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The Pennsylvania State University The Graduate School Eberly College of Science MICROTUBULE POLARITY AND DROSOPHILA SENSORY NEURON RESPONSES TO AXON AND DENDRITE INJURY A Thesis in Genetics by Michelle C. Stone © 2014 Michelle C. Stone Submitted in Partial Fulfillment of the Requirements for the Degree of Master of Science December 2014 The thesis of Michelle C. Stone was reviewed and approved* by the following: Melissa Rolls Associate Professor of Biochemistry and Molecular Biology Thesis Adviser Richard Ordway Professor of Molecular Neuroscience and Genetics Robert Paulson Professor of Veterinary and Biomedical Sciences Program Chair of Genetics Graduate Program *Signatures are on file in the Graduate School ii Abstract Microtubule polarity in axons and dendrites is essential for polarized transport and function of the neuron. It is also a key indicator of axonal or dendritic identity. Maintenance of neuronal microtubule polarity, especially following injury, is crucial for survival of the neuron. This work describes the dramatic microtubule rearrangements that occur following axon injury in Drosophila sensory neurons and the regeneration that results from converting a dendrite into an axon, all of which depend on the DLK/JNK pathway. This is in contrast to the DLK independent regeneration that occurs following dendrite injury in the same cells. This work underscores the importance of understanding how the neuronal cytoskeleton responds to injury, which is fundamental for our overall understanding of neurodegenerative diseases, stroke and traumatic brain injuries. iii Table of Contents List of Figures…………………………………………………………………………………………………….vii List of Tables…………………………………………………………………………………………………….…x Acknowledgements……………………………………………………………………………………………..xi Chapter 1. Introduction……………………………………………………………………………………….1 References………………………………………………………………………………………….7 Chapter 2. Global Up-­‐Regulation of Microtubule Dynamics and Polarity Reversal during Regeneration of an Axon from a Dendrite…………………...9 Abstract, Introduction………………………………………………………………………..10 Materials and Methods……………………………………………………………………….13 Results………………………………………………………………………………………………15 The number of growing microtubules is dramatically up-­‐regulated by axon, but not dendrite, severing…………………………...15 Microtubule polarity switching in dendrites after axon removal………………………………………………………………………………………..18 A dendrite that switches to plus-­‐end-­‐out microtubules can become a growing axon…………………………………………………………..19 Respecification of a dendrite to an axon requires msps-­‐ stimulated microtubule dynamics………………………………………………….23 c-­‐Jun N-­‐terminal Kinase activation is required for all identified responses to axon removal…………………………………………….24 Discussion………………………………………………………………………………………....27 Model for polarity reversal of a dendrite to a regenerating axon……………………………………………………………………………………………..27 iv Microtubule polarity reversal………………………………………………………..29 Microtubule dynamics and stability in regenerating neurons………….29 Control of axon number……………………………………………………………..30 Involvement of JNK signaling in respecification of a dendrite to a regenerating axon………………………………………………….31 Acknowledgements………………………………………………………………………...32 References……………………………………………………………………………………..32 Supporting Data and Supplemental Materials………………………………….35 Chapter 3. Dendrite Injury Triggers DLK-­‐Independent Regeneration…………….…40 Summary, Introduction…………………………………………………………………...41 Results……………………………………………………………………………………………43 Dendrite injury triggers robust regeneration in multiple types of dendritic arborization neurons………………………………….…..43 Late larval and adult neurons retain the capacity to regrow dendrites……………………………………………………………………....46 The conserved DLK axon regeneration pathway is not involved in the response to dendrite injury…………………………….……48 Discussion……………………………………………………………………………………....52 Experimental Procedures………………………………………………………………..53 Acknowledgements………………………………………………………………………...54 References…………………………………………………………………………………......54 Supplemental Information……………………………………………………………....58 Supplemental Figures…………………………………………………………………58 Supplemental Experimental Procedures……………………………………...63 Chapter 4. Conclusions…………………………………………………………………………………….67 v Microtubules and axon versus dendrite injury……………………………..67 Microtubule polarity in sea anenome neurons……………………………..70 Acknowledgements………………………………………………………………………...73 References……………………………………………………………………………………..74 vi List of Figures Chapter 1 Figure 1. Distribution of organelles within a neuron…………………………………..1 Figure 2. Arrangement of microtubules in axons and dendrites of cultured vertebrate neurons………………………………………………………..3 Figure 3. EB3-­‐GFP live imaging………………………………………………………………….4 Figure 4. Comparison of microtubule arrangements in vertebrate and Drosophila neurons……………………………………………………………...5 Figure 5. Model of polarized transport into axons and dendrites Using microtubule motor proteins………………………………………………6 Chapter 2 Figure 1. The number of growing microtubules is upregulated by axon, but not dendrite, severing………………………………………………...16 Figure 2. Axon injury induces orientation switching of dendritic microtubules…………………………………………………………………………….18 Figure 3. Axon, but not dendrite, injury induces extensive tip growth from a dendrite…………………………………………………………….20 Figure 4. Apc2-­‐GFP is excluded from growing processes…………………………..22 Figure 5. RNAi targeting msps blocks regeneration from a dendrite after axon removal……………………………………………………………………24 Figure 6. JNK is required for up-­‐regulation of microtubule number and initiation of growth in response to axon removal…………………25 Figure 7. Reduction of JNK signaling affects microtubule polarity in uninjured neurons……………………………………………………………………27 Figure 8. Model for conversion of a dendrite to a regenerating axon after axon removal……………………………………………………………………28 Figure S1. Additional examples of cells that initiated tip extension from a respecified dendrite……………………………………………………..35 vii Figure S2. Neurons that initiate regeneration have a single process that switches to the axonal microtubule polarity……………………...36 Figure S3. Neurons that express hairpin RNAs to target msps are grossly normal………………………………………………………………………..37 Figure S4..……………………………………………………………………………………………….38 Chapter 3 Figure 1. Dendrites regrow completely after they are removed from larval neurons…………………………………………………………………………44 Figure 2. Adult neurons reinitiate dendrite outgrowth after dendrite injury………………………………………………………………………………………47 Figure 3. Wnd is not required for dendrite regeneration…………………………..49 Figure 4. Downstream components of the DLK pathway are dispensable for dendrite regeneration………………………………………51 Figure S1. Dendrite regeneration in ddaE and ddaC neurons..............................58 Figure S2. Neurons respond to injury of single dendrites, and regenerate when injury is further from the cell body, and regenerate throughout larval life, related to Figure 1…………61 Figure S3. Wnd and hiw RNAi do not block dendrite regeneration, related to Figure 3………………………………………………………………….62 Figure S4. A target of the DLK/JNK/fos pathway is not activated by dendrite injury, related to Figure 4…………………………………………63 Chapter 4 Figure 1. RNAi screen of microtubule severing proteins and gamma tubulin 23C………………………………………………………………………………68 Figure 2. Axon and dendrite injury trigger different regeneration pathways…………………………………………………………………………………69 Figure 3. Ganglion cells imaged in sea anenome polyps…………………………….71 Figure 4. Microtubule arrangements in neurites of different types of sea anenome ganglion cells……………………………………………………71 viii Figure 5. Possible synaptic arrangements in sea anenome neurons…………...73 ix List of Tables Chapter 4 Table 1. Presynaptic and postsynaptic markers to determine synaptic arrangements in sea anenome ganglion neurons…………...72 x Acknowledgements For research acknowledgements, please see the individual chapters. I would first like to thank Melissa Rolls for being a wonderful mentor and boss, and for believing in me and encouraging me to pursue a graduate degree. I have learned so much about cell biology from you! Thank you for your enthusiasm about science and thank you for your invaluable advice on anything from science to gardening to parenting. I would also like to thank my former and present co-­‐
workers whom I have thoroughly enjoyed working alongside for many years: Floyd Mattie, Michelle Nguyen, Li Chen, Juan Tao, Kavitha Rao, Melissa Long, Kyle Gheres, Rich Albertson, Chengye Feng and Dan Goetschius. You have made working in the Rolls’ lab fun and exciting! I would like to thank the members of my thesis committee Richard Ordway and Robert Paulson for their guidance. A special thanks goes to Deborah Murray for all her help with paperwork and for keeping me on track with degree requirements. I would like to thank my family for all their support. A heartfelt thanks goes to my dad Michael Sobeck, a stroke survivor and the motivation behind my desire to study neuronal responses to injury. Your recovery and continued resolve to lead a normal life in the face of physical and mental limitations is an inspiration to our family. Finally, I would like to thank my husband Tim for his continual support and daughter Emma for all the joy she brings to my life. xi Chapter One Introduction Neurons are among the many polarized cells that comprise the vertebrate and invertebrate body. They in fact are highly polarized, typically consisting of a single axon that sends signals to other neurons or muscle cells, and multiple dendrites that receive signals from other neurons. Because of this intrinsic polarity, different proteins and organelles must be positioned and maintained in the axon and dendrites, as well as the soma. Components of the axon, dendrites and soma have been well studied. The distribution of organelles is such that mitochondria and smooth endoplasmic reticulum are found throughout the neuron, while the Golgi complex and ribosomes are found in the soma and dendrites (particularly at dendritic branch points). The soma is known to be the site of most protein synthesis in neurons and contains the rough ER and mRNAs. In addition, presynaptic components are typically found in the axon and postsynaptic components in the dendrites [1] (Figure 1). Figure 1. Distribution of organelles within a neuron. 1 The positioning of organelles and proteins to specific compartments is carried out through polarized trafficking by motor proteins, and this relies heavily on polarized microtubule arrays within the neuron. Microtubules are a key component of the neuronal cytoskeleton and are important for maintaining the individual functions of axons and dendrites. They are comprised of alpha and beta tubulin heterodimers that associate head-­‐to-­‐tail to form protofilaments, which in turn assemble into hollow rods. These rods, or microtubules, have their own intrinsic polarity, consisting of a static minus end and a dynamic plus end. The minus end of the microtubule is capped by gamma tubulin ring complexes, and is where nucleation begins. The plus end of the microtubule undergoes continual phases of polymerization and depolymerization, also known as dynamic instability. This dynamic instability is driven by GTP hydrolysis [2]. Microtubules are located in the axon, dendrites and soma and lay the “tracks” for motor proteins to carry specific cargos to the intended compartment of the neuron. Kinesin family member proteins and dynein are responsible for this movement of proteins and organelles. Kinesins carry cargo toward the plus end of microtubules and dynein carries cargo toward the minus end. Defects in neuronal transport and microtubule dynamics have been shown to contribute to diseases such as Huntingdon’s disease [3], Charcot-­‐Marie-­‐Tooth disease [4], and ALS [5] to name but a few. Therefore, it is extremely important to understand the microtubule arrangement in both axons and dendrites, and how these arrangements are established and maintained. The earliest studies of microtubule polarity in neurons employed a technique called hooking where polarity was determined by incubating nerve explants in a special tubulin assembly buffer. In the presence of this buffer, tubulin protofilaments bind laterally to the surface of existing microtubules creating a “hook”, which curves in either a clockwise or counterclockwise direction. Microtubule polarity is determined by the direction of these hooks. [6], [7]. From these initial hook experiments done with cat sciatic nerves and frog olfactory nerves, it was found that axon microtubules were oriented with their plus ends away from the cell body, or plus-­‐end out [8]. Later studies done in cultured chick 2 dorsal root ganglion neurons, cultured rat hippocampal neurons, and cultured rat sympathetic neurons confirmed this axonal microtubule plus-­‐end out orientation [8]. Much fewer studies have been done examining the microtubule polarity in dendrites. Two studies using the hook technique revealed that dendrites of rat hippocampal neurons [9] and frog mitral neurons [10] have roughly a 50% mixture of plus-­‐end out and minus-­‐end out microtubules. This was substantiated by a later study done in cultured rat sympathetic neurons[11] (Figure 2). From the above findings, it had generally been accepted in the field of neuronal polarity that axonal microtubules have a plus-­‐end out orientation and dendritic microtubules have a mixed orientation. Figure 2. Arrangement of microtubules in axons and dendrites of cultured vertebrate neurons. The hook technique contributed greatly to the understanding of neuronal polarity in vertebrate neurons. However it relies on electron microscopy, which can be labor intensive and technically challenging. The introduction of a new technique which uses plus-­‐end binding proteins, or +TIPS, to track the movement of growing microtubules, has opened the door to studying neuronal polarity in vivo [12]. One category of +TIPS is the EB proteins, which bind to the plus end of the microtubule, and when tagged with GFP, can be seen as moving comets within neurons. When the direction of movement of these comets is tracked using fluorescent microscopy, polarity can be inferred. For example, a comet seen moving away from the cell body in an axon or dendrite would mean there is a plus-­‐end out microtubule (Figure 3). 3 The first reported study using EB3-­‐GFP to study microtubule polarity was done with mouse hippocampal cells and Purkinje neurons. The results were similar to studies done with vertebrate neurons using the hook technique [12]. Figure 3. EB3-­GFP live imaging. A. EB3-­‐GFP binds to the plus end of a growing microtubule. B. A moving comet is visualized by fluorescent microscopy to infer polarity of the microtubule. Vertebrate neurons have laid the foundation for the study of neuronal polarity, in large part because invertebrate neurons were thought to be nonpolar or somehow fundamentally different than vertebrate neurons [1]. This school of thought mainly comes from the fact that most invertebrate neurons are unipolar and therefore, it was assumed the machinery for sorting proteins must be completely different than in multipolar neurons. However, it has more recently been accepted that invertebrate neurons are polarized, and microtubule polarity is key to the identity of their axons and dendrites. In D. melanogaster, using live imaging with EB1-­‐GFP, our lab has shown that axonal microtubule polarity resembles that of vertebrate neurons, and is all plus-­‐end out [13]. However in the proximal dendrites of drosophila sensory neurons, motor neurons, and interneurons, it was found that microtubules are ~90% minus-­‐end out [14], [13] (Figure 4). Similar results have been found in C. elegans sensory and motor neurons, i.e., a plus-­‐end out arrangement in the axon and a minus-­‐end out arrangement in the dendrites[15], [16]. 4 Figure 4. Comparison of microtubule arrangements in vertebrate and Drosophila neurons. This raises the possibility that one of the distinguishing characteristics of a dendrite is the presence of minus-­‐end out microtubules and not necessarily a mixed microtubule arrangement. This fits with a model of polarized transport, where kinesins would be responsible for carrying cargo into the axon and dynein would be responsible for carrying cargo into the dendrites (Figure 5). This is in contrast to previous models suggesting kinesins are the major molecular motors responsible for transporting cargos into both axons and dendrites [17]. However, in studies done in drosophila mushroom body neurons where either Lis-­1 or Dhc64c (dynein heavy chain) were mutated, there was a severe defect in dendrite development [18]. In addition, mutations in dlic (dynein light intermediate chain) caused mislocalization of dendritic golgi and endosomes in drosophila dendritic arborization neurons [19], [20]. Taken together, these results support the idea that dynein is responsible for selective transport into dendrites. 5 Figure 5. Model of polarized transport into axons and dendrites using microtubule motor proteins. Dynein carries cargo into dendrites and kinesins carry cargo into axons along microtubule tracks. Maintenance of neuronal polarity is critical for polarized transport and the function of the neuron. Damage from physical injury or neurodegenerative diseases threatens to disrupt this homeostasis. Therefore when a neuron is injured it is important to understand how neurons respond to the injury at a cellular level and elucidate any mechanisms for regeneration that allow for restoration of function and survival. In Drosophila neurons, axons and dendrites have opposite microtubule arrangements. This raises some interesting questions about how axons and dendrites respond to injury: How is microtubule polarity affected after axon or dendrite injury? Can axons regenerate after injury? Can dendrites regenerate after injury? If they can regenerate, do axons and dendrites use the same molecular machinery for regeneration? My thesis work focuses on answering these questions and shows that in Drosophila sensory neurons, both axons and dendrites can regenerate after injury. However this regeneration is achieved through independent mechanisms, and following axon injury, microtubules are dramatically rearranged. 6 REFERENCES 1. Craig, A.M. and G. Banker, Neuronal polarity. Annu Rev Neurosci, 1994. 17: p. 267-­‐310. 2. Desai, A. and T.J. Mitchison, Microtubule polymerization dynamics. Annu Rev Cell Dev Biol, 1997. 13: p. 83-­‐117. 3. Li, X.J., A.L. Orr, and S. Li, Impaired mitochondrial trafficking in Huntington's disease. Biochim Biophys Acta, 2010. 1802(1): p. 62-­‐5. 4. Zhao, C., et al., Charcot-­Marie-­Tooth disease type 2A caused by mutation in a microtubule motor KIF1Bbeta. Cell, 2001. 105(5): p. 587-­‐97. 5. Baird, F.J. and C.L. Bennett, Microtubule defects & Neurodegeneration. J Genet Syndr Gene Ther, 2013. 4: p. 203. 6. Heidemann, S.R. and J.R. McIntosh, Visualization of the structural polarity of microtubules. Nature, 1980. 286(5772): p. 517-­‐9. 7. Burton, P.R. and J.L. Paige, Polarity of axoplasmic microtubules in the olfactory nerve of the frog. Proc Natl Acad Sci U S A, 1981. 78(5): p. 3269-­‐73. 8. Baas, P.W. and S. Lin, Hooks and comets: The story of microtubule polarity orientation in the neuron. Dev Neurobiol, 2011. 71(6): p. 403-­‐18. 9. Baas, P.W., et al., Polarity orientation of microtubules in hippocampal neurons: uniformity in the axon and nonuniformity in the dendrite. Proc Natl Acad Sci U S A, 1988. 85(21): p. 8335-­‐9. 10. Burton, P.R., Dendrites of mitral cell neurons contain microtubules of opposite polarity. Brain Res, 1988. 473(1): p. 107-­‐15. 11. Sharp, D.J., et al., Identification of a microtubule-­associated motor protein essential for dendritic differentiation. J Cell Biol, 1997. 138(4): p. 833-­‐43. 12. Stepanova, T., et al., Visualization of microtubule growth in cultured neurons via the use of EB3-­GFP (end-­binding protein 3-­green fluorescent protein). J Neurosci, 2003. 23(7): p. 2655-­‐64. 13. Stone, M.C., F. Roegiers, and M.M. Rolls, Microtubules have opposite orientation in axons and dendrites of Drosophila neurons. Mol Biol Cell, 2008. 19(10): p. 4122-­‐9. 14. Rolls, M.M., et al., Polarity and intracellular compartmentalization of Drosophila neurons. Neural Dev, 2007. 2: p. 7. 15. Maniar, T.A., et al., UNC-­33 (CRMP) and ankyrin organize microtubules and localize kinesin to polarize axon-­dendrite sorting. Nat Neurosci, 2012. 15(1): p. 48-­‐56. 16. Goodwin, P.R., J.M. Sasaki, and P. Juo, Cyclin-­dependent kinase 5 regulates the polarized trafficking of neuropeptide-­containing dense-­core vesicles in Caenorhabditis elegans motor neurons. J Neurosci, 2012. 32(24): p. 8158-­‐72. 17. Hirokawa, N. and R. Takemura, Molecular motors and mechanisms of directional transport in neurons. Nat Rev Neurosci, 2005. 6(3): p. 201-­‐14. 18. Liu, Z., R. Steward, and L. Luo, Drosophila Lis1 is required for neuroblast proliferation, dendritic elaboration and axonal transport. Nat Cell Biol, 2000. 2(11): p. 776-­‐83. 7 19. 20. Zheng, Y., et al., Dynein is required for polarized dendritic transport and uniform microtubule orientation in axons. Nat Cell Biol, 2008. 10(10): p. 1172-­‐80. Satoh, D., et al., Spatial control of branching within dendritic arbors by dynein-­
dependent transport of Rab5-­endosomes. Nat Cell Biol, 2008. 10(10): p. 1164-­‐
71. 8 Chapter Two Global Up-­Regulation of Microtubule Dynamics and Polarity Reversal during Regeneration of an Axon from a Dendrite Michelle C. Stone, Michelle M. Nguyen, Juan Tao, Dana L. Allender, and Melissa M. Rolls This chapter was published in Molecular Biology of the Cell 2010, Vol. 21 Author Contributions Michelle C. Stone performed all axotomy and dendrite injury experiments, analysis of EB1-­‐GFP comets for polarity and whirlpool experiments, and APC-­‐2 GFP experiments. Michelle M. Nguyen performed the msps experiments. Juan Tao was involved in the initial screen that identified bsk as a molecular player in axon regeneration. Dana L. Allender performed the EB1-­‐GFP velocity quantifications. Melissa M. Rolls supervised all experiments and was involved in data interpretation and the writing process. 9 ABSTRACT Axon regeneration is crucial for recovery after trauma to the nervous system. For neurons to recover from complete axon removal they must respecify a dendrite into an axon-­‐ a complete reversal of polarity. We show that Drosophila neurons in vivo can convert a dendrite to a regenerating axon, and that this process involves rebuilding the entire neuronal microtubule cytoskeleton. Two major microtubule rearrangements are specifically induced by axon, and not dendrite, removal: 1. 10-­‐
fold upregulation of the number of growing microtubules and 2. microtubule polarity reversal. After one dendrite reverses its microtubules, it initiates tip growth and takes on morphological and molecular characteristics of an axon. Only neurons with a single dendrite that reverses polarity are able to initiate tip growth, and normal microtubule plus end dynamics are required to initiate this growth. In addition, we find that JNK signaling is required for both the upregulation of microtubule dynamics and microtubule polarity reversal initiated by axon injury. We conclude that regulation of microtubule dynamics and polarity in response to JNK signaling is key to initiating regeneration of an axon from a dendrite. INTRODUCTION Neurons are highly polarized cells; many neurons have several dendrites that receive information and a single axon to send information. Axons and dendrites have distinct proteins targeted to them, as well as different cytoskeletal arrangements [1]. Unlike many other cell types, most neurons are not replaced during an animal’s lifetime, yet they can be damaged by physical traumas or immune attack. Complete axon removal is the most severe type of axon injury a neuron can sustain, and it renders the cell nonfunctional as it can no longer send signals. Studies performed in several systems suggest that neurons have a tremendous capacity for axon regeneration, even in response to total axon removal. In sea lampreys, complete removal of axons from hindbrain neurons triggers new growth from dendrites that extend beyond the normal dendritic field [2]. This is in contrast to axon injuries further from the cell body that induce regrowth from the axon as well as increased dendritic growth. Subsequent analysis showed that at 10 the ultrastructural level the new processes emerging from dendrites after axon removal had axonal features, for example abundant neurofilaments [3]. Similar observations of growth of axon-­‐like processes from dendrites in vivo have been made in cat motoneurons and interneurons after proximal axotomy [4-­‐6], as well as several types of rodent neurons [7, 8]. Axon removal experiments have also been performed in cultured neurons. Axons of mouse hippocampal neurons in dissociated and slice culture were severed at different distances from the cell body [9]. As in earlier in vivo studies, close axotomy induced new growth from dendrites. These new processes contain tau immunoreactivity, like axons, and are presynaptic [9]. Thus in several systems, neurons initiate regeneration after axon removal by converting a dendrite to a new axon, or at least growing a new axon from the tip of a dendrite. It is not known how neuronal polarity is completely rearranged after axon removal to allow a dendrite to become or grow a new axon. Microtubules in axons and dendrites have different polarity, and several different approaches have suggested that this arrangement is crucial for overall neuronal polarity. In cultured mammalian neurons, axons have uniform polarity microtubules with plus ends distal to the cell body (plus-­‐end-­‐out), while dendrites have mixed polarity with about half plus-­‐end-­‐out and half minus-­‐end-­‐out microtubules [10, 11]. The minus-­‐
end-­‐out microtubules in dendrites have been linked to dendritic identity. In dissociated hippocampal neurons, dendrite specification and growth occurs at the same time that minus-­‐end-­‐out microtubules enter dendrites [12], and loss of the minus-­‐end-­‐out population of microtubules from dendrites results in overall loss of dendritic character [13]. Thus if a dendrite becomes or grows an axon one might expect that microtubule polarity would have to be rearranged. In fact, the process of regenerating an axon from a dendrite does seem to be accompanied by changing the microtubule cytoskeleton from the dendritic polarity to the axonal one. When cortical neurons from newborn rat are dissociated, the axon is lost, but a region of the apical dendrite that has mixed microtubule polarity can remain attached to the cell body. Thus dissociation of the cells causes total axon removal. Within 24 hours of plating in culture, growth initiates from the tip of the 11 dendritic process, and the process has switched to the axonal plus-­‐end-­‐out microtubule polarity [14]. It was suggested that this shift in microtubule polarity is accomplished by loss of the minus-­‐end-­‐out microtubules from the growing process [14]. It has also been suggested that an increase in microtubule stability in the growing process is key to the conversion of a dendrite to a regenerating axon [9]. So the best current model for conversion of a dendrite to an axon after complete axotomy is loss of the minus-­‐end-­‐out microtubules from the mixed population and a concomitant increase in stability of the remaining plus-­‐end-­‐out ones. As this model derives from examining the arrangement of microtubules in fixed preparations at single time-­‐points, we wished to determine how microtubules are rearranged over time as neurons respond to axon injury. We also wished to examine neurons in vivo in their normal environment to eliminate any confounding changes induced by cell culture. Like vertebrate neurons, Drosophila neurons are very polarized [15, 16], moreover their cytoskeleton can be studied in vivo. We and others have previously used live imaging to study the layout of Drosophila microtubules in neurons in living larvae [16-­‐19], and have found that microtubules in all major classes of Drosophila neurons have opposite polarity in axons and dendrites [18]. The axonal arrangement of microtubules is plus-­‐end-­‐out, as in other animals, but the dendrites have a more polarized microtubule cytoskeleton than cultured mammalian neurons, and have greater than 90% minus-­‐end-­‐out microtubules. This arrangement of dendritic microtubules suggests either that Drosophila neurons will not be able to regenerate an axon from a dendrite because they do not have a significant population of plus-­‐end-­‐out microtubules in dendrites, or that the current model for conversion of a dendrite to a regenerating axon is incomplete. Using this tractable model system, we test whether highly polarized neurons in vivo can really reverse polarity, and identified major microtubule rearrangements that are required for initiation of regeneration of an axon from a dendrite. 12 MATERIALS AND METHODS System to study neuronal responses to injury: Drosophila dendritic arborization (da) neurons tile the larval body wall in a stereotyped manner [20]. Class I da neurons have a simple dendritic tree and exhibit little structural plasticity over the course of larval life [21]. Two class I da neurons are present in the dorsal region of each hemisegment. We used ddaE as our model neuron as it is easy to identify and visualize. EB1-­‐GFP can be used to monitor microtubule polarity as it only binds to growing microtubule plus ends. The direction of EB1-­‐GFP comet movement over time therefore points towards the plus end. We scored direction of comets that were visible for three or more consecutive frames. Comets that moved towards the cell body were at tips of minus-­‐end-­‐out microtubules; comets that moved away from the cell body were indicative of plus-­‐
end-­‐out microtubules. Axons were severed in larvae with a UV laser. This method has previously been used to sever da neuron dendrites [21] and axons of C. elegans neurons in vivo [22]. After axon severing, we acquired movies of EB1-­‐GFP, and then put the animals back in food to recover. They could be mounted for imaging multiple times after this treatment and followed over four days, at which point they initiated pupariation. Genetic background, imaging and culturing conditions: Whole, live larvae heterozygous for the class I da neuron specific driver 221-­‐
Gal4 and UAS-­‐EB1-­‐GFP were collected 48-­‐72hr after egg laying and mounted on a microscope slide with a dry agarose pad and covered with a glass coverslip. Animals containing 221-­‐Gal4, UAS-­‐EB1-­‐RFP and UAS-­‐Apc2-­‐GFP transgenes were collected similarly. Dendrites and axons of class I dendritic arborization neurons were severed in these larvae using a Micropoint UV laser (Photonic Instruments). Live imaging was performed using an LSM510 confocal microscope (Carl Zeiss). Single frames were collected every 2 sec to image EB1 dynamics in the cell bodies, dendrites, and axons of injured and uninjured neurons. Immediately after imaging, animals were recovered from microscope slides by adding Schneider’s insect media 13 to release them from the agarose pad and placed into standard Drosophila media for recovery until further imaging. Images were analyzed using ImageJ software (http://rsb.info.nih.gov/ij/;NIH). Quantitiation of microtubule number and polarity For all experiments one neuron per animal was analyzed, and the number of animals for each condition is indicated in the figures as n’s. To analyze microtubule orientation, EB1-­‐GFP comets were tracked manually in the region between the cell body and the first branch point of the dendrite for wild-­‐type, and throughout the whole dendrite for bskRNAi or DN, and only comets that were tracked over a minimum of three consecutive frames were counted. Quantitation of number of EB1-­‐GFP comets in the cell body and dendrites of injured and uninjured neurons was done by taking the first in-­‐focus frame in a movie and subsequent fifth in-­‐focus frames. This was done until a total of three frames were analyzed. EB1-­‐GFP comets were counted in the still images of these frames. Neuronal RNAi and dominant negative experiments For the RNAi experiments we crossed a tester line, UAS-­‐dicer2; 221-­‐Gal4, UAS-­‐EB1-­‐GFP to lines that express hairpin RNAs under UAS control and analyzed microtubule dynamics as above in the ddaE neuron in larval progeny. The 221-­‐Gal4 drives expression of the GFP, dicer2, and RNA hairpin only in class I dendritic arborization neurons and a few other cells, meaning that most of the animal is normal. The RNA hairpin lines were from the VDRC collection [23]. The VDRC stock numbers used to target proteins of interest were: 34138 (bsk) and 21982 (msps). As a control the tester line was crossed to a line containing a different transgene, UAS-­‐mCD8-­‐RFP. For bskDN expression we crossed the line w1118, P{UAS-­‐bsk.DN}2 (Bloomington Drosophila Stock Center) to 221-­‐Gal4, UAS-­‐EB1-­‐GFP to generate larvae heterozygous for all transgenes. As a control for this experiment, we crossed the 221-­‐Gal4, UAS-­‐EB1-­‐GFP line to a line containing a different transgene: UAS-­‐
14 mCD8-­‐RFP, so that the 221-­‐Gal4 would be driving expression of the same number of transgenes in both cases. RESULTS The number of growing microtubules is dramatically up-­regulated by axon, but not dendrite, severing. To test how neurons in vivo respond to axon removal, we generated Drosophila larvae that express a microtubule plus end-­‐binding protein tagged with GFP, EB1-­‐GFP, in a small subset of dendritic arborization sensory neurons. These neurons have branched dendrites reminiscent of mammalian dendrites [20], and are ideal for live imaging as they lie immediately under the larval cuticle. Moreover, although these cells are sensory, the arrangement of microtubules in their dendrites is similar to that in Drosophila central neurons [18]. To further simplify the analysis, we focused on the ddaE cell on the dorsal side of the larva. This cell is one of the class I dendritic arborization neurons, and has a relatively simple dendrite branching pattern [20]. To sever axons in these neurons in vivo, we mounted larvae on a slide and focused a pulsed UV laser on the region of the cell we wanted to cut. This method has previously been used to sever Drosophila dendrites [21] and Caenorhabditis elegans axons [22]. Using this method, we successfully generated small cuts in axons or dendrites (Figure 1A), that resulted in degeneration of the region distal to the cut site by 24 hours (Figure 1A). After injury larvae were recovered to media and allowed to develop as normal. They were remounted for imaging at 24 hour intervals. 15 Figure 1. The number of growing microtubules is upregulated by axon, but not dendrite, severing. (A.) Images of EB1-­‐GFP in the ddaE neuron were acquired before, immediately after (0 h) and 24h after axon (top row) or dendrite (bottom row) severing. Two frames are shown from each movie from the 24-­‐h time point. Arrows indicate the site of UV laser-­‐mediated severing. Arrowheads point out examples of EB1-­‐GFP comets in the cell body. In all figures dorsal is up. (B.) Panels from movies acquired 24h after axon or dendrite severing are shown. Images were inverted for ease of identifying EB1-­‐GFP comets in dendrites; examples are marked with arrowheads. (C.) The number of EB1-­‐GFP comets in the cell body or a region of dendrite 2 was counted in single frames from movies of uninjured neurons, and from neurons 24h after axon or dendrite severing. Three frames were averaged for each animal, error bars, SD of the average from all animals. n = number of animals scored (one neuron per animal). Unpaired t tests were used to determine whether the number of comets was significantly increased after dendrite or axon cutting. No significant difference between number of dots in the cell body before and after dendrite cutting was found. Significant differences were found for cell bodies and dendrites before and after axon cutting. The first alteration in EB1-­‐GFP dynamics we observed after axon severing was unexpected. Rather than an increase in microtubule stability, which would be detected as fewer growing microtubule ends labeled with EB1-­‐GFP, we saw a large increase in the number of EB1-­‐GFP comets. This increase was initiated in some cells 16 immediately after axon severing (Movie 1), and was always seen at 24 hours (Figure 1 and Movies 1, 2 and 3). Before axotomy, EB1-­‐GFP comets at tips of growing microtubules were seen occasionally in the cell body. 24 hours after axotomy, many EB1-­‐GFP comets were seen swirling like a whirlpool in the neuronal cell body (Figure 1A and Movies 1 and 2). The number of EB1-­‐GFP comets has previously been used as a readout of total number of dynamic microtubules [24], and so we interpret this increase in comet number as an increase in number of growing microtubules, and it is likely that all neuronal microtubules exhibit growth at their plus ends. Comparison of the number of EB1-­‐GFP comets in uninjured neurons with neurons after axotomy revealed a more than 10-­‐fold increase in number of comets in the cell body 24 hours after axon severing (Figure 1C). The number of comets also increased throughout the dendritic tree (Figure 1B and 1C). Although the number of EB1-­‐GFP comets was increased by axon removal, the rate of microtubule growth was not: before severing the average velocity of EB1-­‐GFP comets was 0.225 mm/s (standard deviation, SD, 0.043, data is from 40 microtubules in 6 animals), and after severing it was 0.229 mm/s (SD 0.068, data is from 37 microtubules in 8 animals). Because this increase in number of growing microtubules was unexpected, we wanted to determine whether it was a general response to neuronal injury, or a specific response to axon severing. We therefore severed dendrites close to the cell body and monitored EB1-­‐GFP dynamics after this injury. Dendrite severing did not increase number of EB1-­‐GFP comets in the cell body or remaining dendrites (Figure 1 and Movie 2), indicating that this is a specific response to axon injury. Thus although injury to both axons and dendrites is predicted to cause a breach in the plasma membrane and ion influx, only axon injury triggers a global increase in dynamic microtubule number. 17 Figure 2. Axon injury induces orientation switching of dendritic microtubules. (A) Images of EB1-­‐GFP in a single ddaE neuron at different times before or after axon injury. Several confocal images were projected to give a complete overview of the dendrite arbor. In zoomed in movies, the direction of EB1-­‐GFP comet movement was scored. Comets moving toward the cell body represent minus-­‐end-­‐out microtubules and comets moving away from the cell body represent plus-­‐end-­‐out microtubules. The raw data are shown in the table and represented by arrows on the overview pictures. A green arrow indicates plus-­‐end-­‐out microtubule orientation, a red arrow indicates minus-­‐end-­‐out orientation and double arrow indicates mixed orientation. Movie 3 shows microtubule dynamics in this cell. (B) Microtubule orientation was quantitated in dendrites of uninjured neurons and neurons at different times after axon severing. Comets were scored in each dendrite as in A: EB1-­‐GFP dots in the region of the dendrite between the cell body and first dendrite branch point were counted; dendrites with four or more comets were classified as plus-­‐end-­‐out if 75% or more comets moved away from the cell body and minus-­‐end-­‐out if 75% or more moved toward the cell body. The class in between was classified as mixed, and is not shown explicitly in the table. n= number of dendrites classified for each time point. Microtubule polarity switching in dendrites after axon removal. As well as monitoring the number of EB1-­‐GFP comets after axon removal, we monitored their direction of movement in dendrites to determine whether a change in microtubule polarity might be triggered by loss of the axon (Figure 2A and Movie 3). EB1-­‐GFP binds only to growing microtubule plus ends, so the direction of movement gives a readout of microtubule polarity [11, 18]. To simplify 18 comparisons between neurons, we numbered each dendrite of the ddaE dendritic arborization neuron (Figure 2). The dendrite with a comb-­‐like-­‐branching pattern was numbered 1; the dendrite closest to the axon site was numbered 2. The presence of a third (or fourth) dendrite was variable. If these were present they were numbered 3 and 4. In uninjured neurons, comets in most dendrites moved towards the cell body consistent with minus-­‐end-­‐out microtubule orientation (Figure 2). Minus-­‐end-­‐out microtubule orientation is found in all types of Drosophila dendrites, and distinguishes them from axons which have plus-­‐end-­‐out microtubules [18]. Dendrites with plus-­‐end-­‐out microtubule orientation were never observed before axon injury (Figure 2B). By 6 hours after proximal axon severing, dendrites were frequently observed to have the axonal plus-­‐end-­‐out microtubule orientation (Figure 2B). The time period from 6 to 24 hours seemed to be a transition period, and by 48 hours many of the neurons had one dendrite with plus-­‐end-­‐out microtubules and the remaining dendrites had reverted to minus-­‐end-­‐out orientation (Figure 2B). In most cases dendrite 2 was the one that acquired the axonal microtubule arrangement. Thus microtubule polarity could be completely reversed in a dendrite after axon removal. A dendrite that switches to plus-­end-­out microtubules can become a growing axon. To determine whether the dendrite that acquired the axonal plus-­‐end-­‐out microtubule orientation had other axonal properties and could initiate growth, we performed experiments over a longer time course. At 72 and 96 hours after axon severing, many processes with plus-­‐end-­‐out microtubules had a bulbous structure at the end. The plus-­‐end-­‐out process also initiated extension in many cases (Figures 2A, 3C, 3D and S1B). 6/9 neurons in one experiment initiated growth from a dendrite tip. The neurons that initiated tip growth were distinguished by having a single process that switched to plus-­‐end-­‐out microtubule orientation (Figure S2). In these neurons, the remaining minus-­‐end-­‐out processes did not extend from their tips; they continued the normal behavior of ddaE dendrites, which is to expand all 19 Figure 3. Axon, but not dendrite, injury induces extensive tip growth from a dendrite. The axon or dendrite of the ddaE neuron was severed with a UV laser, and EB1-­‐GFP was imaged at different time points. Overview images were compiled from movies of EB1-­‐GFP, and microtubule orientation was scored as in Figure 2. Yellow arrows, site of laser severing; red arrows, minus-­‐end-­‐
out microtubules; green arrows, plus-­‐end-­‐out microtubules; double arrows, mixed orientation. Stars label tips of processes that have extended by 96 h. Six of nine cells in which the axon was removed initiated tip growth, and 0 of 5 in which a dendrite was removed initiated tip growth. The dendrites are numbered as in Figure 2. Movie frames were Z projected, maximum method, to show the entire dendritic tree. In some cases several frames of a movie had to be assembled next to one another to cover the complete area of the dendrites. The images were also rotated and placed on a black background so that the neuron would be seen in the same orientation at all time points. Scale is the same for all images. 20 over as the animal grows [21]. In this normal, all-­‐over expansion, the distance between the last branch point and dendrite tip does not increase at a greater rate than the distance between internal branch point. Normal dendrite expansion rather than tip growth was also observed in time course experiments after dendrite removal (Figure 3A and B; n=5; note the same V-­‐shape is seen at the remaining dendrite tips at all time points). Thus tip growth is a specific response to axon removal rather than a general response to injury. As well as initiating tip growth, the plus-­‐end-­‐out process generated after axon removal lost key dendritic features. Class I da neuron dendrites normally exhibit tiling behavior with one another and do not cross [25]. Growing processes with plus-­‐end-­‐out microtubules could cross neighboring processes from the same cell (Figure S1B). Dendrites from these cells are also normally restricted to a subregion of the larval body, but growing processes often extended beyond normal dendritic territory (Figures 3C and S1A). To test more rigorously whether dendrites that initiated tip growth had the molecular characteristics of axons, we tracked the behavior of Apc2-­‐GFP in ddaE after axon severing. There are very few markers that localize robustly to axons or dendrites in Drosophila when tagged and overexpressed. One of the most robust dendrite markers is Apc2-­‐GFP [16]. In Drosophila there are two APC (adenomatous polyposis coli) proteins. A tagged version of one of these, Apc2-­‐GFP, expressed in Drosophila central neurons localizes to spots in dendrites, to the cell body, and to the first part of the axon, but is cleanly excluded from distal axons [16]. When we expressed Apc2-­‐GFP in da neurons we saw a similar pattern of fluorescence, with distinct spots of Apc2-­‐GFP in dendrites, Apc2-­‐GFP in some, but not all, proximal axons (Figure 4A-­‐C), but not in distal axons (Figure 4D). At 96 hours after axon removal, when one process with plus-­‐end-­‐out microtubules exhibited significant growth, Apc2-­‐GFP was seen in puncta throughout most of the dendritic tree, but was not seen in the new region (Figure 4B and C; of 12 animals in which tip growth was initiated, Apc2-­‐GFP was never seen in the new region). 21 Figure 4. Apc2-­GFP is excluded from growing processes. Apc2-­‐GFP and EB1-­‐RFP were expressed in ddaE neurons. (A) Uninjured neurons were imaged over the same time course used in other experiments. At all times Apc2-­‐GFP is seen in spots throughout the dendritic arbor. (B and C) Axon-­‐
severing experiments were performed as in Figure 3. In both cells shown tip growth is initiated from dendrite 2. Apc2-­‐GFP is only found in the proximal region of this process at 96h after axon removal, and this pattern was seen in a total of 12of 12 neurons which initiated tip growth. (D) The ddaE neuron extends its axon from the body wall to the ventral ganglion. In live animals expressing Apc2-­‐
GFP and EB1-­‐RFP in class I da neurons, EB1-­‐RFP can be seen in the distal axons that enter the ventral ganglion. Apc2-­‐GFP is not seen in these axons. See Figure S4 for greyscale images of Apc2-­‐GFP alone. Apc2-­‐GFP was seen at the base of the process with plus-­‐end-­‐out microtubule orientation, consistent with localization of Apc2-­‐GFP in proximal axons, or failure to clear Apc2-­‐GFP from the region that was previously a dendrite. This data strongly argues that the process with plus-­‐end-­‐out microtubules has been respecified as a 22 regenerating axon. Thus even a mature dendrite in vivo in a cell that normally exhibits no structural plasticity [21] can be respecified as an axon, and part of this conversion is total reversal of microtubule polarity throughout the entire process, including the region that was previously a dendrite. Respecification of a dendrite to an axon requires msps-­stimulated microtubule dynamics. As the microtubule cytoskeleton was dramatically rearranged after axon removal, we hypothesized that regulation of microtubule dynamics would be essential to respecifying a dendrite as an axon and initiating growth. To test this hypothesis we searched for genetic methods to reduce microtubule dynamics. We found that targeting the msps transcript by RNAi almost completely eliminated EB1-­‐
GFP comets from neurons (Figure 5A and Movie 4). In control (rtnl2) RNAi neurons EB1-­‐GFP comets are seen frequently throughout the dendritic tree (Figure 5A and Movie 5), similar to uninjured neurons that do not express hairpin RNAs or neurons after dendrite severing (Figure 1 and Movies 1 and 3). This result is consistent with models of microtubule growth in which msps (or XMAP215 in vertebrates) acts as a microtubule polymerase [26, 27], and loss of msps in vivo results in increased microtubule pausing [28]. We confirmed that although microtubule plus ends do not behave normally in neurons with reduced msps, stable microtubules are present (Figure S3A), and neuronal structure is quite normal throughout larval life, with the exception that in some larvae increased dendritic branching was observed close to the cell body (Figure S3B). When ddaE neurons expressing msps RNAi hairpins were subjected to axon removal, in all cases they failed to initiate growth from a dendrite tip (Figure 5B, n=5). As in uninjured larvae, the shape of the dendritic tree remained constant through larval life. We therefore conclude that msps-­‐stimulated microtubule dynamics are not required for maintenance of normal dendrite structure during larval life, but are required to convert a dendrite into a regenerating axon, supporting the role of microtubule plus end dynamics in this process. 23 Figure 5. RNAi targeting msps blocks regeneration from a dendrite after axon removal. (A) The cell body and proximal dendrite of ddaE neurons expressing EB1-­‐GFP and hairpin RNAs that target rtnl2 (control) or msps are shown. rtnl2 RNAi was used as a control as its loss has no known consequences in flies. EB1-­‐GFP comets (red arrows) can be seen in control, but not msps, RNAi neurons. (B) An axon-­‐severing experiment as in Figure 3 was performed on a ddaE neuron expressing EB1-­‐GFP and msps hairpin RNA. Cell shape (but not microtubule polarity, as no EB1-­‐GFP comets were present) were tracked over time. No tip growth was observed (n=5). c-­Jun N-­terminal Kinase activation is required for all identified responses to axon removal. We have identified two major types of microtubule reorganization in response to axon removal: increased number of growing microtubules and polarity reversal. To identify the upstream signals that trigger these rearrangements, we took a candidate approach. JNK, or related MAP kinases, have been implicated in axon regeneration in multiple systems, including C. elegans, Drosophila, and mammals [29-­‐31]. In each of these systems these kinases were shown to be important for initiating growth from an axon stump after distal axon severing, and it has been hypothesized that unidentified rearrangements of microtubules might be important for this growth[29]. We therefore hypothesized that JNK signaling could 24 Figure 6. JNK is required for up-­regulation of microtubule number and initiation of growth in response to axon removal. (A) ddaE neurons expressing EB1-­‐GFP and mCD8-­‐RFP (control), RNAi hairpins to target bsk , or bskDN, were imaged 24h after axon removal. Numerous EB1-­‐GFP comets (arrowheads) were seen in control neurons, and many fewer were seen is bsk RNAi or bskDN neurons. (B) EB1-­‐GFP comets in the cell body were quantitated 24h after axon injury. Number of comets in individual frames of movies was counted as in Figure 1C. Genotypes of the larvae in order shown in the table were: (1) UAS-­‐Dicer2/UAS-­‐mCD8-­‐RFP; 221-­‐Gal4, UAS-­‐EB1-­‐GFP/+ (2) UAS-­‐
Dicer2/UAS-­‐bskRNAi; 221-­‐Gal4, UAS-­‐EB1-­‐GFP/+ (3) UAS-­‐mCD8-­‐RFP/+; 221-­‐Gal4, UAS-­‐EB1-­‐GFP/+ (4) UAS-­‐bskDN/+;; 221-­‐Gal4; UAS-­‐EB1-­‐GFP. (C and D) A ddaE neuron expressing EB1-­‐GFP and bskDN was tracked over time. The dendrite arbor of these cells retained the same shape over time as in controls ( n=6). In D, microtubule orientation was determine as in Figure 3, except that comets were quantitated throughout major dendrites because fewer comets were present. Red arrows, minus-­‐end-­‐out polarity (>75% of comets to the cell body); double arrows, mixed polarity (25-­‐75% of comets to the cell body). play a role in the response to proximal axon severing, although this initiates growth from a dendrite rather than extension from the axon stump. 25 We used two methods to block JNK signaling, expression of RNA hairpins to target bsk, the Drosophila JNK, and expression of a non-­‐phosphorylatable dominant negative JNK (bskDN [32]. Control neurons expressing a different hairpin RNA (targeting rtnl2) had normal shape, and the dendrite arbor remained stable through larval life (not shown). After axon severing, the increase in number of growing microtubules was significantly blocked in bsk RNAi and bskDN neurons compared with neurons expressing a different transgene, mCD8-­‐RFP (Figure 6A and B). We conclude that one of the earliest changes in the microtubule cytoskeleton induced by axon injury, upregulation of dynamic microtubule number, requires JNK activity. To determine whether the other intracellular events we mapped during conversion of a dendrite to a regenerating axon were also initiated by JNK signaling, we assayed neuron shape and microtubule polarity for four days after axon removal. Expression of either bsk RNAi or bskDN blocked growth from the tip of a dendrite (n=10 for RNAi, n=6 for DN, Figure 6C and D); the only change in shape after injury was an increase in proximal branching in some animals. Both treatments also blocked most polarity changes in dendrites after axon removal. Out of five ddaE neurons assayed for microtubule polarity, only one had any dendrites that acquired the axonal plus-­‐end-­‐out microtubule orientation, and this one actually had two processes that became plus-­‐end-­‐out. In the rest of the neurons, dendrite microtubule polarity was mixed or minus-­‐end-­‐out at all time points as in Figure 6D. As so many dendrites had mixed polarity, we wondered whether there was any change in polarity in uninjured neurons expressing bskDN. Interestingly, the #2 dendrite, which has a simpler branching pattern than the #1 (or comb) dendrite, was significantly more mixed in cells expressing bskDN (Figure 7). This suggests that JNK signaling may play a role in dendritic microtubule polarity even in uninjured neurons. We hypothesize that the difference was only seen in the #2 dendrite and not the #1 because another mechanism is present that relies on dendrite branching to maintain microtubule polarity (F. Mattie and M. Rolls, unpublished results). Axon polarity was unchanged by expression of bskDN (not shown). We conclude that activation of JNK initiates microtubule rearrangements 26 that precede conversion of a dendrite to a growing axon, and that JNK may also play a role in controlling microtubule polarity in uninjured dendrites. Figure 7. Reduction of JNK signaling affects microtubule polarity in uninjured neurons. Microtubule polarity was assayed by tracking direction of EB1-­‐GFP comet movement in uninjured neurons. Comets were counted in main trunk of the comb dendrite (1), and in dendrite 2 (see Figure 2B). The percent of dots moving toward the cell body is shown for dendrites. The percent was calculated for each cell; error bars, SD. An unpaired t-­‐test was used to calculate the significance of the difference between wild type and bskDN. For axons, the percent of comets moving away from the cell body is shown. DISCUSSION Model for polarity reversal of a dendrite to a regenerating axon. Using this simple system to study neuronal responses to injury, we have found that even mature dendrites that normally exhibit little plasticity can undergo polarity reversal to be respecified as axons in vivo. Vertebrate neurons can also initiate axon formation from a dendrite [2-­‐9, 14], and thus this process is likely to be evolutionarily ancient and widely conserved. Based on our data, we propose a model for respecifying a dendrite as a regenerating axon (Figure 8). First, axon injury elicits a specific signal that is not generated by similar injury to dendrites. This signal turns on a kinase cascade that leads to JNK activation. Phosphorylated JNK upregulates the number of growing microtubules, perhaps by increasing microtubule nucleation or severing. We 27 hypothesize that this results in overall shorter microtubules that may be more likely to undergo complete catastrophe, facilitating polarity changes. After a transition phase during which multiple microtubule orientation switches can occur, a single dendrite takes on the axonal microtubule arrangement. JNK signaling also regulates the acquisition of plus-­‐end-­‐out microtubule polarity in this dendrite. Growth from the tip of the new axon is initiated, presumably as a result of transport of new components along the reoriented microtubules. We think that these changes in microtubule dynamics and polarity are required to initiate regeneration of an axon from a dendrite because only those neurons that switch one dendrite to the axonal microtubule polarity initiate tip growth, and genetically disrupting normal plus end microtubule dynamics blocks tip growth. Figure 8. Model for conversion of a dendrite to a regenerating axon after axon removal. Before axon injury, microtubules in dendrites have minus-­‐end-­‐out polarity (red arrows) and axonal microtubules are plus-­‐end-­‐out. Axon removal up-­‐regulates JNK signaling, which switches microtubule polarity in dendrites, frequently resulting in mixed polarity (purple double arrows) and up-­‐regulates the number of microtubules in the cell body (white circle) and throughout the dendrites. Over several days polarity resolves such that one dendrite takes on the axonal microtubule polarity and the rest return to minus-­‐end-­‐out polarity. After this point the process with axonal microtubule polarity initiates tip growth. 28 Microtubule polarity reversal. While our results suggest that reversing microtubule polarity is crucial for respecifying a dendrite as an axon, we do not yet know how this is accomplished at the molecular level. In the dendrite that switches from minus-­‐end-­‐out to plus-­‐end-­‐
out polarity, it is unlikely existing microtubules could be turned around in the narrow dendrite, and so all the minus-­‐end-­‐out microtubules must be completely depolymerized over the 48 hour time period in which respecification occurs. Depolymerization could be accomplished by severing minus-­‐end-­‐out microtubules or increasing catastrophe rates. Alternately, the normal microtubule turnover rate could be maintained, but addition of new minus-­‐end-­‐out microtubules could be blocked. During this switchover time, new plus-­‐end-­‐out microtubules must be added. One way this could be done is by nucleation of new microtubules in dendrites, although, for this to result in addition of only plus-­‐end-­‐out microtubules, orientation of nucleation sites would need to be controlled. Very little is known about mechanisms that generate uniform microtubule polarity in neurons, and so it will be extremely exciting to test whether oriented nucleation or selective depolymerization could play a role in determining polarity. Microtubule dynamics and stability in regenerating neurons. Massive upregulation of the number of growing microtubules, and most likely overall microtubule number, in neurons after axon injury is extremely surprising. The number of growing microtubules is known to be upregulated at centrosomes upon entry into mitosis [24], but this type of regulation has not been described in terminally differentiated cells like neurons. Our results suggest that neurons maintain a pathway to upregulate microtubule number and activate it when injured. A previous study suggested that increased microtubule stability, rather than an increase in microtubule dynamics, might be key for axon regeneration from a dendrite [9] consistent with a role for microtubule stability in initial axon specification [33, 34]. How can we reconcile this with our finding that increased microtubule dynamics is a robust early step in the response to axon removal? 29 The increase in microtubule number that we observe happens before tip growth is initiated, and number of growing microtubules generally decreases by 72 hours after axon injury (Movie 3); tip growth is typically first obvious at 72 hours, and is extensive by 96 hours. Quantitation of the number of growing microtubules in dendrites confirms this impression: at 24 hours after axon removal an average of 4.9 (SD 1.8) and 5.0 (SD 2.9) comets per 20 mm were observed in the comb and #2 dendrites respectively. At 96 hours after axon removal these numbers were reduced to 2.7 (SD 1.9) and 2.1 (SD 1.5) comets per 20 mm (in an unpaired t test the difference between the number of comets in the comb or #2 dendrite at both time points had p values < 0.05). Microtubule stability thus increases in the days following the initial upregulation of dynamics. Our results support a model in which reduced microtubule stability plays a role in respecification of a dendrite to an axon. After respecification, microtubule stability is increased, and this increase could be important for growth, although we have not tested this idea in the current study. Control of axon number. A previous study on responses to axon severing in slice and dissociated neuronal cultures found that more than one axon could be generated by respecification [9]. In contrast, we find that in vivo the control on axon number is maintained: in cells that initiated tip growth, only one process arising from the cell body acquired axonal microtubule orientation. Importantly, the control on axon number was in the proximal region of the process: only one plus-­‐end-­‐out process arose from the cell body, but more than one tip of this process could have a growth cone and extend (Figure 3C and D). In fact, this was often the case, and the growing tips tended to extend in different directions across the body wall. This could maximize their chance of finding the nerve and route to the target in the brain. We were only able to follow neurons for four days after axon severing due to onset of pupariation, so we could not determine whether extending axons were ever successful in finding the nerve or targets. Most of the time the single dendrite that was respecfied as an axon was the one closest to the site of the previous axon, the #2 dendrite. There are several 30 possible models to explain this. One is that the side of the cell nearest the previous axon experiences some type of inductive signal so that the new axon is closest to the original one, and thus has the best chance of finding the target. We prefer an alternative model as there is no evidence that the respecified process receives any directional information once it initiates growth. In this model, the key factor is the branching complexity of the dendrite. The dendrite opposite the axon has a comb shape with many more branches off the main trunk than the #2 dendrite. It may be more straighforward to generate uniform plus-­‐end-­‐out polarity in a process with a simpler branching pattern. Involvement of JNK signaling in respecification of a dendrite to a regenerating axon. The finding that JNK regulates number of growing microtubules, as well as microtubule polarity and tip growth are significant for several reasons. JNK, or other similar MAP kinases, are established initiators of axon regeneration from an axon stump in other systems [29-­‐31], and the fact that it is also required to respond to axon severing near the cell body suggests these responses may be closely related. In regeneration of an axon from its stump the cytoskeletal rearrangements initiated by JNK signaling have not been identified. Control of microtubule number and polarity are also possible JNK targets in this response. In fact, in aplysia, local polarity reversal of microtubules near axon cut sites are seen [35]. Although the classic role for JNK is modulation of transcription by phosphorylation of transcription factors, it can also phosphorylate cytoskeletal proteins [36]. The fact that we observe extremely rapid (less than five minutes) changes in microtubule dynamics in some cells (Movie 1) suggests that JNK may regulate microtubules in injured neurons by direct regulation of cytoskeletal proteins rather than through transcription changes. Regulation of microtubule number by JNK is also significant because very little is known about what controls the number of growing microtubules in interphase cells. New work suggests that reducing levels of the stathmin protein in interphase cultured cells can increase microtubule nucleation, and thus microtubule number [37], but no circumstances have been identified in which regulation of 31 microtubule number might be important in non-­‐dividing cells. Our results show that studying control of microtubule number will be key for understanding neuronal responses to injury and neuronal polarity. ACKNOWLEDGEMENTS We would like to thank W. Grueber for the 221-­‐Gal4 line and W. Hanna-­‐Rose for sharing her UV laser and for stimulating discussions. We would also like to acknowledge the Bloomington Drosophila Stock Center and the Vienna Drosophila RNAi Center; they are amazing resources. Funding for this work was provided by an American Heart Association Scientist Development Grant and a March of Dimes Basil O’Connor Starter Scholar Award and NIH grant R21NS066216. MMR is a Pew Scholar in the Biomedical Sciences. REFERENCES 1. 2. 3. 4. 5. 6. 7. Craig, A.M. and G. Banker, Neuronal polarity. Annu Rev Neurosci, 1994. 17: p. 267-­‐310. Hall, G.F. and M.J. Cohen, Extensive dendritic sprouting induced by close axotomy of central neurons in the lamprey. Science, 1983. 222(4623): p. 518-­‐
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Itoh, A., et al., Impaired regenerative response of primary sensory neurons in ZPK/DLK gene-­trap mice. Biochem Biophys Res Commun, 2009. 383(2): p. 258-­‐62. Ayaz, D., et al., Axonal injury and regeneration in the adult brain of Drosophila. J Neurosci, 2008. 28(23): p. 6010-­‐21. Adachi-­‐Yamada, T., et al., p38 mitogen-­activated protein kinase can be involved in transforming growth factor beta superfamily signal transduction in Drosophila wing morphogenesis. Mol Cell Biol, 1999. 19(3): p. 2322-­‐9. Witte, H. and F. Bradke, The role of the cytoskeleton during neuronal polarization. Curr Opin Neurobiol, 2008. 18(5): p. 479-­‐87. Conde, C. and A. Caceres, Microtubule assembly, organization and dynamics in axons and dendrites. Nat Rev Neurosci, 2009. 10(5): p. 319-­‐32. Erez, H., et al., Formation of microtubule-­based traps controls the sorting and concentration of vesicles to restricted sites of regenerating neurons after axotomy. J Cell Biol, 2007. 176(4): p. 497-­‐507. Bogoyevitch, M.A. and B. Kobe, Uses for JNK: the many and varied substrates of the c-­Jun N-­terminal kinases. Microbiol Mol Biol Rev, 2006. 70(4): p. 1061-­‐95. Ringhoff, D.N. and L. Cassimeris, Stathmin regulates centrosomal nucleation of microtubules and tubulin dimer/polymer partitioning. Mol Biol Cell, 2009. 20(15): p. 3451-­‐8. 34 SUPPORTING DATA AND SUPPLEMENTAL MATERIALS Figure S1. Additional examples of cells that initiated tip extension from a respecified dendrite. Axons from these cells were severed as in Figure 3, but intermediate time points were not acquired. These examples illustrate the extensive growth that could be initiated by respecified dendrites, and also that these new axons can cross one another. 35 Figure S2. Neurons that initiate regeneration have a single process that switches to the axonal microtubule polarity. Neuronal shape and microtubule polarity were tracked for four days after axon severing using live imaging of EB1-­‐GFP. Movies of EB1-­‐GFP were acquired at 24, 48 and 96 hours (except for one cell which was only imaged at 24 and 96 hours). Low magnification images were acquired at each time point to map cell shape, and higher magnification movies of the region near the cell body were taken to determine microtubule orientation. If a dendrite extended significantly beyond its normal territory at 96 hr it was scored as having initiated tip growth. The direction of EB1-­‐GFP dot movement was scored manually. Data from 24 and 48 hours was averaged for classification of MT polarity in the figure. If 75% or more comets moved away from the cell body, microtubule polarity was classified as plus-­‐end-­‐out (axonal, green arrow). If 75% or more dots moved towards the cell body, microtubule polarity was classified as minus-­‐end-­‐out (dendritic, red arrow). Numbers in between were classified as mixed (purple double arrow). Microtubule orientation before cutting is not from this data set, it is based on uninjured cells. 36 Figure S3. Neurons that express hairpin RNAs to target msps are grossly normal. A. Stable microtubules in neurons were visualized by staining with the 22C10 antibody which recognizes a neuronal microtubule-­‐associated protein, futsch (Hummel et al., 2000). Stable microtubules are seen throughout the dendrites of ddaE (cell body marked with star) in control and msps RNAi neurons, although the futsch staining is reduced in intensity in the dendrite trunk with msps RNAi (right panels). B. An uninjured ddaE neuron expressing EB1-­‐GFP and a hairpin RNA to target msps was imaged over the same time course as for the axon removal experiments. The dendritic tree covers the same region of the body wall as normal, and remains stable over time, increasing in size as the body increases. The only difference in shape of the dendritic tree was an increase in proximal branches (arrow) in some animals. 37 Figure S4. This is a version of Figure 4 in which only the Apc2-­‐GFP channel is shown as greyscale so that the pattern of Apc2-­‐GFP fluorescence can be seen clearly by itself. Movie S01. EB1-­‐GFP dynamics in the same cell before and after axon severing. A ddaE cell expressing EB1-­‐GFP was imaged immediately before proximal axotomy. Immediately after this time series was acquired the axon was severed, and after severing the optics were immediately returned to the confocal mode and a new time series was acquired. The larva was then recovered to food and remounted for further imaging after 24 hours. http://www.molbiolcell.org/content/21/5/767/suppl/DC1 38 Movie S02. The number of growing microtubules is up-­‐regulated by axon, but not dendrite, severing. Time series of EB1-­‐GFP in the ddaE neuron 24 hours after axon or dendrite severing. Images from some of these movies are shown in Figure 1B-­‐ scale bars can be found there. http://www.molbiolcell.org/content/21/5/767/suppl/DC1
Movie S03. Microtubule orientation switches as a dendrite is respecified into an axon. EB1-­‐GFP movies of the same ddaE neuron before axon severing and 24, 48 and 72 hours after axon severing. The exact orientation of the cell is slightly different at the different time points. Dendrite 2 is respecified to an axon, it is at the right or lower right in all movies, and direction of EB1-­‐GFP movements in this dendrite is indicated with arrows during the movie. This is a movie of the cell shown in Figure 2A. http://www.molbiolcell.org/content/21/5/767/suppl/DC1 Movie S04. EB1-­‐GFP comets are extremely rare in neurons expressing a hairpin RNA that targets msps. A ddaE neuron expressing EB1-­‐GFP, dicer2 and a hairpin RNA targeting msps is shown. No distinct comets are visible. For comparison see Movie 5. http://www.molbiolcell.org/content/21/5/767/suppl/DC1 Movie S05. Microtubule dynamics in animals expressing a control (rtnl2) hairpin RNA. Movies of the ddaE neurons were acquired in larvae expressing EB1-­‐GFP, dicer2 and a control RNA hairpin (Rtnl2) that does not show any neuronal phenotypes. EB1-­‐GFP comets are seen moving towards the cell body in the main trunk of the dendrites. http://www.molbiolcell.org/content/21/5/767/suppl/DC1 39 Chapter Three Dendrite Injury Triggers DLK-­Independent Regeneration Michelle C. Stone, Richard M. Albertson, Li Chen, Melissa M. Rolls This chapter was published in Cell Reports 2014, Vol. 6, Issue 2 Author Contributions Michelle C. Stone performed dendriotomy experiments in class I and class IV neurons and quantitated regrowth by branching complexity or coverage. Michelle C. Stone also tracked microtubule polarity in regenerated dendrites, performed all dendrite injury experiments in adult flies, tested components of the DLK pathway, performed puc experiments after axon or dendrite removal, and dendrite removal experiments in 2 day vs. 4 day larvae. Richard M. Albertson performed the dynein heavy chain RNAi and highwire RNAi experiments, APC2 experiments, and performed dendriotomies further from the cell body. Li Chen developed the puc reporter assay used in the paper. Melissa M. Rolls supervised all experiments and was involved in data interpretation and the writing process. 40 SUMMARY Axon injury triggers regeneration through activation of a conserved kinase cascade that includes the dual leucine zipper kinase (DLK). While dendrites are damaged during stroke, traumatic brain injury and seizure, it is not known whether mature neurons monitor dendrite injury and initiate regeneration. We probed the response to dendrite damage using model Drosophila neurons. Two larval neuron types regrew dendrites in distinct ways after all dendrites were removed. Dendrite regeneration was also triggered by injury in adults. We next tested whether dendrite injury was initiated with the same machinery as axon injury. Surprisingly, DLK, JNK and fos were dispensable for dendrite regeneration. Moreover, this MAP kinase pathway was not activated by injury to dendrites. Thus neurons respond to dendrite damage and initiate regeneration without using the conserved DLK cascade that triggers axon regeneration. INTRODUCTION Most neurons need both dendrites and axons to function. The responses of neurons to axon injury are relatively well documented. After severe axon damage, distal regions of axons are cleared by Wallerian degeneration [1, 2], which is followed by reinitiation of axon outgrowth. In mammals, both degeneration and regeneration are more efficient in peripheral than central neurons [3-­‐5]. Axon injury triggers a cascade of signals that travel back from the injury site to the cell body. Within minutes of the initial trauma, a calcium wave travels back to the cell body [6, 7]. Subsequently, microtubule motor-­‐based transport brings signaling molecules between the site of injury and the cell body [8, 9]. This process may take hours to days depending on the distance from the site of injury to the cell body. Several key proteins take this motor-­‐based route. Among these, the mitogen-­‐
activated protein kinase kinase kinase (MAPKKK) dual leucine zipper kinase (DLK) has been shown to be required for efficient regeneration in worms, flies and mammals [10-­‐13], and thus seems to be a core signaling element in the response to 41 axon injury. A major transcriptional response is mounted downstream of DLK to allow reinitiation of axon growth from the damaged cell. While key players in axon regeneration have been identified, it is not known whether neurons have signaling machinery that senses dendrite damage, or if mature neurons can regenerate dendrites. Dendrites are particularly susceptible to damage by excitotoxicity and other environmental changes during stroke, seizure and traumatic brain injury [14-­‐18]. It is difficult to track dendrites from individual cells in mammals over time, so it has not been possible to determine whether dendrite regeneration is possible after damage using these models. Therefore several labs have turned to Drosophila dendritic arborization neurons to ask whether dendrite injury triggers regeneration. Dendritic arborization (da) neurons in Drosophila larvae are highly polarized cells with dendrites that innervate the body wall and axons that extend to the CNS [19]. While their dendrites are sensory, they share many features with dendrites that house postsynaptic sites. For example, minus-­‐end-­‐out microtubules are present in dendrites, but not axons, of all types of mammalian and Drosophila neurons examined, including da neurons [20, 21]. Different types of da neurons exist in Drosophila larvae, and they can be distinguished by arbor complexity and tiling behavior [19]. Several studies have suggested that the most complex of these neurons, the class IV cells, have some capacity to respond to dendrite damage, although this becomes more limited as larvae age [22, 23]. These studies also agree that the simplest da neurons, the class I cells, are unable to reinitiate growth in response to dendrite injury at any point during larval life, and can only do this during embryogenesis. Loss of sensory endings in zebrafish skin also triggers robust reinnervation only very early in development [24]. Thus the evidence so far suggests that, even in the peripheral nervous system where regeneration tends to be most exuberant, only some cells early in development can upregulate dendrite growth in response to injury. However, all studies on responses of single cells to dendrite injury have damaged only one dendrite. To fully test the idea that only subsets of neurons at specific developmental stages can initiate dendrite regeneration, we developed methods to 42 remove all dendrites from neurons and reasoned that the cells would either die or regenerate. RESULTS Dendrite injury triggers robust regeneration in multiple types of dendritic arborization neurons. To test the idea that only subsets of neurons at specific developmental stages can initiate dendrite regeneration, we developed methods to remove all dendrites from da neurons and reasoned that the cells would either die or regenerate. The simplest da neurons, including the ddaE cell, normally have very stable dendrite arbors during larval life and do not add or remove branches (Figure 1A, 1D and [22]). We thus tested whether these neurons would die if all dendrites were removed. We severed all dendrites from larval ddaE neurons in whole animals with a pulsed UV laser. After dendrite removal, larvae were returned to food. We imaged the cells at 5h to make sure dendrites were completely severed, and then on subsequent days to determine if the cells would die. At 24h after injury, the cells were still alive. Moreover in most cases new processes could be seen to emerge from the cell body. By 48h a branched dendrite arbor was visible in almost half (16/34) of the animals tested (Figure 1B and S1A). The remaining neurons grew fine processes alongside the axon (Figure S1B). We speculate that in neurons with peri-­‐axonal growth, the new processes could not reach the layer in which they normally arborize, which is between the epidermal cells and basement membrane [25]; in many of them the cell body was seen to float deeper into the animal after the dendrites were cut. The remaining analysis will focus only on the cells that regrew processes into the region normally occupied by dendrites rather than along the axon. At 96h after dendrite injury, the area of the body wall covered by the new processes was smaller than in cells without injury. However, when we counted dendrite branch points, we found that the complexity had reached a similar level to that before injury (Figure 1E). These neurons, which are part of a circuit that is responsible for coordinated motion [26], and do not normally tile the entire body wall [19] perhaps rely more on complexity of the arbor rather than exact area of 43 coverage to function. We conclude that the ddaE neuron can regenerate a branched arbor after dendrite removal, and that it regrows to the same branching complexity as uninjured neurons. Figure 1: Dendrites regrow completely after they are removed from larval neurons. A. An example of an uninjured ddaE neuron expressing mCD8-­‐GFP and tracked during four days of larval life is shown. B. A ddaE neuron expressing mCD8-­‐GFP had all dendrites removed at their base (arrows) and was then tracked for four days. At 5h the cut dendrites have degenerated. C. A ddaC neuron had its dendrites removed (arrows) and was then imaged over four days. 4h after dendrite severing degenerating remnants are seen. At 48h new dendrites do not reach the edge of the normal territory (double arrows). By 96h the new dendrites directly abut neighboring cells. D and E. Quantitation of branch point number in uninjured (D) ddaE neurons, and ddaE neurons after dendrite removal (E). Averages from 9 cells are shown in D and 7 (5h and 96h) or 8 (0h and 48h) cells in E. related to S1 To test whether other cell types could respond to dendrite damage by growing new processes, we performed dendrite removal on the class IV neuron ddaC. Class IV neurons tile the entire body wall [19] and receive information about painful stimuli [27]. After dendrite removal, new processes initiated growth by 24h. At 48h after injury an elaborate arbor covered the central part of the ddaC territory 44 (Figure 1C). By 96h after injury the entire territory was covered (Figure 1C). Out of 27 neurons tested all except 2 followed this pattern (pooled data from all genotypes used, Figure S1C). Thus, unlike the ddaE neuron, the ddaC neuron regenerated to cover its complete territory. To determine whether the regrown processes were correctly specified as dendrites, we analyzed microtubule polarity in the new neurites that emerged after dendrite removal. The presence of minus-­‐end-­‐out microtubules distinguishes dendrites from axons in Drosophila, C. elegans and mammalian neurons [21, 28, 29], and is a good indicator of compartment identity. The new processes that formed after dendrite removal in the ddaC neuron initially had slightly mixed polarity. By 48h after injury, this had resolved to about 90% minus-­‐end-­‐out microtubules, similar to uninjured ddaC neurons (Figure S1D). This result suggests that the new processes are dendrites, and is in sharp contrast to dendrite growth initiated by damage to the proximal axon. In both mammals [30] and flies [31] proximal axotomy causes a new axon to grow from a dendrite. However, this new axon is easily distinguished from the processes that regrow after dendrite injury; it leaves the normal territory of the dendrites, is unbranched, and has plus-­‐end-­‐out microtubule polarity [31]. To further test the identity of the regrown processes, we used two additional methods. First, we used a marker, Apc2-­‐GFP, that localizes to dendrite branch points, the proximal axon, but not more distal regions of the axon [31, 32]. This marker was expressed in ddaC neurons, and analyzed after dendrite removal. Both 48h and 96h after dendrite removal, it could be seen to occupy branch points in regrown processes, again suggesting these are specified as dendrites (Figure S1E). Second, we tested the effect of dynein on dendrite regrowth. Dynein is known to be required for dendrite development, and ddaC neurons in which it is reduced generate short, bushy arbors [33, 34]. In fact, because of the arrangement of microtubules in Drosophila neurons, it is predicted to be required for the bulk of transport from the cell body into dendrites [35]. Consistent with a role for dynein in transporting building blocks for dendrites, but not axons, axonal regeneration proceeded normally when dynein was reduced by RNAi (Figure S1F). Dendrite regrowth after removal initiated normally, but dendrite arbors were small 45 and bushy (Figure S1F), recapitulating the phenotype observed during dendrite development [33, 34]. Thus several different lines of evidence suggest that the processes that regrow after dendrite removal are correctly specified as dendrites. As previous studies have suggested that larval neurons have limited capacity to regenerate dendrites after a single dendritic injury [22, 23, 31], we removed single dendrites from the ddaE and ddaC neurons. In analyzing the results we kept in mind that after total dendrite removal the ddaE neuron regenerated a dendrite arbor with similar complexity, but not coverage, to control arbors, and the ddaC neuron regenerated to cover the body wall. We therefore analyzed number of branch points for ddaE and visually scored coverage for ddaC (Figure S2A and B). Using this perspective, we found that removal of a single dendrite triggered a similar response in both cells to removal of all dendrites, that is the ddaC neuron regrew until its territory was covered, and the ddaE neuron added branch points to its remaining dendrite until complexity was regained. We also tested whether the position of the injury might affect the outcome and severed some dendrites further from the cell body. Again, we observed robust regrowth (Figure S2C). Late larval and adult neurons retain the capacity to regrow dendrites. Previous studies suggested that the capacity of neurons to respond to dendrite damage decreases during larval life [22, 23]. We therefore aged larvae longer than in the published studies and tested whether neurons could still respond to dendrite injury. We saw no difference in regenerative capacity between young and old larvae (Figure S2D). We conclude that dendrite injury triggers a robust regenerative growth response in larval Drosophila neurons irrespective of age, branching complexity, or whether one or all dendrites are removed. The dendrite arbors of ddaE and ddaC function during larval life and so are, at least in this sense, mature. During pupariation, dendrites from both neurons are pruned and new arbors are regrown into the adult body wall. ddaC and other class 46 Figure 2. Adult neurons reinitiate dendrite outgrowth after dendrite injury. The major dendrite root in the anterior v’ada neuron in the adult abdomen was severed with a laser at 0h (except in top row). Flies were then returned to food vials and aged for 2, 4 or 5 days. They were then remounted and the same neuron was imaged. In all cases the cell remained alive and new dendrites had sprouted. 47 IV neurons, including v’ada, persist through adult life, while ddaE undergoes apoptosis 3-­‐5 days after adults emerge [36]. To address the possibility that larval neurons have some dendrite growth potential not present in adult neurons, we tested whether dendrite removal in adults could also trigger regenerative growth. We developed methods to mount living young adult flies so that we could sever dendrites in intact animals. We chose to use the most anterior v’ada neuron for these experiments as it could be visualized in the abdomen using the ppk-­‐Gal4 driver with relatively little overlap from dendrites of other neurons. As in larval neurons, we severed a major dendrite near its base and determined whether the cell would undergo apoptosis or would reinitiate dendrite outgrowth. We could not repeatedly remount the same adult for imaging, therefore we analyzed different cells at various times after injury. At 48h after dendrite removal 5/6 neurons had new processes sprouting near the cell body (Figure 2). At 96h or 120h after injury 12/13 neurons had regrown complex, branched arbors into their normal territory (Figure 2). Thus adult neurons seem to retain the capacity to respond to dendrite injury and to regenerate dendrites. The conserved DLK axon regeneration pathway is not involved in the response to dendrite injury. In C. elegans, Drosophila and mammals, efficient axon regeneration requires signaling through the dual leucine zipper kinase (DLK, or Wallenda (wnd) in flies) pathway [10-­‐13]. DLK is a mitogen-­‐activated protein kinase kinase kinase (MAPKKK) that initiates a signal transduction cascade after axon injury that includes cJun N-­‐terminal kinase (JNK) in flies [10], and both JNK and the related MAPK, p38, in C. elegans [37]. In Drosophila, the AP-­‐1 transcription factor fos is required for the transcriptional response to axon injury downstream of DLK [10], and in mammals accumulation of the AP-­‐1 transcription factor jun in the nucleus after axon injury requires DLK [12]. Thus DLK is believed to be at the core of a conserved signaling cascade that initiates the cell body response to axon injury. To determine whether dendrite regeneration requires this conserved injury signaling pathway, we used RNAi and mutant approaches to lower levels of DLK and 48 assayed dendrite regeneration after complete dendrite removal in ddaC. As a control for the effectiveness of the RNAi in the ddaC neuron, we assayed axon regeneration. In control ddaC neurons, 7/7 axons initiated regeneration from the axon stump (Figure 3A and D). In contrast, only 1/8 of the wnd (DLK) RNAi neurons or 1/9 of the wnd1/wnd3 loss of function [38] mutants showed any sign of axon outgrowth (Figure 3B and D). Figure 3. Wnd is not required for dendrite regeneration. Axons (A and B) or dendrites (C) of ddaC neurons were severed at 0h. Larvae were remounted for imaging 4-­‐5 hours later to check for complete severing and then at 48h and 96h to assay regeneration. Quantitation is shown in D. For axon injury, cells that were scored as regrowing extended axon stumps as shown in A, and cells that were scored as not regrowing retained a clear stump as in B. For dendrite injury, regrowing cells all had robust dendrite arbors as in C. 49 Thus wnd reduction effectively blocked axon injury signaling in the ddaC neuron. We then tested dendrite regeneration in the same genetic backgrounds. The ddaC neurons with reduced wnd regrew dendrite arbors with similar efficiency and timing to control neurons (Figure 3C and D and S3A). We also tested whether wnd was required for dendrite regeneration in ddaE neurons, and again found that dendrite regeneration was robust when wnd was targeted by RNAi (Figure S3B). This result suggests DLK signaling is not required for dendrite regeneration. In the response to axon injury, DLK levels increase due to reduction in turnover by the ubiquitin ligase highwire (hiw) [10]. Hiw also seems to regulate DLK during dendrite development [39]. However, like DLK, reduction of hiw impaired the response to axon injury, but not dendrite injury (Figure S3D). In Drosophila, downstream components of the DLK pathway include the kinase JNK (called bsk in flies), and the AP-­‐1 transcription factor fos. Dominant negative forms of JNK (bskDN) or fos (fosDN) block the response to axon injury in motor neurons [10] and block axon regeneration in dendritic arborization neurons ([31] and not shown). We therefore expressed both of these transgenes in the ddaC and ddaE neurons. In both genetic backgrounds in both cell types cases, dendrites regrew in the majority of neurons (Figure 4). Thus downstream components of the axon injury signaling pathway do not seem to be required for dendrite regeneration. It is possible that although the DLK/JNK/fos pathway is not required for dendrite regeneration, it could be activated in parallel with another pathway that can compensate for its loss during the injury response. We therefore wished to use a reporter for pathway activation. JNK signaling is regulated in a negative feedback loop by MAP kinase phosphatases (MKPs) that are transcribed in response to activity of the pathway [40]. After motor axon injury, levels of the MKP puckered (puc) can be seen to increase when monitored with a puc-­‐bgal reporter [10]. We wished to use a puc reporter that could be visualized in whole, living animals, so we tested whether a puc-­‐GFP protein trap [41] that reports on JNK signaling in embryos [42] would increase in nuclear fluorescence after axon injury in the ddaE neuron. In uninjured ddaE neurons, low levels of puc-­‐GFP fluorescence were seen in the nucleus. We used ddaE for this experiment as a similar class I da neuron, 50 ddaD, lies next to it and can be used for comparison (Figure S4). By 24h after axon injury the amount of nuclear fluorescence in ddaE compared to its uninjured ddaD neighbor had increased about four-­‐fold (Figure S4). We therefore used this as a reporter for JNK signaling after dendrite injury. When we tracked puc-­‐GFP levels in ddaE neurons after dendrite removal, no increase in fluorescence was observed (Figure S4). We conclude that the DLK/JNK pathway is neither required for the response to dendrite injury nor activated by it. Thus dendrite injury triggers regenerative growth without using the conserved axon injury signal transduction cascade. Figure 4. Downstream components of the DLK pathway are dispensable for dendrite regeneration. Examples of ddaC (A) and ddaE (B) neurons expressing bskDN and fosDN are shown at different times after dendrite removal. Arrows indicate sites of severing at 0h. C. ddaC cells were scored as regrowing dendrite arbors or not. All cells that regrew had robust arbors similar to those shown in A. D. Quantitation of branch point number in ddaE neurons after dendrite removal was performed as in Figure 1. The error bars show the standard deviation. The n’s are numbers of cells analyzed for each genotype at each timepoint following severing and are shown on the bars in the graph. 51 DISCUSSION It has not previously been clear whether mature neurons have a surveillance mechanism that allows them to respond to dendrite injury. We demonstrate that dendrite removal triggers robust reinitiation of dendrite outgrowth in multiple neuron types, even after development of dendrite shape is complete. Unlike previous studies that have examined the response to removal of single dendrites, we find that dendrite regeneration is not restricted to a specific developmental window, or to neurons with complex dendrite arbors. Instead our data suggests that dendrite regeneration may be a property of many neurons throughout life. It is also interesting to note that different cell types seem to complete dendrite regrowth in distinct ways. Not surprisingly, neurons that respond to pain regrew dendrites until their normal territory was covered. Intriguingly, neurons responsible for proprioception regrew dendrites until they had regained the normal number of dendrite branch points. This difference in how regeneration is completed can account for some of the differences between our conclusions and those made in previous studies. Area of coverage was previously assayed as a key aspect of all types of dendrite regeneration [22, 23], and so the response of ddaE to removal of a single dendrite was not noted even though an increase in complexity of the remaining dendrite is clear in the published data [23]. By performing complete dendrite removal, we were able to get much more striking results to demonstrate the ability of multiple cell types to respond to dendrite injury. Our data suggest that neurons possess at least two different injury surveillance systems: (1) signaling through DLK informs the cell body of axon injury, and (2) an independent signal is responsible for informing the cell body of dendrite injury. A previous study suggested that the Akt kinase may be a shared component of axon and dendrite regeneration [23]. Perhaps Akt is generally required to promote cell growth downstream of specific regulators that inform the cell of axon or dendrite injury. If the DLK pathway does not signal dendrite injury, what type of surveillance machinery might monitor dendrites? It seems likely that transcription changes are 52 required to allow growth of a new dendrite arbor. Perhaps a signal is generated at the site of injury and transported back to the cell body in the same way that has been proposed for axons [8, 9]. In this case, however, the retrograde motor should be a kinesin rather than dynein since microtubules have minus-­‐end-­‐out orientation in Drosophila dendrites [21]. Alternately, a change in electrical signals arising from dendrites could directly regulate transcription factors in the cell body. It will be very interesting to explore different types of injury signals in the future. The ability of mature neurons to initiate new growth in response to large-­‐
scale dendrite damage could play an important role in recovery from stroke, seizure and TBI. Considerable neurite sprouting occurs during recovery from these conditions [43] and so it is possible that both axon and dendrite outgrowth are triggered. If so, it seems likely that they are triggered through separate signaling pathways as DLK is required for injury-­‐induced axon outgrowth, but not injury-­‐
induced dendrite outgrowth. By establishing a tractable genetic model for dendrite regeneration, we should be able to identify key players in this process. Having a molecular handle on this uncharacterized type of regeneration will allow us to probe its importance in mammalian models of nervous system trauma. EXPERIMENTAL PROCEDURES Labeling Identified Neurons in Drosophila Larvae and Adults For most experiments, UAS-­‐mCD8-­‐GFP was expressed in neuronal subsets to identify specific neurons in whole animals. UAS-­‐EB1-­‐GFP was used to analyze microtubule polarity and dynamics. GFP markers were expressed in class I neurons with 221-­‐Gal4 and class IV neurons with either 477-­‐Gal4 (larvae) or ppk-­‐Gal4 in adults. For more detailed information about genotypes, please see the Supplemental Experimental Procedures. Axon and Dendrite Injury In both larvae and adults, individual axons or dendrites were severed at their base with a pulsed UV laser (Micropoint, Andor Technology). In all cases, whole living animals were mounted for laser surgery and were recovered to 53 normal growth conditions immediately after severing. Animals were remounted for imaging at later time points. Please see the Supplemental Experimental Procedures for more details about imaging conditions. SUPPLEMENTAL INFORMATION Supplemental Information includes Supplemental Experimental Procedures and four figures and can be found with this article online at http://dx.doi.org/ 10.1016/j.celrep.2013.12.022. ACKNOWLEDGMENTS We are very grateful to the Bloomington Drosophila Stock Center, the Vienna Drosophila RNAi Center and FlyTrap; all are invaluable resources. Catherine Collins, Wesley Grueber and Dirk Bohmann generously provided us with Drosophila stocks. Tadashi Uemura provided helpful information for imaging da neurons in adults. Michelle Nguyen kindly helped collect flies and embryos. This work was funded by the NIH (R01 GM085115) and MMR was a Pew Scholar in the Biomedical Sciences. REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. Wang, J.T., Medress, Z.A., and Barres, B.A. (2012). Axon degeneration: molecular mechanisms of a self-­‐destruction pathway. J Cell Biol 196, 7-­‐18. Coleman, M.P., and Freeman, M.R. (2010). Wallerian degeneration, wld(s), and nmnat. Annu Rev Neurosci 33, 245-­‐267. Vargas, M.E., and Barres, B.A. (2007). Why is Wallerian degeneration in the CNS so slow? Annu Rev Neurosci 30, 153-­‐179. Huebner, E.A., and Strittmatter, S.M. (2009). Axon regeneration in the peripheral and central nervous systems. Results Probl Cell Differ 48, 339-­‐
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69. Wang, X., Kim, J.H., Bazzi, M., Robinson, S., Collins, C.A., and Ye, B. (2013). Bimodal control of dendritic and axonal growth by the dual leucine zipper kinase pathway. PLoS Biol 11, e1001572. Caunt, C.J., and Keyse, S.M. (2012). Dual-­‐specificity MAP kinase phosphatases (MKPs): Shaping the outcome of MAP kinase signalling. The FEBS journal. Morin, X., Daneman, R., Zavortink, M., and Chia, W. (2001). A protein trap strategy to detect GFP-­‐tagged proteins expressed from their endogenous loci in Drosophila. Proc Natl Acad Sci U S A 98, 15050-­‐15055. Taniguchi, K., Hozumi, S., Maeda, R., Ooike, M., Sasamura, T., Aigaki, T., and Matsuno, K. (2007). D-­‐JNK signaling in visceral muscle cells controls the laterality of the Drosophila gut. Dev Biol 311, 251-­‐263. Nudo, R.J. (2006). Mechanisms for recovery of motor function following cortical damage. Curr Opin Neurobiol 16, 638-­‐644. Stone, M.C., Rao, K., Gheres, K.W., Kim, S., Tao, J., La Rochelle, C., Folker, C.T., Sherwood, N.T., and Rolls, M.M. (2012). Normal Spastin Gene Dosage Is Specifically Required for Axon Regeneration. Cell reports. 57 SUPPLEMENTAL INFORMATION Supplemental Figures Figure S1 Figure S1. Dendrite regeneration in ddaE and ddaC neurons. (A) An example of a ddaE neuron 96h after axon severing that regrew new processes (arrows) below the region of the body wall in which dendrites normally reside. These processes are branched like dendrites, and so we suggest they are dendrites that did not gain access to the plane between the epidermal cells and basement membrane. (B) A ddaE neuron expressing EB1-­‐GFP had dendrites severed at the roots at 0h. The larva was remounted for imaging at 5 h and then each subsequent day for 4 days. As with ddaE neurons labeled with mCD8-­‐GFP, the neuron regrew a branched dendrite arbor. (C) Similar numbers of cells regrow dendrites (n=number of cells) whether they are labeled with mCD8-­‐GFP or EB1-­‐GFP, however fine tips of dendrites are not labeled well with EB1-­‐GFP. (D) Microtubule polarity was assayed by tracking EB1-­‐GFP comets in either uncut ddaC neurons, or ddaC neurons at different 58 times during dendrite regrowth. In D, n’s are number of EB1-­‐GFP comets analyzed. (E) Apc2-­‐GFP to mark dendrite branch points and mCD8-­‐RFP to mark cell outlines were expressed in ddaC neurons with 477-­‐Gal4. Green arrowheads indicate Apc2-­‐GFP punctae in dendrites. Dendriotomy was performed at 0h (arrow). The cell was checked at 4h to ensure complete cutting, and regrowth was analyzed at 48h and 96h. (F) Axon and dendrite regeneration were assayed in ddaC neurons expressing mCD8-­‐GFP, Dicer2 and Dhc64C (dynein heavy chain) RNA hairpins. In this background axon regeneration was generally initiated by 48h after injury, as in the left panels, and 5 out of 6 regenerated by 96h. Dendrite regeneration was always initiated (n=33), although sometimes dendrites grew from the initial region of the axon. The new dendrite arbor was short and highly branched near the cell body, consistent with reported dynein phenotypes. Error bars represent SD. 59 Figure S2 Figure S2. Neurons respond to injury of single dendrites, and regenerate when injury is further from the cell body, and regenerate throughout larval life, related to Figure 1. Single dendrite roots in either ddaE (A) or ddaC (B) neurons were severed at time 0h (arrows). (A) For ddaE neurons the dorsal comb-­‐like dendrite was removed in neurons with only one other dendrite 60 emerging from the cell body, which we call the #2 dendrite. The number of branch points along the #2 dendrite was counted in uninjured and injured neurons (right). 9 uninjured and 19 injured neurons were analyzed. The number of branch points along the #2 dendrite increased dramatically after removal of the comb dendrite (arrowheads indicate examples of new branch points), but not over the same time period in control neurons. (B) An example of a ddaC neuron with a single dendrite cut (arrow) is shown. Over time, the area of the body wall left open by degeneration of this dendrite (see 5h image) was covered by dendrite branches growing from the stump or neighboring dendrites. The inset graph summarizes the results for 6 neurons analyzed. All filled in the area left open by dendrite removal. (C) In most of our experiments dendrites were severed very close to the cell body so that the minimum number of laser surgeries per animal was performed. An example of an animal in which cuts were made further from the cell body is shown here. This cell, and similar ones, responded much the same as when cuts were closer to the cell body. The only noticeable pattern was that if a longer dendrite stump was left then regeneration tended to initiate from this. Cut sites are indicated with arrowheads. (D) In most of our experiments larvae were aged for two days after an overnight embryo collection. When larvae were aged for 4 days instead of 2, robust dendrite regrowth was also observed (images to left), and similar numbers of neurons regrew dendrites at both timepoints (right panel, n= number of neurons analyzed). 61 Figure S3 Figure S3. Wnd and hiw RNAi do not block dendrite regeneration, related to Figure 3. (A and B) Examples of ddaC (A) and ddaE (B) neurons expressing Dicer2 and wnd RNAi hairpins are shown at different times after dendrite removal. Arrows indicate sites of severing for ddaE. Graphs in Figure 4 include the wnd RNAi data. (C and D) ddaC neurons expressing hiw RNAi hairpins were subjected to either axon removal (C) or dendrite removal (D). About half of the axons did not initiate regeneration by 96h after injury as in the example. After dendrite injury, dendrites regrew in all cases. The graph summarizes axon and dendrite data, numbers on the bars are the number of animals in each condition. 62 Figure S4 Figure S4. A target of the DLK/JNK/fos pathway is not activated by dendrite injury, related to Figure 4. ddaE neurons in animals expressing puc-­‐GFP and UAS-­‐mCD8-­‐RFP were subjected to axon or dendrite removal. Larvae were remounted for imaging and 6 and 24h. The intensity of puc-­‐GFP fluorescence in nuclei of the injured ddaE neuron (labeled E) and its neighboring ddaD neuron (labeled D) was measured, and ratios of the fluorescence in ddaE/ddaD are shown in the graph. Arrows indicated the sites of severing in 0h images and new dendrite growth in the lower 24h image. Error bars represent SD. Supplemental Experimental Procedures Drosophila stocks The tester lines for dendrite injury experiments in class I neurons were (1) 221-­‐
Gal4, (2) 221-­‐Gal4, UAS-­‐mCD8 GFP and (3) UAS-­‐dicer2; 221-­‐Gal4, UAS-­‐EB1-­‐
GFP/TM6. The tester lines for dendrite injury experiments in class IV neurons were (1) 477-­‐Gal4, UAS-­‐EB1-­‐GFP/CyO, (2) 477-­‐Gal4, UAS-­‐mCD8 GFP and (3) 477 Gal4, UAS-­‐EB1-­‐GFP/CyO; UAS-­‐dicer2/TM6. Tester lines were crossed to UAS-­‐EB1-­‐GFP or YW flies. Adult dendrite injury experiments were done using homozygous ppk-­‐ Gal4, UAS-­‐mCD8-­‐GFP flies. For the DLK RNAi experiment, the tester line 477-­‐Gal4, UAS-­‐mCD8-­‐GFP; UAS-­‐dicer2 was crossed to the UAS-­‐wallenda RNAi line (identification number 103410) from the Vienna Drosophila RNAi Center (VDRC, Vienna, Austria). To test the involvement of downstream components of the DLK/JNK pathway, dominant negative bsk and fos were used. UAS-­‐bskDN was obtained from the Bloomington Drosophila Stock Center (Bloomington, IN) and UAS-­‐
63 fosDN was obtained from Catherine Collins (University of Michigan, Ann Arbor, MI). These lines were crossed to the tester line 477-­‐Gal4, UAS-­‐EB1-­‐GFP/CyO and assayed for dendrite regeneration. Other RNAi lines used for similar experiments were also from VDRC: hiw (#26998) and Dhc64C (#28054). DLK mutants were previously described, and a line was constructed to pair wnd1 with 477-­‐Gal4, UAS-­‐mCD8-­‐GFP. This line was then crossed to a line with the wnd3 mutation. To test for activation of the DLK pathway, a puc-­‐GFP protein trap was used as a reporter. This line, G00462, was obtained from the FlyTrap Project (http://flytrap.med.yale.edu/) and rebalanced with TM6. The rebalanced line was crossed to the tester line UAS-­‐
mCD8-­‐RFP; 221-­‐Gal4. Larval Dendrite and Axon Injury Experiments Fly embryos were collected for 24 hours on apple caps with yeast paste at 20°C. Embryos were then transferred to vials of standard Drosophila food. They were then aged for 2 days at 25°C, or 4 days for the experiment in Figure S3. Whole larvae were mounted on a microscope slide coated with a dry agarose pad to aid in immobilization of the larvae. Dendrites of the class I neuron (ddaE) or the class IV neuron (ddaC) were severed close to the cell body using a Micropoint UV laser (Photonics Instruments). For injury experiments where only one dendrite was severed, one dendrite of ddaE or the comb dendrite of the ddaE was severed close to the cell body. After severing, the neuron was imaged immediately to obtain an overview picture and then the larva was returned to food and kept at 20°C until imaging at later timepoints. Imaging was performed using an LSM510 confocal microscope (Zeiss) at a frame rate of 2 seconds. Dendrite regeneration in the class I neuron was quantitated by counting all visible branchpoints in the dendrite arbor. Dendrite regeneration in the class IV neuron was qualitatively scored by determining whether the dendrite arbor reached the edge of its territory. Distal axotomy experiments were performed using the same methods as dendrite severing, except instead of injuring the base of the dendrites the axon was at least 50μm from the cell body as previously described (Stone et al., 2012). 64 Analysis of Microtubule Polarity Microtubule polarity was assayed in emerging neurites after dendrite injury by tracking the movement of individual EB1-­‐GFP comets. Since EB1-­‐GFP only binds to the growing plus end of the microtubule, tracking the direction of comet movement is a representation of microtubule polarity. Images of EB1-­‐GFP were acquired every 2 seconds with an LSM 510 confocal microscope (Zeiss). Analysis was done using ImageJ software and comets that were visible in at least three consecutive frames were scored as moving towards or away from the cell body. Adult Dendrite Injury Experiments Adult flies were collected just after eclosion and anesthetized on a CO2 pad. They were allowed to recover briefly before being placed laterally on a microscope slide with an agarose pad. A drop of 100% glycerol was placed on a coverslip and then placed over the entire body of the fly. The coverslip was then secured by tape. The class IV v’ada neuron at the anterior of the abdomen was located and its dendrites were severed using a pulsed UV laser. The neuron was immediately imaged with an LSM 510 confocal microscope (Zeiss) followed by recovery of the fly from the microscope slide. Before placing the fly into a food vial, a kimwipe was used to blot the body and remove excess glycerol. The fly was then placed in a food vial with filter paper covering the top of the food to absorb any remaining glycerol. Flies were then imaged at later timepoints including 48h, 96h and 120h. Puc-­GFP Reporter for JNK Signaling Activation To analyze JNK pathway activation after dendrite or axon injury, the puc-­‐GFP protein trap line was crossed to UAS-­‐mCD8-­‐RFP; 221-­‐Gal4 and axons or dendrites of the class I ddaE neuron were severed close to the cell body. Images of the cell bodies of both class I neurons (ddaD and ddaE) were taken at 0h, 6h and 24h after severing using an LSM 510 confocal microscope (Zeiss). Images were analyzed using ImageJ software and a ratio of ddaE/ddaD nuclear fluorescence was calculated. 65 Stone, M.C., Rao, K., Gheres, K.W., Kim, S., Tao, J., La Rochelle, C., Folker, C.T., Sherwood, N.T., and Rolls, M.M. (2012). Normal Spastin Gene Dosage Is Specifically Required for Axon Regeneration. Cell reports. 66 Chapter Four Conclusions Microtubules and axon versus dendrite injury Healthy neurons are essential for an organism to carry out basic and complex life functions. However neurons are susceptible to many types of injury and diseases that can disrupt the flow of information within the nervous system. Axons and dendrites are the polar compartments of the neuron tasked with sending and receiving signals, respectively, and both compartments can be damaged. Therefore, it is imperative that neurons be able to repair themselves or regenerate following injury. This work demonstrates how Drosophila sensory neurons are a powerful system for studying responses to axon and dendrite injury at the cellular level and highlights microtubule polarity as a key indicator of axonal and dendritic identity. Following axon injury, there is a major rearrangement of the cytoskeleton and increase in microtubule dynamics. Microtubule polarity reverses in one of the dendrites, and it is this dendrite that initiates growth from its tip and becomes the new axon. All of these steps require the activation of the JNK pathway. However, it is not yet known what allows for the reversal of polarity in the dendrite that becomes an axon. This is a crucial step in the process of axon regeneration, because without this polarity reversal, regeneration fails. Our lab has hypothesized that microtubule severing proteins or increased nucleation could account for the addition of new plus end-­‐out microtubules needed to convert a dendrite into an axon. Microtubule severing is an ATP-­‐dependent process, which causes a break in the microtubule polymer, thus creating a new plus end. Microtubule severing proteins are members of the AAA protein family and include katanin, spastin, and fidgetin. Previous studies of microtubule severing proteins in Drosophila have revealed their importance in severing spindle microtubules during mitosis [1]. In neurons, spastin and katanin have been shown to be involved in axon branching [2] and spastin to be required for maintenance of the neuromuscular junction [3]. Spastin has also been shown to be important for outgrowth of the dendritic arbor in 67 class IV neurons [4]. A fourth severing protein, katanin 60L1, was shown to be required for dendrite pruning in pupal Drosophila sensory neurons [5] and for dendritic arbor stabilization in larval class IV neurons [6]. Nucleation is the process by which microtubule seeds are formed from gamma tubulin. This in turn creates more plus ends. Gamma tubulin is a highly conserved protein among eukaryotes and has recently been shown to be important for controlling microtubule polarity in Drosophila sensory neurons [7]. To test the possibility of microtubule severing or nucleation being responsible for polarity reversal after axon cutting, an RNAi screen was performed in class I neurons. The number of plus end-­‐out microtubules in dendrites was assayed at 96hr after axon removal (Figure 1). Figure 1. RNAi screen of microtubule severing proteins and gamma tubulin 23C. Loss of each individual severing protein or gamma tubulin 23C did not prevent the addition of plus end-­‐out microtubules to dendrites after axon cutting. These results however do not rule out the possibility that multiple severing proteins could be working together. This would need to be further investigated by either combinations of RNAi or mutants to knock out multiple severing proteins at once. It 68 is important to note that in a separate assay for regeneration, the spastin and katanin 60 RNAi lines cause a significant reduction in regeneration of an axon from a dendrite after axon injury [8]. However this failure to regenerate is not thought to be a result of the inability to add sufficient plus end-­‐out microtubules (as seen in Figure 1.), but rather at a step downstream of the rearrangement of the cytoskeleton. At this point in time, microtubule severing or nucleation do not seem to be responsible for the reversal of polarity required for regeneration of an axon from a dendrite. Further studies are needed to identify the key players involved in this process. The rearrangement of microtubules, especially following proximal axon injury, is critical to the survival of the neuron and its ability to regenerate a new axon. Dendrite injury does not provoke this same dramatic rearrangement of microtubules, however Drosophila sensory neurons are still capable of regenerating their dendrites. In addition, components of the DLK signaling pathway are not involved in dendrite regeneration like they are for axon regeneration. Thus, neurons possess two distinct mechanisms for responding to axon and dendrite injury and triggering regeneration (Figure 2). Being able to understand and identify the components of these mechanisms will be extremely important for our overall understanding of the potential for neurons to recover from neurodegenerative diseases and nervous system trauma. Figure 2. Axon and dendrite injury trigger different regeneration pathways. A. Axon injury induces activation of the DLK pathway. B. Dendrite injury does not trigger activation of the DLK pathway and it is currently unknown what molecular machinery is required for dendrite regeneration. 69 Microtubule polarity in sea anenome neurons Microtubule polarity underlies the identity of axons and dendrites. Key evidence for this comes from the fact that after proximal axon removal in Drosophila sensory neurons, a single dendrite completely flips its polarity in order to become the new axon. Recently, I have taken an evolutionary approach to identify the origin of polarized neurons by using the model organism Nematostella vectensis, or sea anenome. Polarization of compartments is evident among vertebrate neurons and is important for directional signaling and polarized trafficking. This polarization is also present in many neurons of the bilaterian invertebrates. We hypothesize that neuronal polarity arose before the divergence of bilaterians and cnidarians, suggesting a more ancient common ancestor for neuronal polarity. Sea anenomes belong to the cnidarian phylum and consist of an oral-­‐aboral axis and nerve net rather than a centralized nervous system. The sea anenome nerve net is comprised of multiple types of ganglion cells and sensory cells [9]. It is currently not known whether sea anenome neurons are polarized with distinct axonal and dendritic compartments. Using a Nematostella ortholog of EB1 tagged with GFP and a pan neuronal promotor, Elav1, microtubule polarity in the different types of ganglion cells can be examined. These cells were picked to track polarity due to their long extending neurites and the ability to easily identify their cell bodies. In order to obtain transgenic animals to image, sea anenome embryos are injected using a previously described meganuclease-­‐based method [10]. Roughly 10-­‐14 days after injection, embryos have developed into polyps, which can be screened for positive animals and then imaged using a confocal microscope. Ganglion cells that I have imaged reproducibly include cells that have two opposing neurites extending from the cell body, cells that have three neurites extending from the cell body, and cells with two neurites extending from the cell body, where one of the neurites is branched (Figure 3). 70 Figure 3. Ganglion cells imaged in sea anenome polyps. A: cell with two opposing neurites B: cell with three neurites and C: cell with one branched and one unbranched neurite Preliminary results suggest there are two types of microtubule arrangements in the neurites of these cells: 1) neurons that have neurites that are all plus end-­‐out, (Figure 4A) and 2) neurons with a neurite that is plus end-­‐out and a neurite with mixed polarity (Figure 4B). Evidence collected so far suggests that the presence of branching in a neurite coincides with a mixed polarity, however further experiments must be done to confirm this observation. Figure 4. Microtubule arrangements in neurites of different types of sea anenome ganglion cells. 71 Based on microtubule polarity alone, neurites with all plus end-­‐out microtubule polarity could be classified as axon-­‐like and those with a mixed polarity as dendrite-­‐like. In order to verify whether or not a neurite is truly an axon or dendrite, we propose to look at synaptic arrangements using fluorescently tagged presynaptic and postsynaptic markers (Table 1). The rationale for this approach comes from the fact that many axons are presynaptic and many dendrites are postsynaptic. Our hypothesis is that presynaptic markers would segregate to neurites with axon-­‐
like microtubule polarity and postsynaptic markers would segregate to neurites with dendrite-­‐like polarity in cells where both types of neurites exist, and provide evidence for the evolution of polarized neurons before cnidarians and bilaterians diverged (Figure 5A). In the cases where neurons have neurites that are all plus end-­‐out, it is possible that either 1) presynapses and postsynapes are distributed so they are spatially segregated in each neurite, 2) presynapses and postsynapses form bidirectional synapses in each neurite, or 3) presynapses and postsynapses are segregated to separate neurites (Figure 5B). 72 Figure 5. Possible synaptic arrangements in sea anenome neurons. A. Neuron with neurite with axon-­‐like microtubule polarity has presynapses and neurite with dendrite-­‐like microtubule polarity has postsynapses. B. Neuron with neurites that all have axon-­‐like microtubule polarity with presynapses and postsynapses segregated spatially in all neurites (left), presynapses and postsynapses form bidirectional synapses in all neurites (middle), presynapses and postsynapses segregated to separate neurites (right). Using combinations of the fluorescently tagged synaptic markers in Table 1, we should be able to map the synaptic arrangement in the sea anenome ganglion neurons. Results from these experiments combined with results from the microtubule polarity experiments will help us answer the question of when axons and dendrites arose and neurons became polarized. This work may also help us understand the evolutionary origin of the distinct regeneration responses exhibited by axons and dendrites following injury. ACKNOWLEDGEMENTS I would like to thank Timothy Jegla for training and guidance with the sea anenome project. Thanks also to Justin Sturkie, Emma Baker, Liana Trigg, Fortunay Diatta, Sophia Yang, and Aditya Pisupati for making constructs for injecting, helping with injections and general sea anenome maintenance. 73 REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. Zhang, D., et al., Three microtubule severing enzymes contribute to the "Pacman-­flux" machinery that moves chromosomes. J Cell Biol, 2007. 177(2): p. 231-­‐42. Yu, W., et al., The microtubule-­severing proteins spastin and katanin participate differently in the formation of axonal branches. Mol Biol Cell, 2008. 19(4): p. 1485-­‐98. Sherwood, N.T., et al., Drosophila spastin regulates synaptic microtubule networks and is required for normal motor function. PLoS Biol, 2004. 2(12): p. e429. Jinushi-­‐Nakao, S., et al., Knot/Collier and cut control different aspects of dendrite cytoskeleton and synergize to define final arbor shape. Neuron, 2007. 56(6): p. 963-­‐78. Lee, H.H., L.Y. Jan, and Y.N. Jan, Drosophila IKK-­related kinase Ik2 and Katanin p60-­like 1 regulate dendrite pruning of sensory neuron during metamorphosis. Proc Natl Acad Sci U S A, 2009. 106(15): p. 6363-­‐8. Stewart, A., et al., Katanin p60-­like1 promotes microtubule growth and terminal dendrite stability in the larval class IV sensory neurons of Drosophila. J Neurosci, 2012. 32(34): p. 11631-­‐42. Nguyen, M.M., et al., Gamma-­tubulin controls neuronal microtubule polarity independently of Golgi outposts. Mol Biol Cell, 2014. 25(13): p. 2039-­‐50. Stone, M.C., et al., Normal spastin gene dosage is specifically required for axon regeneration. Cell Rep, 2012. 2(5): p. 1340-­‐50. Marlow, H.Q., et al., Anatomy and development of the nervous system of Nematostella vectensis, an anthozoan cnidarian. Dev Neurobiol, 2009. 69(4): p. 235-­‐54. Layden, M.J., et al., Microinjection of mRNA or morpholinos for reverse genetic analysis in the starlet sea anemone, Nematostella vectensis. Nat Protoc, 2013. 8(5): p. 924-­‐34. 74