IMR Press / JIN / Volume 20 / Issue 2 / DOI: 10.31083/j.jin2002055
Open Access Commentary
Potential protective role of astrocytes in the pathogenesis of astrocyte-mediated synaptic plasticity of Parkinson’s disease
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1 2017 Clinical Excellence and Innovation Class of the Second Clinical Medical College of Southern Medical University, 510515 Guangzhou, Guangdong Province, China
2 Department of Histology and Embryology, School of Basic Sciences, Southern Medical University, 510515 Guangzhou, Guangdong Province, China
*Correspondence: (Kangrong Lu)
J. Integr. Neurosci. 2021, 20(2), 515–525;
Submitted: 3 February 2021 | Revised: 8 March 2021 | Accepted: 22 April 2021 | Published: 30 June 2021
Copyright: © 2021 The Author(s). Published by IMR Press.
This is an open access article under the CC BY 4.0 license (

Astrocytes are the most abundant glia in the central nervous system that play a significant role in disease. Recently, it roles of synaptic plasticity in neuropathological damages have been questioned whether the structural and functional plasticity of synapses contributes to the pathogenesis of Parkinson’s disease. The regulation of synaptic plasticity by astrocytes has also been widely researched based on astrocytes regulate synaptic plasticity by releasing Adenosine triphosphate, glutamate, and D-serine. We discuss the possible role of astrocytes in the regulation of synaptic plasticity, which may provide a new direction to Parkinson’s disease treatment.

Parkinson's disease (PD)
Synaptic plasticity
1. Introduction

Parkinson’s disease (PD) is a complex neurodegenerative disease [1, 2, 3] with an unknown etiology. The disease is characterized by several pathological manifestations and symptoms that affect bodily systems [4]. Patients with the disease have irreversible functional alterations in the nerve cells, resulting in selective, progressive loss of dopaminergic neurons in the nigrostriatal pathway [1, 2, 5]. The gradual dopaminergic denervation leads to dopamine (DA) deficit in the striatum over the long disease course, causing complex functional deficits within the basal ganglia network. The progressive degeneration retards the voluntary movement, resulting in symptoms of rigidity, tremor, and bradykinesia [6], which worsen as the dopaminergic denervation advances. Nonmotor symptoms, such as psychiatric and cognitive impairment, sleep disorders, autonomic dysfunctions, and gastrointestinal dysfunction [7], have an earlier onset, and more are disabling than motor symptoms [8, 9, 10, 11]. Extensive research over several years has highlighted oxidative stress [12, 13], inflammation [14, 15], accumulation of altered proteins [16, 17], excitotoxicity [18], endoplasmic reticulum stress [19, 20] and mitochondrial dysfunction [21, 22] as potential molecular mechanisms of the disease (The possible mechanisms are shown in Fig. 1). Recently, the significant role of synaptic plasticity in the pathological processes of PD has been identified and validated. Neurons use most of the energy at the synapse. The onset and progression of PD can be explained by the inability of neurons to experience synaptic plasticity [23]. The ultrastructural features of pre-and post-synaptic neuronal elements at the remaining corticostriatal and thalamostriate axo-spinous synapses appeared to undergo complex remodeling, conceding with increased synaptic activity striatum of 1-methyl-4-phenyl-1, 2, 3, 6-tetrahydropyridine (MPTP)-treated Parkinsonian monkeys [24, 25]. In addition, glial reactions play a crucial role in PD. Astrocytes may confer neuroprotective effects by releasing glial transmitters, such as glial-derived neurotrophic factor (GDNF) [26], mesencephalic astrocyte-derived neurotrophic factor (MANF) [27], and ciliary neurotrophic factor (CNTF) [28]. A relative increase in the level of inflammatory cytokines, senescence markers, and metalloproteinases from astrocytes was observed in post-mortem substantia nigra specimens of five patients with PD compared with that in the specimens of five control subjects [29]. Astrocytes produce increased amounts of proinflammatory cytokines in response to inflammatory stimulation by LPS, IL-1β, or TNF-α, which indicates their involvement in neuroinflammatory processes in PD [30, 31]. The addition of α-syn into a primary culture of astrocytes increases IL-6 and TNF-α expressions [32].

Transmitters released by astrocytes that regulate synaptic activity are concluded: (1) ATP [33, 34, 35, 36, 37, 38, 39, 40], (2) Purines [41, 42], (3) Glutamate [43, 44], (4) D-serine [45, 46, 47, 48, 49, 50, 51], (5) NO [52], (6) BDNF [53, 54], (7) S100b [55], (8) Thrombospondin (TSP-1) [56, 57]. In brain diseases, particularly in neurodegenerative conditions, astrocytes become highly reactive, and the process is known as astrogliosis. In this process, astrocytes undergo genetic [58] and morphological modifications [59]. Two subtypes, A1 reactive astrocytes and A2 reactive astrocytes, are formed [60, 61, 62, 63]. A1 reactive astrocytes lose their normal function and gain toxic functions. Co-culturing of human dopaminergic neurons with A1 astrocytes activates neuronal apoptosis, increasing cell death by 25% [64]. In contrast, A2 reactive astrocytes can upregulate many neurotrophic factors [56, 65, 66, 67, 68], contributing to the survival and growth of neurons or synaptic repair.

Fig. 1.

Possible pathogenetic mechanisms of Parkinson’s disease have been studied.

2. Astrocytes and synaptic plasticity

Synaptic plasticity is the ability of synapses to change the strength of their connections. The strength of a synapse refers to the response generated in a postsynapse due to presynaptic activity [69]. Plasticity can occur in a short time and involves molecular mechanisms that influence the synaptic strength and remove, modify, or add synaptic connections [70, 71, 72, 73, 74]. Two forms of classical synaptic plasticity, namely long-term potentiation (LTP) and long-term depression (LTD) [75], depending on the synaptic activation of N-methyl-D-aspartate receptors (NMDARs), are regarded as the foundation of learning and memory [73, 76]. The highly dynamic astrocytes play various roles in the central neural system (CNS) by regulating the blood-brain barrier (BBB) [77], providing structural and metabolic support [78, 79], maintaining ionic homeostasis [80] and secreting neurotrophic factors, such as brain-derived nutritional factor (BDNF) and glial-derived neurotrophic factor (GDNF). It is estimated that multiple neuronal cell somas, 300–600 dendrites and more than 100,000 synapses can be ensheathed by just a single cortical astrocyte in the mouse brain. The number is even larger in primates and humans, reaching 2 million synapses [81, 82]. Hence, the concept of ʻtripartite synapses’ (TS) was proposed, wherein astrocytes integrate process and exchange information with pre-and post-synaptic neuronal elements [83]. In TS, this type of interaction with neurons requires physical contact between astrocytes and synaptic spines. Electron microscopic reconstructions revealed that 57% of the spines in a mature hippocampus are associated with astrocytes [84]. The size and morphology of the spine regulate the synaptic strength and determine the efficiency of synaptic transmission. For instance, large, mushroom-shaped spines contain functional synapses and are more stable, whereas thin filopodia-shaped spines are relatively unstable and nonfunctional [85]. Alterations in astrocytic processes are coordinated with the stabilization of larger spines [86]. Astrocyte processes are essential to the maturation and regulation of newly forming spines. Thus, the contact between astrocytes and spines improves both morphological maturation and the life of spines [87]. Dendritic spines are vulnerable to structural changes during aging and neurologic diseases [88, 89, 90, 91, 92, 93, 94, 95]. Compared with presynaptic terminals, astrocyte processes prefer to localize near dendritic spines [96]. Thrombospondins (TSPs) released by astrocytes are involved in the formation of excitatory synapses [56]. The reductive levels of TSP-1 can change the dendritic spine structure and decrease the number of spine and synapses [97]. Moreover, astrocytes regulate neurite formation and spine density by releasing BDNF in vitro [53].

In addition, astrocytes can secrete various factors such as glutamate, ATP and D-serine, which directly regulate the formation of synapses [79]. Hence, astrocytes are considered to have a central role in the regulation of synaptic plasticity.

2.1 Calcium

Astrocytes are involved in regulating the synaptic function [98, 99]. The passive homeostatic regulation of synaptic function by astrocytes is well recognized. In addition, astrocytes sense synaptic activity and respond to neurotransmitters released by these synapses and, in turn, release gliotransmitters to regulate synaptic transmission and plasticity [100, 101]. The release of Ca2+ underlies this regulatory function of astrocytes. Moreover, several ion channels and membrane receptors expressed by astrocytes allow them to respond to neuronal activity on a millisecond time scale by increasing the intracellular Ca2+ levels [102, 103, 104, 105, 106, 107]. In astrocytes, the basic steps leading to intracellular Ca2+ waves (ICWs) typically involve the activation of G-protein-coupled receptors and phospholipase C and production of IP3, which lead to Ca2+ release from the endoplasmic reticulum (ER) following IP3R activation [108, 109]. Moreover, these cells can transmit such calcium signals to nonstimulated astrocytes located proximally through ICWs [107, 110]. The first glial transmitter found to be dependent on Ca2+ release is glutamate [111]. Afterward, the release of ATP [112] and D-serine [113] was proved to be associated with Ca2+ signaling in astrocytes. In [114], it was indicated that choline could induce LTP production. The realization of this process requires Ca2+ signaling in astrocytes to stimulate glutamate release, thus activating metabotropic glutamate receptors (mGluRs) on neurons.

2.2 Glutamate

Glutamate is converted to glutamine in astrocytes by glutamine synthetase, and glutamine is then recycled to neurons to synthesize glutamate that facilitates the regulation of synaptic transmission [115]. For instance, astrocytes release glutamate, which acts on the pre- [116, 117, 118] and post-synaptic [119, 120, 121, 122] sites to alter synaptic transmission and neuronal excitability at both excitatory [43] and inhibitory [123] synapses. Synapses are dynamic structures in terms of morphology, biochemistry, and function. Most excitatory glutamatergic synapses terminate at dendritic spines undergoing actin-driven movement [124]. An interesting feature is that glutamate can control the structural characteristics of neurons. For instance, glutamate released from the presynaptic terminals maintains the spine stability [125] and leads to small protrusions from the spine when locally applied [125].

Further, astrocytes induce the remodeling of dendritic spines by regulating the extracellular glutamate level [126]. They were revealed that the synaptic strength could be increased by stimulating astrocytes directly. In addition, direct stimulation of single astrocytes in the hypothalamus [127] and hippocampus [117] induces the sustained potentiation of synapses. This type of astrocyte stimulation in the hippocampus increases glutamate release and leads to NMDA-independent LTP production [116]. Furthermore, evidence indicates that astrocytes can modulate LTP by regulating glutamate levels at the synapse and by releasing cofactors of glutamate receptors at the neurons [128].

Moreover, astrocytes can regulate the rate of uptake and release of glutamate [119, 128, 129, 130], and glutamate combines with the AMPA and NMDA receptors on the post-synaptic neurons to induce LTP [130, 131]. This type of functional expression of LTP is due to increased exocytosis of native AMPA receptors to the membrane surface and their recruitment into excitatory synapses [132]. The AMPA-type glutamate receptors (AMPARs) are glutamate-gated ion channels that induce many fast excitatory synaptic transmissions in the brain [133]. Alteration in the number, composition and biophysical properties of the AMPARs in the post-synaptic membrane is the primary mechanism of synaptic strength regulation [134]. During LTP induction, presynaptic input stimulation on a neuron’s postsynapse terminates post-synaptic Ca2+ influx through the NMDARs. Intracellular Ca2+ then causes AMPAR elevation in the postsynapse, which enhances the synaptic strength [135]. The production and regulation of the LTP require the most suitable extracellular glutamate concentration [136], which is maintained by glutamate transporters in astrocytes. Thus, glutamate transport in astrocytes is the key to induce and sustain hippocampal LTP. In addition, glutamate transport in astrocytes is associated with LTD in the hippocampus, amygdala [137], and cerebellum [138]. LTD can be enhanced by blocking the glutamate uptake [137, 139]. However, excess synaptic glutamate levels lead to post-synaptic glutamate receptor overstimulation, resulting in excitotoxic neuronal death [140]. Generally, astrocytes involved in glutamate uptake can control excitotoxic glutamate levels in the brain to avoid excitotoxicity [141]. However, during an excitatory crisis, the potentially protective functions of reactive astrocytes, such as K+ buffering, glutamate uptake, and elimination of free radicals, can be eventually reduced or even destroyed, which could worsen the neural damage [142, 143].

2.3 Adenosine triphosphate

Adenosine triphosphate (ATP), released by astrocytes and neurons in the pathophysiological environment and during neural activity, modulates the synaptic strength and plasticity by activating ionic P2X and metabolic P2Y receptors. ATP released by astrocytes regulates neuronal excitability by stimulating the purine receptors [144]. Moreover, ATP released by astrocytes is degraded to adenosine, which participates in the regulation of activity-dependent heterosynaptic depression at excitatory synapses [41, 100]. Some studies have indicated that ATP released by astrocytes leads to highly efficient glutamatergic synaptic transmission in the paraventricular nucleus; however, it causes low synaptic efficiency in the CA1 region of the hippocampus [144, 145]. ATP from both glia and neurons can activate the post-synaptic P2X, wherein the synaptic currents mediated by the NMDAR decreased significantly [146]. LTP in CA1 neurons was shown to be decreased in P2X4-/- mice [147]. Under pathological conditions, such as brain injury and ischemia, several ATPs released by astrocytes can provoke synaptic plasticity by activating the P2X receptors [148, 149]. This regulation mechanism inhibits LTP by activating Ca2+-dependent NMDRs [150, 151] or induces LTP in the hippocampus [147, 150]. In addition, ATP release was shown to inhibit LTD in [152]. Associating aging with synaptic plasticity, ATP driven by astrocytes significantly weakens the synaptic inhibition in the pyramidal neurons via Ca2+ interaction between neuronal ATP and γ-aminobutyric acid (GABA) receptors. Meanwhile, ATP from astrocytes contributes to the LTP of synaptic plasticity in the neocortex [36].

2.4 D-serine

NMDA-dependent LTP and LTD have been widely studied, and the NMDA receptors are regarded as the crucial components of LTP [135, 153]. The activation of NMDA receptors requires glycine and D-serine in addition to glutamate [46, 154]. A product of serine racemase, D-serine, was first identified in most astrocytes and was considered an endogenous ligand for the glycine-binding site of the NMDA receptor. Addition of D-amino acid oxidase to the cerebellar slices to deaminate D-serine significantly reduces the NMDA receptor-dependent synaptic transmission [155]. D-serine induces synaptogenesis in embryonic cortical neuron culture induced by TGFβR1 activated by astrocyte-derived TGFβ [156].

The effects of astrocyte-derived D-serine on synaptic plasticity and astrocyte-derived amino acids were shown to participate in the LTP [157]. LTP ability of the cultured neurons was recovered by adding D-serine to the astrocyte-conditioned medium [157], and the recovery process required interaction between the astrocytes and neurons. Yang et al. [157] demonstrated that hippocampal neurons cultured with astrocytes could undergo LTP, whereas LTP could not be induced in neurons cultured in astrocyte-conditioned media without astrocyte contact. However, the induction of NMDA-dependent LTP was limited by D-serine released by the hippocampal astrocytes [48]. Christian Henneberger reported that the clamping of internal Ca2+ in astrocytes could block LTP induction in nearby excitatory synapses. However, this LTP blockade can be reversed by exogenous D-serine [48]. The lack of D-serine and disruption of exocytosis in an individual astrocyte blocks local LTP [48].

Moreover, blocking glial cell activation with fluoroacetate, a metabolic inhibitor, was shown to block LTP in the prefrontal cortex due to decreased extrasynaptic D-serine [158]. In contrast to the findings above, [159] reported that the astrocytes release L-serine, converted to D-serine by serine racemase in the neurons, and the latter modulates synaptic plasticity. To confirm this finding, Benneyworth et al. [160] demonstrated that LTP and NMDAR currents are significantly reduced in neuronal serine racemase conditional knockout mice but not in astrocytic serine racemase conditional knockout mice. Therefore, these findings highlight that astrocytes possibly play direct and indirect roles in the D-serine regulation of LTP.

3. Synaptic plasticity in Parkinson disease

PD is a neurodegenerative disorder characterized by the gradual loss of dopaminergic neurons in the substantia nigra pars compacta (SNpc), which decreases DA input to the striatum and hence causes motor dysfunctions [161]. A decrease in LTP activation has been observed in PD models, alleviated by the DA precursor treatment [162, 163]. Short- and long-term changes in corticostriatal synaptic plasticity may be involved in PD [162, 164]. Some researchers have described two classic forms of synaptic plasticity, LTP and LTD, at the corticostriatal synapse in medium spiny neurons in vivo [165, 166]. In the early stages of PD, extracellular DA was shown to be reduced by 40% in PINK1-knockout heterozygous mice; although these mice did not demonstrate any motor symptoms, LTP was alternatively impaired [167]. At the onset of motor dysfunctions in patients with PD, >50% of SNc DA neurons are already lost and DA is reduced by >60% in the striatum [167, 168, 169]. Meanwhile, both LTP and LTD are impaired [167, 170]. Damage to LTP and LTD is concurrent with the deletion of DA and the appearance of PD symptoms. These findings suggest that changes in synaptic plasticity induced by the loss of DA may play an essential role in PD motor dysfunction. According to the classic pathological mechanism of PD, the loss of dopaminergic neurons is the foundation for the complex structural and functional changes in the striatal projection nerves that may also act as a trigger for changes in synaptic plasticity.

Synaptic plasticity occurs in dendritic spines under physiological conditions [171]. Thus, the alterations of spines in PD are related to synaptic plasticity damage. Significant striatal spine loss induced by nigrostriatal dopaminergic lesions has been observed in various PD animal models [25]. The neurotoxin MPTP causes the alternative loss of dopaminergic neurons in the midbrain area of both humans and animals [172, 173]. The MPTP-treated mouse is a common PD animal model [174]. Some studies have found that MPTP treatment can impair LTP [175, 176]. Toy et al. [177] suggested that MPTP injection decreases, whereas exercise increases the dendritic spine density. Motor training has alleviated PD symptoms by promoting synaptic formation and increasing dendritic arborization in the motor cortex [178, 179]. The classic synaptic plasticity forms of LTD and LTP also influence PD symptoms by mediating changes in dendritic spines. In adult mice with dopaminergic nerve degeneration in the motor cortex, LTP loss was accompanied by an increased rate of spine elimination [180].

In addition to motor dysfunctions, cognitive deficits in PD, including learning and memory impairment, seriously impair patients’ life quality and social functions, which cause a great deal of psychological and economic burden to their families. The hippocampus is crucial for spatial and episodic memory formation and novelty detection [181, 182]. Most studies on PD cognitive impairment have focused on the hippocampus. Abnormalities in the hippocampus’s structure and function were observed in sporadic patients and patients with genetic predisposition with PD, suggesting an association of the organ with memory deficits in PD [183, 184, 185, 186]. In addition, the hippocampus is associated with memory deficits [184, 185, 187, 188, 189]. Like motor regulation, cognitive regulation in the disease is based on DA, which conforms to the classical PD pathology. Reportedly, the changed habituation to a new environment in PD animals due to impaired hippocampal LTP can be reversed by the systemic L-DOPA treatment [190]. Moreover, LTP in the CA1 hippocampus is reduced in both genetic and neurotoxic PD models [190]. This alteration of CA1 LTP is accompanied by hippocampal-dependent learning deficits in both 6-OHDA-lesioned and mutant animals [191].

4. Future

PD is a neurodegenerative disease associated with DA depletion and progressive loss of dopaminergic neurons. The primary treatment strategy involves external DA supplementation to alleviate the functional impairment of the basal ganglia induced by DA deficiency. However, none of the currently available PD therapies can slow down or impede the disease progression without dopaminergic neuron recovery. Therefore, the development of effective therapeutic strategies is essential to prevent disease progression. The cell-replacement therapy has been advancing considerably that uses the fetal midbrain tissue as the source of midbrain dopamine neurons (mDAs) for transplantation [192]. However, technical difficulties in obtaining sufficient graft tissues, ethical considerations, and rejection of cells limit the use of this therapy [193, 194, 195]. Alternative cell sources, such as stem cells or reprogrammed cells, have also been considered [196, 197]. The therapeutic effects of embryonic stem cells (ESCs), neural stem cells (NSCs), and bone marrow mesenchymal stem cells (BMSCs) have been widely studied [198, 199, 200].

Nevertheless, ethical consideration, differentiation, and risk of tumor limit the application of NSCs and ESCs. The complex brain environment also interferes with the survival and function of BMSC-derived transplanted neurons [201, 202, 203]. Astrocytes have been studied as potential candidates to improve the hostile brain environment for neural transplantation. Astrocyte cografting can significantly improve the survival rate of dopaminergic neurons and the behavior of PD rats [201]. Stem cell transplantation and cell reprogramming are also the proposed alternatives. In these processes, induced dopaminergic neurons, which are the phenotypically specific and functional DA neurons obtained directly from somatic cells, are used. The induced dopaminergic neurons share morphological characteristics similar to those of resident dopaminergic neurons in the brain. They produce and release DA, survive in the brain, and stimulate the target regions [204, 205, 206, 207, 208]. Astrocytes are considered potential cells for reprogramming due to their particular role in PD. In vitro studies have demonstrated that astrocytes can be converted into functional neurons [209, 210, 211, 212]. Novel strategies, such as direct reprogramming of resident astrocytes in vivo [212], by combining gene- and cell-based therapy have been developed. These strategies can be applied to convert reactive astrocytes in the injured brain and degenerative diseases into functional neurons in vivo and are being developed further [213]. In addition, compensating dysfunctional astrocytes is an attractive choice. Functional astrocytes can be obtained from embryonic glial restricted precursor cells (GRPCs) and human pluripotent stem cells [214, 215, 216]. In vivo transplantation of embryonic GRPC-derived astrocytes could release trophic factors and antioxidants in the striatum, improved behavioral deficits, and restored TH expression in PD rats [217]. Mouse astrocytes could be transformed into dopaminergic neurons in vivo, and this strategy could be used to treat animal models of PD [218].

In addition, as mentioned earlier, damage to LTP and LTD in PD is parallel to DA depletion and the occurrence of PD dyskinesia. Indeed, damage to LTP precedes the onset of PD symptoms. Astrocytes can affect the synaptic plasticity of neurons and improve LTP and LTD by releasing Ca2+, glutamate, ATP, and D-serine. Therefore, astrocytes may be an essential intervention target in PD therapy, and the progression of PD can be considerably delayed, possibly by targeting the astrocytes. In the past few years, significant progress has been made in understanding the molecular mechanism, signal transduction, and function and morphology of astrocytes and laying a technical foundation for astrocytes [218]. Based on the vital role of astrocytes in neuropathy and broad research prospects, we need more tools to explore astrocyte biology from the molecular level to the level of the system to investigate astrocyte therapy mechanisms for PD.

In conclusion, further research is required to explore effective treatment strategies to delay or stop PD progression. Advances in the transplantation technology and regulation of synaptic plasticity by astrocytes can be leveraged to develop new strategies to prevent disease progression at an early stage. We believe that this approach can provide a potential research direction for future studies on PD.


AMPARs, AMPA-type glutamate receptors; ATP, Adenosine triphosphate; BBB, blood-brain barrier; BDNF, brain-derived nutritional factor; BMSCs, bone-marrow mesenchymal stem cells; CNS, central neural system; CNTF, ciliary neurotrophic factor; DA, dopamine; ER, endoplasmic reticulum; GABA, γ-aminobutyric acid; GRPCs, glial restricted precursor cells; ESCs, embryonic stem cells; GDNF, glial-derived neurotrophic factor; ICWs, intracellular Ca2+ waves; LTD, long-term depression; LTP, long-term potentiation; MANF, mesencephalic astrocyte-derived neurotrophic factor; mGluRs, metabotropic glutamate receptors; MPTP, 1-methyl-4-phenyl-1, 2, 3, 6-tetrahydropyridine; NMDARs, N-methyl-D-aspartate receptors; NSCs, neural stem cells; PD, Parkinson disease; TS, tripartite synapses; TSPs, Thrombospondins.

Author contributions

YQZ and KRL read the literature together and wrote the review. YQZ is the first author. KRL is the corresponding author.

Ethics approval and consent to participate

The content of the article does not involve any experiments, and there are no ethical issues.


We thank three anonymous reviewers for their excellent criticism of the article.


Regulation of GABAA-mediated tonic inhibition on cognitive impairment in Parkinson’s disease and its mechanism 2018A0303130259.

Conflict of interest

The authors declare no conflict of interest.

Hirsch EC, Jenner P, Przedborski S. Pathogenesis of Parkinson’s disease. Movement Disorders. 2013; 28: 24–30.
Obeso JA, Rodriguez-Oroz MC, Goetz CG, Marin C, Kordower JH, Rodriguez M, et al. Missing pieces in the Parkinson’s disease puzzle. Nature Medicine. 2010; 16: 653–661.
Toulouse A, Sullivan AM. Progress in Parkinson’s disease—where do we stand? Progress in Neurobiology. 2008; 85: 376–392.
Braak H, Del Tredici K. Invited article: nervous system pathology in sporadic Parkinson disease. Neurology. 2008; 70: 1916–1925.
Damier P, Hirsch EC, Agid Y, Graybiel AM. The substantia nigra of the human brain. II. Patterns of loss of dopamine-containing neurons in Parkinson’s disease. Brain. 1999; 122: 1437–1448.
Blandini F, Nappi G, Tassorelli C, Martignoni E. Functional changes of the basal ganglia circuitry in Parkinson’s disease. Progress in Neurobiology. 2000; 62: 63–88.
Napier TC, Corvol J, Grace AA, Roitman JD, Rowe J, Voon V, et al. Linking neuroscience with modern concepts of impulse control disorders in Parkinson’s disease. Movement Disorders. 2015; 30: 141–149.
Zesiewicz TA, Sullivan KL, Hauser RA. Nonmotor symptoms of Parkinson’s disease. Expert Review of Neurotherapeutics. 2006; 6: 1811–1822.
O’Sullivan SS, Williams DR, Gallagher DA, Massey LA, Silveira-Moriyama L, Lees AJ. Nonmotor symptoms as presenting complaints in Parkinson’s disease: a clinicopathological study. Movement Disorders. 2008; 23: 101–106.
Lim S, Lang AE. The nonmotor symptoms of Parkinson’s disease-an overview. Movement Disorders. 2010; 25: S123–S130.
Gallagher DA, Lees AJ, Schrag A. What are the most important nonmotor symptoms in patients with Parkinson’s disease and are we missing them? Movement Disorders. 2010; 25: 2493–2500.
Appel SH, Beers DR, Henkel JS. T cell-microglial dialogue in Parkinson’s disease and amyotrophic lateral sclerosis: are we listening? Trends in Immunology. 2010; 31: 7–17.
Wong SS, Li RH, Stadlin A. Oxidative stress induced by MPTP and MPP (+): selective vulnerability of cultured mouse astrocytes. Brain Research. 1999; 836: 237–244.
Koprich JB, Reske-Nielsen C, Mithal P, Isacson O. Neuroinflammation mediated by IL-1β increases susceptibility of dopamine neurons to degeneration in an animal model of Parkinson’s disease. Journal of Neuroinflammation. 2008; 5: 8.
Dawson TM, Ko HS, Dawson VL. Genetic animal models of Parkinson’s disease. Neuron. 2010; 66: 646–661.
Kakoty V, K C S, Tang R, Yang CH, Dubey SK, Taliyan R. Fibroblast growth factor 21 and autophagy: a complex interplay in Parkinson disease. Biomedicine & Pharmacotherapy. 2020; 127: 110145.
Emamzadeh FN. Role of apolipoproteins and α-synuclein in Parkinson’s disease. Journal of Molecular Neuroscience. 2017; 62: 344–355.
Iovino L, Tremblay ME, Civiero L. Glutamate-induced excitotoxicity in Parkinson’s disease: the role of glial cells. Journal of Pharmacological Sciences. 2020; 144: 151–164.
Rozpędek-Kamińska W, Siwecka N, Wawrzynkiewicz A, Wojtczak R, Pytel D, Diehl JA, et al. The PERK-dependent molecular mechanisms as a novel therapeutic target for neurodegenerative diseases. International Journal of Molecular Sciences. 2020; 21: 2108.
Shah A, Chao J, Legido-Quigley C, Chang RC. Oxyresveratrol exerts ATF4- and Grp78-mediated neuroprotection against endoplasmic reticulum stress in experimental Parkinson’s disease. Nutritional Neuroscience. 2021; 24: 181–196.
Malpartida AB, Williamson M, Narendra DP, Wade-Martins R, Ryan BJ. Mitochondrial dysfunction and mitophagy in Parkinson’s disease: from mechanism to therapy. Trends in Biochemical Sciences. 2021; 46: 329–343.
Foo ASC, Soong TW, Yeo TT, Lim KL. Mitochondrial dysfunction and Parkinson’s disease-near-infrared photobiomodulation as a potential therapeutic strategy. Frontiers in Aging Neuroscience. 2020; 12: 89.
Calabresi P, Picconi B, Parnetti L, Di Filippo M. A convergent model for cognitive dysfunctions in Parkinson’s disease: the critical dopamine-acetylcholine synaptic balance. Lancet Neurology. 2006; 5: 974–983.
Villalba RM, Smith Y. Differential structural plasticity of corticostriatal and thalamostriatal axo-spinous synapses in MPTP-treated Parkinsonian monkeys. Journal of Comparative Neurology. 2011; 519: 989–1005.
Villalba RM, Smith Y. Striatal spine plasticity in Parkinson’s disease. Frontiers in Neuroanatomy. 2010; 4: 133.
Aron L, Klein R. Repairing the parkinsonian brain with neurotrophic factors. Trends in Neurosciences. 2011; 34: 88–100.
Shen Y, Sun A, Wang Y, Cha D, Wang H, Wang F, et al. Upregulation of mesencephalic astrocyte-derived neurotrophic factor in glial cells is associated with ischemia-induced glial activation. Journal of Neuroinflammation. 2012; 9: 254.
McGregor MJ, Cohen M, Stocks-Rankin C, Cox MB, Salomons K, McGrail KM, et al. Complaints in for-profit, non-profit and public nursing homes in two Canadian provinces. Open Medicine. 2011; 5: e183–e192.
Chinta SJ, Woods G, Demaria M, Rane A, Zou Y, McQuade A, et al. Cellular senescence is induced by the environmental neurotoxin paraquat and contributes to neuropathology linked to Parkinson’s disease. Cell Reports. 2018; 22: 930–940.
Saijo K, Winner B, Carson CT, Collier JG, Boyer L, Rosenfeld MG, et al. A Nurr1/CoREST pathway in microglia and astrocytes protects dopaminergic neurons from inflammation-induced death. Cell. 2009; 137: 47–59.
Tanaka T, Kai S, Matsuyama T, Adachi T, Fukuda K, Hirota K. General anesthetics inhibit LPS-induced IL-1β expression in glial cells. PLoS ONE. 2013; 8: e82930.
Fellner L, Irschick R, Schanda K, Reindl M, Klimaschewski L, Poewe W, et al. Toll-like receptor 4 is required for α-synuclein dependent activation of microglia and astroglia. Glia. 2013; 61: 349–360.
Coco S, Calegari F, Pravettoni E, Pozzi D, Taverna E, Rosa P, et al. Storage and release of ATP from astrocytes in culture. Journal of Biological Chemistry. 2003; 278: 1354–1362.
Zhang J, Wang H, Ye C, Ge W, Chen Y, Jiang Z, et al. ATP released by astrocytes mediates glutamatergic activity-dependent heterosynaptic suppression. Neuron. 2003; 40: 971–982.
Chen J, Tan Z, Zeng L, Zhang X, He Y, Gao W, et al. Heterosynaptic long-term depression mediated by ATP released from astrocytes. Glia. 2013; 61: 178–191.
Lalo U, Rasooli-Nejad S, Pankratov Y. Exocytosis of gliotransmitters from cortical astrocytes: implications for synaptic plasticity and aging. Biochemical Society Transactions. 2014; 42: 1275–1281.
Lalo U, Palygin O, Verkhratsky A, Grant SGN, Pankratov Y. ATP from synaptic terminals and astrocytes regulates NMDA receptors and synaptic plasticity through PSD-95 multi-protein complex. Scientific Reports. 2016; 6: 33609.
Boué-Grabot E, Pankratov Y. Modulation of central synapses by astrocyte-released ATP and postsynaptic P2X receptors. Neural Plasticity. 2017; 2017: 9454275.
Crosby KM, Murphy-Royal C, Wilson SA, Gordon GR, Bains JS, Pittman QJ. Cholecystokinin switches the plasticity of GABA synapses in the dorsomedial hypothalamus via astrocytic ATP release. The Journal of Neuroscience. 2018; 38: 8515–8525.
Lalo U, Bogdanov A, Pankratov Y. Age- and experience-related plasticity of ATP-mediated signaling in the neocortex. Frontiers in Cellular Neuroscience. 2019; 13: 242.
Pascual O, Casper KB, Kubera C, Zhang J, Revilla-Sanchez R, Sul J, et al. Astrocytic purinergic signaling coordinates synaptic networks. Science. 2005; 310: 113–116.
Ikeda H, Tsuda M, Inoue K, Murase K. Long-term potentiation of neuronal excitation by neuron-glia interactions in the rat spinal dorsal horn. European Journal of Neuroscience. 2007; 25: 1297–1306.
Bezzi P, Carmignoto G, Pasti L, Vesce S, Rossi D, Rizzini BL, et al. Prostaglandins stimulate calcium-dependent glutamate release in astrocytes. Nature. 1998; 391: 281–285.
Falcón-Moya R, Pérez-Rodríguez M, Prius-Mengual J, Andrade-Talavera Y, Arroyo-García LE, Pérez-Artés R, et al. Astrocyte-mediated switch in spike timing-dependent plasticity during hippocampal development. Nature Communications. 2020; 11: 4388.
Schell MJ, Molliver ME, Snyder SH. D-serine, an endogenous synaptic modulator: localization to astrocytes and glutamate-stimulated release. Proceedings of the National Academy of Sciences, USA. 1995; 92: 3948–3952.
Wolosker H, Blackshaw S, Snyder SH. Serine racemase: a glial enzyme synthesizing D-serine to regulate glutamate-N-methyl-D-aspartate neurotransmission. Proceedings of the National Academy of Sciences, USA. 1999; 96: 13409–13414.
Panatier A, Theodosis DT, Mothet J, Touquet B, Pollegioni L, Poulain DA, et al. Glia-derived D-serine controls NMDA receptor activity and synaptic memory. Cell. 2006; 125: 775–784.
Henneberger C, Papouin T, Oliet SHR, Rusakov DA. Long-term potentiation depends on release of D-serine from astrocytes. Nature. 2010; 463: 232–236.
Bodner O, Radzishevsky I, Foltyn VN, Touitou A, Valenta AC, Rangel IF, et al. D-serine signaling and NMDAR-mediated synaptic plasticity are regulated by system a-type of glutamineD-serine dual transporters. Journal of Neuroscience. 2020; 40: 6489–6502.
Hardingham GE. Targeting synaptic NMDA receptor co-agonism as a therapy for Alzheimer’s disease? Cell Metabolism. 2020; 31: 439–440.
ong JM, Folorunso OO, Barragan EV, Berciu C, Harvey TL, Coyle JT, et al. Postsynaptic serine racemase regulates NMDA receptor function. The Journal of Neuroscience. 2020; 40: 9564–9575.
Zhuo M, Small SA, Kandel ER, Hawkins RD. Nitric oxide and carbon monoxide produce activity-dependent long-term synaptic enhancement in hippocampus. Science. 1993; 260: 1946–1950.
de Pins B, Cifuentes-Díaz C, Farah AT, López-Molina L, Montalban E, Sancho-Balsells A, et al. Conditional BDNF delivery from astrocytes rescues memory deficits, spine density, and synaptic properties in the 5xFAD mouse model of Alzheimer disease. Journal of Neuroscience. 2019; 39: 2441–2458.
Lalo U, Bogdanov A, Moss GW, Pankratov Y. Astroglia-derived BDNF and MSK-1 mediate experience- and diet-dependent synaptic plasticity. Brain Sciences. 2020; 10.
Nishiyama H, Knopfel T, Endo S, Itohara S. Glial protein S100B modulates long-term neuronal synaptic plasticity. Proceedings of the National Academy of Sciences, USA. 2002; 99: 4037–4042.
Christopherson KS, Ullian EM, Stokes CCA, Mullowney CE, Hell JW, Agah A, et al. Thrombospondins are astrocyte-secreted proteins that promote CNS synaptogenesis. Cell. 2005; 120: 421–433.
Tran MD, Neary JT. Purinergic signaling induces thrombospondin-1 expression in astrocytes. Proceedings of the National Academy of Sciences, USA. 2006; 103: 9321–9326.
Sofroniew MV. Molecular dissection of reactive astrogliosis and glial scar formation. Trends in Neurosciences. 2009; 32: 638–647.
Wilhelmsson U, Bushong EA, Price DL, Smarr BL, Phung V, Terada M, et al. Redefining the concept of reactive astrocytes as cells that remain within their unique domains upon reaction to injury. Proceedings of the National Academy of Sciences, USA. 2006; 103: 17513–17518.
Karimi-Abdolrezaee S, Billakanti R. Reactive astrogliosis after spinal cord injury-beneficial and detrimental effects. Molecular Neurobiology. 2012; 46: 251–264.
Li X, Yang B, Xiao Z, Zhao Y, Han S, Yin Y, et al. Comparison of subacute and chronic scar tissues after complete spinal cord transection. Experimental Neurology. 2018; 306: 132–137.
Teh DBL, Prasad A, Jiang W, Ariffin MZ, Khanna S, Belorkar A, et al. Transcriptome analysis reveals neuroprotective aspects of human reactive astrocytes induced by interleukin 1β. Scientific Reports. 2017; 7: 13988.
Robel S, Buckingham SC, Boni JL, Campbell SL, Danbolt NC, Riedemann T, et al. Reactive astrogliosis causes the development of spontaneous seizures. Journal of Neuroscience. 2015; 35: 3330–3345.
Liddelow SA, Guttenplan KA, Clarke LE, Bennett FC, Bohlen CJ, Schirmer L, et al. Neurotoxic reactive astrocytes are induced by activated microglia. Nature. 2017; 541: 481–487.
Zador Z, Stiver S, Wang V, Manley GT. Role of aquaporin-4 in cerebral edema and stroke. Handbook of Experimental Pharmacology. 2009; 159–170.
Hayakawa K, Pham LD, Arai K, Lo EH. Reactive astrocytes promote adhesive interactions between brain endothelium and endothelial progenitor cells via HMGB1 and beta-2 integrin signaling. Stem Cell Research. 2014; 12: 531–538.
Arregui L, Benítez JA, Razgado LF, Vergara P, Segovia J. Adenoviral astrocyte-specific expression of BDNF in the striata of mice transgenic for Huntington’s disease delays the onset of the motor phenotype. Cellular and Molecular Neurobiology. 2011; 31: 1229–1243.
Wang L, Lin F, Wang J, Wu J, Han R, Zhu L, et al. Truncated N-terminal huntingtin fragment with expanded-polyglutamine (htt552-100Q) suppresses brain-derived neurotrophic factor transcription in astrocytes. Acta Biochimica Et Biophysica Sinica. 2012; 44: 249–258.
Atwood HL, Karunanithi S. Diversification of synaptic strength: presynaptic elements. Nature Reviews Neuroscience. 2002; 3: 497–516.
Lüscher C, Malenka RC. NMDA receptor-dependent long-term potentiation and long-term depression (LTP/LTD). Cold Spring Harbor Perspectives in Biology. 2012; 4: a005710.
Katz LC, Shatz CJ. Synaptic activity and the construction of cortical circuits. Science. 1996; 274: 1133–1138.
Holtmaat AJGD, Trachtenberg JT, Wilbrecht L, Shepherd GM, Zhang X, Knott GW, et al. Transient and persistent dendritic spines in the neocortex in vivo. Neuron. 2005; 45: 279–291.
Bliss TVP, Collingridge GL. A synaptic model of memory: long-term potentiation in the hippocampus. Nature. 1993; 361: 31–39.
Bear MF. Homosynaptic long-term depression: a mechanism for memory? Proceedings of the National Academy of Sciences, USA. 1999; 96: 9457–9458.
Mulkey RM, Endo S, Shenolikar S, Malenka RC. Involvement of a calcineurin/ inhibitor-1 phosphatase cascade in hippocampal long-term depression. Nature. 1994; 369: 486–488.
Bear MF, Abraham WC. Long-term depression in hippocampus. Annual Review of Neuroscience. 1996; 19: 437–462.
Abbott NJ, Rönnbäck L, Hansson E. Astrocyte-endothelial interactions at the blood-brain barrier. Nature Reviews. Neuroscience. 2006; 7: 41–53.
Scharfman HE, Binder DK. Aquaporin-4 water channels and synaptic plasticity in the hippocampus. Neurochemistry International. 2013; 63: 702–711.
Barker AJ, Ullian EM. Astrocytes and synaptic plasticity. Neuroscientist. 2010; 16: 40–50.
Simard M, Nedergaard M. The neurobiology of glia in the context of water and ion homeostasis. Neuroscience. 2004; 129: 877–896.
Verkhratsky A. Physiology of neuronal-glial networking. Neurochemistry International. 2010; 57: 332–343.
Oberheim NA, Wang X, Goldman S, Nedergaard M. Astrocytic complexity distinguishes the human brain. Trends in Neurosciences. 2006; 29: 547–553.
Araque A, Sanzgiri RP, Parpura V, Haydon PG. Astrocyte-induced modulation of synaptic transmission. Canadian Journal of Physiology and Pharmacology. 1999; 77: 699–706.
Ventura R, Harris KM. Three-dimensional relationships between hippocampal synapses and astrocytes. Journal of Neuroscience. 1999; 19: 6897–6906.
Yoshihara Y, De Roo M, Muller D. Dendritic spine formation and stabilization. Current Opinion in Neurobiology. 2009; 19: 146–153.
Haber M. Cooperative astrocyte and dendritic spine dynamics at hippocampal excitatory synapses. Journal of Neuroscience. 2006; 26: 8881–8891.
Nishida H, Okabe S. Direct astrocytic contacts regulate local maturation of dendritic spines. Journal of Neuroscience. 2007; 27: 331–340.
Ferrante RJ, Kowall NW, Richardson EP. Proliferative and degenerative changes in striatal spiny neurons in Huntington’s disease: a combined study using the section-Golgi method and calbindin D28k immunocytochemistry. Journal of Neuroscience. 1991; 11: 3877–3887.
Dickstein DL, Weaver CM, Luebke JI, Hof PR. Dendritic spine changes associated with normal aging. Neuroscience. 2013; 251: 21–32.
Mostany R, Anstey JE, Crump KL, Maco B, Knott G, Portera-Cailliau C. Altered synaptic dynamics during normal brain aging. Journal of Neuroscience. 2013; 33: 4094–4104.
Le Y, Liu S, Peng M, Tan C, Liao Q, Duan K, et al. Aging differentially affects the loss of neuronal dendritic spine, neuroinflammation and memory impairment at rats after surgery. PLoS ONE. 2014; 9: e106837.
Sato Y, Okabe S. Nano-scale analysis of synapse morphology in an autism mouse model with 15q11-13 copy number variation using focused ion beam milling and scanning electron microscopy. Microscopy. 2019; 68: 122–132.
Pereira AC, Lambert HK, Grossman YS, Dumitriu D, Waldman R, Jannetty SK, et al. Glutamatergic regulation prevents hippocampal-dependent age-related cognitive decline through dendritic spine clustering. Proceedings of the National Academy of Sciences, USA. 2014; 111: 18733–18738.
Dorostkar MM, Zou C, Blazquez-Llorca L, Herms J. Analyzing dendritic spine pathology in Alzheimer’s disease: problems and opportunities. Acta Neuropathologica. 2015; 130: 1–19.
Herms J, Dorostkar MM. Dendritic spine pathology in neurodegenerative diseases. Annual Review of Pathology. 2016; 11: 221–250.
Lehre KP, Rusakov DA. Asymmetry of glia near central synapses favors presynaptically directed glutamate escape. Biophysical Journal. 2002; 83: 125–134.
Eroglu C, Allen NJ, Susman MW, O’Rourke NA, Park CY, Ozkan E, et al. Gabapentin receptor α2δ-1 is a neuronal thrombospondin receptor responsible for excitatory CNS synaptogenesis. Cell. 2009; 139: 380–392.
Volterra A, Meldolesi J. Astrocytes, from brain glue to communication elements: the revolution continues. Nature Reviews Neuroscience. 2005; 6: 626–640.
Eroglu C, Barres BA. Regulation of synaptic connectivity by glia. Nature. 2010; 468: 223–231.
Serrano A, Haddjeri N, Lacaille J, Robitaille R. GABAergic network activation of glial cells underlies hippocampal heterosynaptic depression. Journal of Neuroscience. 2006; 26: 5370–5382.
Di Castro MA, Chuquet J, Liaudet N, Bhaukaurally K, Santello M, Bouvier D, et al. Local Ca2+ detection and modulation of synaptic release by astrocytes. Nature Neuroscience. 2011; 14: 1276–1284.
Barres BA, Koroshetz WJ, Chun LL, Corey DP. Ion channel expression by white matter glia: the type-1 astrocyte. Neuron. 1990; 5: 527–544.
MacVicar BA, Tse FWY. Norepinephrine and cyclic adenosine 3’:5’-cyclic monophosphate enhance a nifedipine-sensitive calcium current in cultured rat astrocytes. Glia. 1988; 1: 359–365.
Marrero H, Astion ML, Coles JA, Orkand RK. Facilitation of voltage-gated ion channels in frog neuroglia by nerve impulses. Nature. 1989; 339: 378–380.
McCarthy KD, Salm AK. Pharmacologically-distinct subsets of astroglia can be identified by their calcium response to neuroligands. Neuroscience. 1991; 41: 325–333.
Salm AK, McCarthy KD. Norepinephrine-evoked calcium transients in cultured cerebral type 1 astroglia. Glia. 1990; 3: 529–538.
Usowicz MM, Gallo V, Cull-Candy SG. Multiple conductance channels in type-2 cerebellar astrocytes activated by excitatory amino acids. Nature. 1989; 339: 380–383.
Golovina VA, Blaustein MP. Unloading and refilling of two classes of spatially resolved endoplasmic reticulum Ca (2+) stores in astrocytes. Glia. 2000; 31: 15–28.
Scemes E. Components of astrocytic intercellular calcium signaling. Molecular Neurobiology. 2000; 22: 167–179.
Charles AC, Merrill JE, Dirksen ER, Sanderson MJ. Intercellular signaling in glial cells: calcium waves and oscillations in response to mechanical stimulation and glutamate. Neuron. 1991; 6: 983–992.
Parpura V, Basarsky TA, Liu F, Jeftinija K, Jeftinija S, Haydon PG. Glutamate-mediated astrocyte-neuron signalling. Nature. 1994; 369: 744–747.
Bal-Price A, Moneer Z, Brown GC. Nitric oxide induces rapid, calcium-dependent release of vesicular glutamate and ATP from cultured rat astrocytes. Glia. 2002; 40: 312–323.
Mothet J, Pollegioni L, Ouanounou G, Martineau M, Fossier P, Baux G. Glutamate receptor activation triggers a calcium-dependent and SNARE protein-dependent release of the gliotransmitter D-serine. Proceedings of the National Academy of Sciences, USA. 2005; 102: 5606–5611.
Navarrete M, Perea G, Fernandez de Sevilla D, Gómez-Gonzalo M, Núñez A, Martín ED, et al. Astrocytes mediate in vivo cholinergic-induced synaptic plasticity. PLoS Biology. 2012; 10: e1001259.
Danbolt NC. Glutamate uptake. Progress in Neurobiology. 2001; 65: 1–105.
Perea G, Araque A. Astrocytes potentiate transmitter release at single hippocampal synapses. Science. 2007; 317: 1083–1086.
Jourdain P, Bergersen LH, Bhaukaurally K, Bezzi P, Santello M, Domercq M, et al. Glutamate exocytosis from astrocytes controls synaptic strength. Nature Neuroscience. 2007; 10: 331–339.
Fiacco TA, McCarthy KD. Intracellular astrocyte calcium waves in situ increase the frequency of spontaneous AMPA receptor currents in CA1 pyramidal neurons. Journal of Neuroscience. 2004; 24: 722–732.
Perea G, Araque A. Properties of synaptically evoked astrocyte calcium signal reveal synaptic information processing by astrocytes. Journal of Neuroscience. 2005; 25: 2192–2203.
Fellin T, Carmignoto G. Neurone-to-astrocyte signalling in the brain represents a distinct multifunctional unit. Journal of Physiology. 2004; 559: 3–15.
Angulo MC, Kozlov AS, Charpak S, Audinat E. Glutamate released from glial cells synchronizes neuronal activity in the hippocampus. Journal of Neuroscience. 2004; 24: 6920–6927.
Parri HR, Gould TM, Crunelli V. Spontaneous astrocytic Ca2+ oscillations in situ drive NMDAR-mediated neuronal excitation. Nature Neuroscience. 2001; 4: 803–812.
Kang J, Jiang L, Goldman SA, Nedergaard M. Astrocyte-mediated potentiation of inhibitory synaptic transmission. Nature Neuroscience. 1998; 1: 683–692.
Bourne JN, Harris KM. Balancing structure and function at hippocampal dendritic spines. Annual Review of Neuroscience. 2008; 31: 47–67.
McKinney RA, Capogna M, Dürr R, Gähwiler BH, Thompson SM. Miniature synaptic events maintain dendritic spines via AMPA receptor activation. Nature Neuroscience. 1999; 2: 44–49.
Reagan LP, Rosell DR, Wood GE, Spedding M, Muñoz C, Rothstein J, et al. Chronic restraint stress up-regulates GLT-1 mRNA and protein expression in the rat hippocampus: reversal by tianeptine. Proceedings of the National Academy of Sciences, USA. 2004; 101: 2179–2184.
Gordon GRJ, Iremonger KJ, Kantevari S, Ellis-Davies GCR, MacVicar BA, Bains JS. Astrocyte-mediated distributed plasticity at hypothalamic glutamate synapses. Neuron. 2009; 64: 391–403.
Ivens S, Çalışkan G, Papageorgiou I, Cesetti T, Malich A, Kann O, et al. Persistent increase in ventral hippocampal long‐term potentiation by juvenile stress: a role for astrocytic glutamine synthetase. Glia. 2019; 67: 2279–2293.
Devaraju P, Sun M, Myers TL, Lauderdale K, Fiacco TA. Astrocytic group I mGluR-dependent potentiation of astrocytic glutamate and potassium uptake. Journal of Neurophysiology. 2013; 109: 2404–2414.
Bonansco C, Couve A, Perea G, Ferradas CÁ, Roncagliolo M, Fuenzalida M. Glutamate released spontaneously from astrocytes sets the threshold for synaptic plasticity. European Journal of Neuroscience. 2011; 33: 1483–1492.
Larson J, Munkácsy E. Theta-burst LTP. Brain Research. 2015; 1621: 38–50.
Lu W, Man H, Ju W, Trimble WS, MacDonald JF, Wang YT. Activation of synaptic NMDA receptors induces membrane insertion of new AMPA receptors and LTP in cultured hippocampal neurons. Neuron. 2001; 29: 243–254.
Shepherd JD, Huganir RL. The cell biology of synaptic plasticity: AMPA receptor trafficking. Annual Review of Cell and Developmental Biology. 2007; 23: 613–643.
Huganir RL, Nicoll RA. AMPARs and synaptic plasticity: the last 25 years. Neuron. 2013; 80: 704–717.
Malenka RC, Bear MF. LTP and LTD: an embarrassment of riches. Neuron. 2004; 44: 5–21.
Katagiri H, Tanaka K, Manabe T. Requirement of appropriate glutamate concentrations in the synaptic cleft for hippocampal LTP induction. European Journal of Neuroscience. 2001; 14: 547–553.
Omrani A, Melone M, Bellesi M, Safiulina V, Aida T, Tanaka K, et al. Up-regulation of GLT-1 severely impairs LTD at mossy fibre-CA3 synapses. Journal of Physiology. 2009; 587: 4575–4588.
Brasnjo G, Otis TS. Neuronal glutamate transporters control activation of postsynaptic metabotropic glutamate receptors and influence cerebellar long-term depression. Neuron. 2001; 31: 607–616.
Yang C, Huang C, Hsu K. Behavioral stress enhances hippocampal CA1 long-term depression through the blockade of the glutamate uptake. Journal of Neuroscience. 2005; 25: 4288–4293.
Karki P, Hong P, Johnson J, Pajarillo E, Son D, Aschner M, et al. Arundic acid increases expression and function of astrocytic glutamate transporter EAAT1 via the ERK, Akt, and NF-κB pathways. Molecular Neurobiology. 2018; 55: 5031–5046.
Anderson CM, Swanson RA. Astrocyte glutamate transport: review of properties, regulation, and physiological functions. Glia. 2000; 32: 1–14.
Takahashi M, Billups B, Rossi D, Sarantis M, Hamann M, Attwell D. The role of glutamate transporters in glutamate homeostasis in the brain. Journal of Experimental Biology. 1997; 200: 401–409.
Parpura V, Scemes E, Spray DC. Mechanisms of glutamate release from astrocytes: gap junction “hemichannels”, purinergic receptors and exocytotic release. Neurochemistry International. 2004; 45: 259–264.
Gordon GRJ, Baimoukhametova DV, Hewitt SA, Rajapaksha WRAKJS, Fisher TE, Bains JS. Norepinephrine triggers release of glial ATP to increase postsynaptic efficacy. Nature Neuroscience. 2005; 8: 1078–1086.
Pougnet J, Toulme E, Martinez A, Choquet D, Hosy E, Boué-Grabot E. ATP P2X receptors downregulate AMPA receptor trafficking and postsynaptic efficacy in hippocampal neurons. Neuron. 2014; 83: 417–430.
Migaud M, Charlesworth P, Dempster M, Webster LC, Watabe AM, Makhinson M, et al. Enhanced long-term potentiation and impaired learning in mice with mutant postsynaptic density-95 protein. Nature. 1998; 396: 433–439.
Sim JA, Chaumont S, Jo J, Ulmann L, Young MT, Cho K, et al. Altered hippocampal synaptic potentiation in P2X4 knock-out mice. Journal of Neuroscience. 2006; 26: 9006–9009.
Pankratov Y, Lalo U, Krishtal OA, Verkhratsky A. P2X receptors and synaptic plasticity. Neuroscience. 2009; 158: 137–148.
Yamazaki Y, Kaneko K, Fujii S, Kato H, Ito K. Long-term potentiation and long-term depression induced by local application of ATP to hippocampal CA1 neurons of the guinea pig. Hippocampus. 2003; 13: 81–92.
Wang Y, Mackes J, Chan S, Haughey NJ, Guo Z, Ouyang X, et al. Impaired long-term depression in P2X3 deficient mice is not associated with a spatial learning deficit. Journal of Neurochemistry. 2006; 99: 1425–1434.
Pankratov YV, Lalo UV, Krishtal OA. Role for P2X receptors in long-term potentiation. Journal of Neuroscience. 2002; 22: 8363–8369.
Guzman SJ, Schmidt H, Franke H, Krügel U, Eilers J, Illes P, et al. P2Y1 receptors inhibit long-term depression in the prefrontal cortex. Neuropharmacology. 2010; 59: 406–415.
Cavazzini M, Bliss T, Emptage N. Ca2+ and synaptic plasticity. Cell Calcium. 2005; 38: 355–367.
Johnson JW, Ascher P. Glycine potentiates the NMDA response in cultured mouse brain neurons. Nature. 1987; 325: 529–531.
Mothet JP, Parent AT, Wolosker H, Brady RO, Linden DJ, Ferris CD, et al. D-serine is an endogenous ligand for the glycine site of the N-methyl-D-aspartate receptor. Proceedings of the National Academy of Sciences, USA. 2000; 97: 4926–4931.
Diniz LP, Almeida JC, Tortelli V, Vargas Lopes C, Setti-Perdigão P, Stipursky J, et al. Astrocyte-induced synaptogenesis is mediated by transforming growth factor β signaling through modulation of D-serine levels in cerebral cortex neurons. Journal of Biological Chemistry. 2012; 287: 41432–41445.
Yang Y, Ge W, Chen Y, Zhang Z, Shen W, Wu C, et al. Contribution of astrocytes to hippocampal long-term potentiation through release of D-serine. Proceedings of the National Academy of Sciences, USA. 2003; 100: 15194–15199.
Fossat P, Turpin FR, Sacchi S, Dulong J, Shi T, Rivet J, et al. Glial D-serine gates NMDA receptors at excitatory synapses in prefrontal cortex. Cerebral Cortex. 2012; 22: 595–606.
Wolosker H, Balu DT, Coyle JT. The rise and fall of the D-serine-mediated gliotransmission hypothesis. Trends in Neurosciences. 2016; 39: 712–721.
Benneyworth MA, Li Y, Basu AC, Bolshakov VY, Coyle JT. Cell selective conditional null mutations of serine racemase demonstrate a predominate localization in cortical glutamatergic neurons. Cellular and Molecular Neurobiology. 2012; 32: 613–624.
Dauer W, Przedborski S. Parkinson’s disease: mechanisms and models. Neuron. 2003; 39: 889–909.
Picconi B, Centonze D, Håkansson K, Bernardi G, Greengard P, Fisone G, et al. Loss of bidirectional striatal synaptic plasticity in L-DOPA-induced dyskinesia. Nature Neuroscience. 2003; 6: 501–506.
Goldberg MS, Pisani A, Haburcak M, Vortherms TA, Kitada T, Costa C, et al. Nigrostriatal dopaminergic deficits and hypokinesia caused by inactivation of the familial Parkinsonism-linked gene DJ-1. Neuron. 2005; 45: 489–496.
Gubellini P, Picconi B, Bari M, Battista N, Calabresi P, Centonze D, et al. Experimental parkinsonism alters endocannabinoid degradation: implications for striatal glutamatergic transmission. Journal of Neuroscience. 2002; 22: 6900–6907.
Reynolds JN, Hyland BI, Wickens JR. A cellular mechanism of reward-related learning. Nature. 2001; 413: 67–70.
Charpier S, Deniau JM. In vivo activity-dependent plasticity at cortico-striatal connections: evidence for physiological long-term potentiation. Proceedings of the National Academy of Sciences, USA. 1997; 94: 7036–7040.
Madeo G, Schirinzi T, Martella G, Latagliata EC, Puglisi F, Shen J, et al. PINK1 heterozygous mutations induce subtle alterations in dopamine-dependent synaptic plasticity. Movement Disorders. 2014; 29: 41–53.
Hilker R, Schweitzer K, Coburger S, Ghaemi M, Weisenbach S, Jacobs AH, et al. Nonlinear progression of Parkinson disease as determined by serial positron emission tomographic imaging of striatal fluorodopa F 18 activity. Archives of Neurology. 2005; 62: 378–382.
Tissingh G, Bergmans P, Booij J, Winogrodzka A, van Royen EA, Stoof JC, et al. Drug-naive patients with Parkinson’s disease in Hoehn and Yahr stages I and II show a bilateral decrease in striatal dopamine transporters as revealed by [123I]β-CIT SPECT. Journal of Neurology. 1998; 245: 14–20.
Kitada T, Pisani A, Porter DR, Yamaguchi H, Tscherter A, Martella G, et al. Impaired dopamine release and synaptic plasticity in the striatum of PINK1-deficient mice. Proceedings of the National Academy of Sciences, USA. 2007; 104: 11441–11446.
Calabresi P, Maj R, Pisani A, Mercuri N, Bernardi G. Long-term synaptic depression in the striatum: physiological and pharmacological characterization. Journal of Neuroscience. 1992; 12: 4224–4233.
Langston JW, Langston EB, Irwin I. MPTP-induced parkinsonism in human and non-human primates—clinical and experimental aspects. Acta Neurologica Scandinavica Supplementum. 1984; 100: 49–54.
Heikkila RE, Hess A, Duvoisin RC. Dopaminergic neurotoxicity of 1-methyl-4-phenyl-1, 2, 5, 6-tetrahydropyridine in mice. Science. 1984; 224: 1451–1453.
Meredith GE, Totterdell S, Potashkin JA, Surmeier DJ. Modeling PD pathogenesis in mice: advantages of a chronic MPTP protocol. Parkinsonism & Related Disorders. 2008; 14: S112–S115.
Zhang Y, He X, Wu X, Lei M, Wei Z, Zhang X, et al. Rapamycin upregulates glutamate transporter and IL-6 expression in astrocytes in a mouse model of Parkinson’s disease. Cell Death & Disease. 2017; 8: e2611.
Zhu G, Li J, He L, Wang X, Hong X. MPTP-induced changes in hippocampal synaptic plasticity and memory are prevented by memantine through the BDNF-TrkB pathway. British Journal of Pharmacology. 2015; 172: 2354–2368.
Toy WA, Petzinger GM, Leyshon BJ, Akopian GK, Walsh JP, Hoffman MV, et al. Treadmill exercise reverses dendritic spine loss in direct and indirect striatal medium spiny neurons in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) mouse model of Parkinson’s disease. Neurobiology of Disease. 2014; 63: 201–209.
Kleim JA, Hogg TM, Vandenberg PM, Cooper NR, Bruneau R, Remple M. Cortical synaptogenesis and motor map reorganization occur during late, but not early, phase of motor skill learning. Journal of Neuroscience. 2004; 24: 628–633.
Kleim JA, Lussnig E, Schwarz ER, Comery TA, Greenough WT. Synaptogenesis and Fos expression in the motor cortex of the adult rat after motor skill learning. Journal of Neuroscience. 1996; 16: 4529–4535.
Hill TC, Zito K. LTP-induced long-term stabilization of individual nascent dendritic spines. Journal of Neuroscience. 2013; 33: 678–686.
Jenkins TA, Amin E, Pearce JM, Brown MW, Aggleton JP. Novel spatial arrangements of familiar visual stimuli promote activity in the rat hippocampal formation but not the parahippocampal cortices: a c-fos expression study. Neuroscience. 2004; 124: 43–52.
Ryan L, Lin C, Ketcham K, Nadel L. The role of medial temporal lobe in retrieving spatial and nonspatial relations from episodic and semantic memory. Hippocampus. 2010; 20: 11–18.
Helton TD, Otsuka T, Lee M, Mu Y, Ehlers MD. Pruning and loss of excitatory synapses by the parkin ubiquitin ligase. Proceedings of the National Academy of Sciences, USA. 2008; 105: 19492–19497.
Joelving FC, Billeskov R, Christensen JR, West M, Pakkenberg B. Hippocampal neuron and glial cell numbers in Parkinson’s disease—a stereological study. Hippocampus. 2006; 16: 826–833.
Summerfield C, Junqué C, Tolosa E, Salgado-Pineda P, Gómez-Ansón B, Martí MJ, et al. Structural brain changes in Parkinson disease with dementia: a voxel-based morphometry study. Archives of Neurology. 2005; 62: 281–285.
Shohamy D, Myers CE, Hopkins RO, Sage J, Gluck MA. Distinct hippocampal and basal ganglia contributions to probabilistic learning and reversal. Journal of Cognitive Neuroscience. 2009; 21: 1821–1833.
Jokinen P, Brück A, Aalto S, Forsback S, Parkkola R, Rinne JO. Impaired cognitive performance in Parkinson’s disease is related to caudate dopaminergic hypofunction and hippocampal atrophy. Parkinsonism & Related Disorders. 2009; 15: 88–93.
Nagano-Saito A, Washimi Y, Arahata Y, Kachi T, Lerch JP, Evans AC, et al. Cerebral atrophy and its relation to cognitive impairment in Parkinson’s disease. Neurology. 2005; 64: 224–229.
Brück A, Kurki T, Kaasinen V, Vahlberg T, Rinne JO. Hippocampal and prefrontal atrophy in patients with early non-demented Parkinson’s disease is related to cognitive impairment. Journal of Neurology, Neurosurgery, and Psychiatry. 2004; 75: 1467–1469.
Costa C, Sgobio C, Siliquini S, Tozzi A, Tantucci M, Ghiglieri V, et al. Mechanisms underlying the impairment of hippocampal long-term potentiation and memory in experimental Parkinson’s disease. Brain. 2012; 135: 1884–1899.
Calabresi P, Castrioto A, Di Filippo M, Picconi B. New experimental and clinical links between the hippocampus and the dopaminergic system in Parkinson’s disease. Lancet Neurology. 2013; 12: 811–821.
Brundin P, Strecker RE, Lindvall O, Isacson O, Nilsson OG, Barbin G, et al. Intracerebral grafting of dopamine neurons: experimental basis for clinical trials in patients with Parkinson’s disease. Annals of the New York Academy of Sciences. 1987; 495: 473–496.
Wang X, Lu Y, Zhang H, Wang K, He Q, Wang Y, et al. Distinct efficacy of pre-differentiated versus intact fetal mesencephalon-derived human neural progenitor cells in alleviating rat model of Parkinson’s disease. International Journal of Developmental Neuroscience. 2004; 22: 175–183.
Hellmann MA, Panet H, Barhum Y, Melamed E, Offen D. Increased survival and migration of engrafted mesenchymal bone marrow stem cells in 6-hydroxydopamine-lesioned rodents. Neuroscience Letters. 2006; 395: 124–128.
Offen D, Barhum Y, Levy YS, Burshtein A, Panet H, Cherlow T, et al. Intrastriatal transplantation of mouse bone marrow-derived stem cells improves motor behavior in a mouse model of Parkinson’s disease. Journal of Neural Transmission Supplementum. 2007; 133–143.
Arenas E, Denham M, Villaescusa JC. How to make a midbrain dopaminergic neuron. Development. 2015; 142: 1918–1936.
Arenas E. Towards stem cell replacement therapies for Parkinson’s disease. Biochemical and Biophysical Research Communications. 2010; 396: 152–156.
Yan M, Sun M, Zhou Y, Wang W, He Z, Tang D, et al. Conversion of human umbilical cord mesenchymal stem cells in Wharton’s jelly to dopamine neurons mediated by the Lmx1a and neurturin in vitro: potential therapeutic application for Parkinson’s disease in a rhesus monkey model. PLoS ONE. 2013; 8: e64000.
Venkataramana NK, Pal R, Rao SAV, Naik AL, Jan M, Nair R, et al. Bilateral transplantation of allogenic adult human bone marrow-derived mesenchymal stem cells into the subventricular zone of Parkinson’s disease: a pilot clinical study. Stem Cells International. 2012; 2012: 931902.
Fu W, Zheng Z, Zhuang W, Chen D, Wang X, Sun X, et al. Neural metabolite changes in corpus striatum after rat multipotent mesenchymal stem cells transplanted in hemiparkinsonian rats by magnetic resonance spectroscopy. International Journal of Neuroscience. 2013; 123: 883–891.
Song J, Oh S, Kwon O, Wulansari N, Lee H, Chang M, et al. Cografting astrocytes improves cell therapeutic outcomes in a Parkinson’s disease model. Journal of Clinical Investigation. 2018; 128: 463–482.
Kim H. Stem cell potential in Parkinson’s disease and molecular factors for the generation of dopamine neurons. Biochimica Et Biophysica Acta. 2011; 1812: 1–11.
Folkerth RD, Durso R. Survival and proliferation of nonneural tissues, with obstruction of cerebral ventricles, in a parkinsonian patient treated with fetal allografts. Neurology. 1996; 46: 1219–1225.
Dell’Anno MT, Caiazzo M, Leo D, Dvoretskova E, Medrihan L, Colasante G, et al. Remote control of induced dopaminergic neurons in parkinsonian rats. Journal of Clinical Investigation. 2014; 124: 3215–3229.
Kim J, Su SC, Wang H, Cheng AW, Cassady JP, Lodato MA, et al. Functional integration of dopaminergic neurons directly converted from mouse fibroblasts. Cell Stem Cell. 2011; 9: 413–419.
Liu X, Huang Q, Li F, Li CY. Enhancing the efficiency of direct reprogramming of human primary fibroblasts into dopaminergic neuron-like cells through p53 suppression. Science China Life Sciences. 2014; 57: 867–875.
Liu X, Li F, Stubblefield EA, Blanchard B, Richards TL, Larson GA, et al. Direct reprogramming of human fibroblasts into dopaminergic neuron-like cells. Cell Research. 2012; 22: 321–332.
Caiazzo M, Dell’Anno MT, Dvoretskova E, Lazarevic D, Taverna S, Leo D, et al. Direct generation of functional dopaminergic neurons from mouse and human fibroblasts. Nature. 2011; 476: 224–227.
Heinrich C, Gascón S, Masserdotti G, Lepier A, Sanchez R, Simon-Ebert T, et al. Generation of subtype-specific neurons from postnatal astroglia of the mouse cerebral cortex. Nature Protocols. 2011; 6: 214–228.
Berninger B, Costa MR, Koch U, Schroeder T, Sutor B, Grothe B, et al. Functional properties of neurons derived from in vitro reprogrammed postnatal astroglia. Journal of Neuroscience. 2007; 27: 8654–8664.
Heins N, Malatesta P, Cecconi F, Nakafuku M, Tucker KL, Hack MA, et al. Glial cells generate neurons: the role of the transcription factor Pax6. Nature Neuroscience. 2002; 5: 308–315.
Heinrich C, Spagnoli FM, Berninger B. In vivo reprogramming for tissue repair. Nature Cell Biology. 2015; 17: 204–211.
Guo Z, Zhang L, Wu Z, Chen Y, Wang F, Chen G. In vivo direct reprogramming of reactive glial cells into functional neurons after brain injury and in an Alzheimer’s disease model. Cell Stem Cell. 2014; 14: 188–202.
Shaltouki A, Peng J, Liu Q, Rao MS, Zeng X. Efficient generation of astrocytes from human pluripotent stem cells in defined conditions. Stem Cells. 2013; 31: 941–952.
Krencik R, Weick JP, Liu Y, Zhang Z, Zhang S. Specification of transplantable astroglial subtypes from human pluripotent stem cells. Nature Biotechnology. 2011; 29: 528–534.
Proschel C, Stripay JL, Shih C, Munger JC, Noble MD. Delayed transplantation of precursor cell-derived astrocytes provides multiple benefits in a rat model of Parkinson’s. EMBO Molecular Medicine. 2014; 6: 504–518.
Rivetti di Val Cervo P, Romanov RA, Spigolon G, Masini D, Martín-Montañez E, Toledo EM, et al. Induction of functional dopamine neurons from human astrocytes in vitro and mouse astrocytes in a Parkinson’s disease model. Nature Biotechnology. 2017; 35: 444–452.
Yu X, Nagai J, Khakh BS. Improved tools to study astrocytes. Nature Reviews Neuroscience. 2020; 21: 121–138.
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