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Cellular and Molecular Life Sciences

, Volume 72, Issue 19, pp 3621–3635 | Cite as

The function of RNA-binding proteins at the synapse: implications for neurodegeneration

  • Chantelle F. SephtonEmail author
  • Gang Yu
Open Access
Review

Abstract

The loss of synapses is a central event in neurodegenerative diseases. Synaptic proteins are often associated with disease neuropathology, but their role in synaptic loss is not fully understood. Of the many processes involved in sustaining the integrity of synapses, local protein translation can directly impact synaptic formation, communication, and maintenance. RNA-binding proteins and their association with RNA granules serve to regulate mRNA transportation and translation at synapses and in turn regulate the synapse. Genetic mutations in RNA-binding proteins FUS and TDP-43 have been linked with causing neurodegenerative diseases: amyotrophic lateral sclerosis and frontotemporal dementia. The observation that mutations in FUS and TDP-43 coincide with changes in RNA granules provides evidence that dysfunction of RNA metabolism may underlie the mechanism of synaptic loss in these diseases. However, we do not know how mutations in RNA-binding proteins would affect RNA granule dynamics and local translation, or if these alterations would cause neurodegeneration. Further investigation into this area will lead to important insights into how disruption of RNA metabolism and local translation at synapses can cause neurodegenerative diseases.

Keywords

Amyotrophic lateral sclerosis Frontotemporal dementia Local translation RNA granules RNP granules Stress granules FUS TDP-43 

Abbreviations

ALS

Amyotrophic lateral sclerosis

AMPA

α-Amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid

CPEB

Cytoplasmic polyadenylation element-binding protein

FTD

Frontotemporal dementia

FMRP

Fragile X mental retardation protein 1

FUS

Fused in sarcoma

hnRNP

Heterogeneous nuclear ribonucleoprotein

mGluR

Metabotropic glutamate receptor

NES

Nuclear export signal

NLS

Nuclear localization signal

PSD-95

Postsynaptic density protein 95

PY-NLS

Proline-tyrosine nuclear localization signal

RISC

RNA-induced silencing complex

RGG

Arginine-glycine-glycine

RNP

Ribonucleoprotein particles

RRM

RNA recognition motif

TDP-43

Transactive response DNA-binding protein 43

tRNP

Transport ribonucleoprotein particles

UTR

Untranslated region

ZBP1

Zipcode-binding protein 1

Introduction

Translational control occurs mostly by homeostatic responses that alter general protein synthesis. However, gene-specific translational control depends on regulatory elements in the mRNA, such as upstream open reading frames, secondary structures or regulatory protein-binding sites [1]. As such, mRNA specificity in translational control can be achieved by the general translation machinery or by RNA-binding proteins, which are the main group of proteins that regulate mRNA transport and protein translation at synapses. In particular, RNA-binding proteins help meet the demands of the ever-changing microenvironment of the neuron which include responses to synaptic depolarization and depression, reduced nutrient availability, oxidative stress, misfolded proteins, and apoptosis. It is both a mystery and a marvel how large complexes of RNA-binding proteins and other core proteins coordinate mRNA transport and translation in response to these cues.

The importance of RNA-binding protein function at synapses is highlighted in patients with neurological and neurodegenerative disorders where genetic mutations or deletions of genes encoding RNA-binding proteins result in loss of synaptic plasticity and neuron function. Genetic mutations or deletions in disorders of autism, fragile X syndrome and Rett syndrome, which also correspond with a loss of synaptic plasticity and function, strongly suggest that disruption of RNA regulation is a central cause of synaptic defects in these brain disorders. Neurodegenerative disorders, such as amyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD), are related by overlapping clinical phenotypes, genetic links and rapid disease progression [2]. Whereas ALS is a motor neuron disease and is caused by selective degeneration of motor neurons, which results in gradual muscle weakness and atrophy and death [3], it also has loss of synapses as part of the disease. FTD is a common form of dementia and is characterized by atrophy of the frontal and temporal lobes, which cause changes in behavior, cognition, and changes in personality and/or language [4]. FTD can be accompanied by loss of motor neuron function, and up to 75 % of ALS patients experience behavior and cognitive impairment [5]. This has led to the conclusion that the cause of ALS and FTD are somehow linked. With the advances in genetic screening, what has emerged is a strong association with several RNA regulatory proteins with causing familial forms of both ALS and FTD [6, 7]. Given this information altered RNA regulation is likely the underlying cause of these diseases and is at the forefront of understanding the mechanism behind ALS and FTD. This review will highlight the role of RNA-binding proteins in regulating local translation, their impact on maintaining synapses and the potential role of disease-linked RNA-binding proteins, Fused in sarcoma (FUS) and Transactive response DNA-binding protein (TDP-43) in the dysregulation of synaptic function and the initiation of neurodegeneration.

The role of RNA granules at synapses

Neurons are highly complex cells with polarized and elaborate processes that extend long distances in the central nervous system. The distance between the synapse and cell body creates a supply and demand challenge for neurons, particularly at synapses. Neurons have the challenge of regulating the local translation of proteins at the synapse in order to meet the rapidly changing environment of neuronal inputs. The solution to these demands requires a local mechanism for controlling transport of mRNA to synapses and regulation of translation to allow local synthesis of new synaptic proteins at a moment’s notice. This is achieved by having all of the necessary components for translation; mRNA, ribosomes and translation factors, present in dendrites and even in the dendritic spines, ready for local protein synthesis [8, 9]. The regulation of local protein synthesis is particularly interesting because it allows neurons to rapidly modulate the production of proteins independent of new transcription or mRNA transport, which can modify the synapse directly. For instance, the localization of mRNAs at synapses and local protein synthesis is demonstrated to be critical for synaptic plasticity and the consolidation and storage of information in the brain [10]. Likewise, mRNA targeting and local protein synthesis have also been shown to influence axon guidance and nerve regeneration [11].

Messenger ribonucleoprotein complexes (mRNPs) form when mRNA associates with protein complexes. In higher eukaryotes, mRNPs comprise more than ten thousand different RNA sequences and hundreds of different RNA-binding proteins. mRNPs can also assemble into more complex structures known as RNA granules (or RNP granules). There are many types of RNA granules, for example, transport RNP granules (tRNP), stress granules, processing bodies, germ granules, and nuclear paraspeckles [12]. The classification of RNA granules is based on their composition, subcellular localization, cell of origin, response to stimuli, dynamic behavior, and proposed functions [12, 13]. RNA granules can form in response to cellular inputs and environmental cues. In turn, they regulate the distribution, translation, and degradation of mRNA transcripts. RNA granules do not function as isolated particles, but instead constantly interact with each other, exchanging mRNPs, cytosolic proteins, and with polysomes (Fig. 1a) (reviewed in [14]). In essence, the formation of different granules regulates mRNA and protein synthesis, which directly impact the neuron’s fate. Common to all RNA granules is the presence of RNA-binding proteins, which associate with mRNA in untranslated regions (5′UTR or 3′UTR) or coding regions [15, 16] and are largely responsible for coordinating thier localization, stability, and translation. Transport RNP granules, stress granules and processing bodies have been linked with the pathology of a variety of diseases and we will focus our discussion on their role in maintaining neuronal function and how disruption of these granules may lead to disease.
Fig. 1

Model of RNA granule dynamics in neurodegenerative disorders. a RNA-binding proteins associate with RNAs to form mRNPs, which assemble into large, diverse multi-mRNP complexes like tRNPs, stress granules, or processing bodies. tRNP granules determine the cytoplasmic localization and fate of the mRNA and keep the mRNA in a translationally dormant state. tRNP granules can associate and exchange mRNPs with stress granules and processing bodies in response to cellular cues such as stress. mRNAs are protected within stress granules during times of stress and serve as sites of mRNA triage where mRNP complexes are monitored for integrity and composition and are then routed to sites of reinitiation, degradation or storage. Once the stress has been removed, stress granules disassemble, mRNAs are repacked into translationally competent mRNAs and proteins are synthesized or are selectively exported to associated processing bodies for degradation. Processing bodies are sites of mRNA degradation, mRNA surveillance, translational repression, RNA-mediated silencing, and may also be involved in storage of select RNAs and recycling/modification of decay factors. Processing bodies can associate with tRNPs, stress granules, and translation machinery. Throughout the different exchanges between mRNP:RNA granules and mRNP:translation machinery, RNA-binding proteins are associated with their target mRNAs. Following translation, mRNPs can assemble back into translationally repressed tRNP granules, degraded or assembled into processing bodies. For a functioning neuron, these dynamic exchanges are important factors in the quality control of local translation at synapses and the maintenance of synaptic communication and plasticity. b Depicted is a model of how ALS/FTD mutations in FUS and TDP-43 affect RNA granule dynamics and the impact on translation and synaptic function. FUS-disease mutations cause an increase in number and size of both tRNP and stress granules. The impact of this may be two-fold: (1) FUS mutations which cause more spontaneous assembly of tRNP granules and increased translational activities would impact the normal processes of the neuron; and (2) FUS mutations which cause tRNP and stress granules to be more insoluble would lead to “seeding” of insoluble pathological inclusions associated with ALS and FTD. However, the insoluble nature of both tRNP and stress granules could also impact translation in a negative manner, which has yet to be determined. Additionally, FUS-disease mutations negatively impact the formation of processing bodies and solubility of stress granules, which would likely alter the normal functions and of these RNA granules. TDP-43-disease mutations on the other hand cause larger and fewer tRNP granules in the dendrites as well as larger and more stress granules to form in response to stress. The consequences of this may be very similar to what occurs with FUS mutations including reduced RNA granule exchanges, altered translational activities and increased “seeding” of insoluble protein aggregates. There have been no changes observed with processing bodies, but the insolubility of stress granules would predict a disturbance in mRNP:RNA granule exchanges. The net impact of FUS- and TDP-43-disease mutations would be altered RNA granule dynamics, leading to misregulation of mRNA and translation, which would impact synaptic function and cause neurodegeneration. (RBP RNA-binding protein, x a mutation in an RNA-binding protein, black arrows RNA granule exchanges, gray arrows RNA granule interactions with translation machinery, broken arrows altered RNA granule dynamics)

Transport RNP granules (tRNP granule) are ribonucleoprotein particles that function in transport and storage of mRNA and can contain miRNA [17, 18]. Several core protein components of tRNP granules include RNA-binding proteins such as Staufen1, Staufen2, Fragile X mental retardation protein 1 (FMR1 commonly known as FMRP), heterogeneous nuclear ribonucleoprotein A2 (hnRNPA2), cytoplasmic polyadenylation element-binding protein (CPEB), survival of motor neuron protein (SMN), zipcode-binding protein 1 (ZBP1), and Purine-Rich Element-Binding Protein alpha (Purα), which participate in dendritic transcript transport [19]. Other RNA-binding proteins that are found in tRNP granules include, Smaug, Paumilio, FUS, and TDP-43 [20, 21, 22, 23, 24]. More than 40 different proteins including motor proteins (i.e., kinesin) have been identified in tRNP granules, many of which are proteins related to RNA transport and regulation, protein synthesis and several of which their function is unknown [23, 25, 26]. These tRNP protein components may or may not be “essential” for tRNP granule formation; however, depending on the context may be essential for regulating mRNA fate.

Stress granules are cytoplasmic aggregates composed of RNA binding proteins, RNA and stalled translation initiation complexes that usually form in a reversible manner upon cellular stress [27]. In some instances stress granule growth will persist after removal of the stressor [28] or will dissolve even before the stress has been removed [29]. Moreover, stress granule clearance in mammalian cells can also occur by autophagy [30]. Stress graules serve as sites of mRNA triage where mRNP complexes are monitored for integrity and composition. Once the stress has been removed, they disassemble and mRNAs can exchange with tRNP granules [21], or be repacked into translationally competent mRNAs and translation can occur [28, 31, 32] or are mRNAs are selectively exported to associated processing bodies for degradation [33]. The primary protein components for stress granules formation include TIA1 cytotoxic granule-associated RNA-binding protein (TIA-1), TIA1 cytotoxic granule-associated RNA-binding protein-like 1 (TIAR), and GTPase Activating Protein (SH3 Domain) Binding Protein 1 (G3BP) along with poly(A)-binding protein (PABP) and the 40S ribosomal subunit [14, 19]. Stress granules also recruit translation initiation complex (i.e., eukaryotic translation initiation factor 2 (eIF2, eiF3, and eiF4E), multiple enzymes and signaling molecules, scaffolding and adaptor proteins, ubiquitin-modifying enzymes, RNA helicases, ribonucleases, ribosyl-, glucosyl- and methyl-transferases (reviewed in [34]). A number of disease-linked proteins are also recruited to stress granules, these proteins include FMRP, SMN, FUS, TDP-43, Ataxin-2 (ATXN2), and other RNA-binding proteins (Table 1) [27, 29, 35, 36, 37, 38, 39, 40, 41]. The role RNA-binding proteins have in the assembly and disassembly of stress granules and subsequent effects on protein translation is not fully understood, but it may be that they bring mRNAs to these granules and help protect and repress translation.
Table 1

Summary of RNA-binding proteins associated with RNA granules and linked to neurological diseases

RNA-binding protein

RNA granule

Link to disease

References

Angiogenin (ANG)

SG

Mutations in ALS and PD

[37, 98, 99]

Ataxin-2 (ATXN2)

SG

PolyQ expansions in ALS and SCA2

[27, 36, 95, 96]

Ewing sarcoma protein (EWS)

SG

Mutations in ALS, inclusions in FTD

[38, 100, 101]

Fragile X mental retardation protein (FMRP)

tRNP, SG, PB

Mutations in FXS

[19, 27, 44, 59]

Fused in sarcoma (FUS)

tRNP, SG

Mutations and inclusions in ALS, FTD & PQE

[6, 24, 35, 38, 104]

Heterogeneous nuclear ribonuclearprotein (hnRNPA2B1)

SG

Mutations in ALS, FTD and PGD

[39]

Heterogeneous nuclear ribonuclearprotein (hnRNPA1)

SG

Mutations in ALS and PGD

[39]

Survival of motor neuron (SMN)

tRNP, SG

Mutations in ALS and SMA

[19, 27, 40, 92, 93, 94]

TATA-binding protein-associated factor 15 (TAF15)

tRNP, SG, PB

Mutations in ALS, inclusions in ALS and FTD

[25, 38, 100, 102, 103]

TAR DNA-binding protein (TDP-43)

tRNP, SG, PB

Mutations in ALS, FTD, inclusions in AD and HD

[7, 22, 29, 105, 106]

AD Alzheimer’s disease, ALS amyotrophic lateral sclerosis, FTD frontotemporal dementia, FXS fragile X syndrome, HD Huntington’s disease, PB processing body, PD Parkinson’s disease, PGD Paget disease, PQE polyQ expansion disease, tRNP transport ribonucleoprotein particle granule, SCA2 spinocerebellar ataxia type 2, SG stress granule, SMA spinal muscular atrophy

Processing bodies are sites enriched with factors involved in mRNA degradation, mRNA surveillance, translational repression, RNA-mediated silencing, and may also be involved in storage of select RNAs and recycling/modification of decay factors. Many processing bodies exchange rapidly with cytoplasmic proteins and contain only a few stable components. Processing bodies have also been shown to interact with tRNPs [17] and are in physically and functionally associated with stress granules; sharing certain proteins, containing the same species of mRNAs and assemble and disassemble in response to drugs that promote or inhibit polysome disassembly [12, 33]. Due to their dynamic nature it is difficult to identify their exact protein composition and fully understand their function (reviewed in [14]). Whereas stress granules form transiently in response to stress, processing bodies are distinct cytoplasmic silencing foci that are present constitutively and can be enhanced by stress. Like stress granules, processing bodies can be induced by stress [32, 42, 43], are composed of several RNA-binding proteins (Table 1) [22, 25, 44] and contain translationally stalled mRNAs that can be targeted for degradation or may return to translation [32, 42, 43, 45]. However, processing bodies can be distinguished from stress granules by the presence of the RNA-binding proteins, decapping mRNA 2 (DCP2), decapping enzyme 1a (DCP1A), and U6 Small Nuclear RNA-Associated (LSM), trinucleotide repeat-containing 6A (TNRC6A also know as GW182) proteins [19, 46]. Moreover, due to their close ties with mRNA degradation, processing bodies also contain proteins involved in mRNA decay (i.e., decapping factors, DCP1/DCP1), RNA degradation (i.e., deadenylase complex CCR4/CAF1/NOT), nonsense-mediated mRNA decay proteins, ARE-mediated decay factors, and RNAi machinery (GW182 and Argonautes) (reviewed in [14]).

Mechanisms of local translational regulation by RNA-binding proteins

mRNP granules form in the nucleus. When not being actively translated, cytoplasmic mRNPs can assemble into large multi-mRNP complexes (i.e., tRNP granules) or be permanently disassemble and degraded (reviewed in [47]). The transportation of mRNAs to dendrites by tRNP granules is thought to occur in a translationally dormant state. Consistent with this model, eukaryotic translation initiation factor 4AIII (eIF4AIII), a protein involved in pre-mRNA splicing in the nucleus, was shown to be associated with dendritic mRNA [48]. Because eIF4AIII would be removed from the mRNA by the first ribosome to read the transcript, this suggests that these dendritic mRNAs have not been previously translated. The mechanism by which tRNP granule protein components can selectively inhibit mRNA translation until the proper cues occur is not known. In the case of mammalian Staufen-1 and -2 proteins, some tRNP granules contain ribosomes, whereas others do not. When fractionated by size, the largest Staufen pools contained ribosomes and endoplasmic reticulum, whereas the smaller tRNP granules were cofractioned with kinesin and were free of ribosomes and endoplasmic reticulum [49]. This evidence suggests that the smaller tRNP granules might represent the translationally repressed pool of this type of RNA granule. Moreover, in response to neuronal activity, mRNAs can be released from tRNP granules to the polyribosome fraction where transcripts are actively translated [50]. However, repression of translation by tRNP granules has also been reported in response to neuronal activity (discussed below). The determining factor between active or repressed translation may be controlled by the protein composition of the granule.

The role of the protein components of tRNP granules is an ongoing question. There is evidence to suggest that RNA-binding proteins which associate with tRNP granules have the ability to regulate mRNA repression and/or translation. The RNA-binding protein, ZBP1 associates with and transports β-actin mRNA to synapses [51]. Once in the cytoplasm ZBP1 can be dissociated from the mRNA by Src phosphorylation, allowing synthesis of β-actin, which is necessary for cell migration and neurite outgrowth [52]. ZBP1 can repress the joining of ribosomal subunits in the cytoplasm, thereby regulating translation initiation [52]. Another regulator of mRNA translation in neurons is the CPEB family of RNA-binding proteins. In this example, CPEB1 functions as both a repressor and activator of translation. Initially, CPEB1 binds near the end of the 3′UTR anchors a complex of proteins that include an eIF4E-binding protein (maskin), a poly(A)-polymerase (Gld2), a scaffolding protein (symplekin), and a deadenylase (poly(A)-specific ribonuclease (PARN) [53, 54, 55]. In oocytes, binding of CPEB1 to the mRNA initially inhibits mRNA translation through the interaction of an maskin and eIF4E; however, CPEB1 phosphorylation leads to the dissociation of PARN from the complex and subsequent polyadenylation of the 3′ tail by Gld2 [55]. This polyadenylation results in the dissociation of maskin from eIF4E and the activation of translation [56]. Neurons likely use a similar process to regulate translation in dendrites [57, 58].

One well-known regulator of translation is the RNA-binding protein FMRP, which regulates translation and mRNA transportation to dendrites. Mutations in the gene encoding FMRP are associated with a loss of function and cause one of the most common inherited forms of autism, Fragile X syndrome [59]. FMRP-associated tRNP granules traffic into dendrites upon activation of group 1 metabotropic glutamate receptors (mGluR) to regulate translation [60]. The binding of FMRP to mRNA is shown to inhibit translation [61]; however, mice that lack FMRP exhibit both up- and down-regulation of FMRP mRNA targets [62, 63]. Moreover, in the absence of FMRP, the over translation of mRNA normally regulated by FMRP in the dendritic spine leads to excess internalization of the α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor and enhanced long-term depression following mGluR activation [64]. Consistent with this multifaceted role in mRNA translation, FMRP can associate with multiple types of RNA granules that contain both actively translating polyribosomes [65, 66] and non-translating RNPs [67, 68]. At the synapse, a model for FMRP function has been proposed whereby mGluR activation results in a disinhibition of FMRP-bound mRNA. Where in acute mGluR stimulation, FMRP is dephosphorylated by protein phosphatase-2A (PP2A), ubiquitinated and degraded, which relieves translation inhibition and enable immediate translation of FMRP-bound mRNAs [60, 69]. In contrast, extended activation of mGluR1 (1 min or more) results in rapamycin (mTOR)-mediated PP2A suppression and FMRP rephosphorylation, which coincides with translation inhibition of select FMRP target transcripts [69].

Several studies indicate that FMRP plays a critical role in regulation of mRNA translation by serving as a link between transport tRNP granules and polyribosomes. This is based on the distribution of FMRP to these fractions. For instance, FMRP is present in polyribosomes and acts to stall ribosomal translocation during elongation of its target mRNAs [61]. The phosphorylated form of FMRP associates with stalled polyribosomes, whereas unphosphorylated FMRP associates with actively translating polyribosomes [70]. Presumably, dephosphorylated FMRP no longer acts as a repressor of translation, allowing the ribosomes to translate mRNA and run-off. FMRP can also regulate translation through an association between miRNAs and the RNA-induced silencing complex (RISC) [71]. FMRP regulates translation by acting on the RISC complex containing miR-125a to modulate translation of postsynaptic density protein 95 (PSD-95) [72]. When FMRP is phosphorylated, FMRP recruits Argonaute 2 (Ago2) complexes containing miR-125a and represses synthesis of proteins, such as PSD-95. In response to mGluR signaling, FMRP dephosphorylation leads to the release of RISC from PSD-95 mRNA, which stimulates translation [72]. In this case, FMRP acts as a bridge to deliver miRNA to complementary mRNAs. Thus, dysregulation of microRNAs is also part of how RNA-binding proteins exert translational control, a potential process that is disrupted in fragile X and diseases involving other RNA-binding proteins. We have only mentioned some of the findings for FMRP function and regulation, for more detailed review see [73]; however, the mentioned body of work highlights the complexity of RNA-binding proteins in regulating translation in the dendrites. Knowing the dynamic nature of RNA granules, the existence of different pools of tRNP granules, and the multi-protein complexes that make up these granules at any given moment, it is still unclear how RNA-binding proteins collectively influence translation, the transition of targeting mRNAs to polyribosomes, and subsequent regulation of translation.

Dendrite and synapse attrition and the link to neurodegenerative disorders

Neuron dendritic branching, synapse formation, and stabilization play significant roles in the structural and functional plasticity of the brain. Precise synapse development and formation is important for accurate neuronal network activity and normal brain function. Therefore, it is not surprising that alterations in dendrite morphology or defects in neuronal development, including changes in dendrite branching patterns, fragmentation of dendrites, retraction or loss of dendrite branching, and changes in spine morphology and number, contribute to disease. In particular, these changes have been observed in several neurodegenerative, neurodevelopmental, and neuropsychiatric disorders, such as, ALS [74], FTD [75], Alzheimer’s disease, Down’s syndrome, autism spectrum disorders, fragile X syndrome, Rett syndrome [76], anxiety and depression [77, 78], schizophrenia [79], and Parkinson’s disease [80]. Various studies report that many neuropsychiatric disorders are characterized by dendritic and synaptic pathology, including abnormal spine density and morphology, synapse loss, and aberrant synaptic signaling and plasticity [81, 82].

Animal models of neurodegenerative diseases also show changes in dendritic branches and abnormal spine morphology including animal models of ALS [83, 84] and FTD [75, 83], as well as in models of mental retardation and fragile X syndrome [85, 86]. In particular, there are several examples of RNA-binding proteins which have been found to affect neuronal morphology and function, and their deficiency are implicated in causing alterations in dendritic branching and spines which underlie the associated neurological diseases. Alterations in dendritic spines in fragile X syndrome and the corresponding FMRP knockout mouse model is characterized by an excess of long and thin filopodial-like spines and a reduction in mature spines [86], which is likely due to dysregulated protein synthesis at synapses. Staufen associates with tRNP granules along with FMRP, TDP-43 and huntingtin [22, 87, 88] and may indirectly contribute to neurological diseases. This is predicted to occur in diseases related to FMRP, TDP-43 and huntingtin where RNA granule biology is affected, which also alters Staufen localization and function. The relevance of Staufens and disease comes from its role in maintaining dendrites and spines. For instance, in hippocampal neurons derived from Staufen-1 knockout mice have deficits in dendritic delivery of β-actin tRNP granules and these neurons have significantly reduced dendritic tree and develop fewer synapses [89]. Staufen-2 is shown to be essential to dendritic spines in mammalian hippocampal neurons, wherein neurons deficient for Staufen-2 have reduced dendritic spines and increased filopodia, which are caused, in part, from impaired β-actin mRNA localization [90]. FUS, which will be discussed in more detail below, is locally translated and localized to spines in response to mGluR5 activation [83, 91]. Neurons cultured from FUS-null mice showed an excess of filopodial-like or thin spines lacking heads and a reduction of mature spines having a mushroom shape [91] and transgenic mice expressing ALS-FUS-associated mutations also have fewer mature spines and reduced dendritic branches [83, 84]. Collectively, these studies demonstrate the role of mRNA regulation by RNA-binding proteins in translation affecting spine development, which likely have important consequences for synaptic plasticity, learning, and memory [10]. There are diverse mechanisms and genetics that underlie the loss of dendritic branching patterns and synapses for these disorders. However, protein translation at synapses has emerged as a central process in the maintenance of dendritic branches and synapses. Thus, altered RNA metabolism at synapses may be the root cause of neurodegeneration observed in ALS and FTD.

RNA-binding proteins associated with neurodegenerative disorders

In the last 10 years more RNA-binding proteins have been identified through genetic studies to be linked with causing neurodegenerative diseases. Genes that encode SMN [92, 93, 94], ATXN2 [95, 96], senataxin (ALS4) [97], angiogenin (ANG) [98, 99], ewing sarcoma protein (EWS) [100, 101], heterogeneous nuclear ribonucleoproteins (hnRNPA2B1 and hnRNPA1) [39], TATA-binding protein-associated factor 15 (TAF15) [100, 102, 103] along with previously discussed FUS [6] and TDP-43 [7], have functions in post-transcriptional regulation of RNA and mutations in these genes cause motor neuron degeneration and other neurological disorders (Table 1). Additionally, several of these proteins have also been linked with neuropathology of diseases (Table 1) [6, 7, 100, 104, 105, 106]. Mutations in the RNA-binding proteins, FUS [6], and TDP-43 [7] are identified as genetic causes of both ALS and FTD. The role in how these proteins contribute to disease is not fully understood, but dysregulation of RNA metabolism as in fragile X syndrome is likely a major factor in the contribution to neurodegeneration. Here, we will discuss the prominent ALS and FTD-associated RNA-binding proteins, FUS, and TDP-43, and their roles in promoting neurodegeneration, potentially through altered protein translational regulation at synapses.

FUS properties and function

FUS is part of the FET family of RNA-binding proteins that include EWSR1 and TAF15 [107]. FUS binds to thousands of cellular RNAs [108, 109, 110, 111] through its two RNA recognition motifs (RRMs), zinc-finger domain, and three arginine-glycine-glycine (RGG) boxes [112, 113, 114]. FUS exists in different ribonucleoprotein complexes involved in pre-mRNA splicing, mRNA stability and mRNA transport [115, 116, 117, 118] and miRNA biogenesis [119]. FUS co-purifies with the spliceosome [120, 121], different pre-mRNA splicing [117, 118] and miRNA Drosha complexes [122]. At its steady state, FUS is localized to the nucleus of cells and in nuclear gemini of coiled bodies (gems) along with TDP-43 and SMN, where all three proteins function in spliceosome maintenance [115]. A feature of FUS is that it binds the whole length of nascent RNA [109, 111], which implies its close association with transcripts from the time of production and supports the idea that it may be involved in transcriptional elongation. This is further supported by the finding that FUS has a close association with the C-terminal domain of RNA polymerase II and its association with that complex can influence phosphorylation and transcriptional activation [123, 124]. FUS binds to pre-RNA and mRNA at introns, coding sequences, 3′UTR and 5′UTRs and also targets noncoding RNAs [108, 109, 111]. The impact of RNA regulation by FUS is demonstrated in FUS-null cells where there is a broad misregulation of RNA processing involving mRNA regulation and pre-mRNA splicing [109, 111]. Functional classification of FUS RNA targets reveals a number of essential cellular processes. Notably, a significant number of their RNA targets encode proteins that function at the synapse, several of which are involved in neuronal development and synaptic transmission [108, 109, 111].

The majority of FUS is localized to the nucleus of cells; however, it can localize to different cellular compartments and RNA granules in response to various stimuli. This is facilitated by its non-classical proline-tyrosine nuclear localization sequence (PY-NLS) and nuclear export sequence (NES) [125]. For example, treatment of acute cortical slices or hippocampal neurons with mGluR1/5 agonists, results in local translation [83], and localization to synapses [91]. FUS has also been found in tRNP granules localized to dendrites [23, 24] and localized to synapses where some of FUS associates with the NMDA receptor [24]. Although it has not been tested, the role of FUS in the spines may control localization, anchoring, or regulating mRNAs at the synapse. The function of FUS in tRNP granules may be to repress or facilitate translation, this has yet to be fully understood. Recently, FUS has been detected in tRNP granules containing the tumor suppressor protein, adenomatous polyposis coli (APC) in hippocampal neurons, and post-mortem tissues from FTD-FUS patients [126]. In APC-tRNP granules, FUS is demonstrated to be required for the efficient translation of associated transcripts. As discussed above, this is an unexpected finding given that tRNP granules are thought to be translationally dormant. Interestingly, overexpression of ALS-FUS mutations causes the spontaneous formation of APC-tRNP granules which are translationally active [126]. FUS has also been purified from polyribosomes [127], which suggests it has an active role in regulating translation and that it could function in a similar way as FMRP to bridge tRNP granules and translation machinery. In response to oxidative stressors like sorbitol and sodium arsenite, FUS associates with cytoplasmic stress granules [35, 38, 41], which indicates it also has the capability to be an active repressor of translation. Finally, the association of FUS with the Drosha complex and miRNA biogenesis [122], suggests that FUS can repress translation by distinct mechanisms. Future work should identify the relationship between FUS and RNA granule dynamics and the role FUS plays in the regulation of translation.

FUS and neurodegeneration

The majority of familial ALS mutations that are identified in the gene encoding FUS occur in its C-terminal PY-NLS [128], altering the cytoplasmic localization of the protein and are aggregate prone [35, 129]. Mutations in the 3′UTR of FUS have also been identified, which results in increased FUS expression and cause ALS [130]. The extent to which ALS-FUS mutations localize to the cytoplasm correlates with disease severity [35, 129]. FTD genetic mutations are identified in the RGG1 and N-terminal region, but as demonstrated in cell culture, cytoplasmic localization of FUS is not as prominent [131]. Post-mortem examination of ALS patients with FUS mutations shows abnormal accumulation of FUS in the cytoplasm or nucleus of motor neurons [6]. Similarly, familial FTD and some sporadic forms of the disease have similar FUS aggregation, but in FTD affected brain regions [6].

Animal models expressing various FUS mutations and wild-type FUS reproduce aspects of the human diseases. For instance, D. melanogaster [132], C. elegans [133], and R. norvegicus [134], models that overexpress ALS-FUS mutations causes motor defects in these models, although it is unclear why overexpression of wild-type FUS in these models has little toxicity given that ALS-FUS 3′UTR mutations cause an increase in FUS expression [130]. Transgenic mouse models harboring human wild-type FUS or the ALS-FUS R521G mutation both develop deficits in motor function, motor neuron denervation, and inflammation die before reaching adulthood [83]. However, persistent reduction of dendritic branching and mature spines is only present in the FUSR521G transgenic mice [135], which has also been shown in the transgenic mouse model containing the R521C mutation [84]. These transgenic mouse models of ALS-FUS show similar deficits in dendritic branches and spines as reported in FUS-null hippocampal cultures [91]. These observations suggest that the mechanism of toxicity of these mutants may be due to a partial loss of function regarding RNA regulation and a gain of function regarding the toxicity the mutation causes at the synapse. Meanwhile, a recent study in D. melanogaster demonstrates that replacement of the endogenous FUS homolog with the human FUS R521C mutation causes defects in synapse structure and function that precedes motor neurodegeneration [136], which is consistent with the observations in FUSR521G transgenic mice [83]. These observations may be explained by the finding that the FUSR521G mutation is not regulated in the same manner as wild-type in response to neuronal stimulation [83]. It is possible that in response to mGluR, FUS is normally translated at the synapses; however, translation of the FUSR521G protein is dysregulated, which may have negative implications at the synapse. A commonality among all animal models is that they do not develop large amounts of pathological FUS aggregates. This could indicate that FUS aggregation is not central to the disease process, and/or it could suggest that overexpressing FUS causes toxicity by altering its other biological processes as indicated in animal models that have altered dendritic branches and spines.

A potential significance of FUS aggregation comes from the finding that several stress granule markers are also deposited in the aggregates [6]. This has led to several studies that examine stress granule dynamics of FUS mutations, these studies have been more easily modeled in cell culture. In general, ALS-FUS PY-NLS mutations which correlate with disease severity are more prominently localized to the cytoplasm and form larger stress granules in the absence and presence of stress [35, 129]. For instance, recruitment of GFP-FUS into perinuclear stress granules is extensive for truncation mutants R495X or G515X compared with R521G or H517Q [35]. Moreover, there is an indication that FUS mutations may delay the assembly of stress granules [137] and irreversibly sequester a variety of RNA-binding proteins and mRNAs [138]. Despite the close contact of processing bodies with stress granules under conditions of acute stress, GFP-FUS and mutant variants do not incorporate into associated processing bodies or affect docking to stress granules [35]. However, FUS mutations impact the number of processing bodies [137, 138]. The functional implications for these findings are still not clear. It has also been shown that R521G and H517Q mutations have reduced binding to RNA [110]. Not all mutations have been tested for their ability to bind RNA, but the altered ability of the mutants so far tested to bind mRNAs and sequester them in RNA granules may be an important factor in disease pathogenesis. Until now, the model put forth is that FUS mutations somehow “seed” for protein aggregates which sequesters more FUS and other proteins, thus depleting the cell of essential proteins which leads to cell death. This model still needs in vivo data to support the observations made in cell culture.

TDP-43 properties and function

FUS and TDP-43 have often been reported as having a similar role in disease and biology, but there are some distinctions and similarities, which we will point out in this section. TDP-43 is a member of the heterogeneous nuclear ribonucleoprotein (hnRNP) family of proteins [139]. Like FUS, TDP-43 is a global regulator of gene expression and binds thousands of RNAs [140, 141, 142] through its two highly conserved RNA recognition motifs (RRM1 and RRM2), wherein the RRM1 is the major domain for binding RNA and DNA [143, 144]. TDP-43 regulates transcription and multiple aspects of RNA processing and function, including pre-mRNA splicing, mRNA stability, transport, translation [140, 142, 145], and miRNA biogenesis [146, 147]. TDP-43 co-localizes with splicing structures in the nucleus [115], Drosha [122] and DICER complexes [147]. Additionally, TDP-43 interacts with many proteins and RNAs and functions in protein:RNA complexes [142, 148]. TDP-43 is ubiquitously expressed and localizes primarily to the nucleus of cells and can localize to splicing structures known as GEMs [115]. TDP-43 targets thousands of RNAs, which has been shown in vivo and in various model systems [140, 141, 142]. Binding RNA through its highly conserved RRMs, TDP-43 has enriched binding to over 4500 RNA species, preferentially localizing to introns, 3′UTR and 5′UTRs and non-coding RNAs [140, 141, 142]. Deletion of TDP-43 in cells leads to broad misregulation of mRNA and pre-mRNA splicing [140, 141]. Similar to FUS, TDP-43 targets RNAs of diverse biological importance. In the context of synaptic biology, it shares similar targets to FUS, but with different binding sites [108, 111, 140, 141, 142].

TDP-43 can localize to different cellular compartments and RNA granules via a classical NLS and NES [149]. It too is responsive to various stimuli which somehow directs the localization of TDP-43 to these specialized areas. Under basal conditions TDP-43 resides in the dendrites of hippocampal neurons and co-localizes with RNA granules, some of which stain positive for processing bodies, and it co-localizes with the post-synaptic protein, PSD-95. These granules contain RNAs including β-actin and calmodulin kinase II α (CaMKIIα) mRNA. Upon depolarization, TDP-43 co-localizes FMRP and Staufen-1 in tRNP granules within dendrites [22]. In response to oxidative stressors, TDP-43 localizes to the cytoplasm and into stress granules [22, 29, 150, 151, 152]. TDP-43 has not been costained with processing bodies in other studies [153]; however, this does not preclude that TDP-43 can associate with these structures under some conditions because many proteins, such as Staufen, FMRP, and HuR are present in processing bodies and stress granules depending on the conditions [154]. TDP-43 has also been found as an integral component of the Dicer complex and is required for the cleavage of pre-miRNAs by Dicer and for the recruitment of Argonaute 2 (Ago2) to the catalytic engine of RISC, to miRNA bound by Dicer [147, 155]. In this study, TDP-43, but not FUS, was found as a component of nuclear Drosha complexes that contain DGCR8, which is indispensable for pri-miRNA processing [146]. TDP-43 is shown to facilitate the production of a subset of precursor miRNAs (pre-miRNAs) by both interacting with the nuclear Drosha complex and binding directly to primary miRNAs (pri-miRNAs) [146]. Furthermore, cytoplasmic TDP-43, which interacts with the Dicer complex, promotes the processing of some of these pre-miRNAs to miRNAs [146]. The implications for TDP-43 involvement in miRNA biogenesis were shown to be indispensable for neuronal outgrowth [146]. Taken together, these findings suggest that the maturation of a subset of miRNAs is modulated at multiple steps by TDP-43, which reveals a unique function of TDP-43 not only in the nucleus but also in the cytoplasm. As demonstrated in this work, TDP-43 is functioning in a similar manner as FMRP in translational repression via miRNA regulation.

TDP-43 and neurodegeneration

TDP-43 is a major component of ubiquitinated inclusions found in the brain and spinal cord of the majority of ALS patients [156, 157]. In approximately 50 % of FTD patients TDP-43 is found to be the pathological hallmark, the remaining cases are TDP-43 negative and most of which have tau-positive neuropathology [2]. There are now more than 40 TDP-43 familial ALS mutations that have been identified and the majority are found in the C-terminal glycine region [128]. Mutations in the 3′UTR of TDP-43 have also been identified, resulting in increased levels of TDP-43 [158]. Familial FTD mutations in TDP-43 are less prevalent [128]. Unlike FUS, the identified ALS-TDP-43 mutations do not occur in the NLS or NES domains and the impact on localization is not as obvious. Most of the mutations occur in the glycine-rich region, which is shown to mediate protein interactions [159] and contains a predicted prion-like domain [160, 161]. Despite extensive research over the last several years, the pathogenesis of these mutations is still unclear.

Animal models of TDP-43 demonstrate that altering endogenous levels of the wild-type protein or expressing ALS-TDP-43 mutations is highly toxic and reproduces aspects of ALS and FTD [135, 162, 163, 164, 165, 166]. Depletion of TDP-43 in D. melanogaster results in reduced lifespan and locomotor defects due to alterations in dendritic branching and synapses [167, 168, 169, 170], overexpression also caused loss of motor function and is accompanied by a decrease in dendrites and synapses [170, 171]. In D. rerio, both overexpression of human TDP-43 and knockdown of TARDBP result in swimming behavior defects caused by defective neuronal axon formation, and premature and excessive branching [172]. Similarly, transgenic mouse models of hTDP-43 expressing either wild-type or ALS-associated mutations cause motor defects [162, 163, 164]. TDP-43M337V transgenic rats expressing mutant proteins in motor neurons recover their motor function when TDP-43M337V expression is turned off [135]. This study suggests that mutant TDP-43 in motor neurons is sufficient to promote the onset and progression of ALS-like degeneration and that, most importantly, its toxic effects are reversible. TDP-43 knockout mice die between embryonic day E3.5 and E8.5 and TDP-43 null-embryonic stem cells are not viable [165]. Conditional knockout of TDP-43 in motor neurons exhibit progressive development of ALS-related motor phenotypes and accumulation of ubiquitinated proteins [166]. Although dendrite and spine morphology have not been extensively studies in murine models, knockdown or overexpression of ALS-TDP-43 mutant proteins (A315T, Q331K, and M337V) in cortical neurons have been shown to cause both abnormal neurites and decreased cell viability [173]. Depletion of TDP-43 leads to an increase number of mature spines in hippocampal neurons, with an increase clustering of AMPA receptors on the dendritic surface which corresponds with increased neuron firing [174]. Overexpression of TDP-43 causes a loss of mature spines [174]. In knockdown cells, these changes correlate with increased level of Rac1 [174], a positive regulator of spinogenesis, suggesting that TDP-43 may be an upstream suppressor of Rac1 translation. These models indicate that the balance of TDP-43 levels is important for normal biological processes, at least in the case of the loss of TDP-43 and given the large number of RNA targets of this protein we can predict that there would be a significant biological impact. However, in the case of overexpression of either wild-type or TDP-43 mutations, the gain of function mechanism is not clear.

Another property of ALS-TDP-43 mutations and wild-type TDP-43 is that they are actively recruited to cytoplasmic stress granules in response to stress [150]. Moreover, TDP-43 ALS-associated mutations (i.e., A315T, G348C) are more sensitive to oxidative stress, and form larger stress granules in the cytoplasm, but do not form stress granules in the absence of stress [29]. The region in which these mutations are located appears to be important for stress granule recruitment because the deletion of the C-terminal glycine-rich domain abolishes TDP-43’s association with these granules [29, 113, 152]. In addition to this, the N-terminal RRM1 is also necessary for TDP-43’s incorporation into stress granules [113]. Together this indicates that stress granule recruitment of TDP-43 requires both RNA-binding and protein–protein interactions. Other consequences of TDP-43 mutations include increased half-life of the protein [116] and the stability of its target mRNAs [175]. As in the case of FUS, TDP-43 and stress granule markers co-stain in ALS and FTLD-U pathological aggregates [113], which suggests that TDP-43 may also “seed” stress granules formation in pathological aggregates [150].

Much of the field has focused on stress granules; however, TDP-43 mutations have been shown to affect tRNP granule formation and migration. TDP-43 is associated with RNA granules that are prevalent throughout the dendritic arbor in neurons. “Aggregation” of TDP-43 is also important for the formation of these neuronal tRNP granules, and it is reasonable to assume disease-linked mutations might alter granule formation. Indeed, ALS-TDP-43 mutations are shown to increase the size of neuronal TDP-43 granules in the dendritic arbor of rat hippocampal neurons under basal conditions [153]. Depolarization of rat hippocampal neurons with KCl stimulates TDP-43 granule migration into dendrites, but A315T and Q343R TDP-43 granules migrate shorter distances and into fewer dendrites than wild-type TDP-43 [153]. The mutations correspondingly reduce the granule density, movement, and mobility of TDP-43 granules. Interestingly, some TDP-43-positive RNA granules also exhibit a close interaction with processing bodies [153]. In another study that examined TDP-43 tRNP granules, TDP-43 mutations impair mRNA transport in stem cell-derived motor neurons from ALS patients bearing any one of three different TDP-43 ALS causing mutations [176]. These findings highlight novel elements of TDP-43 biology that are affected by disease-linked mutations and suggest a neuronally selective mechanism through which TDP-43 mutations might elicit neuronal dysfunction. The functional implications could be the absence of important RNAs to sites of local translation or misregulated mRNA products. Given the close association of TDP-43 with tRNP granules, the functional implications of TDP-43 mutations in tRNP granule formation and trafficking will also need to be examined.

Future perspectives

The role of RNA-binding proteins in translational control and how it relates to neurodegeneration is gaining interest in the field. This has occurred for a number of reasons: (1) identification of genetic mutations in genes encoding proteins that are involved in RNA regulation and linked to neurodegenerative diseases; (2) in vivo disease models do not recapitulate pathological aggregate phenotypes of the diseases and in vitro stress granule models do not fully explain the global impact on RNA misregulation; (3) at least in the case of TDP-43-FTD, cross-linking and immunoprecipitation of TDP-43 and transcriptome analysis reveal very few changes in RNA-binding profiles from control patients [141], and the genes that are altered do not link directly to causes of neurodegeneration; (4) synaptic dysfunction precedes neurodegeneration and recent evidence shows that tRNP granule formation, localization and dynamics are affected by disease mutations of RNA-binding proteins. This suggests that mRNA transportation, processing and translation may be affected, which may lead to defects at the synapse and trigger the earliest events during the neurodegeneration process (Fig. 1b).

If alterations of RNA granule dynamics, transport, and translation are important factors in maintaining the synapse, then how are RNA-binding proteins coordinating these processes? Dynamic exchange of mRNA between tRNP granules, stress granules and processing bodies is in part coordinated by specific RNA-binding proteins. There is evidence that supports RNA-binding proteins are essential in the shuttling of mRNA to and from these specialized granules until such time that the mRNA is translated in polyribosomes. The impact of deletion of RNA-binding proteins can lead to alterations in dendritic branches and spines as well as impact translation. Key questions that arise are: How do RNA-binding proteins target mRNA to each RNA granule? What cues initiate mRNA exchange between RNA granules? What post-translational modifications do RNA-binding proteins undergo to coordinate exchanges between RNA granules? It is clear that stress granule formation and the presence and numbers of processing bodies are affected by ALS-FUS and ALS-TDP-43 mutations. Recent work shows that tRNP granules are also affected by ALS-FUS and ALS-TDP-43 mutations. Putting all this evidence together would strongly suggest that mutations at least in these RNA-binding proteins would affect the dynamics of mRNP–RNA granule exchanges, which we defined here as the transfer of mRNA:Protein complexes between RNA granules in response to cellular stimuli and cues (Fig. 1b). More concerted efforts need to be made to examine the effects on local translation and the downstream consequences at the synapse.

Finally, if RNA-binding proteins affect mRNP–RNA granule exchange, how does this affect local translation and synaptic function, and can the resulting effects lead to neurodegeneration? The strongest precedence by an RNA-binding protein has been set for FMRP in the autism disorder fragile X syndrome. In this case, loss of FMRP causes changes in synaptic morphology and function in fragile X syndrome, but this disease does not involve neurodegeneration. The challenge in the neurodegeneration field will be to mechanistically link dysfunction of mRNP-RNA granule exchange and local translation with initiation of neurodegeneration. This will be difficult to dissect given that RNA-binding proteins like FUS and TDP-43 regulate thousands of RNAs in both the nucleus and cytoplasm. A particular focus will need to be on target RNAs that encode proteins critical for synaptic function at both the steady state and in response to neuron stimulation. As observed in the FUSR521G transgenic model, defects in translation are more prominent at synapses in response to mGluR stimulation. The function of FUS and TDP-43 at synapses is not well understood, but the existing evidence point to a prominent role in maintaining the function and integrity of synapses: these RNA-binding proteins have been purified with synaptic tRNP granules; their expression is elevated upon neuron stimulation; genetic deletion results in altered dendritic spines and branches in primary cultured neurons. This in addition to their genetic link to ALS and FTD strongly implicate RNA-binding proteins as having a major role in causing alterations in RNA metabolism locally at the synapse, which would alter synaptic function and trigger neurodegeneration.

Notes

Acknowledgments

Due to space limitations, we sincerely apologize for the inability to cite all worthy contributions made to the biology of RNA-binding proteins. This work was supported in part by Alzheimer’s Association New Investigator Research Grant (NIRG-14-321584) and Alzheimer Society of Canada Young Investigator Research Grant (15-29) for CFS, and Alzheimer’s Association Grant (IIRG-11-205714) to GY.

References

  1. 1.
    Dever TE (2002) Gene-specific regulation by general translation factors. Cell 108(4):545–556PubMedCrossRefGoogle Scholar
  2. 2.
    Van Langenhove T, van der Zee J, Van Broeckhoven C (2012) The molecular basis of the frontotemporal lobar degeneration-amyotrophic lateral sclerosis spectrum. Ann Med 44(8):817–828PubMedCentralPubMedCrossRefGoogle Scholar
  3. 3.
    Kiernan MC et al (2011) Amyotrophic lateral sclerosis. Lancet 377(9769):942–955PubMedCrossRefGoogle Scholar
  4. 4.
    Snowden JS, Neary D, Mann DM (2002) Frontotemporal dementia. Br J Psychiatry 180:140–143PubMedCrossRefGoogle Scholar
  5. 5.
    Strong MJ et al (2009) Consensus criteria for the diagnosis of frontotemporal cognitive and behavioural syndromes in amyotrophic lateral sclerosis. Amyotroph Lateral Scler 10(3):131–146PubMedCrossRefGoogle Scholar
  6. 6.
    Dormann D, Haass C (2013) Fused in sarcoma (FUS): an oncogene goes awry in neurodegeneration. Mol Cell Neurosci 56:475–486PubMedCrossRefGoogle Scholar
  7. 7.
    Lee EB, Lee VM, Trojanowski JQ (2012) Gains or losses: molecular mechanisms of TDP43-mediated neurodegeneration. Nat Rev Neurosci 13(1):38–50Google Scholar
  8. 8.
    Steward O, Levy WB (1982) Preferential localization of polyribosomes under the base of dendritic spines in granule cells of the dentate gyrus. J Neurosci 2(3):284–291PubMedGoogle Scholar
  9. 9.
    Sutton MA, Schuman EM (2006) Dendritic protein synthesis, synaptic plasticity, and memory. Cell 127(1):49–58PubMedCrossRefGoogle Scholar
  10. 10.
    Klann E, Dever TE (2004) Biochemical mechanisms for translational regulation in synaptic plasticity. Nat Rev Neurosci 5(12):931–942PubMedCrossRefGoogle Scholar
  11. 11.
    Willis D et al (2005) Differential transport and local translation of cytoskeletal, injury-response, and neurodegeneration protein mRNAs in axons. J Neurosci 25(4):778–791PubMedCrossRefGoogle Scholar
  12. 12.
    Anderson P, Kedersha N (2009) RNA granules: post-transcriptional and epigenetic modulators of gene expression. Nat Rev Mol Cell Biol 10(6):430–436PubMedCrossRefGoogle Scholar
  13. 13.
    Buchan JR (2014) mRNP granules. Assembly, function, and connections with disease. RNA Biol 11(8):1019–1030PubMedPubMedCentralCrossRefGoogle Scholar
  14. 14.
    Anderson P, Kedersha N, Ivanov P (2014) Stress granules, P-bodies and cancer. Biochim Biophys Acta. doi: 10.1016/j.bbagrm.2014.11.009 PubMedCentralGoogle Scholar
  15. 15.
    Martin KC, Ephrussi A (2009) mRNA localization: gene expression in the spatial dimension. Cell 136(4):719–730PubMedCentralPubMedCrossRefGoogle Scholar
  16. 16.
    Richter JD, Klann E (2009) Making synaptic plasticity and memory last: mechanisms of translational regulation. Genes Dev 23(1):1–11PubMedCrossRefGoogle Scholar
  17. 17.
    Zeitelhofer M et al (2008) Dynamic interaction between P-bodies and transport ribonucleoprotein particles in dendrites of mature hippocampal neurons. J Neurosci 28(30):7555–7562PubMedCrossRefGoogle Scholar
  18. 18.
    Xing L, Bassell GJ (2013) mRNA localization: an orchestration of assembly, traffic and synthesis. Traffic 14(1):2–14PubMedCentralPubMedCrossRefGoogle Scholar
  19. 19.
    Kiebler MA, Bassell GJ (2006) Neuronal RNA granules: movers and makers. Neuron 51(6):685–690PubMedCrossRefGoogle Scholar
  20. 20.
    Baez MV, Boccaccio GL (2005) Mammalian Smaug is a translational repressor that forms cytoplasmic foci similar to stress granules. J Biol Chem 280(52):43131–43140PubMedCrossRefGoogle Scholar
  21. 21.
    Vessey JP et al (2006) Dendritic localization of the translational repressor Pumilio 2 and its contribution to dendritic stress granules. J Neurosci 26(24):6496–6508PubMedCrossRefGoogle Scholar
  22. 22.
    Wang IF et al (2008) TDP-43, the signature protein of FTLD-U, is a neuronal activity-responsive factor. J Neurochem 105(3):797–806PubMedCrossRefGoogle Scholar
  23. 23.
    Kanai Y, Dohmae N, Hirokawa N (2004) Kinesin transports RNA: isolation and characterization of an RNA-transporting granule. Neuron 43(4):513–525PubMedCrossRefGoogle Scholar
  24. 24.
    Belly A et al (2005) Delocalization of the multifunctional RNA splicing factor TLS/FUS in hippocampal neurones: exclusion from the nucleus and accumulation in dendritic granules and spine heads. Neurosci Lett 379(3):152–157PubMedCrossRefGoogle Scholar
  25. 25.
    Marko M et al (2012) Domains involved in TAF15 subcellular localisation: dependence on cell type and ongoing transcription. Gene 506(2):331–338PubMedCrossRefGoogle Scholar
  26. 26.
    Mikl M, Vendra G, Kiebler MA (2011) Independent localization of MAP2, CaMKIIalpha and beta-actin RNAs in low copy numbers. EMBO Rep 12(10):1077–1084PubMedCentralPubMedCrossRefGoogle Scholar
  27. 27.
    Anderson P, Kedersha N (2008) Stress granules: the Tao of RNA triage. Trends Biochem Sci 33(3):141–150PubMedCrossRefGoogle Scholar
  28. 28.
    Mollet S et al (2008) Translationally repressed mRNA transiently cycles through stress granules during stress. Mol Biol Cell 19(10):4469–4479PubMedCentralPubMedCrossRefGoogle Scholar
  29. 29.
    Dewey CM et al (2011) TDP-43 is directed to stress granules by sorbitol, a novel physiological osmotic and oxidative stressor. Mol Cell Biol 31(5):1098–1108PubMedCentralPubMedCrossRefGoogle Scholar
  30. 30.
    Buchan JR et al (2013) Eukaryotic stress granules are cleared by autophagy and Cdc48/VCP function. Cell 153(7):1461–1474PubMedCentralPubMedCrossRefGoogle Scholar
  31. 31.
    Nover L, Scharf KD, Neumann D (1989) Cytoplasmic heat shock granules are formed from precursor particles and are associated with a specific set of mRNAs. Mol Cell Biol 9(3):1298–1308PubMedCentralPubMedCrossRefGoogle Scholar
  32. 32.
    Kedersha N et al (2000) Dynamic shuttling of TIA-1 accompanies the recruitment of mRNA to mammalian stress granules. J Cell Biol 151(6):1257–1268PubMedCentralPubMedCrossRefGoogle Scholar
  33. 33.
    Kedersha N et al (2005) Stress granules and processing bodies are dynamically linked sites of mRNP remodeling. J Cell Biol 169(6):871–884PubMedCentralPubMedCrossRefGoogle Scholar
  34. 34.
    Kedersha N, Ivanov P, Anderson P (2013) Stress granules and cell signaling: more than just a passing phase? Trends Biochem Sci 38(10):494–506PubMedCrossRefGoogle Scholar
  35. 35.
    Bosco DA et al (2010) Mutant FUS proteins that cause amyotrophic lateral sclerosis incorporate into stress granules. Hum Mol Genet 19(21):4160–4175PubMedCentralPubMedCrossRefGoogle Scholar
  36. 36.
    Nonhoff U et al (2007) Ataxin-2 interacts with the DEAD/H-box RNA helicase DDX6 and interferes with P-bodies and stress granules. Mol Biol Cell 18(4):1385–1396PubMedCentralPubMedCrossRefGoogle Scholar
  37. 37.
    Emara MM et al (2010) Angiogenin-induced tRNA-derived stress-induced RNAs promote stress-induced stress granule assembly. J Biol Chem 285(14):10959–10968PubMedCentralPubMedCrossRefGoogle Scholar
  38. 38.
    Andersson MK et al (2008) The multifunctional FUS, EWS and TAF15 proto-oncoproteins show cell type-specific expression patterns and involvement in cell spreading and stress response. BMC Cell Biol 9:37PubMedCentralPubMedCrossRefGoogle Scholar
  39. 39.
    Kim HJ et al (2013) Mutations in prion-like domains in hnRNPA2B1 and hnRNPA1 cause multisystem proteinopathy and ALS. Nature 495(7442):467–473PubMedCentralPubMedCrossRefGoogle Scholar
  40. 40.
    Hua Y, Zhou J (2004) Survival motor neuron protein facilitates assembly of stress granules. FEBS Lett 572(1–3):69–74PubMedCrossRefGoogle Scholar
  41. 41.
    Shelkovnikova TA et al (2014) Multistep process of FUS aggregation in the cell cytoplasm involves RNA-dependent and RNA-independent mechanisms. Hum Mol Genet 23(19):5211–5226PubMedCentralPubMedCrossRefGoogle Scholar
  42. 42.
    Cougot N, Babajko S, Seraphin B (2004) Cytoplasmic foci are sites of mRNA decay in human cells. J Cell Biol 165(1):31–40PubMedCentralPubMedCrossRefGoogle Scholar
  43. 43.
    Bhattacharyya SN et al (2006) Relief of microRNA-mediated translational repression in human cells subjected to stress. Cell 125(6):1111–1124PubMedCrossRefGoogle Scholar
  44. 44.
    Antar LN et al (2005) Localization of FMRP-associated mRNA granules and requirement of microtubules for activity-dependent trafficking in hippocampal neurons. Genes Brain Behav 4(6):350–359PubMedCrossRefGoogle Scholar
  45. 45.
    Brengues M, Teixeira D, Parker R (2005) Movement of eukaryotic mRNAs between polysomes and cytoplasmic processing bodies. Science 310(5747):486–489PubMedCentralPubMedCrossRefGoogle Scholar
  46. 46.
    van Dijk E et al (2002) Human Dcp2: a catalytically active mRNA decapping enzyme located in specific cytoplasmic structures. EMBO J 21(24):6915–6924PubMedCentralPubMedCrossRefGoogle Scholar
  47. 47.
    Singh G, Pratt G, Yeo GW, Moore MJ (2015) The clothes make the mRNA: past and present trends in mRNP fashion. Annu Rev Biochem 84:325–354Google Scholar
  48. 48.
    Giorgi C et al (2007) The EJC factor eIF4AIII modulates synaptic strength and neuronal protein expression. Cell 130(1):179–191PubMedCrossRefGoogle Scholar
  49. 49.
    Mallardo M et al (2003) Isolation and characterization of Staufen-containing ribonucleoprotein particles from rat brain. Proc Natl Acad Sci USA 100(4):2100–2105PubMedCentralPubMedCrossRefGoogle Scholar
  50. 50.
    Krichevsky AM, Kosik KS (2001) Neuronal RNA granules: a link between RNA localization and stimulation-dependent translation. Neuron 32(4):683–696PubMedCrossRefGoogle Scholar
  51. 51.
    Zhang HL et al (2001) Neurotrophin-induced transport of a beta-actin mRNP complex increases beta-actin levels and stimulates growth cone motility. Neuron 31(2):261–275PubMedCrossRefGoogle Scholar
  52. 52.
    Huttelmaier S et al (2005) Spatial regulation of beta-actin translation by Src-dependent phosphorylation of ZBP1. Nature 438(7067):512–515PubMedCrossRefGoogle Scholar
  53. 53.
    Stebbins-Boaz B et al (1999) Maskin is a CPEB-associated factor that transiently interacts with elF-4E. Mol Cell 4(6):1017–1027PubMedCrossRefGoogle Scholar
  54. 54.
    Barnard DC et al (2004) Symplekin and xGLD-2 are required for CPEB-mediated cytoplasmic polyadenylation. Cell 119(5):641–651PubMedCrossRefGoogle Scholar
  55. 55.
    Kim JH, Richter JD (2006) Opposing polymerase-deadenylase activities regulate cytoplasmic polyadenylation. Mol Cell 24(2):173–183PubMedCrossRefGoogle Scholar
  56. 56.
    Cao Q, Richter JD (2002) Dissolution of the maskin-eIF4E complex by cytoplasmic polyadenylation and poly(A)-binding protein controls cyclin B1 mRNA translation and oocyte maturation. EMBO J 21(14):3852–3862PubMedCentralPubMedCrossRefGoogle Scholar
  57. 57.
    Wu L et al (1998) CPEB-mediated cytoplasmic polyadenylation and the regulation of experience-dependent translation of alpha-CaMKII mRNA at synapses. Neuron 21(5):1129–1139PubMedCrossRefGoogle Scholar
  58. 58.
    Wells DG et al (2001) A role for the cytoplasmic polyadenylation element in NMDA receptor-regulated mRNA translation in neurons. J Neurosci 21(24):9541–9548PubMedGoogle Scholar
  59. 59.
    Garber K et al (2006) Transcription, translation and fragile X syndrome. Curr Opin Genet Dev 16(3):270–275PubMedCrossRefGoogle Scholar
  60. 60.
    Nalavadi VC et al (2012) Dephosphorylation-induced ubiquitination and degradation of FMRP in dendrites: a role in immediate early mGluR-stimulated translation. J Neurosci 32(8):2582–2587PubMedCentralPubMedCrossRefGoogle Scholar
  61. 61.
    Darnell JC et al (2011) FMRP stalls ribosomal translocation on mRNAs linked to synaptic function and autism. Cell 146(2):247–261PubMedCentralPubMedCrossRefGoogle Scholar
  62. 62.
    Miyashiro KY et al (2003) RNA cargoes associating with FMRP reveal deficits in cellular functioning in Fmr1 null mice. Neuron 37(3):417–431PubMedCrossRefGoogle Scholar
  63. 63.
    Brown V et al (2001) Microarray identification of FMRP-associated brain mRNAs and altered mRNA translational profiles in fragile X syndrome. Cell 107(4):477–487PubMedCrossRefGoogle Scholar
  64. 64.
    Bassell GJ, Warren ST (2008) Fragile X syndrome: loss of local mRNA regulation alters synaptic development and function. Neuron 60(2):201–214PubMedCentralPubMedCrossRefGoogle Scholar
  65. 65.
    Khandjian EW et al (2004) Biochemical evidence for the association of fragile X mental retardation protein with brain polyribosomal ribonucleoparticles. Proc Natl Acad Sci USA 101(36):13357–13362PubMedCentralPubMedCrossRefGoogle Scholar
  66. 66.
    Stefani G et al (2004) Fragile X mental retardation protein is associated with translating polyribosomes in neuronal cells. J Neurosci 24(33):7272–7276PubMedCrossRefGoogle Scholar
  67. 67.
    Zalfa F et al (2003) The fragile X syndrome protein FMRP associates with BC1 RNA and regulates the translation of specific mRNAs at synapses. Cell 112(3):317–327PubMedCrossRefGoogle Scholar
  68. 68.
    Siomi MC et al (1996) Specific sequences in the fragile X syndrome protein FMR1 and the FXR proteins mediate their binding to 60S ribosomal subunits and the interactions among them. Mol Cell Biol 16(7):3825–3832PubMedCentralPubMedCrossRefGoogle Scholar
  69. 69.
    Narayanan U et al (2007) FMRP phosphorylation reveals an immediate-early signaling pathway triggered by group I mGluR and mediated by PP2A. J Neurosci 27(52):14349–14357PubMedCrossRefGoogle Scholar
  70. 70.
    Ceman S et al (2003) Phosphorylation influences the translation state of FMRP-associated polyribosomes. Hum Mol Genet 12(24):3295–3305PubMedCrossRefGoogle Scholar
  71. 71.
    Jin P et al (2004) Biochemical and genetic interaction between the fragile X mental retardation protein and the microRNA pathway. Nat Neurosci 7(2):113–117PubMedCrossRefGoogle Scholar
  72. 72.
    Muddashetty RS et al (2011) Reversible inhibition of PSD-95 mRNA translation by miR-125a, FMRP phosphorylation, and mGluR signaling. Mol Cell 42(5):673–688PubMedCentralPubMedCrossRefGoogle Scholar
  73. 73.
    Darnell JC, Klann E (2013) The translation of translational control by FMRP: therapeutic targets for FXS. Nat Neurosci 16(11):1530–1536PubMedCentralPubMedCrossRefGoogle Scholar
  74. 74.
    Fischer LR et al (2004) Amyotrophic lateral sclerosis is a distal axonopathy: evidence in mice and man. Exp Neurol 185(2):232–240PubMedCrossRefGoogle Scholar
  75. 75.
    Petkau TL et al (2012) Synaptic dysfunction in progranulin-deficient mice. Neurobiol Dis 45(2):711–722PubMedCrossRefGoogle Scholar
  76. 76.
    Kaufmann WE, Moser HW (2000) Dendritic anomalies in disorders associated with mental retardation. Cereb Cortex 10(10):981–991PubMedCrossRefGoogle Scholar
  77. 77.
    Eiland L, McEwen BS (2012) Early life stress followed by subsequent adult chronic stress potentiates anxiety and blunts hippocampal structural remodeling. Hippocampus 22(1):82–91PubMedCrossRefGoogle Scholar
  78. 78.
    Soetanto A et al (2010) Association of anxiety and depression with microtubule-associated protein 2- and synaptopodin-immunolabeled dendrite and spine densities in hippocampal CA3 of older humans. Arch Gen Psychiatry 67(5):448–457PubMedCentralPubMedCrossRefGoogle Scholar
  79. 79.
    Stephan KE, Baldeweg T, Friston KJ (2006) Synaptic plasticity and dysconnection in schizophrenia. Biol Psychiatry 59(10):929–939PubMedCrossRefGoogle Scholar
  80. 80.
    Calabresi P et al (2006) A convergent model for cognitive dysfunctions in Parkinson’s disease: the critical dopamine-acetylcholine synaptic balance. Lancet Neurol 5(11):974–983PubMedCrossRefGoogle Scholar
  81. 81.
    Blanpied TA, Ehlers MD (2004) Microanatomy of dendritic spines: emerging principles of synaptic pathology in psychiatric and neurological disease. Biol Psychiatry 55(12):1121–1127PubMedCrossRefGoogle Scholar
  82. 82.
    Bourne JN, Harris KM (2008) Balancing structure and function at hippocampal dendritic spines. Annu Rev Neurosci 31:47–67PubMedCentralPubMedCrossRefGoogle Scholar
  83. 83.
    Sephton CF et al (2014) Activity-dependent FUS dysregulation disrupts synaptic homeostasis. Proc Natl Acad Sci USA 111(44):E4769–E4778PubMedCentralPubMedCrossRefGoogle Scholar
  84. 84.
    Qiu H et al (2014) ALS-associated mutation FUS-R521C causes DNA damage and RNA splicing defects. J Clin Invest 124(3):981–999PubMedCentralPubMedCrossRefGoogle Scholar
  85. 85.
    Dolen G, Bear MF (2008) Role for metabotropic glutamate receptor 5 (mGluR5) in the pathogenesis of fragile X syndrome. J Physiol 586(6):1503–1508PubMedCentralPubMedCrossRefGoogle Scholar
  86. 86.
    Irwin SA, Galvez R, Greenough WT (2000) Dendritic spine structural anomalies in fragile-X mental retardation syndrome. Cereb Cortex 10(10):1038–1044PubMedCrossRefGoogle Scholar
  87. 87.
    Savas JN et al (2010) A role for huntington disease protein in dendritic RNA granules. J Biol Chem 285(17):13142–13153PubMedCentralPubMedCrossRefGoogle Scholar
  88. 88.
    Villace P, Marion RM, Ortin J (2004) The composition of Staufen-containing RNA granules from human cells indicates their role in the regulated transport and translation of messenger RNAs. Nucleic Acids Res 32(8):2411–2420PubMedCentralPubMedCrossRefGoogle Scholar
  89. 89.
    Vessey JP et al (2008) A loss of function allele for murine Staufen1 leads to impairment of dendritic Staufen1-RNP delivery and dendritic spine morphogenesis. Proc Natl Acad Sci USA 105(42):16374–16379PubMedCentralPubMedCrossRefGoogle Scholar
  90. 90.
    Goetze B et al (2006) The brain-specific double-stranded RNA-binding protein Staufen2 is required for dendritic spine morphogenesis. J Cell Biol 172(2):221–231PubMedCentralPubMedCrossRefGoogle Scholar
  91. 91.
    Fujii R et al (2005) The RNA binding protein TLS is translocated to dendritic spines by mGluR5 activation and regulates spine morphology. Curr Biol 15(6):587–593PubMedCrossRefGoogle Scholar
  92. 92.
    Lefebvre S et al (1995) Identification and characterization of a spinal muscular atrophy-determining gene. Cell 80(1):155–165PubMedCrossRefGoogle Scholar
  93. 93.
    Corcia P et al (2002) Abnormal SMN1 gene copy number is a susceptibility factor for amyotrophic lateral sclerosis. Ann Neurol 51(2):243–246PubMedCrossRefGoogle Scholar
  94. 94.
    Coovert DD et al (1997) The survival motor neuron protein in spinal muscular atrophy. Hum Mol Genet 6(8):1205–1214PubMedCrossRefGoogle Scholar
  95. 95.
    Elden AC et al (2010) Ataxin-2 intermediate-length polyglutamine expansions are associated with increased risk for ALS. Nature 466(7310):1069–1075PubMedCentralPubMedCrossRefGoogle Scholar
  96. 96.
    Pulst SM et al (1996) Moderate expansion of a normally biallelic trinucleotide repeat in spinocerebellar ataxia type 2. Nat Genet 14(3):269–276PubMedCrossRefGoogle Scholar
  97. 97.
    Chen YZ et al (2004) DNA/RNA helicase gene mutations in a form of juvenile amyotrophic lateral sclerosis (ALS4). Am J Hum Genet 74(6):1128–1135PubMedCentralPubMedCrossRefGoogle Scholar
  98. 98.
    Greenway MJ et al (2006) ANG mutations segregate with familial and ‘sporadic’ amyotrophic lateral sclerosis. Nat Genet 38(4):411–413PubMedCrossRefGoogle Scholar
  99. 99.
    van Es MA et al (2011) Angiogenin variants in Parkinson disease and amyotrophic lateral sclerosis. Ann Neurol 70(6):964–973PubMedCrossRefGoogle Scholar
  100. 100.
    Neumann M et al (2011) FET proteins TAF15 and EWS are selective markers that distinguish FTLD with FUS pathology from amyotrophic lateral sclerosis with FUS mutations. Brain 134(Pt 9):2595–2609PubMedCentralPubMedCrossRefGoogle Scholar
  101. 101.
    Couthouis J et al (2012) Evaluating the role of the FUS/TLS-related gene EWSR1 in amyotrophic lateral sclerosis. Hum Mol Genet 21(13):2899–2911PubMedCentralPubMedCrossRefGoogle Scholar
  102. 102.
    Couthouis J et al (2011) A yeast functional screen predicts new candidate ALS disease genes. Proc Natl Acad Sci USA 108(52):20881–20890PubMedCentralPubMedCrossRefGoogle Scholar
  103. 103.
    Ticozzi N et al (2011) Mutational analysis reveals the FUS homolog TAF15 as a candidate gene for familial amyotrophic lateral sclerosis. Am J Med Genet B Neuropsychiatr Genet 156B(3):285–290PubMedCrossRefGoogle Scholar
  104. 104.
    Doi H et al (2010) The RNA-binding protein FUS/TLS is a common aggregate-interacting protein in polyglutamine diseases. Neurosci Res 66(1):131–133PubMedCrossRefGoogle Scholar
  105. 105.
    Schwab C et al (2008) Colocalization of transactivation-responsive DNA-binding protein 43 and huntingtin in inclusions of Huntington disease. J Neuropathol Exp Neurol 67(12):1159–1165PubMedCrossRefGoogle Scholar
  106. 106.
    Amador-Ortiz C et al (2007) TDP-43 immunoreactivity in hippocampal sclerosis and Alzheimer’s disease. Ann Neurol 61(5):435–445PubMedCentralPubMedCrossRefGoogle Scholar
  107. 107.
    Tan AY, Manley JL (2009) The TET family of proteins: functions and roles in disease. J Mol Cell Biol 1(2):82–92PubMedCentralPubMedCrossRefGoogle Scholar
  108. 108.
    Ishigaki S et al (2012) Position-dependent FUS-RNA interactions regulate alternative splicing events and transcriptions. Sci Rep 2:529PubMedCentralPubMedCrossRefGoogle Scholar
  109. 109.
    Rogelj B et al (2012) Widespread binding of FUS along nascent RNA regulates alternative splicing in the brain. Sci Rep 2:603PubMedCentralPubMedCrossRefGoogle Scholar
  110. 110.
    Hoell JI et al (2011) RNA targets of wild-type and mutant FET family proteins. Nat Struct Mol Biol 18(12):1428–1431PubMedCentralPubMedCrossRefGoogle Scholar
  111. 111.
    Lagier-Tourenne C et al (2012) Divergent roles of ALS-linked proteins FUS/TLS and TDP-43 intersect in processing long pre-mRNAs. Nat Neurosci 15(11):1488–1497PubMedCentralPubMedCrossRefGoogle Scholar
  112. 112.
    Lerga A et al (2001) Identification of an RNA binding specificity for the potential splicing factor TLS. J Biol Chem 276(9):6807–6816PubMedCrossRefGoogle Scholar
  113. 113.
    Bentmann E et al (2012) Requirements for stress granule recruitment of fused in sarcoma (FUS) and TAR DNA-binding protein of 43 kDa (TDP-43). J Biol Chem 287(27):23079–23094PubMedCentralPubMedCrossRefGoogle Scholar
  114. 114.
    Iko Y et al (2004) Domain architectures and characterization of an RNA-binding protein. TLS. J Biol Chem 279(43):44834–44840PubMedCrossRefGoogle Scholar
  115. 115.
    Tsuiji H et al (2013) Spliceosome integrity is defective in the motor neuron diseases ALS and SMA. EMBO Mol Med 5(2):221–234PubMedCentralPubMedCrossRefGoogle Scholar
  116. 116.
    Ling SC et al (2010) ALS-associated mutations in TDP-43 increase its stability and promote TDP-43 complexes with FUS/TLS. Proc Natl Acad Sci USA 107(30):13318–13323PubMedCentralPubMedCrossRefGoogle Scholar
  117. 117.
    Calvio C et al (1995) Identification of hnRNP P2 as TLS/FUS using electrospray mass spectrometry. RNA 1(7):724–733PubMedCentralPubMedGoogle Scholar
  118. 118.
    Kameoka S, Duque P, Konarska MM (2004) p54(nrb) associates with the 5′ splice site within large transcription/splicing complexes. EMBO J 23(8):1782–1791PubMedCentralPubMedCrossRefGoogle Scholar
  119. 119.
    Morlando M et al (2012) FUS stimulates microRNA biogenesis by facilitating co-transcriptional Drosha recruitment. EMBO J 31(24):4502–4510PubMedCentralPubMedCrossRefGoogle Scholar
  120. 120.
    Rappsilber J et al (2002) Large-scale proteomic analysis of the human spliceosome. Genome Res 12(8):1231–1245PubMedCentralPubMedCrossRefGoogle Scholar
  121. 121.
    Zhou Z et al (2002) Comprehensive proteomic analysis of the human spliceosome. Nature 419(6903):182–185PubMedCrossRefGoogle Scholar
  122. 122.
    Gregory RI et al (2004) The Microprocessor complex mediates the genesis of microRNAs. Nature 432(7014):235–240PubMedCrossRefGoogle Scholar
  123. 123.
    Kwon I et al (2013) Phosphorylation-regulated binding of RNA polymerase II to fibrous polymers of low-complexity domains. Cell 155(5):1049–1060PubMedCentralPubMedCrossRefGoogle Scholar
  124. 124.
    Schwartz JC et al (2013) RNA seeds higher-order assembly of FUS protein. Cell Rep 5(4):918–925PubMedCentralPubMedCrossRefGoogle Scholar
  125. 125.
    Lee BJ et al (2006) Rules for nuclear localization sequence recognition by karyopherin beta 2. Cell 126(3):543–558PubMedCentralPubMedCrossRefGoogle Scholar
  126. 126.
    Yasuda K et al (2013) The RNA-binding protein Fus directs translation of localized mRNAs in APC-RNP granules. J Cell Biol 203(5):737–746PubMedCentralPubMedCrossRefGoogle Scholar
  127. 127.
    Reschke M et al (2013) Characterization and analysis of the composition and dynamics of the mammalian riboproteome. Cell Rep 4(6):1276–1287PubMedCrossRefGoogle Scholar
  128. 128.
    Bentmann E, Haass C, Dormann D (2013) Stress granules in neurodegeneration–lessons learnt from TAR DNA binding protein of 43 kDa and fused in sarcoma. FEBS J 280(18):4348–4370PubMedCrossRefGoogle Scholar
  129. 129.
    Dormann D et al (2010) ALS-associated fused in sarcoma (FUS) mutations disrupt Transportin-mediated nuclear import. EMBO J 29(16):2841–2857PubMedCentralPubMedCrossRefGoogle Scholar
  130. 130.
    Sabatelli M et al (2013) Mutations in the 3′ untranslated region of FUS causing FUS overexpression are associated with amyotrophic lateral sclerosis. Hum Mol Genet 22(23):4748–4755PubMedCrossRefGoogle Scholar
  131. 131.
    Wang WY et al (2013) Interaction of FUS and HDAC1 regulates DNA damage response and repair in neurons. Nat Neurosci 16(10):1383–1391PubMedCrossRefGoogle Scholar
  132. 132.
    Lanson NA Jr et al (2011) A Drosophila model of FUS-related neurodegeneration reveals genetic interaction between FUS and TDP-43. Hum Mol Genet 20(13):2510–2523PubMedCentralPubMedCrossRefGoogle Scholar
  133. 133.
    Murakami T et al (2012) ALS mutations in FUS cause neuronal dysfunction and death in Caenorhabditis elegans by a dominant gain-of-function mechanism. Hum Mol Genet 21(1):1–9PubMedCentralPubMedCrossRefGoogle Scholar
  134. 134.
    Huang C et al (2011) FUS transgenic rats develop the phenotypes of amyotrophic lateral sclerosis and frontotemporal lobar degeneration. PLoS Genet 7(3):e1002011PubMedCentralPubMedCrossRefGoogle Scholar
  135. 135.
    Huang C et al (2012) Mutant TDP-43 in motor neurons promotes the onset and progression of ALS in rats. J Clin Invest 122(1):107–118PubMedCentralPubMedCrossRefGoogle Scholar
  136. 136.
    Shahidullah M et al (2013) Defects in synapse structure and function precede motor neuron degeneration in Drosophila models of FUS-related ALS. J Neurosci 33(50):19590–19598PubMedCentralPubMedCrossRefGoogle Scholar
  137. 137.
    Baron DM et al (2013) Amyotrophic lateral sclerosis-linked FUS/TLS alters stress granule assembly and dynamics. Mol Neurodegener 8:30PubMedCentralPubMedCrossRefGoogle Scholar
  138. 138.
    Takanashi K, Yamaguchi A (2014) Aggregation of ALS-linked FUS mutant sequesters RNA binding proteins and impairs RNA granules formation. Biochem Biophys Res Commun 452(3):600–607PubMedCrossRefGoogle Scholar
  139. 139.
    Krecic AM, Swanson MS (1999) hnRNP complexes: composition, structure, and function. Curr Opin Cell Biol 11(3):363–371PubMedCrossRefGoogle Scholar
  140. 140.
    Polymenidou M et al (2011) Long pre-mRNA depletion and RNA missplicing contribute to neuronal vulnerability from loss of TDP-43. Nat Neurosci 14(4):459–468PubMedCentralPubMedCrossRefGoogle Scholar
  141. 141.
    Tollervey JR et al (2011) Characterizing the RNA targets and position-dependent splicing regulation by TDP-43. Nat Neurosci 14(4):452–458PubMedCentralPubMedCrossRefGoogle Scholar
  142. 142.
    Sephton CF et al (2011) Identification of neuronal RNA targets of TDP-43-containing ribonucleoprotein complexes. J Biol Chem 286(2):1204–1215PubMedCentralPubMedCrossRefGoogle Scholar
  143. 143.
    Ou SH et al (1995) Cloning and characterization of a novel cellular protein, TDP-43, that binds to human immunodeficiency virus type 1 TAR DNA sequence motifs. J Virol 69(6):3584–3596PubMedCentralPubMedGoogle Scholar
  144. 144.
    Buratti E, Baralle FE (2001) Characterization and functional implications of the RNA binding properties of nuclear factor TDP-43, a novel splicing regulator of CFTR exon 9. J Biol Chem 276(39):36337–36343PubMedCrossRefGoogle Scholar
  145. 145.
    Buratti E, Baralle FE (2010) The multiple roles of TDP-43 in pre-mRNA processing and gene expression regulation. RNA Biol 7(4):420–429PubMedCrossRefGoogle Scholar
  146. 146.
    Kawahara Y, Mieda-Sato A (2012) TDP-43 promotes microRNA biogenesis as a component of the Drosha and Dicer complexes. Proc Natl Acad Sci USA 109(9):3347–3352PubMedCentralPubMedCrossRefGoogle Scholar
  147. 147.
    Chendrimada TP et al (2005) TRBP recruits the Dicer complex to Ago2 for microRNA processing and gene silencing. Nature 436(7051):740–744PubMedCentralPubMedCrossRefGoogle Scholar
  148. 148.
    Freibaum BD et al (2010) Global analysis of TDP-43 interacting proteins reveals strong association with RNA splicing and translation machinery. J Proteome Res 9(2):1104–1120PubMedCentralPubMedCrossRefGoogle Scholar
  149. 149.
    Winton MJ et al (2008) Disturbance of nuclear and cytoplasmic TAR DNA-binding protein (TDP-43) induces disease-like redistribution, sequestration, and aggregate formation. J Biol Chem 283(19):13302–13309PubMedCentralPubMedCrossRefGoogle Scholar
  150. 150.
    Dewey CM et al (2012) TDP-43 aggregation in neurodegeneration: are stress granules the key? Brain Res 1462:16–25PubMedCentralPubMedCrossRefGoogle Scholar
  151. 151.
    Moisse K et al (2009) Divergent patterns of cytosolic TDP-43 and neuronal progranulin expression following axotomy: implications for TDP-43 in the physiological response to neuronal injury. Brain Res 1249:202–211PubMedCrossRefGoogle Scholar
  152. 152.
    Colombrita C et al (2009) TDP-43 is recruited to stress granules in conditions of oxidative insult. J Neurochem 111(4):1051–1061PubMedCrossRefGoogle Scholar
  153. 153.
    Liu-Yesucevitz L et al (2014) ALS-linked mutations enlarge TDP-43-enriched neuronal RNA granules in the dendritic arbor. J Neurosci 34(12):4167–4174PubMedCentralPubMedCrossRefGoogle Scholar
  154. 154.
    Barbee SA et al (2006) Staufen- and FMRP-containing neuronal RNPs are structurally and functionally related to somatic P bodies. Neuron 52(6):997–1009PubMedCentralPubMedCrossRefGoogle Scholar
  155. 155.
    Gregory RI et al (2005) Human RISC couples microRNA biogenesis and posttranscriptional gene silencing. Cell 123(4):631–640PubMedCrossRefGoogle Scholar
  156. 156.
    Neumann M et al (2006) Ubiquitinated TDP-43 in frontotemporal lobar degeneration and amyotrophic lateral sclerosis. Science 314(5796):130–133PubMedCrossRefGoogle Scholar
  157. 157.
    Mackenzie IR et al (2007) Pathological TDP-43 distinguishes sporadic amyotrophic lateral sclerosis from amyotrophic lateral sclerosis with SOD1 mutations. Ann Neurol 61(5):427–434PubMedCrossRefGoogle Scholar
  158. 158.
    Gitcho MA et al (2009) TARDBP 3′-UTR variant in autopsy-confirmed frontotemporal lobar degeneration with TDP-43 proteinopathy. Acta Neuropathol 118(5):633–645PubMedCentralPubMedCrossRefGoogle Scholar
  159. 159.
    Ayala YM et al (2008) Structural determinants of the cellular localization and shuttling of TDP-43. J Cell Sci 121(Pt 22):3778–3785PubMedCrossRefGoogle Scholar
  160. 160.
    Fuentealba RA et al (2010) Interaction with polyglutamine aggregates reveals a Q/N-rich domain in TDP-43. J Biol Chem 285(34):26304–26314PubMedCentralPubMedCrossRefGoogle Scholar
  161. 161.
    Cushman M et al (2010) Prion-like disorders: blurring the divide between transmissibility and infectivity. J Cell Sci 123(Pt 8):1191–1201PubMedCentralPubMedCrossRefGoogle Scholar
  162. 162.
    Swarup V et al (2011) Pathological hallmarks of amyotrophic lateral sclerosis/frontotemporal lobar degeneration in transgenic mice produced with TDP-43 genomic fragments. Brain 134(Pt 9):2610–2626PubMedCrossRefGoogle Scholar
  163. 163.
    Wils H et al (2010) TDP-43 transgenic mice develop spastic paralysis and neuronal inclusions characteristic of ALS and frontotemporal lobar degeneration. Proc Natl Acad Sci U S A 107(8):3858–3863PubMedCentralPubMedCrossRefGoogle Scholar
  164. 164.
    Wegorzewska I et al (2009) TDP-43 mutant transgenic mice develop features of ALS and frontotemporal lobar degeneration. Proc Natl Acad Sci USA 106(44):18809–18814PubMedCentralPubMedCrossRefGoogle Scholar
  165. 165.
    Sephton CF et al (2010) TDP-43 is a developmentally regulated protein essential for early embryonic development. J Biol Chem 285(9):6826–6834PubMedCentralPubMedCrossRefGoogle Scholar
  166. 166.
    Wu LS, Cheng WC, Shen CK (2012) Targeted depletion of TDP-43 expression in the spinal cord motor neurons leads to the development of amyotrophic lateral sclerosis-like phenotypes in mice. J Biol Chem 287(33):27335–27344PubMedCentralPubMedCrossRefGoogle Scholar
  167. 167.
    Fiesel FC et al (2010) Knockdown of transactive response DNA-binding protein (TDP-43) downregulates histone deacetylase 6. EMBO J 29(1):209–221PubMedCentralPubMedCrossRefGoogle Scholar
  168. 168.
    Feiguin F et al (2009) Depletion of TDP-43 affects Drosophila motoneurons terminal synapsis and locomotive behavior. FEBS Lett 583(10):1586–1592PubMedCrossRefGoogle Scholar
  169. 169.
    Lu Y, Ferris J, Gao FB (2009) Frontotemporal dementia and amyotrophic lateral sclerosis-associated disease protein TDP-43 promotes dendritic branching. Mol Brain 2:30PubMedCentralPubMedCrossRefGoogle Scholar
  170. 170.
    Lin MJ, Cheng CW, Shen CK (2011) Neuronal function and dysfunction of Drosophila dTDP. PLoS One 6(6):e20371PubMedCentralPubMedCrossRefGoogle Scholar
  171. 171.
    Li Y et al (2010) A Drosophila model for TDP-43 proteinopathy. Proc Natl Acad Sci USA 107(7):3169–3174PubMedCentralPubMedCrossRefGoogle Scholar
  172. 172.
    Kabashi E et al (2010) Gain and loss of function of ALS-related mutations of TARDBP (TDP-43) cause motor deficits in vivo. Hum Mol Genet 19(4):671–683PubMedCrossRefGoogle Scholar
  173. 173.
    Han JH et al (2013) ALS/FTLD-linked TDP-43 regulates neurite morphology and cell survival in differentiated neurons. Exp Cell Res 319(13):1998–2005PubMedCrossRefGoogle Scholar
  174. 174.
    Majumder P et al (2012) TDP-43 regulates the mammalian spinogenesis through translational repression of Rac1. Acta Neuropathol 124(2):231–245PubMedCrossRefGoogle Scholar
  175. 175.
    Colombrita C et al (2012) TDP-43 and FUS RNA-binding proteins bind distinct sets of cytoplasmic messenger RNAs and differently regulate their post-transcriptional fate in motoneuron-like cells. J Biol Chem 287(19):15635–15647PubMedCentralPubMedCrossRefGoogle Scholar
  176. 176.
    Alami NH et al (2014) Axonal transport of TDP-43 mRNA granules is impaired by ALS-causing mutations. Neuron 81(3):536–543PubMedCentralPubMedCrossRefGoogle Scholar

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Authors and Affiliations

  1. 1.Department of Psychiatry and Neuroscience, Institut Universitaire En Santé Mentale de QuébecUniversité LavalQuebecCanada
  2. 2.Department of NeuroscienceUniversity of Texas Southwestern Medical CenterDallasUSA

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