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Motor defects in a Drosophila model for spinal muscular atrophy result from SMN depletion during early neurogenesis [1]

['Stuart J. Grice', 'Medical Research Council Functional Genomics Unit', 'Department Of Physiology', 'Anatomy', 'Genetics', 'University Of Oxford', 'Oxford', 'United Kingdom', 'Ji-Long Liu', 'School Of Life Science']

Date: 2022-09

Abstract Spinal muscular atrophy (SMA) is the most common autosomal recessive neurodegenerative disease, and is characterised by spinal motor neuron loss, impaired motor function and, often, premature death. Mutations and deletions in the widely expressed survival motor neuron 1 (SMN1) gene cause SMA; however, the mechanisms underlying the selectivity of motor neuron degeneration are not well understood. Although SMA is degenerative in nature, SMN function during embryonic and early postnatal development appears to be essential for motor neuron survival in animal models and humans. Notwithstanding, how developmental defects contribute to the subversion of postnatal and adult motor function remains elusive. Here, in a Drosophila SMA model, we show that neurodevelopmental defects precede gross locomotor dysfunction in larvae. Furthermore, to specifically address the relevance of SMN during neurogenesis and in neurogenic cell types, we show that SMN knockdown using neuroblast-specific and pan-neuronal drivers, but not differentiated neuron or glial cell drivers, impairs adult motor function. Using targeted knockdown, we further restricted SMN manipulation in neuroblasts to a defined time window. Our aim was to express specifically in the neuronal progenitor cell types that have not formed synapses, and thus a time that precedes neuromuscular junction formation and maturation. By restoring SMN levels in these distinct neuronal population, we partially rescue the larval locomotor defects of Smn mutants. Finally, combinatorial SMN knockdown in immature and mature neurons synergistically enhances the locomotor and survival phenotypes. Our in-vivo study is the first to directly rescue the motor defects of an SMA model by expressing Smn in an identifiable population of Drosophila neuroblasts and developing neurons, highlighting that neuronal sensitivity to SMN loss may arise before synapse establishment and nerve cell maturation.

Author summary Spinal muscular atrophy (SMA) is the most common genetic cause of infant mortality and leads to the degeneration of the nerves that control muscle function. Loss-of-function mutations in the widely expressed survival motor neuron 1 (SMN1) gene cause SMA, but how low levels of SMN protein cause the neuronal dysfunction is not known. Although SMA is a disease of nerve degeneration, SMN function during nerve cell development may be important, particularly in severe forms of SMA. Nevertheless, how the defects during development and throughout early life contribute to the disease is not well understood. We have previously demonstrated that SMN protein becomes enriched in neuroblasts, which are the cells that divide to produce neurons. In the present study, motor defects observed in our fly model for SMA could be rescued by restoring SMN in neuroblasts alone. In addition, we show that knocking down SMN in healthy flies within the same cell type causes impaired motor function. The present study shows that the manipulation of SMN in a developmentally important cell type can cause motor defects, indicating that a period of abnormal neurodevelopment may contribute to SMA.

Citation: Grice SJ, Liu J-L (2022) Motor defects in a Drosophila model for spinal muscular atrophy result from SMN depletion during early neurogenesis. PLoS Genet 18(7): e1010325. https://doi.org/10.1371/journal.pgen.1010325 Editor: Gregory A. Cox, The Jackson Laboratory, UNITED STATES Received: December 13, 2021; Accepted: July 5, 2022; Published: July 25, 2022 Copyright: © 2022 Grice, Liu. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited. Data Availability: All relevant data are within the manuscript and its Supporting Information files. Funding: The authors received no specific funding for this work. Competing interests: The authors have declared that no competing interests exist.

Introduction Survival motor neuron (SMN) is an essential protein that functions in the biogenesis of spliceosomal small nuclear ribonucleoproteins (snRNPs), which subsequently mediate pre-mRNA splicing [1]. Loss-of-function mutations in the SMN1 gene cause the disease spinal muscular atrophy (SMA), which is characterised by the selective loss of alpha motor neurons of the spinal cord, muscle wasting and, in most severe cases, premature death in infancy [2]. Since the identification of the disease-associated gene SMN1 in 1995 [2], the drive to uncover the mechanisms underlying SMA pathogenesis has been complicated by the pleiotropic nature of the SMN locus [3], coupled with the varied levels of SMN protein in human and animal models [4–6]. There has been considerable debate about how aberrations in both the canonical and non-canonical motor neuron-specific functions of SMN may lead to the observed motor neuron selectivity [7]. SMN has been shown to play a fundamental role in snRNP and messenger ribonucleoprotein (mRNP) biogenesis [8], whilst also being involved in mRNA trafficking and local translation, cytoskeletal dynamics, endocytosis and ubiquitin homeostasis (reviewed in [3,9]). In addition, the nature of SMA pathology, and of the animal models engineered to study the disease, are greatly affected by the systemic, temporal and spatial levels of SMN protein [5,6]. In humans, in addition to SMN1, SMN is also encoded by a second paralogous gene called SMN2, which, owing to a mutation affecting exon 7 splicing, generates comparatively low levels of full-length SMN protein [10,11]. Due to the fact that SMN2 copy number can also vary between individuals, there is a broad spectrum of disease severity that, at the population level, correlates with the amount of SMN2-derived wild-type SMN protein [4]. As SMN levels decrease, disease severity increases, the motor defects become more pronounced, and many more cell and tissue types present with phenotypes caused by loss of the protein [12,13]. Classed as a ubiquitous protein, SMN localises to the cytoplasm and nucleus, and can be observed in many RNP-enriched subcellular foci, such as Gems [14], nucleoli [15], U bodies [16] and Cajal bodies [14]. Cells do not necessarily require organised Gems, U Bodies and Cajal bodies to survive; however, evidence shows that these celluar foci promote the efficient clustering of the RNA processing factors required in embryonic, dividing and metabolically active cells [17]. SMN protein level and associated snRNP assembly are highest during embryonic development, and are substantially downregulated postnatally [18] and as cells differentiate and mature [19–21]. Furthermore, severe SMN loss can lead to developmental defects, with a hierarchy of cell types, many of which are uncharacterised, having differing sensitivities to a reduction in the level of the protein [6,12]. Undoubtedly, the alpha motor neurons are particularly sensitive to SMN reduction. Notwithstanding, it is not known how this selectivity manifests in its entirety (i.e., whether it is a result of aberrations in set-up or degeneration, or if it is through a non-cell autonomous mechanism). Previous research has shown that the selective loss of SMN in motoneuronal progenitors is sufficient to cause SMA like phenotypes [22]. Furthermore, restoration of SMN in mature motoneurons only rescued the SMA phenotype partially [23,24], whereas motor neuron-specific SMN reduction in wild type mice fails to recapitulate the entirety of the disease phenotypes, highlighting the importance of neuronal development. Motor neuron loss is also a relatively late feature in SMN patients and mammalian models [25, 26], although patients with type 1 SMA present with neuromuscular junction (NMJ) maturation defects during fetal development [27]. Importantly, when performing rescue studies using mouse and Drosophila SMA models, early stage ubiquitous restoration of SMN results in the greatest improvement in motor function and animal survival [26,28,29]. This is supported by evidence from patient clinical trials [30] and early versus later treatment of SMA mice [6,31–37]. To complement these findings, SMN reduction in young adult mice caused more modest phenotypes when compared with mice in which SMN was knocked down at an earlier developmental time point [6, 26,28]. Furthermore, Drosophila studies using Smn mutant models have reported severe growth defects and considerable developmental retardation, in addition to motor and NMJ dysfunction [5,21,38–41]. In mouse embryos, although no overt developmental outgrowth defects have been observed [42], defective radial outgrowth and poor Schwann ensheathment led to some axons degenerating postnatally [43]. Altogether, this research highlights that the cause of SMA may not be solely through classical neurodegenerative processes, but via a combinatory multi-cell type mechanism that may be sensitised by neurodevelopmental abnormalities. An understanding of the precise nature of the developmental requirements of SMN, and how perturbations in SMN protein level leads to defects that manifest in progenitor and non-differentiated neuronal cell types, is important for SMA treatment. The aim of the present study was to understand how manipulation of SMN protein level during specific periods of neurogenesis can cause and modify the phenotypes present in Drosophila models for SMA. The aim was to restrict SMN manipulation to the neuronal progenitor cell types that have not yet formed synapses, and to a period that precedes NMJ maturation. To achieve this, knockdown and rescue studies were used during the waves of proliferation and differentiation in the larval and pupal central nervous system (CNS). The classical GAL4 and the more targeted GAL80 repression systems were used to allow for spatiotemporal transgene expression [44]. The reduction of SMN in neuroblasts and undifferentiated neurons, but not subsequently in differentiated subpopulations of neurons, caused motor defects. In the reciprocal experiment, neurodevelopmental and motor phenotypes are partially rescued by expressing SMN in neuroblasts and immature neurons. Finally, combinatorial SMN knockdown was carried out in immature and mature neurons, which synergistically enhanced the locomotor and survival phenotypes in the present model. This in-vivo study contributes to the understanding of how developmental abnormalities can contribute to the motor defects synonymous with the pathology of SMA. Furthermore, by selectively manipulating SMN in an identifiable population of neuroblasts and developing neurons, we highlight the importance of SMN in Drosophila neurodevelopment.

Discussion In this study, we show that depleting SMN in neuroblasts and their immature daughter cells can predispose larval and adult Drosophila to locomotor dysfunction. In addition, we can partially rescue the larval motor defects of Smn mutants by restoring SMN in the neuroblasts and immature developing neurons using targeted expression systems. Finally, we highlight that the combination of presumptive and mature nervous system SMN reduction increases the severity of SMA model phenotypes. We show that the reduction of SMN in cells that are not synapse forming, and thus precede NMJ and sensory-motor network maturation, cause SMA-like phenotypes in the fly. Although motor neuron loss is also a relatively late feature in SMA patients and mammalian models [25], it is believed that defects in synapse formation and maintenance may be central to the neurological phenotypes observed in SMA patients [59]. Mouse model rescue studies highlight that the therapeutic success of administered rescue constructs generally become progressively diminished only a few days after birth [31–36]. This pre- and peri-natal period coincides with a higher requirement of SMN level in the CNS, a phenomenon also observed in Drosophila [5,39,40]. It is difficult to compare the Drosophila life cycle with the vertebrate progression of disease; however, the mechanistic and cellular readouts from invertebrate models can offer some degree of translation. Our knockdowns and rescues are limited to neuronal stem cells and their immature progeny. Drosophila neuronal stem cells progress through a cascade of transcriptionally distinct identities before permanently differentiating or dying [45,46,61]. During division, this developmental cascade leads to a diversity of developmentally plastic immature daughter cells that undergo further pre- and post-transcriptionally regulated maturation steps, which precede the formation of synapses and ultimately action potentials. Although SMA was classically thought to be a disease of aberrant splicing, the broad requirement for SMN in the regulation of post-transcriptional gene expression is compelling, with roles encompassing snRNP biogenesis [1], mRNP biogenesis [62], mRNA transport [63], ribosomal dynamics [64], chromatin dynamics [65] and translational control [66]. It is probable that deficits in any one of these pathways could lead to stem cell or daughter cell sensitivity to conditions of low SMN. As a stem cell divides and creates a differentiating daughter cell, large changes in alternative splicing drive identity from one of pluripotency to that of an identifiable neuronal lineage with a defined cell biology and physiology [67,68]. It may be that higher SMN levels are required for the temporal–spatial regulation of the alternative splicing events that occur during this switch. Provisional data has shown that both adult flies and larvae display fewer synapses and synaptic boutons respectively, when SMN is knocked down in neuroblasts and the corresponding immature progeny. However, the relevance of bouton number changes has been partly called into question, and these alterations may only be casually linked to the movement defects physiological alterations, and death, observed in Drosophila SMA models [49,69]. It may be that upstream functional changes in motor neurons, interneurons, or other neuronal cell types may ultimately lead to the degeneration of the motor neuron or the neuromuscular junction. To this end, the temporal transcription factor cascades that generate the molecular and physiological diversity of the neurons in the developing CNS may be of interest [46, 61]. In future work, we would like to see, when SMN levels are low, if molecular changes in the developing neurons lead to defects in motor neuron physiology, or alterations in the different neuronal classes. We can speculate that changes at this level could alter, in a subtle manner, their molecular identity sensitising neurons to degeneration in certain conditions, or over time. We have previously reported that SMN overexpression affects developmental timing in Drosophila [21] and protects embryonic stem cells from retinol-induced differentiation [19]. In in-vivo SMN mutant neuroblast clones, the levels of both major and minor spliceosome snRNPs (U5 and U2) are reduced in the nucleus of the neuroblast. We have also shown that SMN loss in neuroblasts perturbs cell division and alters the topology of the daughter cell cluster. Furthermore, gene expression analysis conducted on the spinal cord from SMN deficient mice detected changes in proliferative pathways, and identified morphological changes in the dividing cells in the ventral horn [70]. In both mouse and Drosophila models, SMN reduction promotes the untimely differentiation of neurons and spermatogonia [19–21], suggesting that high SMN is required for the fidelity of the developmental processes key to cellular differentiation and maturation. Alternatively, or in-combination, SMN loss may affect downstream translational control. We have previously shown that SMN-deficient neuroblasts display a mislocalisation of a cortical scaffolding protein that binds asymmetrically localised RNP complexes. Drosophila neuronal stem cells and neurons alike are energetically demanding, polarised and metabolically distinct [71]; therefore, the presence of highly clustered sites for RNP maturation and processing may be necessary for the correct function of metabolically active and dividing cell types [17]. Nevertheless, it is important to note that cell types and tissue beyond the nervous system are affected by SMN loss [72]. Within our model, although we rescue motor phenotypes and expand lifespan, neuroblast SMN rescue cannot support full larval development, pupariation and pupation to adulthood. In addition, SMN depletion in mature neurons also leads to locomotor dysfunction. Mutations in many widely expressed genes cause selective neuropathies and motor neuron diseases [73]. How we come to think about the nature of these disease genes, particularly their pleiotropy and spatial and temporal requirements, will be key to the eventual revealing of the mechanisms that lead to the selectivity of cell type degeneration. It seems that, at least in the Drosophila model, SMN reduction in many presumptive and mature neuronal cell types can cause locomotor defects and reduce lifespan. To understand the temporal and spatial requirement of SMN during nervous system development, we used Drosophila cell and time-specific GAL4 drivers. To date, a large number of UAS/GAL4 studies have investigated how to identify the fly tissue and cell types sensitive of SMN loss [5]. Consistent with other models, ubiquitous SMN knockdown is the most severe, leading to larval lethality, whereas ubiquitous rescue using high level expression drivers confers adult survival [39,40]. Second to this, combinatorial experiments expressing SMN, both early stage pan-neuronal and mesoderm drivers, partially rescues at the adult stage, whereas knockdown using the same driver combination causes larval–pupal lethality [40]. In comparison, pan-neuronal knockdown alone leads to modest adult lethality, neurophysiological and behavioural defects [39,40,74], whereas a subset of other drivers, including those expressed in cholinergic neurons and glutamatergic neurons, have shown specific neuromuscular phenotypes or rescue profiles [41,74]. It is important to highlight that GAL4 drivers will vary in temporal specificity and level. Due to the non-synchronous correlation between transcriptome and proteome [75], enhancer drivers derived from known neuronally expressed genes may generally express earlier or more broadly (at least at lower levels) than expected. These issues highlight the importance of the GAL80TS system used in this study to confine transgene expression to the cell type and time period. However, it is important to note that although the target system offers a high degree of temporal and special control, our study does not fully eliminate the role of low level SMN knockdown and expression in other tissues enhancing the phenotypes and rescues observed, during the period of GAL4 expression. To summarise, the present research supports that the idea that SMA is caused by a combination of defects that impact motor neuron development, maturation, and maintenance. Moreover, although motor neurons seem to be particularly sensitive to SMN loss, the complex background of multiple tissue defects makes it difficult to unveil the precise timing and nature of the causative defects. The present study directly shows that SMN is required during a window of neurogenesis that precedes synapse formation and neuromuscular junction maturation, and that the motor defects observed in Drosophila SMA models can be, in part, be caused by SMN reduction in these cell types. To this end, further study should address how an improper set-up of neuronal networks may compound any motor neuron cell autonomous defects that may arise in SMN-deficient motor neurons.

Materials and methods Drosophila husbandry and stocks Smnx7 null, SmnA, P[UAS-Smn-RNAi]N4, P[UAS-Smn-RNAi]C25 line have been previously described [39, 40]. All stock were backcrossed onto w1118 wild type background. Larvae were grown on apple juice plates with yeast and rich food added. Low population density was maintained for all crosses prior to experimentation. For classical UAS/GAL4 experiments, all crosses were carried out at 25°C to generate extensive but not complete knock-down. GAL4 drivers 1032-GAL, D42-GAL4, OK371-GAL4, Cha-GAL4, Repo-GAL4, and CG-GAL4 drivers, were obtained from Bloomington (Indiana). Pros-GAL4 was obtained from the putative-enhancer collection (Bloomington Drosophila Stock Centre [BDSC] at Indiana University, USA). Insu-GAL4; Tub84B-GAL4TS was a gift from Jürgen Knoblich. Drivers were characterised using UAS-CD8-GFP, UAS-H2B-YFP (Andrea Brand). EdU staining Dissected CNS were incubated for 1.5 h in 10 μM EdU/Grace’s medium, fixed for 10 min in 4% paraformaldehyde, followed by detection of Alexa Fluor azide according to the Click-iT EdU Imaging Kit (Invitrogen, Waltham, MA, USA) and washing in 0.2% Triton X-100 in phosphate buffered saline. Immunofluorescence was carried out as previously described [21]. Larval hatching assays A 2-h lay was carried out on apple juice plates and embryos were lined up in sets of 10. The number of embryos that hatched into larvae was scored for each genotype and was expressed as a percentage of that expected from the lay. Smnx7/TM6B-GFP and SmnA/TM6B-GFP were crossed and the number of embryos and larvae with Smn heterozygotes) and without GFP expression (homozygous smn mutants) were scored as a percentage. Larval locomotion assays Measurement of motor function involved placing individual age-matched third instar larvae at the centre of a 0.7% (weight by volume) agar plate and counting the forward body wall contractions exhibited over 1 min. Larvae were left to acclimatise for 30 s before analysis. Larval survival assay Flies performed a 2hr lay on apple juice plates for 2-h with minimal yeast. Embryos were counted, additional yeast was added, and larval development and death was recorded every 24 h. Adult locomotor function assay Age-matched (1- and 7-day old) male flies were placed individually in a 5-mm glass activity tube containing a food source (5% sucrose [Sigma-Aldrich, St Luis, MO, USA] and 2% Bacto agar [BD Diagnostics, Franklin Lakes, NJ, USA] in distilled water) at one side and a plastic cover with an air hole at the other. The individual glass tubes were placed into the activity monitor (Trikinetics monitors DAM2) (Trikinetics Inc., Waltham, MA, USA) and supported with rubber bands to hold them in place. Locomotor activity was recorded when the flies crossed the infrared light beam at the middle of the glass tubes. Thirty flies were used per genotype and kept under controlled conditions (12-h light–dark cycle at 25°C) for 2 days, day 1 being excluded for habituation. The DAM System collection software was used for collecting data. The raw binary data were processed using DAM Filescan102X (Trikinetics Inc., Waltham, MA, USA) and summed into 1-h bins. Adult flight assay The flight assay was carried out in accordance with a modified protocol originally designed by Benzer [57]. A total of 1000-ml graduated cylinder divided into five sectors was coated internally with mineral oil. Flies were introduced into the top of the cylinder through a funnel and the flies stuck in each sector were counted. The height that flies stick in the cylinder is indicative of their flight capabilities. TubGAL80TS TARGET analysis For larval rescue analysis, GAL80TS analysis, embryos were reared at 29°C (GAL80TS inactive; GAL4 active) and after 24 h (GAL80TS active; GAL4 repressed) and then switched to 19°C during larval life. TubGAL80TS, Insc-GAL4/UAS-dSMN; Smnx7/SmnA stock was used and compared with the mutant TubGAL80TS, Insc-GAL4/+; Smnx7/SmnA and control TubGAL80TS, Insc-GAL4/UAS-GFP backgrounds. For adult analysis, Drosophila larvae were reared at 29°C (GAL80TS inactive; GAL4 active) and then switched to 19°C (GAL80TS active; GAL4 repressed) after pupariation formation. Two non-overlapping RNAi construct was used (SMN-RNAiN4 and SMN-RNAiC25) and expressed using a TubGAL80TS, Insc-GAL4 stock line. Drosophila motor behaviour was analysed using activity monitoring, which was carried out at 19°C at days 1 and 7 after hatching. Flight testing was carried out at 8 days. qRT-PCR We determined the levels of GFP mRNA using qPCR methods as described previously [76], using Fast SYBR Green Master Mix (Applied Biosystems Cat. no. 4385612) and the 500 Fast Real-Time PCR System (Applied Biosystems). Statistical analysis A Kruskal–Wallis test and subsequent Dunn’s multiple comparison testing were carried out unless otherwise stated. GraphPad Prism software was used for all data analysis.

Supporting information S1 Fig. Movement defects present at late larval stages. (A) Control (w1118) and Smnx7/SmnA larvae were monitored at approximately 24, 48, and 72 ± 1 h after egg laying. Acclimatised larvae were filmed for 1 min, and the distance travelled was traced and measured in cm. Smnx7/SmnA larvae displayed significant movement defects at 72 h (***P < 0.001, n = 20); (B) example superimposed larval locomotion path traces from control and Smnx7/SmnA mutants for each time point. https://doi.org/10.1371/journal.pgen.1010325.s001 (TIF) S2 Fig. Characterisation of Insc-Gal4 expression and targeted SMN knockdown. (A) During larval life, a second wave of larval neuroblast division occurs. The majority of neuroblasts in the ventral ganglion reside at the surface of the larval CNS. (B) Representative Inscu-GAL4 expression is seen exclusively in neuroblasts and immature neurones in the ventral ganglion and brain lobes. Insc-Gal4 expression was examined using UAS-mCherry. The ventral and medial regions of a third instar larval central nervous system is shown. (C) The larval CNS were co-stained with SMN. The zoom (Box in B) shows SMN staining overlaps with UAS-mCherry immunofluorescence. (D) Expressing UAS-SMN-RNAiN4 exclusively in neuroblasts and immature neurones significantly reduces, but does not eliminate, SMN levels. Edu staining highlights a population of dividing neuroblasts and ganglion mother cells that no longer show SMN enrichment. https://doi.org/10.1371/journal.pgen.1010325.s002 (TIF) S3 Fig. Relative expression of GFP mRNA normalised to rp49 in Tub-GAL80TS; Insc-GAL4/UAS-GFP during the larval and adults experimental time courses. (A) The GAL80TS system was used to eliminate any adult GAL4 expression. For larval experiments, a temperature sensitive GAL80 (GAL80TS) represses GAL4 at 19°C but becomes inactive at 29°C was used. Embryos were reared for 24 h at 29°C, during which GAL4 is expressed, then switched to 19°C to eliminate expression. (B) GFP RNA was measured in whole embryos and larval CNS over the time course analogous to that used in the locomotor and pupation assays. GFP expression was seen to diminish by 0 hrs. We detected no further GFP expression throughout the course of the experimental period. (C) For adult studies, larvae were reared at 29°C (GAL80TS is inactive; GAL4 is active) and then switched to 19°C (GAL80TS is active; GAL4 is repressed) at the start of pupation. (D) Relative expression of GFP mRNA normalised to rp49 in Tub-GAL80TS; Insc-GAL4/UAS-GFP larvae, pupae and adults. GFP RNA was measured in larval, pupae and adults over the time course analogous to that used in the adult activity and flight assays. The Larvae were switched from 29 to 19°C at the late L3 stage. GFP expression was seen to diminished during larval growth and maturation. We detected no GFP expression throughout the pupal and adult periods studied. (L2, 2nd Instar Larvae; L3, 3rd Instar Larvae). https://doi.org/10.1371/journal.pgen.1010325.s003 (TIF)

Acknowledgments We thank Zillah Deussen and Mayte Siswick for maintaining the Drosophila Stocks; Andrea Brand, Jurgen Knoblich, Marcel van den Heuvel and the Developmental Studies Hybridoma Bank for providing fly stocks and antibodies; James N. Sleigh, Gabriel Aughey, and Kay Davies, for reading the manuscript.

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