5. Wilson BA
    (2024) NCBI Gene Expression Omnibus
    ID GSE263308. Loss of dihydroceramide desaturase drives neurodegeneration by disrupting endoplasmic reticulum and lipid droplet homeostasis in glial cells.
    https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE263308
    1. Cho K
    2. Patti GJ
    3. Skeath JB
    (2024) Metabolics Workbench
    Loss of dihydroceramide desaturase drives neurodegeneration by disrupting endoplasmic reticulum and lipid droplet homeostasis in glial cells.
    https://doi.org/10.21228/M8G72D
The following previously published data sets were used
    1. Lacin H
    2. Zhu Y
    3. DiPaola JT
    4. Wilson BA
    5. Zhu Y
    6. Skeath JB
    (2024) NCBI BioProject
    ID PRJNA1128589. A Genetic Screen for regulators of CNS morphology in Drosophila.
    https://www.ncbi.nlm.nih.gov/bioproject/?term=PRJNA1128589

References

Article and author information

Author details

  1. Yuqing Zhu

    Department of Genetics, Washington University School of Medicine, St. Louis, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8693-4741

  2. Kevin Cho

    1. Department of Chemistry, Washington University in St. Louis, St. Louis, United States
    2. Department of Medicine, Washington University School of Medicine, St. Louis, United States
    3. Center for Mass Spectrometry and Metabolic Tracing, Washington University in St. Louis, St. Louis, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared

  3. Haluk Lacin

    Division of Biological and Biomedical Systems, University of Missouri-Kansas City, Kansas City, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2468-9618

  4. Yi Zhu

    Department of Genetics, Washington University School of Medicine, St. Louis, United States
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared

  5. Jose T DiPaola

    Department of Genetics, Washington University School of Medicine, St. Louis, United States
    Contribution
    Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared

  6. Beth A Wilson

    Department of Genetics, Washington University School of Medicine, St. Louis, United States
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Methodology
    Competing interests
    No competing interests declared

  7. Gary Patti

    1. Department of Chemistry, Washington University in St. Louis, St. Louis, United States
    2. Department of Medicine, Washington University School of Medicine, St. Louis, United States
    3. Center for Mass Spectrometry and Metabolic Tracing, Washington University in St. Louis, St. Louis, United States
    Contribution
    Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration
    Competing interests
    Collaborative research agreement with Agilent Technologies and Thermo Fisher. Chief Scientific Officer of Panome Bio
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3748-6193

  8. James B Skeath

    Department of Genetics, Washington University School of Medicine, St. Louis, United States
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    jskeath@wustl.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1179-4857

Funding

National Institute of Neurological Disorders and Stroke (NS036570)

  • James B Skeath

National Institute of Neurological Disorders and Stroke (NS122903)

  • Haluk Lacin

National Institute of Environmental Health Sciences (ES2028365)

  • Gary Patti

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank the Iowa Developmental Studies Hybridoma Bank for antibodies, and the Bloomington Stock Center, Vienna Drosophila Research Center, FlyORF, and the National Institutes of Genetics stock center in Japan for countless fly lines. We thank Drs. Christian Klambt, Chih-Chiang Chan, and Dion Dickman for reagents. We thank Dr. Tristan Qingyuan Li for comments on the manuscript. We thank the Genome Technology Access Center at Washington University for next-generation sequencing and analysis of RNA-seq samples. We thank Dr. Sanja Sviben, Gregory Strout, and John Wulf II for assistance in TEM studies conducted at the Washington University Center for Cellular Imaging, which is supported in part by Washington University School of Medicine, The Children’s Discovery Institute of Washington University and St. Louis Children’s Hospital (CDI-CORE-2015-505 and CDI-CORE-2019-813), the Foundation for Barnes-Jewish Hospital (3770), and the Washington University Diabetes Research Center (NIH P30 DK020579).

Version history

  1. Preprint posted:
  2. Sent for peer review:
  3. Reviewed Preprint version 1:
  4. Reviewed Preprint version 2:
  5. Version of Record published:

Cite all versions

You can cite all versions using the DOI https://doi.org/10.7554/eLife.99344. This DOI represents all versions, and will always resolve to the latest one.

Copyright

© 2024, Zhu 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.

Metrics

  • 1,226
    views
  • 21
    downloads
  • 1
    citation

Views, downloads and citations are aggregated across all versions of this paper published by eLife.

Citations by DOI

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Open citations (links to open the citations from this article in various online reference manager services)

Cite this article (links to download the citations from this article in formats compatible with various reference manager tools)

  1. Yuqing Zhu
  2. Kevin Cho
  3. Haluk Lacin
  4. Yi Zhu
  5. Jose T DiPaola
  6. Beth A Wilson
  7. Gary Patti
  8. James B Skeath
(2025)
Loss of dihydroceramide desaturase drives neurodegeneration by disrupting endoplasmic reticulum and lipid droplet homeostasis in glial cells
eLife 13:RP99344.
https://doi.org/10.7554/eLife.99344.3

Share this article

https://doi.org/10.7554/eLife.99344

Categories and tags

Research organism