Neural Plasticity
Volume 2016 (2016), Article ID 7969272, 19 pages
http://dx.doi.org/10.1155/2016/7969272

Review Article

Emerging Link between Alzheimer’s Disease and Homeostatic Synaptic Plasticity

Sung-Soo Jang1,2 and Hee Jung Chung1,2

¹Department of Molecular and Integrative Physiology, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
²Neuroscience Program, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA

Received 14 December 2015; Accepted 31 January 2016

Academic Editor: Victor Anggono

Copyright © 2016 Sung-Soo Jang and Hee Jung Chung. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Alzheimer’s disease (AD) is an irreversible brain disorder characterized by progressive cognitive decline and neurodegeneration of brain regions that are crucial for learning and memory. Although intracellular neurofibrillary tangles and extracellular senile plaques, composed of insoluble amyloid-β (Aβ) peptides, have been the hallmarks of postmortem AD brains, memory impairment in early AD correlates better with pathological accumulation of soluble Aβ oligomers and persistent weakening of excitatory synaptic strength, which is demonstrated by inhibition of long-term potentiation, enhancement of long-term depression, and loss of synapses. However, current, approved interventions aiming to reduce Aβ levels have failed to retard disease progression; this has led to a pressing need to identify and target alternative pathogenic mechanisms of AD. Recently, it has been suggested that the disruption of Hebbian synaptic plasticity in AD is due to aberrant metaplasticity, which is a form of homeostatic plasticity that tunes the magnitude and direction of future synaptic plasticity based on previous neuronal or synaptic activity. This review examines emerging evidence for aberrant metaplasticity in AD. Putative mechanisms underlying aberrant metaplasticity in AD will also be discussed. We hope this review inspires future studies to test the extent to which these mechanisms contribute to the etiology of AD and offer therapeutic targets.

1. Introduction

Neurons communicate with each other at specialized intercellular junctions, called synapses. The strength of synaptic transmission can be dynamically and persistently altered in response to changes in neuronal activity. In the book The Organization of Behavior, Donald Hebb postulated that connections between neurons that are simultaneously active are strengthened [1]. Such “Hebbian plasticity” was first demonstrated at excitatory glutamatergic synapses in rabbit hippocampus by the seminal work of Bliss and Lomo [2]. High frequency stimulation of presynaptic axons in the perforant pathway induces stronger and long-lasting excitatory postsynaptic potentials (EPSPs) in neurons of the postsynaptic dentate gyrus [2]. This long-term potentiation (LTP) of excitatory synaptic strength lasts hours to months [2] and can be induced electrically in brain slices as well as in vivo in behaving animals [2, 3]. Hence, the associative and input-specific synaptic plasticity such as LTP and its counterpart long-term depression (LTD) is thought to underlie cellular correlates of learning and memory [4–7].

Hebbian plasticity also represents a positive feedback mechanism. Once LTP is induced, saturated synapses undergo further potentiation with greater ease than before the LTP induction, leading to unstable runaway excitation [8–10]. Similarly, continuous synaptic depression during LTD could result in unnecessary synaptic silencing and elimination [8–10]. In order to sense and counteract destabilizing effects of LTP and LTD, neurons employ negative feedback processes called homeostatic synaptic plasticity [11–14]. This adaptive plasticity offers a compensatory refinement of synaptic strength to maintain the stability of network activity within a physiologic limit [13–15]. For example, prolonged elevation of neuronal activity results in compensatory downscaling of synaptic strength to prevent hyperexcitation, whereas prolonged suppression of neuronal activity leads to compensatory upscaling of synaptic strength to prevent synapse silencing and elimination [13–15]. Without this homeostatic mechanism, the capacity of an active synapse would get saturated due to unconstrained potentiation, limiting its ability to store information (i.e., memory). Homeostatic synaptic plasticity is therefore a vital partner of Hebbian synaptic plasticity.

Defects in homeostatic synaptic plasticity could, in principle, cause abnormal Hebbian plasticity at synapses, leading to pathologic levels of synaptic potentiation or elimination in neurologic diseases. For example, Alzheimer’s disease (AD) is characterized by progressive and irreversible memory impairment [16] and associated with inhibition of LTP and enhancement of LTD in the hippocampus [17–27]. While physiologic levels of soluble amyloid-β (Aβ) oligomers have been shown to enhance synaptic activity and LTP [28, 29], pathologic levels of soluble Aβ oligomers impair LTP and enhance LTD in acute hippocampal slices [30–33]. Such impairment in Hebbian synaptic plasticity correlates strongly with memory impairment in early AD when Aβ plaques and neuronal degeneration are minimal [34–36]. Recent studies suggest that this abnormal Hebbian plasticity is due to pathologic engagement or disruption of metaplasticity [27, 32, 37], a form of homeostatic synaptic plasticity that controls the induction threshold of LTP and LTD [38]. Interestingly, Aβ-induced aberrant hyperexcitability is found in cortical and hippocampal neuronal networks of human AD and mouse models of AD [39–45]. Further, epileptiform electrical seizures and neuronal activity stimulate Aβ synthesis and its release from the neurons in the hippocampus [46–48]. Indeed, a pathologic positive feedback loop between Aβ production and neuronal hyperexcitability would favor LTP inhibition and LTD induction.

These studies have provided a possible link between abnormal metaplasticity and cognitive dysfunction in AD pathogenesis, although our knowledge on the underlying mechanisms is limited. An understanding of the molecular mechanisms through which altered metaplasticity contributes to AD synaptopathology will be crucial in decoding the etiology of AD and may facilitate “correcting metaplasticity” as a putative novel therapy to restore Hebbian synaptic plasticity and treat cognitive dysfunction in early AD. In this paper, we review recent studies demonstrating aberrant metaplasticity in AD and discuss the possible underlying mechanisms focused on glutamate receptor regulation.

2. Abnormal Hebbian Synaptic Plasticity in AD

AD is a neurodegenerative disorder characterized by progressive and irreversible cognitive decline [16]. It is the 6th leading cause of death in the United States and the most common cause of dementia, which affects over 44 million people worldwide [49]. The molecular hallmarks of AD are amyloid plaques (extracellular deposits consisting of aggregated insoluble Aβ) and neurofibrillary tangles (intracellular filamentous aggregates of hyperphosphorylated tau) in the hippocampus and cortices [50–52], the brain regions critical for learning and memory. Interestingly, genetic suppression of endogenous tau blocks cognitive dysfunction in AD animal models, in which Aβ expression has been increased using a transgene [53–55], suggesting that tau acts downstream of Aβ in AD pathogenesis.

Importantly, soluble Aβ peptides rather than insoluble amyloid plaques have emerged to play critical roles in the early stages of AD pathogenesis. First, amyloid plaques are found at later stages after memory loss is already evident in humans and AD animal models with genetically elevated Aβ level [17–27]. Second, intracranial injection of soluble Aβ oligomers is sufficient to cause memory loss [29, 33, 56–59]. Third, rare early-onset autosomal dominant familial AD (FAD) is associated with increased levels of soluble Aβ due to mutations in genes whose protein products are involved in Aβ production and processing [60, 61]. Aβ peptides are generated by successive proteolysis of amyloid-β precursor protein (APP), a large transmembrane glycoprotein that is initially cleaved by the β-site APP-cleaving enzyme 1 (BACE1) and subsequently by γ-secretase in the transmembrane domain [62–64]. The FAD mutations are found in APP and presenilins [60, 61], which are catalytic components of γ-secretase [65]. Lastly, a major genetic risk factor for most AD (i.e., sporadic AD) is polymorphic ε4 allele of apolipoprotein E [66, 67]. The encoded ApoE4 is less efficient in clearing Aβ than the common ApoE3, suggesting a strong association between sporadic AD and increased levels of soluble Aβ [68].

How could pathologic levels of soluble Aβ oligomers cause cognitive dysfunction? The first clue came from the studies in AD mouse models with genetically elevated Aβ [17–27]. Before the development of amyloid plaques is evident, these AD mouse models display severe impairment of hippocampal LTP [17–27]. Furthermore, LTD is induced in these AD mouse hippocampi with subthreshold stimulations, which normally cannot induce LTD in wild-type control mice [17–27]. Subsequent studies have shown that direct application of soluble Aβ oligomers (synthetic, cell-culture secreted, or AD brain-derived) at pathologic levels inhibits LTP and enhances LTD in acute hippocampal slice [30–33]. A persistent and unchecked decrease in synaptic strength is expected to lead to the pathologic elimination of synapses [69–71]. Indeed, decreases in synapse density are evident in hippocampi of patients with early AD [72–75]. Therefore, abnormal Hebbian synaptic plasticity is thought to be the basis of memory loss in early AD when amyloid plaques and neuronal degeneration are minimal [34–36].

3. Is Abnormal Hebbian Synaptic Plasticity due to Defective Homeostatic Synaptic Plasticity in AD?

Decades of studies cited above have compared the magnitudes of LTP and LTD in AD transgenic mouse models to determine if pathologic levels of soluble Aβ oligomers affect Hebbian synaptic plasticity. However, the absolute changes in the LTP and LTD magnitudes vary with age and AD mouse model [147], suggesting age- and strain-dependent differences for the induction threshold of LTP and LTD in these animals. The induction thresholds of LTP and LTD can be modified as a consequence of previous postsynaptic neuronal activity (Figure 1) [10, 148, 149]. LTP induction is favorable in neurons whose previous synaptic and intrinsic activities were low, whereas LTD induction is preferred when the previous activities were high [10, 148, 149]. Such compensatory adjustment of the induction thresholds for LTP and LTD, called “metaplasticity,” occurs as a form of homeostatic synaptic plasticity and provides stability to neuronal networks and supports Hebbian synaptic plasticity [10].

(a)

(b)

Figure 1: Metaplasticity. The induction threshold of LTP and LTD can be modified as a consequence of overall past synaptic or intrinsic activity of postsynaptic neurons. Such compensatory adjustment called “metaplasticity” provides stability to neuronal networks that support Hebbian synaptic plasticity. (a) LTP induction by conventional high frequency stimulation (HFS) is favorable in the neurons whose previous synaptic and intrinsic activities were low. (b) LTD induction by conventional low frequency stimulation (LFS) is favorable in the neurons whose previous synaptic and intrinsic activities were high.

Hence, it is possible that the abnormal Hebbian synaptic plasticity in AD could arise from the defects in metaplasticity. Several studies have provided supporting evidence for this hypothesis. Aberrant neuronal hyperexcitability has been observed in cortical and hippocampal neuronal networks of patients with early AD [150] and FAD AD mouse models with heightened levels of APP and Aβ [21, 41–44, 151]; this is also consistent with reports that patients with early AD and FAD animal models exhibit epileptic seizures [21, 45, 152–160]. Pharmacological inhibition of epileptic seizures inhibits memory loss in AD mouse models [156], implicating critical roles of aberrant neuronal hyperexcitability in cognitive dysfunction presented early in AD pathogenesis [39, 40]. Hence, Aβ-induced cognitive dysfunction in early AD may result from the inability of neurons to adapt to persistent increases in overall neural network activity rather than the absolute changes in LTP and LTD magnitudes.

Additional support for this hypothesis comes from the report that soluble Aβ oligomers result in excessive activation of N-methyl D-aspartate receptors (NMDARs) containing GluN2B subunits, causing LTP inhibition and LTD facilitation via ERK and CREB signaling pathways [161]. GluN2B-selective antagonists effectively prevent Aβ-induced LTP inhibition [161–163], suggesting that early activation of extrasynaptic NMDARs primes the synapse to inhibit LTP induction and facilitate LTD induction. Consistent with this notion, GluN2B-selective antagonists prevent priming-induced inhibition of LTP [164]. The beneficial effects of the partial NMDAR antagonist memantine in AD also support the possible role of metaplasticity in AD-associated synaptic dysfunction because memantine does not block LTP acutely but restores LTP induction impaired by tonic NMDAR activation [165, 166].

A recent study by Megill et al. has provided direct evidence for impaired metaplasticity in an AD transgenic mouse model [27]. This study examined frequency- and age-dependent synaptic plasticity in the APP/PS1 AD mouse model [27], which has two FAD-linked mutations (a Swedish mutation in APP and a deletion FAD mutation in exon 9 of presenilin-1) [167]. These mutations increase total Aβ production, resulting in a higher level of aggregation-prone Aβ42 peptides, and accelerated AD pathology [168] and age-dependent cognitive deficits [169, 170]. While the wild-type mice show a shift of the induction threshold to favor LTP and suppress LTD at the hippocampal CA1 Schaffer collateral synapses with age, the APP/PS1 transgenic mice fail to undergo this normal developmental metaplasticity [27]. As a result, the magnitudes of LTP and LTD remained the same in the APP/PS1 transgenic mice from when they were young (1 month of age) until they were adult (6 months of age). When the absolute magnitudes of LTP and LTD were compared, the adult APP/PS1 mice display LTP inhibition and LTD facilitation compared to age-matched wild-type mice [27]. Although electrophysiological characterization of other AD mouse models with elevated Aβ levels should be performed to see if impaired developmental metaplasticity is a general phenomenon for AD, these findings suggest that the Hebbian synaptic plasticity defects in AD could be due to the inability of neurons to undergo developmental metaplasticity (Figure 2).

Figure 2: Aberrant metaplasticity in AD. Aβ increases the activity of excitatory neurons, which in turn stimulates synthesis and release of Aβ in a positive feedback loop, leading to pathologic accumulation of Aβ. Neuronal hyperexcitability or early activation of GluN2B-containing NMDAR by heightened Aβ expression induces aberrant metaplasticity, leading to inhibition of LTP by HFS and enhancement of LTD in the hippocampus by LFS or normal LTD induction by subthreshold LFS.

4. Putative Mechanisms Underlying Defective Homeostatic Synaptic Plasticity in AD

How can pathologic levels of soluble Aβ oligomers cause aberrant metaplasticity in AD? One way to mediate metaplasticity is to alter the induction mechanisms of LTP and LTD by regulating the function of NMDARs [10, 171–174] because calcium (Ca²⁺) influx through NMDARs at the postsynaptic density (PSD) is critical for inductions of NMDAR-dependent LTP and LTD [175–177]. An effective means to alter Ca²⁺ current per unit charge through NMDAR is to change subunit composition of NMDAR [177]. Such a change influences Ca²⁺/calmodulin-dependent protein kinase II (CaMKII) interaction with NMDARs and has been shown to control Hebbian synaptic plasticity [178]. For example, GluN2B-containing NMDARs bind to CaMKII with high affinity whereas those containing GluN2A interact with CaMKII with low affinity. Consistent with their decreased affinity for CaMKII, altering synaptic NMDARs from GluN2B-containing receptors to GluN2A-containing ones markedly reduces LTP induction [178]. Developmental metaplasticity in the visual cortex has also been suggested to involve experience-dependent changes in the GluN2 subunit composition of NMDARs that influence the induction thresholds of LTP and LTD [173, 179].

Soluble Aβ oligomers have been shown to decrease glutamate reuptake and subsequently increase extracellular glutamate levels [32, 180]. Such glutamate spillover would activate extracellular NMDARs, which are mostly composed of GluN2B-containing NMDARs at mature synapses [175, 176]. Indeed, soluble Aβ oligomers enhance activation of GluN2B-containing NMDARs more rapidly than synaptic depression and such actions would prime excitatory synapses to inhibit LTP induction and favor LTD induction [161]. However, NMDAR subunit composition and current are similar between wild-type mice and APP/PS1 mice at all ages [27], suggesting that developmental metaplasticity defect in the APP/PS1 mice is not due to altered NMDAR function during the induction of LTP and LTD.

Another way to induce metaplasticity is to alter the expression mechanisms of LTP and LTD by regulating α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors (AMPARs) [181], which mediate the majority of excitatory synaptic current upon glutamate binding [182]. A major postsynaptic expression mechanism for LTP is the synaptic recruitment of AMPARs from a perisynaptic reserve pool and their subsequent stabilization at excitatory synapses, whereas that for LTD is the removal and internalization of synaptic AMPARs [175, 182]. Insertion and removal of synaptic AMPARs during the expression of LTP and LTD, respectively, are tightly regulated processes by phosphorylation of AMPAR subunit GluA1 at Ser-845 and Ser-831 [183–185]. Phosphorylation of GluA1 at Ser-845 by protein kinase A (PKA) is necessary for synaptic targeting of GluA1 driven by CaMKII [186], whereas dephosphorylation at Ser-845 mediates GluA1 internalization [183, 187, 188] and NMDAR-dependent LTD [185]. In addition, Ser845 phosphorylation of GluA1 mediates synaptic insertion of Ca²⁺-permeable GluA1-containing AMPARs during synaptic scaling in cultured dissociated cortical neurons upon chronic activity deprivation [92] and homeostatic synaptic scaling in the visual cortex upon sensory deprivation [189, 190]. Phosphorylation of GluA1 at Ser831 by protein kinase C (PKC) [191] and CaMKII [192, 193] increases following LTP induction [184, 194] and supports LTP expression [183–185]. Although GluA1 phosphorylation at Ser-845 and Ser-831 has been shown to reduce the induction threshold for LTP [195, 196], adult APP/PS1 mice display normal levels of GluA1 phosphorylation and perisynaptic AMPARs compared to those of wild-type mice [27]. Hence, the developmental metaplasticity defect in APP/PS1 mice is not due to insufficient AMPAR availability for synaptic insertion; rather, it is due to regulation of AMPAR trafficking by means other than GluA1 phosphorylation.

Metaplasticity is a form of homeostatic synaptic plasticity in which the magnitude and polarity of synaptic plasticity are adjusted accordingly based on the past history of synaptic and neural activity [38]. Since metaplasticity can occur at a single synapse [197, 198], it is tempting to speculate that pathologic levels of Aβ may impair developmental metaplasticity by altering postsynaptic expression mechanisms of homeostatic synaptic plasticity (Figure 3). Homeostatic synaptic plasticity has been extensively investigated using primary dissociated culture of neocortical and hippocampal neurons (Table 1). In these studies, prolonged blockade of network activity for 48 hours (h) with the sodium channel blocker tetrodotoxin (TTX) induces a significant increase in AMPAR-mediated miniature excitatory postsynaptic current (mEPSC) amplitude and synaptic AMPAR density, indicating the postsynaptic expression of homeostatic synaptic scaling [78, 80, 94, 138, 199–201]. Conversely, mEPSC amplitudes are scaled down in dissociated neuronal culture after prolonged enhancement of network activity by KCl depolarization or blocking inhibitory neurotransmission with antagonists for A-type gamma-aminobutyric acid () receptors, such as bicuculline [78, 80, 90, 199, 202, 203]. Interestingly, many of the crucial mediators of homeostatic synaptic plasticity have also been implicated in AMPAR regulation during LTP and LTD expression and AD pathology (Table 1). Taken together, these correlated functional roles raise an intriguing possibility that pathologic accumulation of Aβ may impair molecular mechanisms involved in homeostatic synaptic plasticity, which manifests as disruption of Hebbian synaptic plasticity in AD (Figure 3).

Table 1: Molecular mechanisms and players involved in AD and the expression of homeostatic synaptic plasticity.

Roles in synaptic scaling
Involvement in AD
References

AMPAR scaffolding proteins

GRIP1
Synaptic accumulation and removal of GRIP1 mediate synaptic scaling and downscaling, respectively, by regulating synaptic AMPAR targeting.

[76, 77]

PICK1
PICK1 degradation mediates synaptic scaling.
PICK1 interaction with GluA2 mediates A-induced synaptic depression.
[78, 79]

Regulators of AMPAR trafficking

Arc/Arg3.1
Downregulation of Arc/Arg3.1 mediates synaptic scaling by increasing surface AMPAR density. Upregulation of Arc/Arg3.1 mediates synaptic downscaling by promoting AMPAR endocytosis.
Arc/Arg3.1 expression is elevated in AD and mediates activity-dependent generation of A by binding to presinilin-1 and regulating γ-secretase trafficking.
[80–82]

Homer1a
Downregulation of Homer1a mediates synaptic scaling, whereas upregulation of Homer1a mediates synaptic downscaling by regulating surface AMPAR density and Tyr-phosphorylation.

[83]

Regulators of synaptic AMPAR density

PSD-95
Synaptic accumulation of PSD-95 mediates synaptic scaling, whereas its interaction with TARP mediates synaptic downscaling.
Pathological level of A leads to PSD-95 degradation.
[84–89]

PSD-93
PSD-93 mediates synapticscaling.

[84]

GKAP
Synaptic accumulation and removal of GKAP mediate synaptic scaling and downscaling, respectively, by regulating surface AMPAR density.
Pathological level of A leads to GKAP degradation.
[90, 91]

Posttranslation modification of AMPAR

Calcineurin
Reduced calcineurin activity mediates synaptic scaling via GluA1-Ser845 dephosphorylation and subsequent synaptic trafficking of Ca²⁺-permeable AMPARs.
In AD mouse model, increased activity of calcineurin induces dephosphorylation and synaptic removal of the GluR1 subunit of AMPAR.
[92, 93]

STEP₆₁
Downregulation of STEP₆₁ mediates synaptic scaling, whereas enhanced STEP₆₁ upon chronic activity induces dephosphorylation of GluN2B and GluA2.
STEP₆₁ expression is elevated in AD and mediates A-induced dephosphorylation and internalization of NMDARs and AMPARs, whereas inhibition of STEP₆₁ prevents cognitive deficits and impaired hippocampal LTP in AD mouse models.
[94–98]

PP1
Downregulation of PP1 inhibitor-2 (I-2) mediates synaptic downscaling by reducing surface AMPARs.
Inhibition of PP1 blocks A-induced impairment in hippocampal LTP.
[99, 100]

DHHC2
Translocation of DHHC2 to PSD mediates synaptic scaling by enhancing synaptic targeting of PSD95 and AMPAR.

[86]

Nedd4-1
Upregulation of Nedd4-1 mediates synaptic downscaling by reducing surface AMPAR density.
Nedd4-1 expression is elevated in AD.
[101, 102]

SUMO-1 and Ubc9
SUMOylation of Arc/Arg3.1 mediates synaptic scaling.
SUMO-conjugating enzyme, Ubc9, enhances SUMOylation and rescues A-induced deficits in hippocampal LTP and learning and memory.
[103, 104]

Local dendritic translation of AMPAR

eEF2
Increased eEF2 activity mediates synaptic scaling by stimulating local dendritic synthesis.

[105]

miRNA-92a
Inhibition of miRNA-92A mediates synaptic scaling by stimulating local dendritic synthesis of GluA1.

[106]

Retinoic acid (RA)
Increased RA activity mediates synaptic scaling by stimulating local dendritic synthesis of GluA1 through RA receptor.
RA regulates the expression of APP processing genes, attenuates A deposition, and rescues memory deficits in AD mouse model.
[107–114]

Secreted factors

BDNF
Downregulation of BDNF mediates synaptic scaling.
Downregulation of BDNF levels is associated with the degree of synaptic and cognitive deficits during the progression of AD.
[81, 115–117]

TNFα
TNFα mediates synaptic scaling in primary neuronal culture and visual cortex upon activity deprivation.
TNFα contributes to AD-related brain neuroinflammation and amyloidogenesis via -secretase regulation.
[118–127]

Cell adhesion molecules

3 integrin
Enhanced surface expression of 3 integrin inhibits the small GTPase Rap1 and mediates synaptic scaling by stabilizing synaptic.

[128, 129]

MHC-1
MHC-1 mediates TTX-induced synaptic scaling in hippocampal cultured neurons.

[130]

N-Cadherin
N-Cadherin interaction with -catenin mediates synaptic scaling and downscaling by regulating GluA1-containing AMPARs.
Inhibition of N-Cadherin interaction with -catenin accelerates A-induced synaptic impairments.
[131–135]

EphA4
Increased Eph4 activity mediates synaptic downscaling by stimulating ubiquitin-dependent proteasome degradation of GluA1.
Soluble A oligomers upregulate EphA4 whereas genetic ablation or inhibition of EphA4 prevents hippocampal LTP impairment in AD transgenic model mice.
[136, 137]

Transcriptional regulation

CaMKK-CaMK4
Reduced activity of the CaMKK/CaMK4 signaling pathway mediates synaptic scaling, whereas its stimulation mediates synaptic downscaling.

[138–140]

MSK1
MSK1 mediates TTX-induced synaptic scaling in hippocampal neurons by increasing surface AMPAR density.
MSK1 activity is elevated in AD.
[141, 142]

MeCP2
MeCP2 mediates synaptic scaling in visual cortex upon visual deprivation in vivo.

[143]

Other proteins

Plk2
Increase in Plk2 activity mediates synaptic downscaling.

[144, 145]

Cdk5
Increase in Cdk5 activity mediates synaptic downscaling.
Enhanced Cdk5 activity in AD contributes to Tau phosphorylation and toxicity.
[144, 146]

(a)

(b)

Figure 3: Postsynaptic expression mechanisms in normal and AD synapses. (a) In normal synapses, chronic activity blockade leads to synaptic scaling expressed by a compensatory increase in synaptic AMPAR density and current, whereas chronic activity elevation leads to synaptic downscaling expressed by a compensatory decrease in synaptic AMPAR density and current. (b) In AD, Aβ increases neuronal excitability and/or synaptic activity, leading to induction of synaptic downscaling. Because pathologic feedback loop continues to produce and release Aβ, synaptic downscaling becomes persistent and opposes the postsynaptic expression mechanisms for LTP.

4.1. AMPAR Scaffolding Proteins

Glutamate receptor-interacting protein 1 (GRIP1) and PICK1 (protein interacting with C-kinase 1) are PDZ (postsynaptic density 95/discs large/zona occludens) domain-containing proteins that regulate AMPAR trafficking by binding to the same intracellular C-terminus of GluA2 [204, 205]. GRIP1 binding to the unphosphorylated GluA2 C-terminus promotes synaptic targeting of AMPARs [206] whereas PICK1 can bind to both phosphorylated and unphosphorylated GluA2 [206] and mediates activity-dependent endocytosis of GluA2-containing AMPARs and stabilizes them in intracellular pools [207–209]. Recent studies have reported that chronic activity deprivation increases GRIP1 abundance at excitatory synapses and its interaction with GluA2, leading to synaptic targeting of AMPARs in cortical cultured neurons [76, 77]. In contrast, chronic enhancement of neuronal activity removes GRIP1 from excitatory synapses, which decreases surface AMPARs at synapses [76]. Compared to bidirectional modulation of synaptic GRIP1 expression in homeostatic synaptic plasticity, PICK1 expression is only altered by chronic activity blockade [78]. The TTX-induced synaptic scaling accompanies lysosome-mediated PICK1 degradation and can be occluded by genetic knock-out or shRNA knock-down of PICK1 [78]. Interestingly, pathologic levels of Aβ oligomers fail to reduce surface GluA2 expression and excitatory synaptic transmission in PICK knock-out neurons [79], indicating that GluA2 interaction with PICK1 mediates Aβ-induced synaptic depression. Thus, Aβ-dependent modulation of PICK1 and GRIP1 levels may likely contribute to aberrant developmental metaplasticity in AD.

AMPARs at excitatory synapses are also regulated by scaffolding proteins of the membrane associated guanylate kinase (MAGUK) family, which includes PSD-95, PSD-93, and SAP102 [210]. Chronic activity blockade increases synaptic accumulation of PSD95 and SAP102, whereas chronic activity enhancement decreases synaptic accumulation of PSD95 alone [84–86]. Double knock-down of PSD95/PSD93 or triple knock-down of PSD95/PSD93/SAP102 completely blocks chronic inactivity-induced increase in mEPSC amplitude [84], suggesting that PSD95 and PSD93 mediate synaptic scaling. In contrast, synaptic downscaling requires the PDZ1/2 domains of PSD-95 [84], which interact with transmembrane AMPAR regulatory proteins [TARPs] [211, 212]. Since TARPs link PSD-95 to AMPARs and promote synaptic insertion and stabilization of AMPARs [211, 212], these findings raise the possibility that reduced PSD95-TARP interaction may contribute to synaptic downscaling. Importantly, decreased PSD-95 expression is evident in AD mouse models [87] and Aβ application in cortical neuronal culture leads to downregulation of PSD-95 expression and dispersal of Shank1 [88, 89], another scaffolding protein enriched in excitatory glutamatergic synapses [213]. Interestingly, synaptic accumulation of guanylate kinase-associated protein (GKAP), which links Shank1 to PSD95 [214, 215], is increased upon chronic inhibition of neural activity and decreased by chronic excitation [90]. Such regulation of synaptic GKAP targeting contributes to bidirectional homeostatic scaling of excitatory synaptic strength [90]. Consistent with reports that pathological levels of Aβ increase degradation of PSD-95 and GKAP [89, 91], diminished interactions between PSD-95, TARP, and GKAP could dysregulate homeostatic synaptic plasticity in AD.

4.2. AMPAR Trafficking Regulators

Multiple proteins regulate synaptic AMPAR density by controlling their trafficking. One of them is Arc/Arg3.1, which is an immediate early gene. Arc/Arg3.1 mRNAs accumulate at excitatory synapses, where they are locally translated following synaptic simulation [216, 217]. Arc/Arg3.1 protein facilitates AMPAR internalization from the postsynaptic membrane by interacting with endocytosis mediators, endophilin2/3 and dynamin [218]. Chronic activity blockade of hippocampal or cortical cultured neurons has been shown to decrease mRNA and protein expression of Arc/Arg3.1 [80, 81]. Further, genetic ablation of Arc/Arg3.1 increases basal mEPSC amplitude and surface density of GluA1 and occludes TTX-induced increase in synaptic scaling [80]. Conversely, chronic elevation of neuronal activity increases Arc/Arg3.1 levels and decreases surface density of GluA1, whereas this regulation is absent in Arc/Arg3.1 knock-out neurons [80]. In addition to the critical roles of Arc/Arg3.1 in homeostatic synaptic plasticity, Arc/Arg3.1 expression is elevated in the medial prefrontal cortex of human AD patients [82], suggesting that elevated Arc/Arg3.1 expression may likely lead to AMPAR internalization during AD pathogenesis. In support of this notion, Arc/Arg3.1 is required for metabotropic glutamate receptor- (mGluR-) dependent LTD [219], a form of LTD that is also induced by application of Aβ oligomers [33]. Furthermore, Arc/Arg3.1 has been shown to mediate activity-dependent generation of Aβ by binding to presinilin-1 and regulating γ-secretase trafficking [82]. Based on these reports, persistent elevated Arc/Arg3.1 expression may act in multiple ways to disrupt synaptic homeostasis in AD by enhancing Aβ production and reducing synaptic AMPAR density.

Another immediate early gene, Homer1a, also contributes to homeostatic synaptic plasticity in Arc/Arg3.1-independent pathway [83]. Homer1a interrupts crosslinking action of constitutively expressed forms of Homer [220], thereby activating group I mGluRs in the absence of glutamate [221]. Chronic elevation of activity enhances Homer1a mRNA and protein expression, whereas chronic inactivity reduces Homer1a expression in cortical cultured neurons [83]. Importantly, mGluR inhibition or genetic ablation of Homer1a blocks bidirectional scaling of mEPSC amplitude and surface AMPAR density [83], implicating mGluR signaling and Homer1a in homeostatic synaptic plasticity. Interestingly, elevated tyrosine phosphorylation of GluA2 has been observed in Homer1a knock-out neurons [83] whereas tyrosine phosphorylation of GluA2 is decreased following group 1 mGluR stimulation through striatal enriched protein phosphatase (STEP₆₁) [95]. Although the specific Tyr residues on GluA2 regulated by STEP₆₁ are unknown, the downregulation of GluA2 tyrosine phosphorylation decreases surface expression of GluA2-containing AMPARs [222, 223]. Our recent study has demonstrated that chronic activity deprivation decreases protein and mRNA expression of STEP₆₁ and increases Tyr-phosphorylation of its substrates, including the NMDAR subunit GluN2B and the AMPAR subunit GluA2 in hippocampal cultured neurons [94]. Increasing STEP₆₁ activity blocks the increases in mEPSC amplitude and Tyr-phosphorylation of GluN2B and GluA2 induced by chronic activity blockade [94], suggesting that downregulation of STEP₆₁ is crucial for mediating homeostatic synaptic scaling. Conversely, chronic activity enhancement increases STEP₆₁ expression and decreases Tyr-phosphorylation of GluN2B and GluA2 [94]. Interestingly, elevated STEP₆₁ expression is observed in cortices of human AD patients and causes dephosphorylation and internalization of AMPARs in AD mouse models [95–97]. Further, genetic ablation or pharmacologic inhibition of STEP₆₁ prevents cognitive deficits and impaired hippocampal LTP in AD mouse models [96–98]. Given that STEP₆₁ may also participate in metaplasticity [224], persistent elevation of STEP₆₁ and Homer1a may disrupt developmental metaplasticity in AD.

In addition, alterations in Ca²⁺ influx modulates Ca²⁺-dependent activation of kinases such as Polo-like kinase 2 (Plk2) and Cyclin D kinase 5 (Cdk5) as well as protein phosphatases including calcineurin and protein phosphatase-1 (PP1) during homeostatic synaptic plasticity [93, 144, 145]. The increases in Plk2 and Cdk5 activity are thought to contribute to synaptic downscaling [144, 145] and AD pathogenesis [146]. Calcineurin-induced dephosphorylation of GluA1 at Ser845 has also been implicated in homeostatic synaptic plasticity [92] and AD [93]. Since PP1 activity downstream of calcineurin stimulation is required for LTD [225, 226], calcineurin may contribute to metaplasticity by regulating phosphorylation status of proteins which alters synaptic AMPAR density and function. In addition, Ca²⁺ influx through L-type voltage-gated Ca²⁺ channels (VGCCs) has been shown to increase PP1 activity via Ser43-phosphorylation of PP1 inhibitor-2 (I-2) following chronic activity elevation in hippocampal cultured neurons [99]. Furthermore, selective inhibition of PP1 blocks downscaling of surface AMPAR expression and mEPSC amplitude induced by chronic activity [99] as well as Aβ-induced impairment in hippocampal LTP [100], providing PP1 as another candidate signaling protein that may contribute to aberrant metaplasticity in AD.

4.3. Posttranslational Modification of AMPAR

Recent studies have revealed posttranslational modifications in addition to phosphorylation as important regulatory mechanisms of AMPAR expression during homeostatic synaptic plasticity. One such modification is palmitoylation, which mediates covalent attachment of palmitic acid [227]. The TTX-induced chronic silencing of network activity causes palmitoylation enzyme DHHC2 to be translocated from the dendrite to the postsynaptic density, resulting in homeostatic accumulation of PSD-95 and AMPARs at excitatory synapses [86]. Given that AMPAR trafficking is dynamically regulated by subunit-selective palmitoylation [228–230], these studies implicate palmitoylation of AMPAR subunits in the mechanism of synaptic scaling. Synaptic scaling also involves SUMOylation [103], which mediates covalent attachment of small ubiquitin-like modifiers (SUMO) [231]. Although there is no direct evidence for SUMOylation of AMPAR subunits [232], the TTX-induced elevation of surface AMPAR expression requires SUMOylation of Arc/Arg3.1 [103], a known regulator of AMPAR endocytosis [218]. Interestingly, reduced SUMOylation is observed in adult AD model mice [104]. While inhibition of SUMOylation blocks hippocampal LTP and hippocampal-dependent learning and memory in wild-type mice, the upregulation of SUMOylation by supplying its conjugating enzyme, Ubc9, rescues Aβ-induced deficits in hippocampal LTP and learning and memory [104]. Hence, reduced SUMOylation of Arc/Arg3.1 may contribute to defective developmental metaplasticity in AD.

Lastly, AMPARs are subjected to activity-dependent ubiquitination by the E3 ubiquitin ligase Nedd4-1, leading to their internalization and degradation in lysosomes [233–235]. Chronic elevation of neuronal activity increases Nedd4-1 protein levels, whereas shRNA-mediated knock-down of Nedd4-1 blocks the homeostatic reduction of surface AMPAR expression and mEPSC amplitudes induced by chronic activity [101], indicating that Nedd4-1 is required for homeostatic downscaling of excitatory synaptic strength. Given that elevated Nedd4-1 expression is found in human AD brain tissues [102], dysregulation of Nedd4-1 levels and subsequent impairment in homeostatic synaptic plasticity may play a role in AD etiology.

4.4. Regulation of Transcription and Translation

Ca²⁺ influx through NMDARs or L-type VGCCs activates signaling pathways that regulate transcriptions of genes important for neural development and plasticity. Consistent with this assertion, downscaling of excitatory synaptic strength induced by prolonged excitation of hippocampal CA1 neurons requires Ca²⁺ influx through L-type VGCCs and transcription activated downstream of CaMKK/CaMK4 signaling pathways [139]. TTX-induced synaptic scaling also requires transcription and translation; however, the mechanism involves a decrease in somatic Ca²⁺ influx through L-type VGCCs and subsequent reduction in CaMKK/CaMK4 signaling pathways [138, 140]. Recently, chronic inactivity was shown to increase transcription of genes encoding AMPARs and proteins that regulate AMPAR trafficking by decreasing cytosine methylation of genes [236]. Consistently, inhibition of DNA methylation alone induces synaptic scaling [236]. Furthermore, loss of methyl-CpG-binding protein-2 (MeCP2) prevents synaptic scaling in the visual cortex upon visual deprivation in vivo [143]. Taken together, these studies suggest that bidirectional homeostatic synaptic plasticity involves epigenetic modulation of genes whose protein products regulate excitatory synaptic transmission.

Our laboratory recently identified genes regulated by chronic alterations of neuronal activity in hippocampal neurons using unbiased gene expression analysis [81]. We identified several immediate early genes as well as genes associated with gene ontology terms “synaptic transmission” and “regulation of synaptic plasticity” [81]. One of the immediate early genes encodes brain-derived neurotrophic factor (BDNF) [81]. BDNF, which is secreted in an activity-dependent manner [237], regulates synaptic transmission and plasticity and promotes neuronal survival and transcription [238, 239]. We showed that BDNF mRNA expression is decreased in cultured hippocampal neurons upon chronic activity blockade using TTX treatment [81]. Importantly, inhibition of TrkB receptor signaling alone causes synaptic scaling in a similar extent as prolonged TTX treatment [115] whereas exogenous BDNF application prevents TTX-induced synaptic scaling [115] presumably through activation of mitogen- and stress-activated kinase 1 (MSK1) [141]. Interestingly, downregulation of BDNF is associated with the degree of synaptic and cognitive deficits during AD progression [116, 117, 240] and MSK1 activity is also elevated in AD [142]. These studies raise a possibility that aberrant BDNF-TrkB-MSK1 signaling pathway may disrupt synaptic homeostasis in AD.

In addition to the importance of transcriptional regulation, dendritic protein synthesis may serve as a mechanism to locally maintain the stability of synaptic strength. Chronic silencing of excitatory synaptic inputs stimulates dendritic protein synthesis by increasing the activity of eukaryotic elongation factor-2 (eEF2) [105]. Furthermore, simultaneous treatment of hippocampal neurons with TTX (to block action potentials) and APV (to block NMDAR-mediated miniature synaptic transmission) increases the expression of GluA1 homomers by stimulating local dendritic translation of GluA1 mRNAs [107, 241–243]. This synaptic scaling is mediated by microRNA-92a, which is a small noncoding RNA that inhibits translation of GluA1 mRNAs by binding to their 3′ untranslated region (UTR) [106]. Other studies have also reported that retinoic acid (RA) signaling via RA receptor-α (RARα) interaction with the 5′ UTR of GluA1 mRNA contributes to synaptic scaling following prolonged cotreatment with TTX and APV by stimulating local dendritic synthesis of GluA1 [107–109]. Importantly, RAR signaling has been shown to regulate the expression of genes related to APP processing [110–113], attenuate Aβ deposition, and rescue memory deficits in AD mouse models [114], suggesting that alteration of RA signaling pathways may contribute to impaired metaplasticity in AD.

4.5. Cell Adhesion Molecules (CAMs)

β3 integrin is a cell adhesion molecule (CAM) enriched in excitatory synapses [244, 245] and controls synaptic currents mediated by GluA2-containing AMPARs [128]. Synaptic scaling induced by chronic activity blockade is associated with enhanced surface expression of β3 integrin in hippocampal neurons and is absent in β3 integrin knock-out neurons [128, 129]. Pharmacological perturbation of β3 integrin enhances GluA2 internalization and reduces synaptic AMPAR currents by activating the small GTPase Rap1 [128], which has been implicated in homeostatic downscaling of excitatory synapses [144, 246]. In addition to β3 integrin, class I major histocompatibility complex (MHC-1) proteins, which are found postsynaptically at excitatory synapses, also contribute to synaptic scaling following chronic activity blockade [130]. Although the role of β3 integrin and MHC-1 in AD pathogenesis remains unknown, their neuronal expression is regulated by glia-derived tumor necrosis factor α (TNFα) [128, 247, 248], which is involved in AD pathology in humans and AD mouse models [118–124]. TNFα elevates AMPAR-mediated mEPSC amplitude through activation of TNFα receptor during synaptic scaling [125, 126] whereas TNFα knock-out mice lack synaptic scaling in their visual cortex [127] but display normal LTP [127, 249, 250]. Hence, TNFα may influence metaplasticity through β3 integrin and MHC-1, and such a signaling pathway may be disrupted in AD.

N-Cadherin is another CAM that is enriched at excitatory synapses and has been implicated in AD as well as homeostatic synaptic plasticity. N-Cadherin promotes APP dimerization, modulates Aβ secretion, and reduces surface expression of presinilin-1 [131, 132]. N-Cadherin also binds to the extracellular domains of GluA1 in a Ca²⁺-dependent manner and regulates GluA1 surface expression [251, 252]. Although N-Cadherin interaction with the actin cytoskeleton [133, 253] contributes to dendritic spine enlargement during LTP expression [253–257], the interaction between N-Cadherin and β-catenin mediates bidirectional homeostatic synaptic plasticity by regulating GluA1-containing AMPARs [133, 134]. Given that inhibition of N-Cadherin interaction with β-catenin accelerates Aβ-induced synaptic impairments [135], dysregulation of N-Cadherin may likely impair homeostatic synaptic plasticity in AD. Ephrin receptor tyrosine kinase subfamily EphA4 is another CAM implicated in homeostatic synaptic plasticity. Increased activity of EphA4 mediates homeostatic downscaling by stimulating ubiquitin-dependent proteasome degradation of GluA1 [136]. Interestingly, soluble Aβ oligomers induce EphA4 activation, whereas genetic ablation or inhibition of EphA4 prevents hippocampal LTP impairment in AD transgenic model mice [137], raising an interesting possibility that Aβ-induced enhancement in EphA4 activity may impair metaplasticity in AD by regulating AMPAR degradation.

5. Conclusions

Recent studies have uncovered an exciting link between pathologic accumulation of Aβ and aberrant metaplasticity, a form of homeostatic synaptic plasticity that controls the induction threshold for LTP and LTD. Specifically, these studies have suggested a novel hypothesis that aberrant metaplasticity may contribute to LTP inhibition and LTD enhancement in AD. However, the molecular mechanism underlying Aβ-dependent alteration of metaplasticity remains largely unknown. Since many molecular players involved in homeostatic synaptic plasticity have been shown to regulate synaptic AMPAR density in Hebbian synaptic plasticity, it is tempting to speculate that pathologic levels of Aβ mediate their effect via a common mechanism shared between Hebbian and homeostatic plasticity at excitatory synapses. Challenges lie ahead in understanding how the molecular players and pathways reviewed here work together to express homeostatic plasticity at excitatory synapses and how Aβ disrupts homeostatic synaptic plasticity in AD. Future studies designed to tackle these challenges should offer substantial insights into the homeostatic control of excitatory synaptic strength in normal brain and AD brain. These studies may also facilitate the search for targeted therapeutic interventions to correct aberrant metaplasticity in AD, thus reversing persistent synaptic weakening and cognitive dysfunction in AD.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

Acknowledgments

The authors thank Dr. Eric Bolton at the University of Illinois at Urbana Champaign for his insightful comments on the paper. This work is supported by ICR start-up funding from the University of Illinois at Urbana Champaign and National Institute of Health funding (NS083402) to Hee Jung Chung.

References

D. O. Hebb, The Organization of Behavior: A Neuropsychological Theory, John Wiley & Sons, New York, NY, USA, 1949.

T. V. P. Bliss and T. Lomo, “Long-lasting potentiation of synaptic transmission in the dentate area of the anaesthetized rabbit following stimulation of the perforant path,” The Journal of Physiology, vol. 232, no. 2, pp. 331–356, 1973.

J. R. Whitlock, A. J. Heynen, M. G. Shuler, and M. F. Bear, “Learning induces long-term potentiation in the hippocampus,” Science, vol. 313, no. 5790, pp. 1093–1097, 2006.

M. A. Lynch, “Long-term potentiation and memory,” Physiological Reviews, vol. 84, no. 1, pp. 87–136, 2004.

R. C. Malenka and M. F. Bear, “LTP and LTD: an embarrassment of riches,” Neuron, vol. 44, no. 1, pp. 5–21, 2004.

G. Neves, S. F. Cooke, and T. V. P. Bliss, “Synaptic plasticity, memory and the hippocampus: a neural network approach to causality,” Nature Reviews Neuroscience, vol. 9, no. 1, pp. 65–75, 2008.

D. E. Feldman, “Synaptic mechanisms for plasticity in neocortex,” Annual Review of Neuroscience, vol. 32, pp. 33–55, 2009.

L. F. Abbott and S. B. Nelson, “Synaptic plasticity: taming the beast,” Nature Neuroscience, vol. 3, pp. 1178–1183, 2000.

G. G. Turrigiano and S. B. Nelson, “Hebb and homeostasis in neuronal plasticity,” Current Opinion in Neurobiology, vol. 10, no. 3, pp. 358–364, 2000.

L. N. Cooper and M. F. Bear, “The BCM theory of synapse modification at 30: interaction of theory with experiment,” Nature Reviews Neuroscience, vol. 13, no. 11, pp. 798–810, 2012.

E. Marder and A. A. Prinz, “Current compensation in neuronal homeostasis,” Neuron, vol. 37, no. 1, pp. 2–4, 2003.

G. W. Davis, “Homeostatic control of neural activity: from phenomenology to molecular design,” Annual Review of Neuroscience, vol. 29, pp. 307–323, 2006.

G. G. Turrigiano, “The self-tuning neuron: synaptic scaling of excitatory synapses,” Cell, vol. 135, no. 3, pp. 422–435, 2008.

K. Pozo and Y. Goda, “Unraveling mechanisms of homeostatic synaptic plasticity,” Neuron, vol. 66, no. 3, pp. 337–351, 2010.

G. Turrigiano, “Homeostatic synaptic plasticity: local and global mechanisms for stabilizing neuronal function,” Cold Spring Harbor Perspectives in Biology, vol. 4, no. 1, Article ID a005736, 2012.

G. McKhann, D. Drachman, and M. Folstein, “Clinical diagnosis of Alzheimer's disease: report of the NINCDS-ADRDA Work Group under the auspices of Department of Health and Human Services Task Force on Alzheimer's disease,” Neurology, vol. 34, no. 7, pp. 939–944, 1984.

P. F. Chapman, G. L. White, M. W. Jones et al., “Impaired synaptic plasticity and learning in aged amyloid precursor protein transgenic mice,” Nature Neuroscience, vol. 2, no. 3, pp. 271–276, 1999.

S. M. Fitzjohn, R. A. Morton, F. Kuenzi et al., “Age-related impairment of synaptic transmission but normal long-term potentiation in transgenic mice that overexpress the human APP695SWE mutant form of amyloid precursor protein,” The Journal of Neuroscience, vol. 21, no. 13, pp. 4691–4698, 2001.

J. S. Jacobsen, C.-C. Wu, J. M. Redwine et al., “Early-onset behavioral and synaptic deficits in a mouse model of Alzheimer's disease,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 13, pp. 5161–5166, 2006.

A. Y. Hsia, E. Masliah, L. McConlogue et al., “Plaque-independent disruption of neural circuits in Alzheimer's disease mouse models,” Proceedings of the National Academy of Sciences of the United States of America, vol. 96, no. 6, pp. 3228–3233, 1999.

J. J. Palop, J. Chin, E. D. Roberson et al., “Aberrant excitatory neuronal activity and compensatory remodeling of inhibitory hippocampal circuits in mouse models of Alzheimer's disease,” Neuron, vol. 55, no. 5, pp. 697–711, 2007.

J. Larson, G. Lynch, D. Games, and P. Seubert, “Alterations in synaptic transmission and long-term potentiation in hippocampal slices from young and aged PDAPP mice,” Brain Research, vol. 840, no. 1-2, pp. 23–35, 1999.

S. Roder, L. Danober, M. F. Pozza, K. Lingenhoehl, K.-H. Wiederhold, and H.-R. Olpe, “Electrophysiological studies on the hippocampus and prefrontal cortex assessing the effects of amyloidosis in amyloid precursor protein 23 transgenic mice,” Neuroscience, vol. 120, no. 3, pp. 705–720, 2003.

I. Dewachter, L. Ris, S. Croes et al., “Modulation of synaptic plasticity and Tau phosphorylation by wild-type and mutant presenilin1,” Neurobiology of Aging, vol. 29, no. 5, pp. 639–652, 2008.

I. Gureviciene, S. Ikonen, K. Gurevicius et al., “Normal induction but accelerated decay of LTP in APP + PS1 transgenic mice,” Neurobiology of Disease, vol. 15, no. 2, pp. 188–195, 2004.

F. Trinchese, S. Liu, F. Battaglia, S. Walter, P. M. Mathews, and O. Arancio, “Progressive age-related development of Alzheimer-like pathology in APP/PS1 mice,” Annals of Neurology, vol. 55, no. 6, pp. 801–814, 2004.

A. Megill, T. Tran, K. Eldred et al., “Defective age-dependent metaplasticity in a mouse model of Alzheimer's disease,” The Journal of Neuroscience, vol. 35, no. 32, pp. 11346–11357, 2015.

D. Puzzo, L. Privitera, E. Leznik et al., “Picomolar amyloid-β positively modulates synaptic plasticity and memory in hippocampus,” The Journal of Neuroscience, vol. 28, no. 53, pp. 14537–14545, 2008.

D. Puzzo, L. Privitera, and A. Palmeri, “Hormetic effect of amyloid-beta peptide in synaptic plasticity and memory,” Neurobiology of Aging, vol. 33, no. 7, pp. 1484–e24, 2012.

H. Hsieh, J. Boehm, C. Sato et al., “AMPAR removal underlies Abeta-induced synaptic depression and dendritic spine loss,” Neuron, vol. 52, no. 5, pp. 831–843, 2006.

G. M. Shankar, B. L. Bloodgood, M. Townsend, D. M. Walsh, D. J. Selkoe, and B. L. Sabatini, “Natural oligomers of the Alzheimer amyloid-β protein induce reversible synapse loss by modulating an NMDA-type glutamate receptor-dependent signaling pathway,” The Journal of Neuroscience, vol. 27, no. 11, pp. 2866–2875, 2007.

S. Li, S. Hong, N. E. Shepardson, D. M. Walsh, G. M. Shankar, and D. Selkoe, “Soluble oligomers of amyloid β protein facilitate hippocampal long-term depression by disrupting neuronal glutamate uptake,” Neuron, vol. 62, no. 6, pp. 788–801, 2009.

G. M. Shankar, S. Li, T. H. Mehta et al., “Amyloid-β protein dimers isolated directly from Alzheimer's brains impair synaptic plasticity and memory,” Nature Medicine, vol. 14, no. 8, pp. 837–842, 2008.

G. M. Shankar and D. M. Walsh, “Alzheimer's disease: synaptic dysfunction and Aβ,” Molecular Neurodegeneration, vol. 4, article 48, 2009.

M. S. Parihar and G. J. Brewer, “Amyloid-β as a modulator of synaptic plasticity,” Journal of Alzheimer's Disease, vol. 22, no. 3, pp. 741–763, 2010.

I. Klyubin, W. K. Cullen, N.-W. Hu, and M. J. Rowan, “Alzheimer's disease Aβ assemblies mediating rapid disruption of synaptic plasticity and memory,” Molecular Brain, vol. 5, no. 1, article 25, 2012.

C. F. Zorumski and Y. Izumi, “NMDA receptors and metaplasticity: mechanisms and possible roles in neuropsychiatric disorders,” Neuroscience and Biobehavioral Reviews, vol. 36, no. 3, pp. 989–1000, 2012.

W. C. Abraham, “Metaplasticity: tuning synapses and networks for plasticity,” Nature Reviews Neuroscience, vol. 9, no. 5, pp. 387–399, 2008.

J. J. Palop and L. Mucke, “Amyloid-beta-induced neuronal dysfunction in Alzheimer's disease: from synapses toward neural networks,” Nature Neuroscience, vol. 13, no. 7, pp. 812–818, 2010.

J. J. Palop and L. Mucke, “Synaptic depression and aberrant excitatory network activity in Alzheimer's disease: two faces of the same coin?” NeuroMolecular Medicine, vol. 12, no. 1, pp. 48–55, 2010.

M. A. Busche, X. Chen, H. A. Henning et al., “Critical role of soluble amyloid-β for early hippocampal hyperactivity in a mouse model of Alzheimer's disease,” Proceedings of the National Academy of Sciences of the United States of America, vol. 109, no. 22, pp. 8740–8745, 2012.

R. Lalonde, K.-I. Fukuchi, and C. Strazielle, “Neurologic and motor dysfunctions in APP transgenic mice,” Reviews in the Neurosciences, vol. 23, no. 4, pp. 363–379, 2012.

J. L. Jankowsky, H. H. Slunt, V. Gonzales et al., “Persistent amyloidosis following suppression of Aβ production in a transgenic model of Alzheimer disease,” PLoS Medicine, vol. 2, no. 12, article e355, 2005.

D. L. Vogt, D. Thomas, V. Galvan, D. E. Bredesen, B. T. Lamb, and S. W. Pimplikar, “Abnormal neuronal networks and seizure susceptibility in mice overexpressing the APP intracellular domain,” Neurobiology of Aging, vol. 32, no. 9, pp. 1725–1729, 2011.

J. C. Amatniek, W. A. Hauser, C. DelCastillo-Castaneda et al., “Incidence and predictors of seizures in patients with Alzheimer's disease,” Epilepsia, vol. 47, no. 5, pp. 867–872, 2006.

J. R. Cirrito, K. A. Yamada, M. B. Finn et al., “Synaptic activity regulates interstitial fluid amyloid-β levels in vivo,” Neuron, vol. 48, no. 6, pp. 913–922, 2005.

J. R. Cirrito, J.-E. Kang, J. Lee et al., “Endocytosis is required for synaptic activity-dependent release of amyloid-β in vivo,” Neuron, vol. 58, no. 1, pp. 42–51, 2008.

F. Kamenetz, T. Tomita, H. Hsieh et al., “APP processing and synaptic function,” Neuron, vol. 37, no. 6, pp. 925–937, 2003.

R. S. Wilson, E. Segawa, P. A. Boyle, S. E. Anagnos, L. P. Hizel, and D. A. Bennett, “The natural history of cognitive decline in Alzheimer's disease,” Psychology and Aging, vol. 27, no. 4, pp. 1008–1017, 2012.

K. Blennow, M. J. de Leon, and H. Zetterberg, “Alzheimer's disease,” The Lancet, vol. 368, no. 9533, pp. 387–403, 2006.

J. Hardy and D. J. Selkoe, “The amyloid hypothesis of Alzheimer's disease: progress and problems on the road to therapeutics,” Science, vol. 297, no. 5580, pp. 353–356, 2002.

R. E. Tanzi and L. Bertram, “Twenty years of the Alzheimer's disease amyloid hypothesis: a genetic perspective,” Cell, vol. 120, no. 4, pp. 545–555, 2005.

E. D. Roberson, K. Scearce-Levie, J. J. Palop et al., “Reducing endogenous tau ameliorates amyloid β-induced deficits in an Alzheimer's disease mouse model,” Science, vol. 316, no. 5825, pp. 750–754, 2007.

E. D. Roberson, B. Halabisky, J. W. Yoo et al., “Amyloid-β/fyn-induced synaptic, network, and cognitive impairments depend on tau levels in multiple mouse models of Alzheimer's disease,” The Journal of Neuroscience, vol. 31, no. 2, pp. 700–711, 2011.

L. M. Ittner, Y. D. Ke, F. Delerue et al., “Dendritic function of tau mediates amyloid-β toxicity in Alzheimer's disease mouse models,” Cell, vol. 142, no. 3, pp. 387–397, 2010.

J. P. Cleary, D. M. Walsh, J. J. Hofmeister et al., “Natural oligomers of the amyloid-β protein specifically disrupt cognitive function,” Nature Neuroscience, vol. 8, no. 1, pp. 79–84, 2005.

A. Nitta, A. Itoh, T. Hasegawa, and T. Nabeshima, “β-amyloid protein-induced Alzheimer's disease animal model,” Neuroscience Letters, vol. 170, no. 1, pp. 63–66, 1994.

T. Maurice, B. P. Lockhart, and A. Privat, “Amnesia induced in mice by centrally administered β-amyloid peptides involves cholinergic dysfunction,” Brain Research, vol. 706, no. 2, pp. 181–193, 1996.

Y.-F. Cheng, C. Wang, H.-B. Lin et al., “Inhibition of phosphodiesterase-4 reverses memory deficits produced by Aβ25-35 or Aβ1-40 peptide in rats,” Psychopharmacology, vol. 212, no. 2, pp. 181–191, 2010.

S. J. Webster, A. D. Bachstetter, P. T. Nelson, F. A. Schmitt, and L. J. Van Eldik, “Using mice to model Alzheimer's dementia: an overview of the clinical disease and the preclinical behavioral changes in 10 mouse models,” Frontiers in Genetics, vol. 5, artcile 88, Article ID Article 88, 2014.

D. Puzzo, L. Lee, A. Palmeri, G. Calabrese, and O. Arancio, “Behavioral assays with mouse models of Alzheimer's disease: practical considerations and guidelines,” Biochemical Pharmacology, vol. 88, no. 4, pp. 450–467, 2014.

B. De Strooper, P. Saftig, K. Craessaerts et al., “Deficiency of presenilin-1 inhibits the normal cleavage of amyloid precursor protein,” Nature, vol. 391, no. 6665, pp. 387–390, 1998.

R. Vassar, B. D. Bennett, S. Babu-Khan et al., “β-Secretase cleavage of Alzheimer's amyloid precursor protein by the transmembrane aspartic protease BACE,” Science, vol. 286, no. 5440, pp. 735–741, 1999.

D. Edbauer, E. Winkler, J. T. Regula, B. Pesold, H. Steiner, and C. Haass, “Reconstitution of γ-secretase activity,” Nature Cell Biology, vol. 5, no. 5, pp. 486–488, 2003.

B. Passer, L. Pellegrini, C. Russo et al., “Generation of an apoptotic intracellular peptide by α-secretase cleavage of Alzheimer's amyloid β protein precursor,” Journal of Alzheimer's Disease, vol. 2, no. 3-4, pp. 289–301, 2000.

E. H. Corder, A. M. Saunders, W. J. Strittmatter et al., “Gene dose of apolipoprotein E type 4 allele and the risk of Alzheimer's disease in late onset families,” Science, vol. 261, no. 5123, pp. 921–923, 1993.

A. J. C. Slooter, M. Cruts, S. Kalmijn et al., “Risk estimates of dementia by apolipoprotein E genotypes from a population-based incidence study: the Rotterdam study,” Archives of Neurology, vol. 55, no. 7, pp. 964–968, 1998.

J. M. Castellano, J. Kim, F. R. Stewart et al., “Human apoE isoforms differentially regulate brain amyloid-β peptide clearance,” Science Translational Medicine, vol. 3, no. 89, Article ID 89ra57, 2011.

Q. Zhou, K. J. Homma, and M.-M. Poo, “Shrinkage of dendritic spines associated with long-term depression of hippocampal synapses,” Neuron, vol. 44, no. 5, pp. 749–757, 2004.

N. Bastrikova, G. A. Gardner, J. M. Reece, A. Jeromin, and S. M. Dudek, “Synapse elimination accompanies functional plasticity in hippocampal neurons,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 8, pp. 3123–3127, 2008.

R. Lamprecht and J. LeDoux, “Structural plasticity and memory,” Nature Reviews Neuroscience, vol. 5, no. 1, pp. 45–54, 2004.

E. Masliah, M. Mallory, M. Alford et al., “Altered expression of synaptic proteins occurs early during progression of Alzheimer's disease,” Neurology, vol. 56, no. 1, pp. 127–129, 2001.

S. W. Scheff, D. A. Price, F. A. Schmitt, S. T. Dekosky, and E. J. Mufson, “Synaptic alterations in CA1 in mild Alzheimer disease and mild cognitive impairment,” Neurology, vol. 68, no. 18, pp. 1501–1508, 2007.

S. W. Scheff and D. A. Price, “Alzheimer's disease-related alterations in synaptic density: neocortex and hippocampus,” Journal of Alzheimer's Disease, vol. 9, supplement 3, pp. 101–115, 2006.

S. W. Scheff, D. A. Price, F. A. Schmitt, and E. J. Mufson, “Hippocampal synaptic loss in early Alzheimer's disease and mild cognitive impairment,” Neurobiology of Aging, vol. 27, no. 10, pp. 1372–1384, 2006.

H. L. Tan, B. N. Queenan, and R. L. Huganir, “GRIP1 is required for homeostatic regulation of AMPAR trafficking,” Proceedings of the National Academy of Sciences, vol. 112, no. 32, pp. 10026–10031, 2015.

M. A. Gainey, V. Tatavarty, M. Nahmani, H. Lin, and G. G. Turrigiano, “Activity-dependent synaptic GRIP1 accumulation drives synaptic scaling up in response to action potential blockade,” Proceedings of the National Academy of Sciences of the United States of America, vol. 112, no. 27, pp. E3590–E3599, 2015.

V. Anggono, R. L. Clem, and R. L. Huganir, “PICK1 loss of function occludes homeostatic synaptic scaling,” The Journal of Neuroscience, vol. 31, no. 6, pp. 2188–2196, 2011.

S. Alfonso, H. W. Kessels, C. C. Banos et al., “Synapto-depressive effects of amyloid beta require PICK1,” The European Journal of Neuroscience, vol. 39, no. 7, pp. 1225–1233, 2014.

J. D. Shepherd, G. Rumbaugh, J. Wu et al., “Arc/Arg3.1 mediates homeostatic synaptic scaling of AMPA receptors,” Neuron, vol. 52, no. 3, pp. 475–484, 2006.

K. Lee, S. E. Royston, M. O. Vest et al., “N-methyl-D-aspartate receptors mediate activity-dependent down-regulation of potassium channel genes during the expression of homeostatic intrinsic plasticity,” Molecular Brain, vol. 8, article 4, 2015.

J. Wu, R. S. Petralia, H. Kurushima et al., “Arc/Arg3.1 regulates an endosomal pathway essential for activity-dependent beta-amyloid generation,” Cell, vol. 147, no. 3, pp. 615–628, 2011.

J.-H. Hu, J. M. Park, S. Park et al., “Homeostatic scaling requires group I mGluR activation mediated by Homer1a,” Neuron, vol. 68, no. 6, pp. 1128–1142, 2010.

Q. Sun and G. G. Turrigiano, “PSD-95 and PSD-93 play critical but distinct roles in synaptic scaling up and down,” The Journal of Neuroscience, vol. 31, no. 18, pp. 6800–6808, 2011.

M. J. Kim, K. Futai, J. Jo, Y. Hayashi, K. Cho, and M. Sheng, “Synaptic accumulation of PSD-95 and synaptic function regulated by phosphorylation of serine-295 of PSD-95,” Neuron, vol. 56, no. 3, pp. 488–502, 2007.

J. Noritake, Y. Fukata, T. Iwanaga et al., “Mobile DHHC palmitoylating enzyme mediates activity-sensitive synaptic targeting of PSD-95,” The Journal of Cell Biology, vol. 186, no. 1, pp. 147–160, 2009.

A.-M. Simon, L. Schiapparelli, P. Salazar-Colocho et al., “Overexpression of wild-type human APP in mice causes cognitive deficits and pathological features unrelated to Aβ levels,” Neurobiology of Disease, vol. 33, no. 3, pp. 369–378, 2009.

F. Roselli, M. Tirard, J. Lu et al., “Soluble β-amyloid₁₋₄₀ induces NMDA-dependent degradation of postsynaptic density-95 at glutamatergic synapses,” The Journal of Neuroscience, vol. 25, no. 48, pp. 11061–11070, 2005.

F. Roselli, P. Hutzler, Y. Wegerich, P. Livrea, and O. F. X. Almeida, “Disassembly of shank and homer synaptic clusters is driven by soluble beta-amyloid1–40 through divergent NMDAR-dependent signalling pathways,” PLoS ONE, vol. 4, no. 6, Article ID e6011, 2009.

S. M. Shin, N. Zhang, J. Hansen et al., “GKAP orchestrates activity-dependent postsynaptic protein remodeling and homeostatic scaling,” Nature Neuroscience, vol. 15, no. 12, pp. 1655–1666, 2012.

F. Roselli, P. Livrea, and O. F. X. Almeida, “CDK5 is essential for soluble amyloid β-induced degradation of GKAP and remodeling of the synaptic actin cytoskeleton,” PLoS ONE, vol. 6, no. 7, Article ID e23097, 2011.

S. Kim and E. B. Ziff, “Calcineurin mediates synaptic scaling via synaptic trafficking of Ca²⁺-permeable AMPA receptors,” PLoS Biology, vol. 12, no. 7, Article ID e1001900, pp. 1–15, 2014.

M. D'Amelio, V. Cavallucci, S. Middei et al., “Caspase-3 triggers early synaptic dysfunction in a mouse model of Alzheimer's disease,” Nature Neuroscience, vol. 14, no. 1, pp. 69–76, 2011.

S. S. Jang, S. E. Royston, J. Xu et al., “Regulation of STEP61 and tyrosine-phosphorylation of NMDA and AMPA receptors during homeostatic synaptic plasticity,” Molecular Brain, vol. 8, no. 1, article 55, 2015.

Y. Zhang, D. V. Venkitaramani, C. M. Gladding et al., “The tyrosine phosphatase STEP mediates AMPA receptor endocytosis after metabotropic glutamate receptor stimulation,” The Journal of Neuroscience, vol. 28, no. 42, pp. 10561–10566, 2008.

P. Kurup, Y. Zhang, J. Xu et al., “Aβ-mediated NMDA receptor endocytosis in alzheimer's disease involves ubiquitination of the tyrosine phosphatase STEP61,” The Journal of Neuroscience, vol. 30, no. 17, pp. 5948–5957, 2010.

Y. Zhang, P. Kurup, J. Xu et al., “Genetic reduction of striatal-enriched tyrosine phosphatase (STEP) reverses cognitive and cellular deficits in an Alzheimer's disease mouse model,” Proceedings of the National Academy of Sciences of the United States of America, vol. 107, no. 44, pp. 19014–19019, 2010.

J. Xu, M. Chatterjee, T. D. Baguley et al., “Inhibitor of the tyrosine phosphatase STEP reverses cognitive deficits in a mouse model of Alzheimer's disease,” PLoS Biology, vol. 12, no. 8, Article ID e1001923, 2014.

B. A. Siddoway, H. F. Altimimi, H. Hou et al., “An essential role for inhibitor-2 regulation of protein phosphatase-1 in synaptic scaling,” The Journal of Neuroscience, vol. 33, no. 27, pp. 11206–11211, 2013.

M. Knobloch, M. Farinelli, U. Konietzko, R. M. Nitsch, and I. M. Mansuy, “Aβ oligomer-mediated long-term potentiation impairment involves protein phosphatase 1-dependent mechanisms,” The Journal of Neuroscience, vol. 27, no. 29, pp. 7648–7653, 2007.

S. L. Scudder, M. S. Goo, A. E. Cartier et al., “Synaptic strength is bidirectionally controlled by opposing activity-dependent regulation of Nedd4-1 and USP8,” The Journal of Neuroscience, vol. 34, no. 50, pp. 16637–16649, 2014.

Y.-D. Kwak, B. Wang, J. J. Li et al., “Upregulation of the E3 ligase NEDD4-1 by oxidative stress degrades IGF-1 receptor protein in neurodegeneration,” The Journal of Neuroscience, vol. 32, no. 32, pp. 10971–10981, 2012.

T. J. Craig, N. Jaafari, M. M. Petrovic, P. P. Rubin, J. R. Mellor, and J. M. Henley, “Homeostatic synaptic scaling is regulated by protein SUMOylation,” The Journal of Biological Chemistry, vol. 287, no. 27, pp. 22781–22788, 2012.

L. Lee, E. Dale, A. Staniszewski et al., “Regulation of synaptic plasticity and cognition by SUMO in normal physiology and Alzheimer's disease,” Scientific Reports, vol. 4, article 7190, 2014.

M. A. Sutton, A. M. Taylor, H. T. Ito, A. Pham, and E. M. Schuman, “Postsynaptic decoding of neural activity: eEF2 as a biochemical sensor coupling miniature synaptic transmission to local protein synthesis,” Neuron, vol. 55, no. 4, pp. 648–661, 2007.

M. Letellier, S. Elramah, M. Mondin et al., “MiR-92a regulates expression of synaptic GluA1-containing AMPA receptors during homeostatic scaling,” Nature Neuroscience, vol. 17, no. 8, pp. 1040–1042, 2014.

J. Aoto, C. I. Nam, M. M. Poon, P. Ting, and L. Chen, “Synaptic signaling by all-trans retinoic acid in homeostatic synaptic plasticity,” Neuron, vol. 60, no. 2, pp. 308–320, 2008.

M. M. Poon and L. Chen, “Retinoic acid-gated sequence-specific translational control by RARα,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 51, pp. 20303–20308, 2008.

B. Maghsoodi, M. M. Poon, C. I. Nam, J. Aoto, P. Ting, and L. Chen, “Retinoic acid regulates RARα-mediated control of translation in dendritic RNA granules during homeostatic synaptic plasticity,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 41, pp. 16015–16020, 2008.

D. K. Lahiri and C. Nall, “Promoter activity of the gene encoding the beta-amyloid precursor protein is up-regulated by growth factors, phorbol ester, retinoic acid and interleukin-1,” Molecular Brain Research, vol. 32, no. 2, pp. 233–240, 1995.

Y. Yang, W. W. Quitschke, and G. J. Brewer, “Upregulation of amyloid precursor protein gene promoter in rat primary hippocampal neurons by phorbol ester, IL-1 and retinoic acid, but not by reactive oxygen species,” Molecular Brain Research, vol. 60, no. 1, pp. 40–49, 1998.

J. G. Culvenor, G. Evin, M. A. Cooney et al., “Presenilin 2 expression in neuronal cells: induction during differentiation of embryonic carcinoma cells,” Experimental Cell Research, vol. 255, no. 2, pp. 192–206, 2000.

J.-I. Satoh and Y. Kuroda, “Amyloid precursor protein β-secretase (BACE) mRNA expression in human neural cell lines following induction of neuronal differentiation and exposure to cytokines and growth factors,” Neuropathology, vol. 20, no. 4, pp. 289–296, 2000.

Y. Ding, A. Qiao, Z. Wang et al., “Retinoic acid attenuates β-amyloid deposition and rescues memory deficits in an Alzheimer's disease transgenic mouse model,” The Journal of Neuroscience, vol. 28, no. 45, pp. 11622–11634, 2008.

L. C. Rutherford, S. B. Nelson, and G. G. Turrigiano, “BDNF has opposite effects on the quantal amplitude of pyramidal neuron and interneuron excitatory synapses,” Neuron, vol. 21, no. 3, pp. 521–530, 1998.

H. S. Phillips, J. M. Hains, M. Armanini, G. R. Laramee, S. A. Johnson, and J. W. Winslow, “BDNF mRNA is decreased in the hippocampus of individuals with Alzheimer's disease,” Neuron, vol. 7, no. 5, pp. 695–702, 1991.

B. Connor, D. Young, Q. Yan, R. L. M. Faull, B. Synek, and M. Dragunow, “Brain-derived neurotrophic factor is reduced in Alzheimer's disease,” Molecular Brain Research, vol. 49, no. 1-2, pp. 71–81, 1997.

A. Alvarez, R. Cacabelos, C. Sanpedro, M. Garcia-Fantini, and M. Aleixandre, “Serum TNF-alpha levels are increased and correlate negatively with free IGF-I in Alzheimer disease,” Neurobiology of Aging, vol. 28, no. 4, pp. 533–536, 2007.

J. S. Collins, R. T. Perry, B. Watson Jr. et al., “Association of a haplotype for tumor necrosis factor in siblings with late-onset Alzheimer disease: the NIMH Alzheimer disease genetics initiative,” American Journal of Medical Genetics, vol. 96, no. 6, pp. 823–830, 2000.

H. Fillit, W. Ding, L. Buee et al., “Elevated circulating tumor necrosis factor levels in Alzheimer's disease,” Neuroscience Letters, vol. 129, no. 2, pp. 318–320, 1991.

S. L. Ma, N. L. S. Tang, L. C. W. Lam, and H. F. K. Chiu, “Association between tumor necrosis factor-α promoter polymorphism and Alzheimer's disease,” Neurology, vol. 62, no. 2, pp. 307–309, 2004.

R. Paganelli, A. Di Iorio, L. Patricelli et al., “Proinflammatory cytokines in sera of elderly patients with dementia: levels in vascular injury are higher than those of mild-moderate Alzheimer's disease patients,” Experimental Gerontology, vol. 37, no. 2-3, pp. 257–263, 2002.

Z. S. Tan, A. S. Beiser, R. S. Vasan et al., “Inflammatory markers and the risk of Alzheimer disease: the Framingham study,” Neurology, vol. 68, no. 22, pp. 1902–1908, 2007.

E. Tobinick, H. Gross, A. Weinberger, and H. Cohen, “TNF-alpha modulation for treatment of Alzheimer's disease: a 6-month pilot study,” MedGenMed—Medscape General Medicine, vol. 8, no. 2, article 25, 2006.

D. Stellwagen and R. C. Malenka, “Synaptic scaling mediated by glial TNF-α,” Nature, vol. 440, no. 7087, pp. 1054–1059, 2006.

C. C. Steinmetz and G. G. Turrigiano, “Tumor necrosis factor-α signaling maintains the ability of cortical synapses to express synaptic scaling,” The Journal of Neuroscience, vol. 30, no. 44, pp. 14685–14690, 2010.

M. Kaneko, D. Stellwagen, R. C. Malenka, and M. P. Stryker, “Tumor necrosis factor-α mediates one component of competitive, experience-dependent plasticity in developing visual cortex,” Neuron, vol. 58, no. 5, pp. 673–680, 2008.

L. A. Cingolani, A. Thalhammer, L. M. Y. Yu et al., “Activity-dependent regulation of synaptic AMPA receptor composition and abundance by beta3 integrins,” Neuron, vol. 58, no. 5, pp. 749–762, 2008.

L. A. Cingolani and Y. Goda, “Differential involvement of β3 integrin in pre- and postsynaptic forms of adaptation to chronic activity deprivation,” Neuron Glia Biology, vol. 4, no. 3, pp. 179–187, 2008.

C. A. Goddard, D. A. Butts, and C. J. Shatz, “Regulation of CNS synapses by neuronal MHC class I,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 16, pp. 6828–6833, 2007.

K. Uemura, C. M. Lill, M. Banks et al., “N-cadherin-based adhesion enhances Aβ release and decreases Aβ 42/40 ratio,” Journal of Neurochemistry, vol. 108, no. 2, pp. 350–360, 2009.

M. Asada-Utsugi, K. Uemura, Y. Noda et al., “N-cadherin enhances APP dimerization at the extracellular domain and modulates Aβ production,” Journal of Neurochemistry, vol. 119, no. 2, pp. 354–363, 2011.

T. Okuda, L. M. Y. Yu, L. A. Cingolani, R. Kemler, and Y. Goda, “β-catenin regulates excitatory postsynaptic strength at hippocampal synapses,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 33, pp. 13479–13484, 2007.

N. Vitureira, M. Letellier, I. J. White, and Y. Goda, “Differential control of presynaptic efficacy by postsynaptic N-cadherin and β-catenin,” Nature Neuroscience, vol. 15, no. 1, pp. 81–89, 2012.

A. Andreyeva, K. Nieweg, K. Horstmann et al., “C-terminal fragment of N-cadherin accelerates synapse destabilization by amyloid-β,” Brain, vol. 135, no. 7, pp. 2140–2154, 2012.

A. K. Y. Fu, K.-W. Hung, W.-Y. Fu et al., “APC(Cdh1) mediates EphA4-dependent downregulation of AMPA receptors in homeostatic plasticity,” Nature Neuroscience, vol. 14, no. 2, pp. 181–191, 2011.

A. K. Y. Fu, K.-W. Hung, H. Huang et al., “Blockade of EphA4 signaling ameliorates hippocampal synaptic dysfunctions in mouse models of Alzheimer's disease,” Proceedings of the National Academy of Sciences of the United States of America, vol. 111, no. 27, pp. 9959–9964, 2014.

K. Ibata, Q. Sun, and G. G. Turrigiano, “Rapid synaptic scaling induced by changes in postsynaptic firing,” Neuron, vol. 57, no. 6, pp. 819–826, 2008.

C. P. Goold and R. A. Nicoll, “Single-cell optogenetic excitation drives homeostatic synaptic depression,” Neuron, vol. 68, no. 3, pp. 512–528, 2010.

M. A. Gainey, J. R. Hurvitz-Wolff, M. E. Lambo, and G. G. Turrigiano, “Synaptic scaling requires the GluR2 subunit of the AMPA receptor,” The Journal of Neuroscience, vol. 29, no. 20, pp. 6479–6489, 2009.

S. A. L. Corrêa, C. J. Hunter, O. Palygin et al., “MSK1 regulates homeostatic and experience-dependent synaptic plasticity,” The Journal of Neuroscience, vol. 32, no. 38, pp. 13039–13051, 2012.

K. M. Webber, M. A. Smith, H.-G. Lee et al., “Mitogen- and stress-activated protein kinase 1: convergence of the ERK and p38 pathways in Alzheimer's disease,” Journal of Neuroscience Research, vol. 79, no. 4, pp. 554–560, 2005.

M. P. Blackman, B. Djukic, S. B. Nelson, and G. G. Turrigiano, “A critical and cell-autonomous role for MeCP2 in synaptic scaling up,” The Journal of Neuroscience, vol. 32, no. 39, pp. 13529–13536, 2012.

D. P. Seeburg, M. Feliu-Mojer, J. Gaiottino, D. T. S. Pak, and M. Sheng, “Critical role of CDK5 and Polo-like kinase 2 in homeostatic synaptic plasticity during elevated activity,” Neuron, vol. 58, no. 4, pp. 571–583, 2008.

D. P. Seeburg and M. Sheng, “Activity-induced polo-like kinase 2 is required for homeostatic plasticity of hippocampal neurons during epileptiform activity,” The Journal of Neuroscience, vol. 28, no. 26, pp. 6583–6591, 2008.

A. Wilkaniec, G. A. Czapski, and A. Adamczyk, “Cdk5 at crossroads of protein oligomerization in neurodegenerative diseases: facts and hypotheses,” Journal of Neurochemistry, vol. 136, no. 2, pp. 222–233, 2016.

M. Marchetti and H. Marie, “Hippocampal synaptic plasticity in Alzheimer's disease: what have we learned so far from transgenic models?” Reviews in the Neurosciences, vol. 22, no. 4, pp. 373–402, 2011.

E. L. Bienenstock, L. N. Cooper, and P. W. Munro, “Theory for the development of neuron selectivity: orientation specificity and binocular interaction in visual cortex,” The Journal of Neuroscience, vol. 2, no. 1, pp. 32–48, 1982.

M. F. Bear, “Mechanism for a sliding synaptic modification threshold,” Neuron, vol. 15, no. 1, pp. 1–4, 1995.

R. A. Sperling, B. C. Dickerson, M. Pihlajamaki et al., “Functional alterations in memory networks in early Alzheimer's disease,” NeuroMolecular Medicine, vol. 12, no. 1, pp. 27–43, 2010.

M. A. Busche, G. Eichhoff, H. Adelsberger et al., “Clusters of hyperactive neurons near amyloid plaques in a mouse model of Alzheimer's disease,” Science, vol. 321, no. 5896, pp. 1686–1689, 2008.

A. J. Larner and M. Doran, “Reply to Dr Raux et al.: molecular diagnosis of autosomal dominant early onset Alzheimer's disease: an update (J Med Genet 2005;42:793–5),” Journal of Medical Genetics, vol. 43, no. 8, article e44, 2006.

S. Jayadev, J. B. Leverenz, E. Steinbart et al., “Alzheimer's disease phenotypes and genotypes associated with mutations in presenilin 2,” Brain, vol. 133, no. 4, pp. 1143–1154, 2010.

C. J. Westmark, P. R. Westmark, A. M. Beard, S. M. Hildebrandt, and J. S. Malter, “Seizure susceptibility and mortality in mice that over-express amyloid precursor protein,” International Journal of Clinical and Experimental Pathology, vol. 1, no. 2, pp. 157–168, 2008.

R. Lalonde, M. Dumont, M. Staufenbiel, and C. Strazielle, “Neurobehavioral characterization of APP23 transgenic mice with the SHIRPA primary screen,” Behavioural Brain Research, vol. 157, no. 1, pp. 91–98, 2005.

P. E. Sanchez, L. Zhu, L. Verret et al., “Levetiracetam suppresses neuronal network dysfunction and reverses synaptic and cognitive deficits in an Alzheimer's disease model,” Proceedings of the National Academy of Sciences of the United States of America, vol. 109, no. 42, pp. E2895–E2903, 2012.

L. Verret, E. O. Mann, G. B. Hang et al., “Inhibitory interneuron deficit links altered network activity and cognitive dysfunction in Alzheimer model,” Cell, vol. 149, no. 3, pp. 708–721, 2012.

B. F. Corbett, S. C. Leiser, H.-P. Ling et al., “Sodium channel cleavage is associated with aberrant neuronal activity and cognitive deficits in a mouse model of Alzheimer's disease,” The Journal of Neuroscience, vol. 33, no. 16, pp. 7020–7026, 2013.

R. Minkeviciene, S. Rheims, M. B. Dobszay et al., “Amyloid β-induced neuronal hyperexcitability triggers progressive epilepsy,” The Journal of Neuroscience, vol. 29, no. 11, pp. 3453–3462, 2009.

S. Ziyatdinova, K. Gurevicius, N. Kutchiashvili et al., “Spontaneous epileptiform discharges in a mouse model of Alzheimer's disease are suppressed by antiepileptic drugs that block sodium channels,” Epilepsy Research, vol. 94, no. 1-2, pp. 75–85, 2011.

S. Li, M. Jin, T. Koeglsperger, N. E. Shepardson, G. M. Shankar, and D. J. Selkoe, “Soluble Aβ oligomers inhibit long-term potentiation through a mechanism involving excessive activation of extrasynaptic NR2B-containing NMDA receptors,” The Journal of Neuroscience, vol. 31, no. 18, pp. 6627–6638, 2011.

N.-W. Hu, I. Klyubin, R. Anwy, and M. J. Rowan, “GluN2B subunit-containing NMDA receptor antagonists prevent Aβ-mediated synaptic plasticity disruption in vivo,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 48, pp. 20504–20509, 2009.

R. Rönicke, M. Mikhaylova, S. Rönicke et al., “Early neuronal dysfunction by amyloid β oligomers depends on activation of NR2B-containing NMDA receptors,” Neurobiology of Aging, vol. 32, no. 12, pp. 2219–2228, 2011.

Q. Yang, Z.-H. Liao, Y.-X. Xiao, Q.-S. Lin, Y.-S. Zhu, and S.-T. Li, “Hippocampal synaptic metaplasticity requires the activation of NR2B-containing NMDA receptors,” Brain Research Bulletin, vol. 84, no. 2, pp. 137–143, 2011.

T. Frankiewicz and C. G. Parsons, “Memantine restores long term potentiation impaired by tonic N-methyl-D-aspartate (NMDA) receptor activation following reduction of Mg²⁺ In hippocampal slices,” Neuropharmacology, vol. 38, no. 9, pp. 1253–1259, 1999.

W. Zajaczkowski, T. Frankiewicz, C. G. Parsons, and W. Danysz, “Uncompetitive NMDA receptor antagoists attenuate NMDA-induced impairment of passive avoidance learning and LTP,” Neuropharmacology, vol. 36, no. 7, pp. 961–971, 1997.

J. L. Jankowsky, H. H. Slunt, T. Ratovitski, N. A. Jenkins, N. G. Copeland, and D. R. Borchelt, “Co-expression of multiple transgenes in mouse CNS: a comparison of strategies,” Biomolecular Engineering, vol. 17, no. 6, pp. 157–165, 2001.

J. L. Jankowsky, D. J. Fadale, J. Anderson et al., “Mutant presenilins specifically elevate the levels of the 42 residue β-amyloid peptide in vivo: evidence for augmentation of a 42-specific γ secretase,” Human Molecular Genetics, vol. 13, no. 2, pp. 159–170, 2004.

A. Savonenko, G. M. Xu, T. Melnikova et al., “Episodic-like memory deficits in the APPswe/PS1dE9 mouse model of Alzheimer's disease: relationships to beta-amyloid deposition and neurotransmitter abnormalities,” Neurobiology of Disease, vol. 18, no. 3, pp. 602–617, 2005.

A. Volianskis, R. Køstner, M. Mølgaard, S. Hass, and M. S. Jensen, “Episodic memory deficits are not related to altered glutamatergic synaptic transmission and plasticity in the CA1 hippocampus of the APPswe/PS1ΔE9-deleted transgenic mice model of β-amyloidosis,” Neurobiology of Aging, vol. 31, no. 7, pp. 1173–1187, 2010.

E. M. Quinlan, D. H. Olstein, and M. F. Bear, “Bidirectional, experience-dependent regulation of N-methyl-D-aspartate receptor subunit composition in the rat visual cortex during postnatal development,” Proceedings of the National Academy of Sciences of the United States of America, vol. 96, no. 22, pp. 12876–12880, 1999.

E. M. Quinlan, D. Lebel, I. Brosh, and E. Barkai, “A molecular mechanism for stabilization of learning-induced synaptic modifications,” Neuron, vol. 41, no. 2, pp. 185–192, 2004.

B. D. Philpot, A. K. Sekhar, H. Z. Shouval, and M. F. Bear, “Visual experience and deprivation bidirectionally modify the composition and function of NMDA receptors in visual cortex,” Neuron, vol. 29, no. 1, pp. 157–169, 2001.

B. D. Philpot, J. S. Espinosa, and M. F. Bear, “Evidence for altered NMDA receptor function as a basis for metaplasticity in visual cortex,” The Journal of Neuroscience, vol. 23, no. 13, pp. 5583–5588, 2003.

A. Citri and R. C. Malenka, “Synaptic plasticity: multiple forms, functions, and mechanisms,” Neuropsychopharmacology, vol. 33, no. 1, pp. 18–41, 2008.

O. A. Shipton and O. Paulsen, “GluN2A and GluN2B subunit-containing NMDA receptors in hippocampal plasticity,” Philosophical Transactions of the Royal Society B: Biological Sciences, vol. 369, no. 1633, Article ID 20130163, 2014.

C. G. Lau and R. S. Zukin, “NMDA receptor trafficking in synaptic plasticity and neuropsychiatric disorders,” Nature Reviews Neuroscience, vol. 8, no. 6, pp. 413–426, 2007.

A. Barria and R. Malinow, “NMDA receptor subunit composition controls synaptic plasticity by regulating binding to CaMKII,” Neuron, vol. 48, no. 2, pp. 289–301, 2005.

E. M. Quinlan, B. D. Philpot, R. L. Huganir, and M. F. Bear, “Rapid, experience-dependent expression of synaptic NMDA receptors in visual cortex in vivo,” Nature Neuroscience, vol. 2, no. 4, pp. 352–357, 1999.

M. Matos, E. Augusto, C. R. Oliveira, and P. Agostinho, “Amyloid-beta peptide decreases glutamate uptake in cultured astrocytes: involvement of oxidative stress and mitogen-activated protein kinase cascades,” Neuroscience, vol. 156, no. 4, pp. 898–910, 2008.

S. Huang, M. Trevino, K. He et al., “Pull-push neuromodulation of LTP and LTD enables bidirectional experience-induced synaptic scaling in visual cortex,” Neuron, vol. 73, no. 3, pp. 497–510, 2012.

V. Anggono and R. L. Huganir, “Regulation of AMPA receptor trafficking and synaptic plasticity,” Current Opinion in Neurobiology, vol. 22, no. 3, pp. 461–469, 2012.

H.-K. Lee, K. Takamiya, J.-S. Han et al., “Phosphorylation of the AMPA receptor GluR1 subunit is required for synaptic plasticity and retention of spatial memory,” Cell, vol. 112, no. 5, pp. 631–643, 2003.

H.-K. Lee, M. Barbarosie, K. Kameyama, M. F. Bear, and R. L. Huganir, “Regulation of distinct AMPA receptor phosphorylation sites during bidirectional synaptic plasticity,” Nature, vol. 405, no. 6789, pp. 955–959, 2000.

H.-K. Lee, K. Takamiya, K. He, L. Song, and R. L. Huganir, “Specific roles of AMPA receptor subunit GluR1 (GluA1) phosphorylation sites in regulating synaptic plasticity in the CA1 region of hippocampus,” Journal of Neurophysiology, vol. 103, no. 1, pp. 479–489, 2010.

J. A. Esteban, S.-H. Shi, C. Wilson, M. Nuriya, R. L. Huganir, and R. Malinow, “PKA phosphorylation of AMPA receptor subunits controls synaptic trafficking underlying plasticity,” Nature Neuroscience, vol. 6, no. 2, pp. 136–143, 2003.

H.-Y. Man, Y. Sekine-Aizawa, and R. L. Huganir, “Regulation of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor trafficking through PKA phosphorylation of the Glu receptor 1 subunit,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 9, pp. 3579–3584, 2007.

D. Holman, M. Feligioni, and J. M. Henley, “Differential redistribution of native AMPA receptor complexes following LTD induction in acute hippocampal slices,” Neuropharmacology, vol. 52, no. 1, pp. 92–99, 2007.

A. Goel, L. W. Xu, K. P. Snyder et al., “Phosphorylation of AMPA receptors is required for sensory deprivation-induced homeostatic synaptic plasticity,” PLoS ONE, vol. 6, no. 3, Article ID e18264, 2011.

K. He, L. Song, L. W. Cummings, J. Goldman, R. L. Huganir, and H.-K. Lee, “Stabilization of Ca²⁺-permeable AMPA receptors at perisynaptic sites by GluR1-S845 phosphorylation,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 47, pp. 20033–20038, 2009.

K. W. Roche, R. J. O'Brien, A. L. Mammen, J. Bernhardt, and R. L. Huganir, “Characterization of multiple phosphorylation sites on the AMPA receptor GluR1 subunit,” Neuron, vol. 16, no. 6, pp. 1179–1188, 1996.

A. Barria, V. Derkach, and T. Soderling, “Identification of the Ca²⁺/calmodulin-dependent protein kinase II regulatory phosphorylation site in the α-amino-3-hydroxyl-5-methyl-4-isoxazole-propionate-type glutamate receptor,” The Journal of Biological Chemistry, vol. 272, no. 52, pp. 32727–32730, 1997.

A. L. Mammen, K. Kameyama, K. W. Roche, and R. L. Huganir, “Phosphorylation of the alpha-amino-3-hydroxy-5-methylisoxazole4-propionic acid receptor GluR1 subunit by calcium/calmodulin-dependent kinase II,” The Journal of Biological Chemistry, vol. 272, no. 51, pp. 32528–32533, 1997.

A. Barria, D. Muller, V. Derkach, L. C. Griffith, and T. R. Soderling, “Regulatory phosphorylation of AMPA-type glutamate receptors by CaM-KII during long-term potentiation,” Science, vol. 276, no. 5321, pp. 2042–2045, 1997.

X.-D. Hu, Q. Huang, X. Yang, and H. Xia, “Differential regulation of AMPA receptor trafficking by neurabin-targeted synaptic protein phosphatase-1 in synaptic transmission and long-term depression in hippocampus,” The Journal of Neuroscience, vol. 27, no. 17, pp. 4674–4686, 2007.

Y. Makino, R. C. Johnson, Y. Yu, K. Takamiya, and R. L. Huganir, “Enhanced synaptic plasticity in mice with phosphomimetic mutation of the GluA1 AMPA receptor,” Proceedings of the National Academy of Sciences of the United States of America, vol. 108, no. 20, pp. 8450–8455, 2011.

M.-C. Lee, R. Yasuda, and M. D. Ehlers, “Metaplasticity at single glutamatergic synapses,” Neuron, vol. 66, no. 6, pp. 859–870, 2010.

B. D. Philpot and R. S. Zukin, “Synapse-specific metaplasticity: to be silenced is not to silence 2B,” Neuron, vol. 66, no. 6, pp. 814–816, 2010.

G. G. Turrigiano, K. R. Leslie, N. S. Desai, L. C. Rutherford, and S. B. Nelson, “Activity-dependent scaling of quantal amplitude in neocortical neurons,” Nature, vol. 391, no. 6670, pp. 892–896, 1998.

A. J. Watt, M. C. W. Van Rossum, K. M. MacLeod, S. B. Nelson, and G. G. Turrigiano, “Activity coregulates quantal AMPA and NMDA currents at neocortical synapses,” Neuron, vol. 26, no. 3, pp. 659–670, 2000.

T. C. Thiagarajan, M. Lindskog, and R. W. Tsien, “Adaptation to synaptic inactivity in hippocampal neurons,” Neuron, vol. 47, no. 5, pp. 725–737, 2005.

D. V. Lissin, S. N. Gomperts, R. C. Carroll et al., “Activity differentially regulates the surface expression of synaptic AMPA and NMDA glutamate receptors,” Proceedings of the National Academy of Sciences of the United States of America, vol. 95, no. 12, pp. 7097–7102, 1998.

Z. Qiu, E. L. Sylwestrak, D. N. Lieberman, Y. Zhang, X.-Y. Liu, and A. Ghosh, “The Rett syndrome protein MeCP2 regulates synaptic scaling,” The Journal of Neuroscience, vol. 32, no. 3, pp. 989–994, 2012.

H. Dong, P. Zhang, I. Song, R. S. Petralia, D. Liao, and R. L. Huganir, “Characterization of the glutamate receptor-interacting proteins GRIP1 and GRIP2,” The Journal of Neuroscience, vol. 19, no. 16, pp. 6930–6941, 1999.

J. Xia, X. Zhang, J. Staudinger, and R. L. Huganir, “Clustering of AMPA receptors by the synaptic PDZ domain-containing protein PICK1,” Neuron, vol. 22, no. 1, pp. 179–187, 1999.

H. J. Chung, J. Xia, R. H. Scannevin, X. Zhang, and R. L. Huganir, “Phosphorylation of the AMPA receptor subunit GluR2 differentially regulates its interaction with PDZ domain-containing proteins,” The Journal of Neuroscience, vol. 20, no. 19, pp. 7258–7267, 2000.

J. L. Perez, L. Khatri, C. Chang, S. Srivastava, P. Osten, and E. B. Ziff, “PICK1 targets activated protein kinase Cα to AMPA receptor clusters in spines of hippocampal neurons and reduces surface levels of the AMPA-type glutamate receptor subunit 2,” The Journal of Neuroscience, vol. 21, no. 15, pp. 5417–5428, 2001.

A. Terashima, L. Cotton, K. K. Dev et al., “Regulation of synaptic strength and AMPA receptor subunit composition by PICK1,” The Journal of Neuroscience, vol. 24, no. 23, pp. 5381–5390, 2004.

D.-T. Lin and R. L. Huganir, “PICK1 and phosphorylation of the glutamate receptor 2 (GluR2) AMPA receptor subunit regulates GluR2 recycling after NMDA receptor-induced internalization,” The Journal of Neuroscience, vol. 27, no. 50, pp. 13903–13908, 2007.

E. Kim and M. Sheng, “PDZ domain proteins of synapses,” Nature Reviews Neuroscience, vol. 5, no. 10, pp. 771–781, 2004.

L. Chen, D. M. Chetkovich, R. S. Petralia et al., “Stargazin regulates synaptic targeting of AMPA receptors by two distinct mechanisms,” Nature, vol. 408, no. 6815, pp. 936–943, 2000.

R. A. Nicoll, S. Tomita, and D. S. Bredt, “Auxiliary subunits assist AMPA-type glutamate receptors,” Science, vol. 311, no. 5765, pp. 1253–1256, 2006.

Y.-H. Jiang and M. D. Ehlers, “Modeling autism by SHANK gene mutations in mice,” Neuron, vol. 78, no. 1, pp. 8–27, 2013.

E. Kim, S. Naisbitt, Y.-P. Hsueh et al., “GKAP, a novel synaptic protein that interacts with the guanylate kinase- like domain of the PSD-95/SAP90 family of channel clustering molecules,” The Journal of Cell Biology, vol. 136, no. 3, pp. 669–678, 1997.

S. Naisbitt, E. Kim, J. C. Tu et al., “Shank, a novel family of postsynaptic density proteins that binds to the NMDA receptor/PSD-95/GKAP complex and cortactin,” Neuron, vol. 23, no. 3, pp. 569–582, 1999.

J. F. Guzowski, J. A. Timlin, B. Roysam, B. L. McNaughton, P. F. Worley, and C. A. Barnes, “Mapping behaviorally relevant neural circuits with immediate-early gene expression,” Current Opinion in Neurobiology, vol. 15, no. 5, pp. 599–606, 2005.

O. Steward, C. S. Wallace, G. L. Lyford, and P. F. Worley, “Synaptic activation causes the mRNA for the IEG Arc to localize selectively near activated postsynaptic sites on dendrites,” Neuron, vol. 21, no. 4, pp. 741–751, 1998.

S. Chowdhury, J. D. Shepherd, H. Okuno et al., “Arc/Arg3.1 interacts with the endocytic machinery to regulate AMPA receptor trafficking,” Neuron, vol. 52, no. 3, pp. 445–459, 2006.

S. Park, J. M. Park, S. Kim et al., “Elongation factor 2 and fragile X mental retardation protein control the dynamic translation of Arc/Arg3.1 essential for mGluR-LTD,” Neuron, vol. 59, no. 1, pp. 70–83, 2008.

P. R. Brakeman, A. A. Lanahan, R. O'Brien et al., “Homer: a protein that selectively binds metabotropic glutamate receptors,” Nature, vol. 386, no. 6622, pp. 284–288, 1997.

F. Ango, L. Prézeau, T. Muller et al., “Agonist-independent activation of metabotropic glutamate receptors by the intracellular protein Homer,” Nature, vol. 411, no. 6840, pp. 962–965, 2001.

T. Hayashi and R. L. Huganir, “Tyrosine phosphorylation and regulation of the AMPA receptor by SRC family tyrosine kinases,” The Journal of Neuroscience, vol. 24, no. 27, pp. 6152–6160, 2004.

G. Ahmadian, W. Ju, L. Liu et al., “Tyrosine phosphorylation of GluR2 is required for insulin-stimulated AMPA receptor endocytosis and LTD,” The EMBO Journal, vol. 23, no. 5, pp. 1040–1050, 2004.

K. A. Pelkey, R. Askalan, S. Paul et al., “Tyrosine phosphatase STEP is a tonic brake on induction of long-term potentiation,” Neuron, vol. 34, no. 1, pp. 127–138, 2002.

R. M. Mulkey, S. Endo, S. Shenolikar, and R. C. Malenka, “Involvement of a calcineurin/inhibitor-1 phosphatase cascade in hippocampal long-term depression,” Nature, vol. 369, no. 6480, pp. 486–488, 1994.

W. Morishita, J. H. Connor, H. Xia, E. M. Quinlan, S. Shenolikar, and R. C. Malenka, “Regulation of synaptic strength by protein phosphatase 1,” Neuron, vol. 32, no. 6, pp. 1133–1148, 2001.

G. M. Thomas and R. L. Huganir, “Palmitoylation-dependent regulation of glutamate receptors and their PDZ domain-containing partners,” Biochemical Society Transactions, vol. 41, no. 1, pp. 72–78, 2013.

G. Yang, W. Xiong, L. Kojic, and M. S. Cynader, “Subunit-selective palmitoylation regulates the intracellular trafficking of AMPA receptor,” The European Journal of Neuroscience, vol. 30, no. 1, pp. 35–46, 2009.

T. Hayashi, G. Rumbaugh, and R. L. Huganir, “Differential regulation of AMPA receptor subunit trafficking by palmitoylation of two distinct sites,” Neuron, vol. 47, no. 5, pp. 709–723, 2005.

D.-T. Lin, Y. Makino, K. Sharma et al., “Regulation of AMPA receptor extrasynaptic insertion by 4.1N, phosphorylation and palmitoylation,” Nature Neuroscience, vol. 12, no. 7, pp. 879–887, 2009.

J. B. Hoppe, C. G. Salbego, and H. Cimarosti, “SUMOylation: novel neuroprotective approach for Alzheimer's disease?” Aging and Disease, vol. 6, no. 5, pp. 322–330, 2015.

S. Martin, A. Nishimune, J. R. Mellor, and J. M. Henley, “SUMOylation regulates kainate-receptor-mediated synaptic transmission,” Nature, vol. 447, no. 7142, pp. 321–325, 2007.

L. A. Schwarz, B. J. Hall, and G. N. Patrick, “Activity-dependent ubiquitination of GluA1 mediates a distinct AMPA receptor endocytosis and sorting pathway,” The Journal of Neuroscience, vol. 30, no. 49, pp. 16718–16729, 2010.

A. Lin, Q. Hou, L. Jarzylo et al., “Nedd4-mediated AMPA receptor ubiquitination regulates receptor turnover and trafficking,” Journal of Neurochemistry, vol. 119, no. 1, pp. 27–39, 2011.

M. P. Lussier, Y. Nasu-Nishimura, and K. W. Roche, “Activity-dependent ubiquitination of the AMPA receptor subunit GluA2,” The Journal of Neuroscience, vol. 31, no. 8, pp. 3077–3081, 2011.

J. P. Meadows, M. C. Guzman-Karlsson, S. Phillips et al., “DNA methylation regulates neuronal glutamatergic synaptic scaling,” Science Signaling, vol. 8, no. 382, p. ra61, 2015.

A. Balkowiec and D. M. Katz, “Cellular mechanisms regulating activity-dependent release of native brain-derived neurotrophic factor from hippocampal neurons,” The Journal of Neuroscience, vol. 22, no. 23, pp. 10399–10407, 2002.

J. Budni, T. Bellettini-Santos, F. Mina, M. Lima Garcez, and A. Ioppi Zugno, “The involvement of BDNF, NGF and GDNF in aging and Alzheimer's disease,” Aging and Disease, vol. 6, no. 5, pp. 331–341, 2015.

G. Leal, P. M. Afonso, I. L. Salazar, and C. B. Duarte, “Regulation of hippocampal synaptic plasticity by BDNF,” Brain Research, vol. 1621, pp. 82–101, 2015.

S. Peng, D. J. Garzon, M. Marchese et al., “Decreased brain-derived neurotrophic factor depends on amyloid aggregation state in transgenic mouse models of Alzheimer's disease,” The Journal of Neuroscience, vol. 29, no. 29, pp. 9321–9329, 2009.

M. A. Sutton, H. T. Ito, P. Cressy, C. Kempf, J. C. Woo, and E. M. Schuman, “Miniature neurotransmission stabilizes synaptic function via tonic suppression of local dendritic protein synthesis,” Cell, vol. 125, no. 4, pp. 785–799, 2006.

I. J. Cajigas, G. Tushev, T. J. Will, S. Tom Dieck, N. Fuerst, and E. M. Schuman, “The local transcriptome in the synaptic neuropil revealed by deep sequencing and high-resolution imaging,” Neuron, vol. 74, no. 3, pp. 453–466, 2012.

W. Ju, W. Morishita, J. Tsui et al., “Activity-dependent regulation of dendritic synthesis and trafficking of AMPA receptors,” Nature Neuroscience, vol. 7, no. 3, pp. 244–253, 2004.

A. B. McGeachie, L. A. Cingolani, and Y. Goda, “A stabilising influence: integrins in regulation of synaptic plasticity,” Neuroscience Research, vol. 70, no. 1, pp. 24–29, 2011.

S. Mortillo, A. Elste, Y. Ge et al., “Compensatory redistribution of neuroligins and N-cadherin following deletion of synaptic β1-integrin,” Journal of Comparative Neurology, vol. 520, no. 9, pp. 2041–2052, 2012.

D. T. S. Pak, S. Yang, S. Rudolph-Correia, E. Kim, and M. Sheng, “Regulation of dendritic spine morphology by SPAR, a PSD-95-associated RapGAP,” Neuron, vol. 31, no. 2, pp. 289–303, 2001.

K. Kubota, K. Inoue, R. Hashimoto et al., “Tumor necrosis factor receptor-associated protein 1 regulates cell adhesion and synaptic morphology via modulation of N-cadherin expression,” Journal of Neurochemistry, vol. 110, no. 2, pp. 496–508, 2009.

P. D. Drew, M. Lonergan, M. E. Goldstein, L. A. Lampson, K. Ozato, and D. E. McFarlin, “Regulation of MHC class I and β2-microglobulin gene expression in human neuronal cells. Factor binding to conserved cis-acting regulatory sequences correlates with expression of the genes,” The Journal of Immunology, vol. 150, no. 8, part 1, pp. 3300–3310, 1993.

B. C. Albensi and M. P. Mattson, “Evidence for the involvement of TNF and NF-κB in hippocampal synaptic plasticity,” Synapse, vol. 35, no. 2, pp. 151–159, 2000.

M. P. Mattson, B. Cheng, D. Davis, K. Bryant, I. Lieberburg, and R. E. Rydel, “β-Amyloid peptides destabilize calcium homeostasis and render human cortical neurons vulnerable to excitotoxicity,” The Journal of Neuroscience, vol. 12, no. 2, pp. 376–389, 1992.

M. Nuriya and R. L. Huganir, “Regulation of AMPA receptor trafficking by N-cadherin,” Journal of Neurochemistry, vol. 97, no. 3, pp. 652–661, 2006.

L. Saglietti, C. Dequidt, K. Kamieniarz et al., “Extracellular interactions between GluR2 and N-cadherin in spine regulation,” Neuron, vol. 54, no. 3, pp. 461–477, 2007.

H. Togashi, K. Abe, A. Mizoguchi, K. Takaoka, O. Chisaka, and M. Takeichi, “Cadherin regulates dendritic spine morphogenesis,” Neuron, vol. 35, no. 1, pp. 77–89, 2002.

L. Tang, C. P. Hung, and E. M. Schuman, “A role for the cadherin family of cell adhesion molecules in hippocampal long-term potentiation,” Neuron, vol. 20, no. 6, pp. 1165–1175, 1998.

O. Bozdagi, W. Shan, H. Tanaka, D. L. Benson, and G. W. Huntley, “Increasing numbers of synaptic puncta during late-phase LTP: N-cadherin is synthesized, recruited to synaptic sites, and required for potentiation,” Neuron, vol. 28, no. 1, pp. 245–259, 2000.

O. Bozdagi, X.-B. Wang, J. S. Nikitczuk et al., “Persistence of coordinated long-term potentiation and dendritic spine enlargement at mature hippocampal CA1 synapses requires N-cadherin,” The Journal of Neuroscience, vol. 30, no. 30, pp. 9984–9989, 2010.

P. Mendez, M. De Roo, L. Poglia, P. Klauser, and D. Muller, “N-cadherin mediates plasticity-induced long-term spine stabilization,” The Journal of Cell Biology, vol. 189, no. 3, pp. 589–600, 2010.