Capturing activated neurons and synapses

Journal Pre-proof Capturing activated neurons and synapses Jung-Eun Choi, Jiwon Kim, Jinhyun Kim

PII:

S0168-0102(19)30656-X

DOI:

https://doi.org/10.1016/j.neures.2019.12.020

Reference:

NSR 4353

To appear in:

Neuroscience Research

Received Date:

4 December 2019

Accepted Date:

27 December 2019

Please cite this article as: Choi J-Eun, Kim J, Kim J, Capturing activated neurons and synapses, Neuroscience Research (2019), doi: https://doi.org/10.1016/j.neures.2019.12.020

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Capturing activated neurons and synapses

Jung-Eun Choi1,3, Jiwon Kim1,3, and Jinhyun Kim1,2,* 1Center

for Functional Connectomics, Korea Institute of Science and Technology (KIST) and 2Division of Bio-Medical Science & Technology, KIST-School, University of Science and Technology, Seoul, Korea 3 Co-first

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author *Correspondence should be addressed to Jinhyun Kim, Center for Functional Connectomics (CFC), L7-7211, Brain Science Institute at the Korea Institute of Science & Technology (KIST), 39-1 Hawolgokdong, Seongbukgu, Seoul 02792, Korea; phone: +82-2-958-7225, [email protected]

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Number of Pages: 28 Figures: 3 Tables: 1

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Advances in IEG-based gene expression system in response to neural activity enabled access to activated neuronal ensemble. Photoactivatable gene expression system in response to neural activity also had great advances recently. Advances in active synapse labeling system allow capturing of activated synapses. New approaches for manipulation of activated synapses have been developed.

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Highlights

Abstract

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An important yet very challenging goal of neuroscience is to understand how brain activity drives cognition and behavior. Many useful tools have been developed to study neurons and synapses, the fundamental units of brain activity. Here, we review recently developed methods to visualize and manipulate active neurons and synapses, providing useful and compelling information about functional neuronal circuitry.

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Keywords Cell ensemble, Active synapse, IEG expression, Ca2+, Light, Genetic tools

1. Introduction

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Mapping active neural circuitry is a fundamental necessity and a challenge for understanding the brain activity mediating cognition and complex behaviors. Because brain activity is shaped by the functional profiles of neurons and synapses, it is essential to understand how a given set of neural ensembles and its synapses contribute to particular functional outcomes within a given timeframe (Asok et al., 2019; Poo et al., 2016; Tonegawa et al., 2018). Therefore, many researchers are working to develop neurotechnologies to take snapshots of neural activity as it progresses and changes (Miesenböck et al., 1998; Reijmers et al., 2007; Tian et al., 2009; Wang et al., 2006). The methods most commonly used to investigate neuronal activity are based on (1) activity-driven expression of immediate early genes (IEGs), (2) transient increases in intracellular Ca2+ concentration, and (3) synaptic events such as vesicle release.

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First, IEGs, such as Fos and Arc, are transiently expressed in cells following input that causes neural activity, allowing their expression patterns to provide an enduring readout of recent patterns of activity within neural networks (Link et al., 1995; Lyford et al., 1995; Morgan et al., 1987; Saffen et al., 1988; Sagar et al., 1988; Worley et al., 1991). Traditionally, IEG promoters have been used to drive either directly a reporter gene such as lacZ and GFP (Barth et al., 2004; Smeyne et al., 1992; Wang et al., 2006) or a further genetic switch such as tTA and Cre (Guenthner et al., 2013; Reijmers et al., 2007) to study a variety of neuronal activities participating in a variety of functions, such as the encoding of memory engrams (DeNardo and Luo, 2017; He et al., 2018; Luo et al., 2018; Mayford and Reijmers, 2015). More recently, new IEG promoter-based methods have been developed to map patterns of neuronal activity with high temporal precision and low background noise. We review these below. Second, neural activity causes rapid changes in intracellular free calcium levels. By imaging and quantifying these changes, one can identify active neurons and characterize their responses, including spike number, timing, and frequency (Luo et al., 2018). Advanced calcium indicators (e.g. genetically encoded calcium indicators, GECIs) allow quantitative optical recording of cellular-resolution activity in awake and behaving animals with cell-type specificity (Akerboom et al., 2012; Chen et al., 2013; Dana et al., 2019; Inoue et al., 2019; Tian et al., 2009). However, calcium imaging requires real-time monitoring, typically within a limited field of view covering only a small fraction of a microcircuit. Even the recently-developed wide field-of-view approach of twin-region two-photon microscopy (Stirman et al., 2016) is restricted

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mostly to the cortical surface a couple of millimeters wide, making it difficult to trace neuronal activity brain-wide and over long periods of time. Last, neuronal activity triggers synaptic events, such as vesicle release, that are mediated by key synaptic proteins such as synaptophysin, vesicle-associated membrane protein 2 (VAMP2), postsynaptic density protein-95 (PSD-95), calcium/calmodulin-dependent kinase II (CaMKII), α-amino-hydroxy-5-methyl-4isoxazolepropionic acid receptor (AMPAR) and others (Edelmann et al., 1995; Lisman et al., 2002; Rizo and Rosenmund, 2008). Changes in these synaptic proteins can be monitored by tagging fluorescence proteins (FPs), including pH-sensitive FPs, providing another way to map synaptic activity on a relatively fine level of detail (Ahmari et al., 2000; Lee et al., 2016; Li and Tsien, 2012; Miesenböck et al., 1998; Ueda et al., 2013; Y asuda, 2006). Recently, there have been notable advances in synaptic protein-based tools for mapping structural and functional synapses. In this review, we describe new methods to visualize and manipulate active neurons and synapses, providing useful and compelling information about functional neuronal circuitry (Table 1).

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2. Advances in activity-dependent gene expression system

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Many genetic strategies have been developed to translate neuronal activation into gene expression, allowing neuroscientists not only to visualize active neuronal populations but also to genetically access, and in some cases even manipulate, recently activated neurons. These approaches utilize the expression of IEGs or the transient rise in intracellular Ca2+ concentration as reporters of neuronal activity (Figure 1 and 2).

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2-1. IEG promoter-based approaches

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TetTag and TRAP: IEG promoters were originally used to control the expression of visually detectable reporters (e.g. EGFP), yet the relatively high basal level and transient time window of IEG expression made it difficult to mark active neuronal populations. To clearly and specifically label activated neurons, inducible genetic switches such as tetracycline-controlled trans-activator (tTA) and Cre recombinase (Cre) were introduced, greatly enhancing IEG-based methods. This approach uses inducible tTA and Cre under the control of an IEG promoter, combined with the expression of a transgene (e.g. EGFP, AMPAR, etc.) regulated by the time-specific administration of specific drugs, such as doxycycline (Dox) and tamoxifen, respectively (Allen et al., 2017; Garner et al., 2012; Guenthner et al., 2013; Liu et al., 2012) The combination of Fos promoter and tTA system in a transgenic line, together called TetTag, has been a successful example of tagging neurons activated during a given time window (Groves et al., 2018; Ohkawa et al., 2015; Okuyama et al., 2016; Reijmers et al., 2007; Tanaka et al., 2014; Tayler et al., 2013) (Figure 1). TetTag mice

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were used to demonstrate that distinct neuronal populations in the basolateral amygdala are activated during fear learning and memory retrieval (Reijmers et al., 2007). However, this technique can be improved, for example, by reducing the relatively high level of non-specific labeling and increasing the precision of temporal regulation caused by the slow metabolism of Dox. Furthermore, combination of TetTag with opto-/chemo-genetic tools have demonstrated the manipulation of activated cell ensembles in various behavioral paradigms (Garner et al., 2012; Liu et al., 2012; Ramirez et al., 2013). Recently developed Targeted Recombination in Active Populations (TRAP) is analogous to TetTag, except it uses tamoxifen-inducible Cre (CreERT2) rather than tTA, controlled by Fos and Arc promoters, called FosTRAP or ArcTRAP, respectively (Guenthner et al., 2013). The TRAP system has been further optimized with an improved knock-in method: it uses an improved Cre (iCre) and the highly effective inducer 4-hydroxytamoxifen (4-OHT) instead of tamoxifen. TRAP2, an optimized version of FosTRAP, was recently used to demonstrate that neuronal ensembles in prelimbic cortex are active during learning and memory retrieval (DeNardo et al., 2019). In line with TetTag, TRAP can also be combined with opto-/chemo-genetic approaches to investigate the function of TRAPed cells (Girasole et al., 2018; Ishii et al., 2017; Kim and Cho, 2017).

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vGATE: A useful virus-mediated combinatorial method has recently been reported (Hasan et al., 2019). Taking advantage of well-established genetic switches such as tTA (Gossen and Bujard, 1992) and Cre (Sauer and Henderson, 1988) systems in multisteps, this new approach, called virus-delivered Genetic Activity-induced Tagging of cell Ensembles (vGATE), selectively labels activated neuron ensemble in a cell typespecific manner. This system is composed of three AAV-mediated transgenes (i.e. Fos promoter-driven rtTA, rtTA-driven Cre, and a Cre-driven functional gene), which are delivered as a mixture of the viruses. Like TetTag, reverse tTA (rtTA) is expressed under the control of a Fos promoter to target active neurons. Then, in the presence of Dox, rtTA turns on Cre expression. Once optimized, this technique provides a tightlycontrolled capturing time window regulated by the Dox injection in vivo. After the injection, Cre switches a functional gene on under the control of a cell-type-specific promoter. In addition to Cre, a second transgene, for instance a genetically-encoded calcium indicator, can be expressed using the bidirectional Tet promoter, providing the useful option of direct monitoring of the changes in intracellular calcium levels as an additional readout of neural activity. These sequential, multi-step strategies were recently used effectively to investigate a fear memory engram (Hasan et al., 2019). CANE: So far, we have described methods using genetic switches controlled by IEG promoters. Another approach, Capturing Activated Neuronal Ensembles (CANE), also a combinatorial method, uses an exogenous avian-specific viral receptor (TVA) under

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the control of Fos promoter (Sakurai et al., 2016). In CANE, destabilized TVA (dsTVA) is knocked-in to the endogenous Fos locus, resulting in rapid but brief expression of TVA on the surface membrane of activated Fos+ neurons. Next, these activated, dsTVAexpressing neurons are specifically labeled with lentiviruses or trans-synaptic rabies viruses coated with a surface glycoprotein (EnvA), which mediates TVA-dependent virus entry to the neuron. The brief half-life of dsTVA, combined with the experimenter’s choice of time points for EnvA-expressing viral injection, allows for great temporal precision. CANE appears to effectively capture small subsets of activated neurons in various brain regions at distinct time points. Further, it allows the manipulation of specific neuronal populations via delivery of visually detectable FP reporters and manipulators such as channelrhodopsin and tetanus toxin (Jiang-Xie et al., 2019; Rodriguez et al., 2017; Sakurai et al., 2016; Tschida et al., 2019).

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E-SARE and RAM: While endogenous IEG promoters have been combined with various genetic switches and delivery systems, their recombinant promoters have recently been optimized in several ways to improve the efficiency and specificity of capturing activated neurons. One example is enhanced synaptic activity-responsive element (ESARE), a 975 base pair short engineered promoter composed of a minimal Arc promoter and five repeats of the synaptic activity-responsive element (SARE), a neuronal activity enhancer (Kawashima et al., 2013). SARE is known to bind to activitydependent transcription factors (CREB, MEF2, and SRF) (Kawashima et al., 2009). The tandem SARE elements, which regulate Arc induction through interactions with these transcription factors, augment reporter gene expression. Like TetTag, TRAP, and vGATE, E-SARE can be combined with CreERT2 to permanently label active neurons with rapid induction rate, a combination successfully used to investigate neuronal ensembles in visual cortex and hippocampus (Attardo et al., 2018; Kawashima et al., 2013). Another example, Robust Activity Marking system (RAM), is also a synthetic short activity-dependent promoter (Sørensen et al., 2016). The RAM promoter (199 base pairs) is comprised of a minimal Fos promoter and four repeats of an enhancer module containing the activator protein 1 (AP-1) site and a Npas4 binding motif. The AP-1 site was found to be the most highly enriched motif in neuronal activity-regulated enhancers and a consensus sequence for the Fos/Jun family transcription factors. The binding motif of the neuron-specific activity-dependent gene Npas4 was used to increase the efficiency and specificity of labeling activated neurons. RAM can also be combined with the tTA system. Because the RAM promoter is small, all necessary gene cassettes (tTA under RAM promoter and a tTA-driven reporter gene) can be packaged into a single AAV, the most widely used vector by neuroscientists. The RAM system has been used successfully to label engram cells in hippocampus activated during contextual fear conditioning (Sørensen et al., 2016). 2-2. Photoactivatable approaches

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IEG-based activity labeling tools have been used extensively to identify neuronal ensembles activated during behavioral experiences, particularly to determine engram cells in learning and memory. Yet, despite the recent advances in IEG-based approaches described above, methods using IEG expression as assays of neural activity share several disadvantages inherent with IEGs, including their relatively slow temporal resolution of hours to days and their relatively low signal compared to baseline expression. Further, certain cell types and brain regions show little IEG expression, restricting the utility of these techniques (Applegate et al., 1995; Labiner et al., 1993). Moreover, a recent study demonstrated that only a fraction of active hippocampal place cells are Fos+ engram neurons participating in memory encoding and specific cell ensembles (Tanaka et al., 2018). However, other approaches are available. For example, a number of tools have been recently developed to take snapshots of neural responses based on activity-dependent transient increases in intracellular Ca2+ concentration (Lin and Schnitzer, 2016; Weisenburger and Vaziri, 2018), including activity-dependent tools that are switched on by the coincidence of calcium elevation and light stimulation (Lee et al., 2017; Wang et al., 2017) (Figure 2).

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Cal-Light and FLARE: The Ca2+-binding protein calmodulin (CaM) and Ca2+/CaM-binding motif, e.g. M13, provided the foundation for GECIs, which have been useful for monitoring changes in intracellular Ca2+ concentration in various brain regions and behavioral paradigms (Broussard et al., 2014; Mank and Griesbeck, 2008). Recently, advances in light-activatable tools have made it possible to manipulate neuronal activity and molecular behaviors of certain key proteins with great spatial and temporal precision (Boyden et al., 2005; Zhang et al., 2007). Two such tools using Ca2+- and lightcontrolled genetic switches, called Cal-Light and Fast Light- and Activity-Regulated Expression (FLARE), were designed to convert neuronal activity into gene expression (Lee et al., 2017; Wang et al., 2017). Both Cal-Light and FLARE use the cleavable tTA in Ca2+- and light-dependent manners, based on five key components: CaM, Ca2+/CaM-binding domain, a lightoxygen-voltage sensing (LOV) domain, tobacco etch virus proteinase (TEVp), and tTA fused with TEVp cleavage sequence (TEVseq). The Ca2+- and photo-cleavable tTA combination was designed to be functionally blocked by being tethered to the plasma membrane far from the nucleus, the location of transcription. When Ca2+ rises (indicating neuronal activity) and light is flashed (providing control over timing), the tTA is released and translocates into its transcription site, mediated by two consequential events: (1) light-sensitive uncaging of the cleavage site (TEVseq) by a conformational change of the LOV domain; and (2) Ca2+-sensitive recruitment of the protease (TEVp) by intermolecular interactions between CaM and the Ca2+/CaMbinding domain. The cleaved and translocated tTA then switches on expression of the gene of interest. Cal-Light and FLARE share common working principles with small

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technical differences that can be adjusted to optimize the efficiency, sensitivity, and other factors for any given application. By using an optogenetic approach to trigger gene expression, both FLARE and Cal-Light achieve high temporal precision in activity mapping in vivo. These tools are also versatile and can be combined with other genetic strategies for specific applications.

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PA-Cre and -Flp: Other new light-controllable gene expression systems are available, too. Here, we describe two photoactivatable (PA) Cre and Flp recombinase systems that are both based on light-sensitive intermolecular dimerization. One, derived from Arabidopsis thaliana, exploits blue light-induced interactions between Cryptochrome 2 (CRY2) and CIB1 (Liu et al., 2008). The other, called Magnets, derived from the filamentous fungus Neurospora crassa, is based on blue light-induced homodimerization of the photoreceptor VVD (Kawano et al., 2015). To accomplish light-controllable gene expression, split N- and C-fragments of Cre and Flp were fused with CRY2, CIBN (1-170 amino acid residues of CIB1 (Liu et al., 2008)) (Kennedy et al., 2010), and a dimerization pair of Magnet (Jung et al., 2019), respectively. Blue light illumination causes CRY2 to bind to CIBN and causes monomers of Magnet to pair with each other; both reactions result in the reconstitution of functional recombinases. These photoactivatable genetic switches have been used to demonstrate that gene expression is controlled in the brain with great spatial and temporal precision (Schindler et al., 2015). These techniques have recently been extended with newly engineered photo-sensitive molecules to allow the study of deep brain regions with noninvasive LED opto-stimulation (Jung et al., 2019). Further improvements in photoactivatable genetic switches are under development (Meador et al., 2019; Taslimi et al., 2016). Although the current versions of PA-Cre and Flp systems have been demonstrated only as a general genetic switch, these systems seem to have a great potential to capture and manipulate neural activity. For instance, these systems can provide better temporal precision than previous drug-inducible gene expression systems, when combined with IEG promoter likely TRAP. Creative combinations of PAgenetic switches with readily available techniques for labeling or manipulating neuronal and synaptic activity are contributing to our understanding of brain functions.

3. Advances in active synapse labeling system The synapse is a primary structural and functional unit underlying brain activity associated with behaviors and learning mechanisms. Mapping cell ensembles will not provide enough information to understand brain activity—it is important to map active synapses, as well. Historically, the main sensors of synaptic activity have been pHsensitive FP-fused synaptic proteins, such as synaptophysin and VAMP2, as previously reviewed (Lee et al., 2016; Li and Tsien, 2012; Miesenböck et al., 1998). Recently, these

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sensors have fallen out of favor because of their relatively poor signal-to-noise characteristics and low sensitivity. But, synaptic proteins are currently providing the basis for updated molecular tools for visualizing synaptic events in vivo (Figure 3).

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SynaptoZIP: VAMP2 (also known as Synaptobrevin-2) is the most abundant synaptic vesicle trans-membrane protein (Takamori et al., 2006) and plays a critical role in vesicle fusion and neurotransmitter release at the presynaptic terminus. A new method, called SynaptoZIP, was recently developed to label active synapses by visualizing VAMP2-mediated synaptic events (Ferro et al., 2017). The key molecular event in the function of SynaptoZIP is the Velcro coiled-coil heteromerization between ZIP and Synbond (SB), similar to leucine zipper dimerization (O'Shea et al., 1993). Unor GFP-tagged VAMP2 is fused with ZIP at its C-terminus, facing the inside of synaptic vesicles. Upon stimulation, ZIP is exposed to the extracellular space where it forms a Velcro coiled-coil heterodimer with soluble fluorophore-conjugated SB. The resulting SynaptoZIP-SB complex is internalized into the vesicle lumen, enabling the visualization of endocytic vesicles. Once generated, the complex is stable and will not undergo substitution with other color fluorophore-conjugated SB molecules up to 48 hours after initial vesicular uptake, and thereby reports vesicular endocytosis with a persistent signal. Due to its long-term stability and multiple color availability, retrospective analysis and multiple chasing of synaptic activity using SB with different fluorescence are possible. SynaptoZIP has been successfully used in vivo to label active synapses in cortical neurons (Ferro et al., 2017). Although this technique leaves room for improvement (for instance, by reducing background signals from remaining SynaptoZIP-SB complexes at the synaptic cleft), SynaptoZIP offers a powerful tool for measuring neural activity in vivo, providing a direct readout at the synaptic level.

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SynTagMA (Synaptic CaMPARI): Another useful tool to label active synapses is Synaptic Tag for Mapping Activity (SynTagMA) (Perez-Alvarez et al., 2019). SynTagMA is based on the synapse-targeted Calcium-Modulated Photoactivatable Ratiometric Integrator (CaMPARI) to map synaptic activity within a time window on the order of one second (Fosque et al., 2015). CaMPARI is an advanced Ca2+ indicator using CaM and M13 motif; it is similar to GCaMP series, but uses a circularly permuted photoconvertible FP, i.e. mEOS2, instead. When experimenter-controlled light illumination coincides with Ca2+ binding, CaMPARI undergoes irreversible photoconversion from green to red fluorescence. Thereby, CaMPARI creates a temporally precise activity snapshot of a large brain area. Recently, CaMPARI has been improved by the introduction of several mutations to optimize brightness, kinetics, activity-dependent photoconversion, and Ca2+ affinity, resulting in CaMPARI2 (Moeyaert et al., 2018). CaMPARI2 (F391W_L398V) is targeted to either pre- or postsynaptic compartments by fusing it with synaptophysin (preSynTagMA) or an intrabody against PSD-95 (postSynTagMA), respectively (Perez-Alvarez et al., 2019). For

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preSynTagMA, the intact presynaptic component, synaptophysin, is used directly. For postSynTagMA, additional recombinant elements (such as an intrabody against PSD95 (PSD-95.FingR) (Gross et al., 2013), zinc finger domain, and the KRAB(A) transcription repressor) are needed to avoid unwanted synaptic changes caused by the overexpression of endogenous PSD-95, and to reduce background, cytoplasmic fluorescence. This synapse-localized CaMPARI, SynTagMA, together with a computational analysis has been successfully demonstrated to map synaptic activity (Perez-Alvarez et al., 2019).

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Syb:GRASP and eGRASP: Because synapses are bilateral microstructures involving both the presynaptic terminal and postsynaptic density, dual component labeling can provide more information about active circuitry than tools that label only single components. The introduction of proximity-based synapse labeling techniques such as GFP Reconstitution Across Synaptic Partners, GRASP, has allowed static and accurate mapping of synapses in a variety of model systems (Feinberg et al., 2008; Gordon and Scott, 2009; Kim et al., 2011). GRASP is based on reconstitution of two complementary membrane-tethered split GFP fragments across the synaptic cleft. In the optimized synaptic version, mGRASP, split GFP fragments are specifically targeted to pre- and post-synaptic compartments using specially designed synapse-specific membrane carriers (Kim et al., 2011). mGRASP has recently been used for comprehensive, finescale, and cell type-specific mapping of connectivity in the hippocampus (Druckmann et al., 2014; Kwon et al., 2018). Furthermore, combining mGRASP and optogenetics enables functional connectome mapping (Song et al., 2018).

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Two additional GRASP-based active synapse mapping tools have recently become available (Choi et al., 2018; Macpherson et al., 2015). First, using the same principle as SynaptoZIP, the presynaptic vesicle protein synaptobrevin is fused with one of the split GFP fragments, now called syb:GRASP. This complex is initially targeted to the vesicle lumen, but following neural activity, is exposed to the extracellular synaptic cleft. There, it joins with the other split GFP fragment which is tethered to the postsynaptic membrane, thus reconstituting fluorescence (Macpherson et al., 2015). This system has so far been used effectively in Drosophila but would require modifications including combinatorial approaches before it could be applied in vertebrates. The developers of syb:GRASP have broadened its utility by introducing differently-colored varieties known as X-RASPs (i.e. yellow Y-RASP and cerulean C-RASP), thereby enabling simultaneous multi-color labeling of active synapses in different locations (Macpherson et al., 2015). Another GRASP-based tool was created by combining GRASP with the IEG promoterbased expression system. This enhanced version of GRASP (eGRASP) has several improvements: it was made by placing a SH3 binding motif near the split GFP fragments

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to facilitate reconstitution, and with mutations of the FP molecules to increase brightness. Also, expression of eGRASP is under the control of the Fos-tTA system, as described above. And, as with X-RASP, yellow and cyan versions of eGRASP exist, allowing multiple simultaneous labeling. eGRASP has been used to label synapses between engram cells in hippocampus such that CA3-CA1 synapses were labeled in yellow when expression of the yellow version is controlled by the Fos-tTA system while structurally connected synapses was labeled in cyan when that of the cyan version under an activity-independent control (Choi et al., 2018). Due to color detection limits, projecting axons are not visible in the current eGRASP system. Nevertheless, these GRASP-based techniques provide powerful active synapse detection tools particularly when further optimized and combined in sophisticated strategies.

4. Activated synapse manipulation

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Thus far, we have described tools for visualizing activated neurons and synapses. Next, we focus on new tools for manipulating specifically activated synapses. Traditionally, neurotoxins and pharmacological blockers or enhancers, and, more recently, opto- or chemo-genetic gated channels, have been used to manipulate neurons and have contributed greatly to our knowledge of brain physiology (Boyden et al., 2005; Brooks et al., 1957; Floresco and Jentsch, 2011; Urban and Roth, 2015; Zhang et al., 2007). And recently, extending the resolution of these methods, improved and newly developed molecular tools have succeeded in using light-controlled synaptic inhibition as a fine molecular tweezer, enabling detailed dissections of synaptic activity (Figure 3).

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PA-BoNT: As synaptic vesicle release is known as one of neuronal activity-triggered synaptic events, neurotoxins that cleave vesicle-associated proteins and block vesicle release have been used to examine functions of active synapses. Using light-triggered dimerization, similar to PA-Cre described above, photoactivatable botulinum neurotoxin (PA-BoNT) was engineered through a clever molecular recombination (Liu et al., 2019) to inhibit excitatory neurotransmission. It is built upon the catalytic light chain of botulinum neurotoxin (BoNT), which is known to disrupt neurotransmitter release by cleaving soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) family proteins, such as VAMP2 (Blasi et al., 1993; Schiavo et al., 19 92). To employ PA-BoNT with useful temporal resolution and selectivity, optimized split BoNT fragments were generated and fused to a carefully chosen photodimerizer pair, improved light-induced dimerizer (iLID) and bacterial SspB (Guntas et al., 2015; Zimm erman et al., 2016). Soluble and synaptophysin-fused vesicular PA-BoNT variants (sPABoNT and vPA-BoNT, respectively) have been tested, and vPA-BoNT showed improved efficacy. Upon light illumination, vPA-BoNT effectively and locally inhibits excitatory

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neurotransmission in both rodent and nematode systems. Unlike opsin-based tools that are suitable for short-term inhibition of neural activity, PA-BoNT exhibits longterm inhibitory effect. Since PA-BoNT functions to cleave VAMP2 at synaptic terminal in a defined manner and does not require constant light stimulation, PA-BoNT has little off-target effect and reduced photodamage of tissue. Also, efficiency of PA-BoNT can be easily accessed using antibodies against VAMP2. Furthermore, PA-BoNT is applicable for inhibition of diverse secreted biomolecules with high versatility.

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AS-PaRac1: Another new synapse manipulator is Activated Synapse Targeting Photoactivatable Rac1 (AS-PaRac1) (Hayashi-Takagi et al., 2015). It was constructed upon small GTPase Rac1, which is known to induce spine shrinkage when persistently activated, and was made photoactivatable (PA-Rac1) by fusing it to the photo-sensitive LOV domain which, upon illumination, sterically blocks and uncages Rac1 (Wu et al., 2009). PA-Rac1 was enhanced by introducing two mutations (L514K and L531E) to reduce background Rac1 activity in the dark. Localization specificity was provided by fusing this improved PaRac1 to truncated post-synaptic protein PSD-95 (PSD∆1.2) to target the postsynaptic density, or to dendritic targeting element (DTE) of Arc mRNA to target neuronal input. DTE is known to be selectively targeted and translated in activated dendritic segments upon synaptic activation. The combination of PSD∆1.2 and DTE on each side of PaRac1 has been shown to label activated synapses (AS). Because sustained activation of Rac1 induces spine shrinkage, light stimulation can selectively erase AS-PaRac1 expressing spines. AS-PaRac1, deployed with a dual task learning protocol, has been used to distinguish and visualize task-specific neural ensembles (Hayashi-Takagi et al., 2015). Recently, AS-PaRac1 has been applied to unraveling mechanism of action of antidepressant ketamine (Moda-Sava et al., 2019). AS-PaRac1 further offers a powerful tool for not only labeling recently potentiated dendritic spines but also optically erasing specific synapses.

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5. Conclusion and future perspectives

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We have described recently developed genetic tools for taking snapshots of activated neurons and synapses, particularly with improved temporal resolution, signal-to-noise ratio, sensitivity, and the possibility of further manipulations (Table 1). These new advanced tools certainly provide more precise measurements of neural and synaptic activity, but they also leave some room for improvement; as is often the case with new techniques, they offer relatively large baseline signals and subset targeted biases. Furthermore, to extend these tools to brain-wide utility, specifically tailored computational analyses will be necessary (Rah et al., 2015). Continuous upgraded versions of these new methods joined with integrative and combinatorial approaches

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will provide a deeper understanding of brain activity underlying mechanisms of cognitive function and behavior.

Declarations of interest: None

Acknowledgements & Funding

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This work is supported by the Korea Institute of Science and Technology (KIST) Institutional Program (2E29180) and the Samsung Science and Technology Foundation (SSTF-BA1502-11).

References

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Figure legends

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Figure 1. Diagram and schematic illustrations of IEG promotor-based gene expression system. Abbreviations: TRE, Tet response element; Dox, doxycycline; tTA*, tTA with Dox-insensitive mutation for long-lasting expression; 4-OHT, 4-hydroxytamoxifen; TetO, tetracycline operator sequence; Tetbi, bidirectional Tet promoter; POxt, oxytocin promoter as a cell type specific promoter; dsTVA, destabilized avian tumor virus receptor A (TVA); PhSyn, human synapsin promoter; EnvA, envelope protein of the avian sarcoma and leukosis virus; SARE, synaptic activity-responsive element; and EM, enhancer module. For more information, refer to the main text.

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Figure 2. Diagram and schematic illustrations of photoactivatable gene expression system. Abbreviations: TEVp, tobacco etch virus proteinase; TEVp-N, split N-fragment of TEVp; TEVp-C, split C-fragment of TEVp; TEVseq, TEVp cleavage sequence; LOV, lightoxygen-voltage sensing domain; CaM, calmodulin; CaMbp, Ca2+/CaM-binding motif; CRY2*, Cryptochrome 2 with single amino acid mutation for long photocycling; CIBN, truncated version of CIB1; Cre-N, split N-fragment of Cre; Cre-C, split C-fragment of Cre; Flp-N, split N-fragment of flippase; Flp-C, split C-fragment of flippase; nMagH, negative Magnet with mutations for increased dimerization; and pMagH, positive Magnet variant with mutations for increased dimerization. For more information, refer to the main text.

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Figure 3. Diagram and schematic illustrations of active synapse-specific labeling and manipulation system. Abbreviations: SB, synbond; PhSyn, human synapsin promoter; CaM; calmodulin; CaMPARI2*, nuclear export sequence (NES) deletion form of CaMPARI2; ZF, zinc finger DNA binding domain; TRE, Tet response element; BoNT-N, split N-fragment of botulinum neurotoxin serotype B (BoNT); BoNT-C, split C-fragment of BoNT; iLIDV416I, improved light-induced dimerizer with mutation; PaRac1, photoactivatable Rac1; and DTE, dendritic targeting element of Arc mRNA. For more information, refer to the main text.

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Table 1. Summary of tools for visualizing active neural ensembles and active synapses Tools

Activity-capturing components Other regulatory components

Available resources

Refs*

[ IEG promoter-based gene expression ] TetTag

Fos promoter

tTA-TRE

JAX 018306, 008344; MMRRC 031756-MU

1

vGATE

Fos promoter

rtTA-TRE, Cre-LoxP

N/A

2

TRAP

Fos or Arc promoter

CreERt2-LoxP

JAX 021881, 021882, 022357, 030323

CANE

Fos promoter

dsTVA, EnvA-coated virus, Cre- JAX 027831; Addgene 86641, LoxP 86666, 87221

E-SARE

SARE enhancers, Arc promoter

RAM

AP-1 site, Npas4 motif, minitTA-TRE Fos promoter

mini-

[ Photoactivatable gene expression ] Cal-Light

7

Addgene 63931

8

CaM, Ca2+/CaM-domain, light

TEVp-TEVseq, LOV, tTA-TRE

Addgene 92391, 92392

9

Ca2+/CaM-domain,

TEVp-TEVseq, LOV, tTA-TRE

Addgene 92214, 92213

10

Addgene 26888-26889, 7536711-13 75368

CaM,

light

PA-Cre

Split Cre, light

Photo-dimers CRY2-CIBN

PA-Flp

Split Flp, light

Photo-dimers Magnet

[ Activated synapse visualization ] ZIP-Synbond

Ca2+/CaM-domain

CaM, CaMPARI2

syb:GRASP

Synaptobrevin

eGRASP

Fos promoter

14

N/A

15

in Synaptophysin (pre), PSD-95 Addgene intrabody (post) 119738

119723,

119736,

16

Addgene 65830, 73698, 73699, 73700

17

tTA-TRE, GRASP

Addgene 111579-111590, 111597-111598, 120309

18

Rac1, LOV

N/A

19

Photodimer iLID-SspB

Addgene 122981

GRASP (split GFP)

lP

SynTagMA

re

Synaptobrevin

N/A

-p

FLARE

SynaptoZIP

6

N/A

ro of

Cre- or CreERt2-LoxP

3-5

[ Activated synapse manipulation ] PSD∆1.2, light

DTE

of Arc mRNA,

PA-BoNT

Split botulinum synaptophysin

na

AS-PaRac1

toxin,

122976,

122979,

ur

* 1, Reijmers et al., 2007; 2, Hasan et al., 2019; 3, Guenthner et al., 2013; 4, Allen et al., 2017; 5, DeNardo et al.,

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2019; 6, Sakurai et al., 2016; 7, Kawashima et al., 2013; 8, Sørensen et al., 2016; 9, Lee et al., 2017; 10, Wang et al., 2017; 11, Kennedy et al., 2010; 12, Schindler et al., 2015; 13, Taslimi et al., 2016; 14, Jung et al., 2019; 15, Ferro et al., 2017; 16, Perez-Alvarez et al., 2019; 17, Macpherson et al., 2015; 18, Choi et al., 2018; 19, Hayashi-Takagi et al., 2015; 20, Liu et al., 2019.

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