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Curr Opin Neurobiol. Author manuscript; available in PMC 2013 February 1.
Published in final edited form as:
PMCID: PMC3294064
NIHMSID: NIHMS342205

Optical highlighter molecules in neurobiology

Abstract

The development of advanced optical methods has played a key role in propelling progress in neurobiology. Genetically-encoded fluorescent molecules found in nature have enabled labeling of individual neurons to study their physiology and anatomy. Here we discuss the recent use of both native and synthetic optical highlighter proteins to address key problems in neurobiology, including questions relevant to synaptic function, neuroanatomy, and the organization of neural circuits.

Introduction

GFP and other conventional fluorescent proteins have had a tremendous impact on biology, and their subsequent development into optical highlighters (photoactivatable, photoconvertible, and photoswitchable fluorescent proteins) has opened even more avenues of experimentation. Respectively, these molecules are initially synthesized as molecules lacking or having low fluorescence, molecules initially fluorescing at wavelengths other than their "activated" wavelength, and molecules exhibiting fluorescence which can be switched "on" and "off" repeatedly. These features enable experiments not possible through conventional photobleaching of constitutively fluorescent proteins: by definition, photobleaching experiments track the properties of those molecules not subject to the photobleaching itself, whereas optical highlighting enables researchers to track precisely those molecule that have been photostimulated. Recounting the >20 variations of these molecules is outside the purview of this review, but interested readers are directed to reviews detailing their characteristics. [1,2]

Optical highlighters at the synapse

Chemical and electrical stimulation protocols induce defined forms of plasticity in vitro whose in vivo correlates are thought to instantiate network-level processes such as learning and memory. These experimental manipulations trigger both local alterations in protein translation, localization and biochemical activity and neuron-wide changes in transcription; together these events respecify the molecular landscape within individual synapses, culminating in altered synapse anatomy and function [35].

The use of live imaging techniques has promised to enable visualization of the anatomic and molecular changes coincident with the induction of synaptic plasticity in real-time. However many of the techniques traditionally deployed by researchers — such as the use of GFP-fusion proteins to track single molecules — do not afford sufficient temporal or spatial resolution to effectively address questions about molecular dynamics at single synapses on the timeframes relevant to plasticity. By conferring tight control over molecular labeling both in space and in time, optical highlighters have circumvented many of the technical limitations of conventional approaches, enabling key experiments to characterize synapses and the individual molecules within them.

For example, the observation that isolated synapses on a single neuron exhibit distinct functional properties suggests that the anatomic structure of dendritic spines — whose bulbuous heads are connected to the dendritic shaft by narrow necks only a few hundred nanometers across — may act as electrical or chemical compartments, enabling the spine to act as an isolated computational unit [6]. By using multiphoton techniques to photoactivate PA-GFP within individual spines in hippocampal slice culture and then monitoring the rate of PA-GFP exit into the dendrite, Bloodgood et al [7] could establish that spines are diffusionally coupled to the dendritic shaft. (Fig. 1) Further, by manipulating levels of electrical activity in the slice and then photoactivating PA-GFP within spines it was shown that degree of effective continuity between spines and dendrites could be dynamically altered [7].

Figure 1
Using PA-GFP to measure diffusional coupling between spines and dendrites

The observation of regulated diffusional coupling between the spine head and the dendrite suggests that molecular mediators of plasticity may be differentially trapped within or released from spines in response to trans-synaptic cues[8]. As PA-GFP diffusion may not reflect the biophysics of native spine-localized molecules, researchers have measured the diffusion constants of various synaptic molecules fused with optical highlighters. Although the molecular weight of fusion proteins significantly differs from that of the native proteins, diffusion coefficients are only weakly related to total molecular weight (approx. MW1/3)[9], enabling fusion proteins to provide an adequate estimate. Several groups have fused PA-GFP to PSD-95, a core protein within the postsynaptic density (PSD) whose local concentration may scale with synapse size and strength. [913] Photoactivation of PSD-95-PA-GFP in spines in vivo has revealed that PSD-95 leaves unstimulated synpases on timescales of 10s of minutes to hours (instead of seconds as measured for free PA-GFP) and that this rate increases in response to activity. Highlighter proteins have also been fused with other adapter proteins, enzymes and ion channels resident in synaptic spines, including Shank2, Shank3, CaMKIIα, CaMKIIβ, GluR2, stargazin, Ras, Rho, and Cdc42 [10,1417] revealing a diversity of diffusion rates that constrain molecular models of plasticity. These experiments have also shown that the spine concentration of certain molecules (PSD-95, CamKII) scales with spine size. Importantly, while protein diffusion may facilitate the spread of signals from activated spines to adjacent spines (as is the case for PSD-95), in at least some cases diffusion may also promote enzyme inactivation. For example, PA-GFP-tagged cdc42 diffuses rapidly out of activated spines, but (as revealed by 2P-FLIM) its enzymatic activity is disabled upon entry into the dendrite [15].

Optical highlighters have also been used to probe actin dynamics within spines, where actin plays a critical role in synaptic function by acting both as a tether (through interactions with signaling molecules) and as a strut[18]. Expression and photoactivation within spines of PA-GFP-β-actin fusion proteins in hippocampal neurons revealed two populations of filamentous F-actin: a treadmilling pool of actin that flows from the spine periphery towards the center, and a stable pool of actin at the spine base whose size is proportional to the size of the spine itself [12,13]. Stimulation via single spine uncaging of MNI-glutamate causes formation of a third unlocalized pool of actin that may play a role in structural plasticity [12]. Photoconvertible protein actin chimeras have also been used in a super-resolution microscopy technique, Photoactivatable Localization Microscopy (PALM) [19]. PALM and other molecular localization microscopy techniques, such as Fluorescence-PALM [20], Stochastic Optical Reconstruction Microscopy (STORM) [21], PALM with independent running acquisition (PALMIRA) [22] and many others [23], rely on the precise localization of single molecules by imaging and fitting their fluorescence signals to two-dimensional Gaussian distributions. These experiments revealed that most actin fibers are short (<300 nm) and that the most dynamic fibers are irregularly distributed within the spine in numbers proportional to spine size [13,24]. As an alternative to direct actin tagging, Izeddin et al [25] developed an actin binding protein (ABP)-tdEosFP fusion for PALM with which they have visualized actin redistribution after exposure to AMPA agonists in hippocampal neurons [25].

Like their postsynaptic counterparts, presynaptic boutons have been proposed to serve as compartments that differentially isolate second messengers and specific pools of synaptic vesicles. By expressing and locally photoactivating PA-GFP fused to presynaptic active zone components, an active exchange of presynaptic material between adjacent boutons was observed; the exchange of certain molecules (such as synapsin) was promoted by activity, whereas the exchange of other molecules (such as bassoon) was not [17,26]. Similar synaptophysin-Dendra2 experiments demonstrated that presynaptic vesicles decorated with synaptophysin are also shared in “packets” between adjacent presynaptic boutons [27]. These results indicate that local exchange of multiple components occurs within presynaptic structures, and that optical highlighters afford sufficient temporal and spatial resolution to track this process.

Tracking translation and transcription with highlighters

In addition to enabling molecule tracking at the synapse, optical highlighters can enable real-time tracking of de novo protein synthesis. Certain forms of synaptic plasticity likely require the local translation of specific mRNAs into protein via synthetic machinery located within the dendrite [28,29]. Although pharmacological and molecular studies (often utilizing pulse-chase approaches) have strongly suggested that protein translation both occurs locally and may be causally involved in synaptic plasticity, visualization of this process has been difficult. The generation of translational reporters consisting of the cDNAs for two-color photoconvertible proteins fused to the UTRs of translationally-regulated messages has facilitated the identification of both the signaling pathways that impinge upon the local translational machinery and the cis-acting elements that confer translational regulation upon specific messages. Typically, in these experiments a photoconvertible fluor such as Kaede is converted to its active red color, and then the stimulus-induced rate of translation of the reporter is revealed as de novo green fluorescence. This strategy has been used to examine the dendrite-specific translation of Kv1.1, CaMKII and Lypla1, and the axon-specific translation of β-actin [3037]. Recently Martin and colleagues used a Sensorin-Dendra2 reporter to test synapse-specific mRNA translation in a culture system in which single, defined synapses between Aplysia sensory and motor neurons can be both manipulated and imaged [38]. These experiments demonstrated that stimuli known to initiate facilitation (such as five pulses of serotonin) trigger synapse-specific translation, whereas stimuli that are ineffective (such as a single pulse of serotonin, or a pulse of FMRFamide) fail to generate new protein. It has also recently been observed that certain forms of synaptic plasticity (such as homeostatic plasticity [39]) may require targeted protein degradation; because photoconversion allows photoconvertible proteins to be used as optical pulse-chase reagents, these molecules can track stimulus-induced protein destruction. For example, degradation reporters have been generated in which PA-GFP is fused to a variety of protein motifs that can target peptides to the proteasome, and used to show that the chronic manipulation of activity can alter dendritic protein turnover [34,35].

Optical highlighters have also been used to reveal mechanisms of new gene expression, which is thought to be essential to the induction of synaptic plasticity over long time scales[36,37]. This process canonically involves recruitment of cellular signal transduction molecules that culminate in the regulation of transcription factors in the nucleus. However it has recently been suggested that some transcription factors are held in abeyance in the dendrite, but that upon stimulation these factors are released and imported into the nucleus where they alter gene expression to promote synaptic modification [40]. For example in the Aplysia system the transcription factor CREB2 has been suggested to translocate from dendrites to the nucleus, where in response to specific types of synaptic stimulation it initiates new gene expression required for long term depression. By tagging CREB2 with Dendra2, Lai et al [41] demonstrated that CREB2 translocates to the nucleus in response to FMRFamide in an importin-dependent manner [41]. Similarly, tagging of the p65 subunit of NF-kB with PA-GFP has been used to demonstrate that dendritically-localized NF-kB is actively transported from the synapse to the nucleus [42].

Axonal transport and long range protein movement

The long distances between the soma and the tips of an axon present challenges to the cell in moving proteins to and from these distal points. In addition to translating mRNAs at peripheral sites (as discussed earlier), neurons deliver proteins to axons via least three other mechanisms, each of which has been clarified through the use of optical highlighter-cargo fusion proteins. Neurofilament fusion proteins, for instance, have been shown to move along microtubules [43] and rely on myosin Va to keep them on microtubule tracks. [44] Fusions of at least two soluble cytosolic proteins, synapsin and CamKII, have been found to be transported via a microtubule-based mechanism in which large multimeric complexes interact with motor proteins. [45] Diffusion can also be a primary mode of transport. For instance, highlightable fusions of the microtubule binding protein tau undergo rapid binding to and unbinding from microtubules during transit to the end of an axon, with transport kinetics indicative of free diffusion to the tip of the neurite. [46,47] Fusions of even slowly diffusing transmembrane proteins, such as the beta-adrenoceptor [48] and voltage-gated potassium channel isoforms 1.3, 1.4, and 2.1 [49], have been shown to move by this mode. These studies of diffusive mechanisms have particularly benefited from using highlighting methods since measurements can be made at many distal points in the cell in addition to the irradiation spot. Consequently, mobility fraction and diffusion coefficient maps over large areas of the membrane and various axon segments can be easily established.

Finally, optical highlighters are also well suited for studying trafficking in the secretory pathway. Since new proteins are often continuously produced over the course of an experiment, photoactivated or photoconverted pools effectively provide a time stamp for a given population of molecules. Transport studies of UNC-2, the C. elegans voltage-gated calcium channel, CaV2, found that a newly discovered endoplasmic reticulum (ER) localized chaperone, CALF-1, is required in conjunction with a calcium channel subunit, alpha2 delta, for proper exit of UNC-2 from the ER. [50] Photoconverted (and thus time stamped) Dendra2-tagged UNC-2 exhibited transport from the ER to synapses, but only in the presence of calf-1, indicating that preexisting protein within the ER exits through interactions with this newly-described chaperone.

Cell fate Mapping

Mapping the fates of individual cells or small numbers of cells during development requires the ability to introduce a label into a sparse number of cells at a specific point in time. Traditionally this has been either achieved through laborious single cell methods like DNA electroporation, through the use of genetic chimeras, or through clever genetic approaches such as MARCM and MADM [51]. However these physical or genetic methods are often impractical or impossible (i.e. because of physical access issues or the absence of effective mitotic recombination in the chosen model organism). The ability to generate focal signals with high contrast ratios through precision photoactivation — particularly via multiphoton methods — potentially circumvents these limitations, and has led to the widespread use of optical highlighters to track cell fates during development. This approach was first used to track the fate of neural crest precursors in chick embryos; chicks were electroporated in ovo with plasmids expressing PA-GFP under the control of a ubiquitous promoter, and then single neural crest cells (NCCs, located on the dorsal surface of the neural plate) were photoactivated and imaged [52].

Since this initial demonstration, a number of different highlighters have been successfully deployed in the chick [5257], zebrafish [5862], xenopus [63], ciona intestinalis [64] and in mammals [52,54,57,61,63,65,66] to track the fates of neural precursors during development. These experiments have correlated the initial positions of neural precursors with the final positions of differentiated neurons [52,54,57,61], visualized interactions between migrating neurons [65], assessed proliferation during migration [57,65], followed the fate of cells during metamorphosis [64] and tested the role of individual signaling molecules and transcription factors in cell migration processes [54,63,66]. By restricting the pattern of optical highlighter expression through the use of cell-type specific promoters researchers can precisely characterize the tracked neuron's function. For example, interneuron subtype-specific promoters driving Kaede expression in zebrafish have been used to address the relative role of CiD and MCoD interneuron types in fast and slow forms of swimming, respectively. [61]. Furthermore, a combinatorial technique called BAPTISM, which combines promoter-specific expression of a photoconvertible fluorescent protein (huc::kaede, for example) with EGFP expression under the control of a second cell-type specific promoter (p2x3b::egfp and trpa1b::egfp, for examples), has enabled researchers to birthdate multiple different cell types was enabled within a specific pool of neurons (Fig. 2) [60].

Figure 2
BAPTISM allows birthdate analysis of genetically-marked subpopulations of neurons

Circuit Mapping

Freely-diffusing optical highlighters not targeted to specific subcellular compartments have been used to great advantage in measuring diffusional coupling between spines and dendrites (see above)[7]. To a first approximation, highlighted molecules within other subcompartments of the neuron also have free access to the rest of the neuron. Because the anatomy of the neuron is regionally specialized based upon connectivity — with input-receiving dendrites spatially segregated from output-transmitting axons —researchers have used photoactivation methods to focally activate a protein within the dendrite or axon and then utilize its de novo fluorescence to visualize (via diffusion) the contiguous cell bodies, axonal arbors, and dendrites. While labeling is restricted to contiguous structures within a given neuron, the complete morphology of targeted neurons and potentially the direction and nature of information flow (at least as constrained by anatomy) within a neural circuit can be revealed with this approach.

This strategy was first deployed in a paper in which Kaede was expressed under the control of the neural HuC promoter in zebrafish followed by focal photoconversion to reveal the complete anatomy of targeted trigeminal neurons and Rohan-Beard cells in the ventral spinal cord [67]. Aramaki et al [59] performed a similar experiment in zebrafish, but with a twist: by using the “rewritable” photoswitchable fluor Dronpa (expressed in neurons using the Gal4-UAS system), these researchers were able to interrogate a targeted neuron’s anatomy, erase the fluorescence, and then target and image a second neuron (and so on with multiple neurons in series) [59]. In this iterative manner, structural features of a number different neurons presumed to be within a single neural network were identified.

While these two proof-of-principle papers demonstrated that optical highlighters could be used as long-range neural tracers, subsequent papers in the fruit fly Drosophila melanogaster illustrate the power of this approach to dissect the structure and function of behaviorally-relevant neural circuits. One such circuit is responsible for sexually-dimorphic behaviors in response to the pheromone cis-vaccenyl acetate (cVA), which (in particular contexts) elicits aversive behaviors in males but is attractive to females [68]. To address whether differences in downstream circuits are responsible for sexually dimorphic responses to cVA, Datta et al [69] expressed PA-GFP in most projection neurons in the antennal lobe, and then took advantage of the spatial specificity afforded by multiphoton techniques to photoactivate the PA-GFP specifically within the DA1 glomerulus; the PA-GFP then diffused, labeling the connected cell bodies and revealing sexual dimorphism in the DA1 projection neuron axonal arbors (Fig. 3) [69]. Furthermore, the induced fluorescence enabled electrophysiological characterization of DA1 projection neurons via cell-attached recordings.

Figure 3
Neural Tracing Using PA-GFP

Because this approach depends upon the inherent diffusional characteristics of PA-GFP, the total neurite length that can be traced is limited, and effective tracing requires photoactivation of a significant amount of PA-GFP (through repeated cycles of activation over relatively large target regions). These constraints have been partially circumvented through the development of two variants of PA-GFP (C3PA and SPA) that exhibit improved apparent rates of diffusion in vivo [70]. By expressing C3PA and SPA in neurons potentially connected in a circuit, Ruta et al [70] traced a single neuron and identified its axonal arbor, and then irradiated the newly-labeled axonal arbor, thereby taking advantage of the spatial overlap with the connected dendritic spine to highlight the downstream neuron. By iteratively repeating this process, a behaviorally-relevant neural circuit was traced across multiple synapses and through multiple neurons [70].

As work in neural circuits becomes increasingly focused on interrogating the role of individual neurons in driving behaviors, researchers are finding new roles for optical highlighter proteins in disambiguating functionally-relevant neurons from bystander cells. For example, a common difficulty is that the genetic drivers that are used to functionally manipulate the activity of neurons promote gene expression within large numbers of cells, making it difficult to identify the particular neurons responsible for a specific behavior. Claridge-Chang and colleagues used optical highlighters to identify which subset of cells labeled by their driver was relevant to behavior on the basis of connectivity revealed by targeted photoactivation of specific neurons [71]. Similarly, a number of researchers have begun co-expressing various photoconvertible proteins along with optogenetic reagents; optical stimuli then both trigger changes in electrical activity via the light-gated channel, and photoconvert and label the stimulated cells, enabling verification of their identity [7274]. Photoconvertible fluors are thus proving to be useful adjuncts in optogenetic experiments aimed at testing neural function.

Cell and organelle continuities

Similar to mapping circuits, the advantages of an optical highlighter become apparent when determining organelle continuities as well as both intracellular and intercellular continuities. Axonal regeneration [75] has been confirmed by photoconverting freely diffusing protein either in the cell body or distal axon and monitoring the presence of red and green signal mixtures. Similarly, continuity between cells has been established by observing photoconverted KikGR move bidirectionally into neighboring cells within the neural crest of chick embryos via cytoplasmic bridges. [76] Alternatively, axon degeneration has been confirmed by highlighting fragmented axon segments to ensure that subsequent re-fusion of fragments did not take place. [77]

Summary and outlook

Some of the experiments discussed here represent approaches that are difficult or impossible with conventional fluorescent protein imaging, demonstrating that optical highlighters have moved from being neat little fluorescent protein tricks to bona fide neurobiology tools. Yet, the challenges of imaging in the nervous system will likely require further optimization of these molecules, with emphasis on brightness, contrast, photostability, monomeric behavior, red-shifted wavelengths, improved folding efficiency, faster folding kinetics, and higher photoactivation quantum efficiency. Improvements in any and all of these parameters will assist in neurobiology experiments that seem to be delving deeper into tissue, imaging with faster kinetics, and observing over longer timescales. If history provides any reasonable forecast, the rapid development of highlighter molecules over the past 10 years is likely to continue and provide tools to facilitate new and innovative approaches.

Highlights

[arrowhead]
Optical highlighter proteins circumvent limits of conventional fluors
[arrowhead]
Photoconversion enables spatially-localized pulse-chase experiments in neurons
[arrowhead]
Neurobiological applications (from synapses to circuits) are discussed

Acknowledgements

This work was funded in part by the Intramural Research Program of the National Institutes of Health including the National Institute of Biomedical Imaging and Bioengineering (GHP), and by grants from the National Institutes of Deafness and Communication Disorders (RO1DC011558), the NIH Office of the Director’s New Innovator Program (DP2OD007109), the McKnight Foundation, the Searle Foundation and from a Career Award in Biomedical Sciences from the Burroughs Wellcome Foundation (SRD).

Footnotes

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References

* paper of special interest

** paper of outstanding interest

1. Patterson GH. Highlights of the optical highlighter fluorescent proteins. J Microsc. 2011;243:1–7. [PMC free article] [PubMed]
2. Chudakov DM, Matz MV, Lukyanov S, Lukyanov KA. Fluorescent proteins and their applications in imaging living cells and tissues. Physiol Rev. 2010;90:1103–1163. [PubMed]
3. Barco A, Bailey CH, Kandel ER. Common molecular mechanisms in explicit and implicit memory. J Neurochem. 2006;97:1520–1533. [PubMed]
4. Feldman DE. Synaptic mechanisms for plasticity in neocortex. Annu Rev Neurosci. 2009;32:33–55. [PMC free article] [PubMed]
5. Redondo RL, Morris RG. Making memories last: the synaptic tagging and capture hypothesis. Nat Rev Neurosci. 2011;12:17–30. [PubMed]
6. Bourne JN, Harris KM. Balancing structure and function at hippocampal dendritic spines. Annu Rev Neurosci. 2008;31:47–67. [PMC free article] [PubMed]
7. Bloodgood BL, Sabatini BL. Neuronal activity regulates diffusion across the neck of dendritic spines. Science. 2005;310:866–869. [PubMed]
8. Sheng M, Hoogenraad CC. The postsynaptic architecture of excitatory synapses: a more quantitative view. Annu Rev Biochem. 2007;76:823–847. [PubMed]
9. Gray NW, Weimer RM, Bureau I, Svoboda K. Rapid redistribution of synaptic PSD-95 in the neocortex in vivo. PLoS Biol. 2006;4:e370. [PMC free article] [PubMed]
10. Sturgill JF, Steiner P, Czervionke BL, Sabatini BL Distinct domains within PSD-95 mediate synaptic incorporation, stabilization, and activity-dependent trafficking. J Neurosci. 2009;29:12845–12854. [PubMed]
. PSD-95, CaMKII alpha and beta, GluR2, and stargazin were tagged with PA-GFP to determine their relative diffusion kinetics. Each is found to have different kinetics of turnover in individual spines in hippocampal slice CA1 neurons, with PSD-95 being the slowest and retaining ~90% fluorescence after 30 minutes. Palmitoylation of PSD-95 was found to be responsible for the slow kinetics and a structure/function analysis showed that only the two N-terminal PDZ domains, which interact with NMDA and shaker channels, are required for stabilization of photoactivated PSD95 within the spine. On the other hand, a third PDZ domain and the C-terminal portion of the protein were implicated in establishing the PSD lattice within the spine.
11. Blanpied TA, Kerr JM, Ehlers MD Structural plasticity with preserved topology in the postsynaptic protein network. Proc Natl Acad Sci U S A. 2008;105:12587–12592. [PubMed]
. Photobleaching of PSD-95-GFP and photoactivation of PSD-95-PA-GFP was used to track PSD-95 dynamics within single spines. Optically tracking PSD-95 revealed that each spine had structural dynamics that were independent from those observed in adjacent spines, and that the post-synaptic density followed those dynamics. However, little exchange of PSD-95 was observed within the PSD (on timescales ranging from 60 second to 30 minutes), leading the authors to suggest a model in which the PSD is largely stable but capable of flexible conformational changes to accommodate alterations in spine shape.
12. Honkura N, Matsuzaki M, Noguchi J, Ellis-Davies GC, Kasai H The subspine organization of actin fibers regulates the structure and plasticity of dendritic spines. Neuron. 2008;57:719–729. [PubMed]
. Photoactivation of actin-PA-GFP was used to observe treadmilling and revealed that two pools of actin are normally present in spines. The actin toward the spine tip shows high turnover (tau=40 sec) while actin toward the base shows low turnover (tau=17 min) and represents ~15% of actin in spine. Activation with caged glutamate results in a third stable pool that is confined to the spine and is necessary for spine enlargement. Photoactivation at spine tips show a retrograde stream of actin-PA-GFP, which does not join the stable pool at the base.
13. Frost NA, Shroff H, Kong H, Betzig E, Blanpied TA Single-molecule discrimination of discrete perisynaptic and distributed sites of actin filament assembly within dendritic spines. Neuron. 2010;67:86–99. [PubMed]
. This study uses two-photon excitation microscopy to photoconvert and image single actin molecules in spines. Diffusion was found to be slow, consistent with treadmilling in the spine. By differentially targeting regions of spine, treadmilling was noted to be faster in the tip than in the core. PALM using one-photon photoconversion of actin-mEos2 revealed numerous single molecule trajectories whose measured velocities were faster than those measured by bulk flow; an inverse relationship between velocity and total distance traveled was noted. Blocking polymerization with jasplakinlide arrested most movement. Two color PALM on fixed samples with Dronpa-GKAP to mark the PSD showed that actin abuts the PSD. Single color PALM on actin plus regular diffraction-limited imaging with PSD95-cerulean showed that actin density is maximal at edges of the PSD.
14. Harvey CD, Yasuda R, Zhong H, Svoboda K The spread of Ras activity triggered by activation of a single dendritic spine. Science. 2008;321:136–140. [PubMed]
. This study used a fluorescence resonance energy transfer (FRET) based sensor to follow active Ras in spines. The sensor consists of H-Ras tagged with EGFP and the Ras binding domain (RBD) of Raf tagged with two mRFP1 molecules. Ras activation increases the affinity between the two parts of the sensor which can be observed by FRET from EGFP to mRFP1. The major finding from this work is that Ras activity spills over into adjacent synapses (~10 µm or 10–20 spines) after local activation by two-photon uncaging of glutamate. This motile activity was found to be unrelated to Ras mobility per se since a Ras-PA-GFP fusion was shown to freely diffuse similar to a membrane-tagged PA-GFP.
15. Murakoshi H, Wang H, Yasuda R Local, persistent activation of Rho GTPases during plasticity of single dendritic spines. Nature. 2011;472:100–104. [PubMed]
. This study used a fluorescence resonance energy transfer (FRET)-based sensor to follow Rho activity in spines. The sensor consists of RhoA/Cdc42 tagged with EGFP and the Rho GTPase binding domain (RBD) of Rhotekin/Pak3 tagged with two mCherry molecules. Rho activation increases the affinity between the two parts of the sensor, which can be observed by FRET from EGFP to mCherry. Specific activation of single spines was performed by local uncaging of glutamate. Rho activity was found to diffuse out of spines, while Cdc42 activity stays localized to active spines; photoactivation experiments using PA-GFP fusions revealed that this was not due to intrinsic differences in mobility.
16. Asrican B, Lisman J, Otmakhov N. Synaptic strength of individual spines correlates with bound Ca2+-calmodulin-dependent kinase II. J Neurosci. 2007;27:14007–14011. [PubMed]
17. Tsuriel S, Geva R, Zamorano P, Dresbach T, Boeckers T, Gundelfinger ED, Garner CC, Ziv NE. Local sharing as a predominant determinant of synaptic matrix molecular dynamics. PLoS Biol. 2006;4:e271. [PMC free article] [PubMed]
18. Hotulainen P, Hoogenraad CC. Actin in dendritic spines: connecting dynamics to function. J Cell Biol. 2010;189:619–629. [PMC free article] [PubMed]
19. Betzig E, Patterson GH, Sougrat R, Lindwasser OW, Olenych S, Bonifacino JS, Davidson MW, Lippincott-Schwartz J, Hess HF. Imaging intracellular fluorescent proteins at nanometer resolution. Science. 2006;313:1642–1645. [PubMed]
20. Hess ST, Girirajan TP, Mason MD. Ultra-high resolution imaging by fluorescence photoactivation localization microscopy. Biophys J. 2006;91:4258–4272. [PubMed]
21. Rust MJ, Bates M, Zhuang X. Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM) Nat Methods. 2006;3:793–795. [PMC free article] [PubMed]
22. Geisler C, Schonle A, Von Middendorff C, Bock H, Eggeling C, Egner A, Hell SW. Resolution of λ/10 in fluorescence microscopy using fast single molecule photo-switching. Applied Physics A. 2007;88:223–226.
23. Patterson GH. Fluorescence microscopy below the diffraction limit. Semin Cell Dev Biol. 2009;20:886–893. [PMC free article] [PubMed]
24. Tatavarty V, Kim EJ, Rodionov V, Yu J Investigating sub-spine actin dynamics in rat hippocampal neurons with super-resolution optical imaging. PLoS One. 2009;4:e7724. [PubMed]
. Single particle tracking PALM was used to monitor actin molecules within spines. These experiments found that filamentous-actin in hippocampal neuron dendritic spines either sits still, randomly walks, or goes vectorially anterograde or retrograde. On average, however, the flow was found to be retrograde at ~138 nm/minute. The resulting model is a system of largely disorganized filaments with barbed ends toward the spine tip. The filaments are short, not well aligned, and have numerous barbed ends distributed throughout the spine.
25. Izeddin I, Specht CG, Lelek M, Darzacq X, Triller A, Zimmer C, Dahan M Super-resolution dynamic imaging of dendritic spines using a low-affinity photoconvertible actin probe. PLoS One. 2011;6:e15611. [PubMed]
. Single particle tracking PALM was used to monitor actin dynamics within spines. Rather than rely on labeled actin, this study uses an actin-binding peptide (which binds with low affinity to actin) fused to tdEosFP. Point activation showed that the signal diffuses quickly, due to the low probe affinity promoting efficient exchange. Using this probe actin was shown not to fill the whole spine, but to form a filamentous network. In addition, using dual color STORM (Alexa647 coupled to Shank2) only a partial overlap between Shank2 (presumably labeling PSD) and actin was observed. To study spine morhpology, super-resolution imaging of sparse antibody-coupled Quantum dots was used to outline the neuron membrane. These data showed that the boundaries of actin matched the boundary of the spine itself.
26. Tsuriel S, Fisher A, Wittenmayer N, Dresbach T, Garner CC, Ziv NE. Exchange and redistribution dynamics of the cytoskeleton of the active zone molecule bassoon. J Neurosci. 2009;29:351–358. [PMC free article] [PubMed]
27. Staras K, Branco T, Burden JJ, Pozo K, Darcy K, Marra V, Ratnayaka A, Goda Y. A vesicle superpool spans multiple presynaptic terminals in hippocampal neurons. Neuron. 2010;66:37–44. [PMC free article] [PubMed]
28. Sutton MA, Schuman EM. Dendritic protein synthesis, synaptic plasticity, and memory. Cell. 2006;127:49–58. [PubMed]
29. Wang DO, Martin KC, Zukin RS. Spatially restricting gene expression by local translation at synapses. Trends Neurosci. 2010;33:173–182. [PMC free article] [PubMed]
30. Raab-Graham KF, Haddick PC, Jan YN, Jan LY. Activity- and mTOR-dependent suppression of Kv1.1 channel mRNA translation in dendrites. Science. 2006;314:144–148. [PubMed]
31. Banerjee S, Neveu P, Kosik KS. A coordinated local translational control point at the synapse involving relief from silencing and MOV10 degradation. Neuron. 2009;64:871–884. [PubMed]
32. Leung KM, Holt CE. Live visualization of protein synthesis in axonal growth cones by microinjection of photoconvertible Kaede into Xenopus embryos. Nat Protoc. 2008;3:1318–1327. [PubMed]
33. Vogelaar CF, Gervasi NM, Gumy LF, Story DJ, Raha-Chowdhury R, Leung KM, Holt CE, Fawcett JW. Axonal mRNAs: characterisation and role in the growth and regeneration of dorsal root ganglion axons and growth cones. Mol Cell Neurosci. 2009;42:102–115. [PubMed]
34. Djakovic SN, Schwarz LA, Barylko B, DeMartino GN, Patrick GN. Regulation of the proteasome by neuronal activity and calcium/calmodulin-dependent protein kinase II. J Biol Chem. 2009;284:26655–26665. [PubMed]
35. Jakawich SK, Neely RM, Djakovic SN, Patrick GN, Sutton MA. An essential postsynaptic role for the ubiquitin proteasome system in slow homeostatic synaptic plasticity in cultured hippocampal neurons. Neuroscience. 2010;171:1016–1031. [PMC free article] [PubMed]
36. Cohen S, Greenberg ME. Communication between the synapse and the nucleus in neuronal development, plasticity, and disease. Annu Rev Cell Dev Biol. 2008;24:183–209. [PMC free article] [PubMed]
37. Greer PL, Greenberg ME. From synapse to nucleus: calcium-dependent gene transcription in the control of synapse development and function. Neuron. 2008;59:846–860. [PubMed]
38. Wang DO, Kim SM, Zhao Y, Hwang H, Miura SK, Sossin WS, Martin KC Synapse- and stimulus-specific local translation during long-term neuronal plasticity. Science. 2009;324:1536–1540. [PubMed]
. Photoconvertible Dendra2 fused with the 5’ and 3’ UTRs of sensorin was used as a translation reporter to study induction of dendritic-specific translation due to exogenous stimuli, such as KCl and BDNF in cultured neurons. The reporter mRNA was synaptically localized by FISH, and after addition of 5x5HT green Dendra signal was found at synapses. These responses were not observed after treatment with 1x5HT or FMRFamide. In addition, swapping the 5’ SV40 UTR for that of sensorin eliminated 5x5HT-mediated translation although the reporter mRNA still localized to the synapse.
39. Turrigiano G. Too many cooks? Intrinsic and synaptic homeostatic mechanisms in cortical circuit refinement. Annu Rev Neurosci. 2011;34:89–103. [PubMed]
40. Ch'ng TH, Martin KC. Synapse-to-nucleus signaling. Curr Opin Neurobiol. 2011;21:345–352. [PMC free article] [PubMed]
41. Lai KO, Zhao Y, Ch'ng TH, Martin KC. Importin-mediated retrograde transport of CREB2 from distal processes to the nucleus in neurons. Proc Natl Acad Sci U S A. 2008;105:17175–17180. [PubMed]
42. Marcora E, Kennedy MB. The Huntington's disease mutation impairs Huntingtin's role in the transport of NF-kappaB from the synapse to the nucleus. Hum Mol Genet. 2010;19:4373–4384. [PMC free article] [PubMed]
43. Trivedi N, Jung P, Brown A. Neurofilaments switch between distinct mobile and stationary states during their transport along axons. J Neurosci. 2007;27:507–516. [PMC free article] [PubMed]
44. Alami NH, Jung P, Brown A. Myosin Va increases the efficiency of neurofilament transport by decreasing the duration of long-term pauses. J Neurosci. 2009;29:6625–6634. [PMC free article] [PubMed]
45. Scott DA, Das U, Tang Y, Roy S Mechanistic logic underlying the axonal transport of cytosolic proteins. Neuron. 2011;70:441–454. [PubMed]
. Axonal transport of at least two cytosolic proteins, synapsin and CamKII, was attributed to a microtubule-based mechanism in which large multimeric complexes interact with motor proteins. PA-GFP-tagged versions of synapsin and CamKII were observed to undergo slow movement in a manner biased in the anterograde direction that was not indicative of vesicular transport-like movement or diffusion. However, small punctate fluorescent speckles moving in a vectorial manner were observed, suggesting formation of dynamic multimeric complexes.
46. Weissmann C, Reyher HJ, Gauthier A, Steinhoff HJ, Junge W, Brandt R. Microtubule binding and trapping at the tip of neurites regulate tau motion in living neurons. Traffic. 2009;10:1655–1668. [PubMed]
47. Gauthier-Kemper A, Weissmann C, Golovyashkina N, Sebo-Lemke Z, Drewes G, Gerke V, Heinisch JJ, Brandt R. The frontotemporal dementia mutation R406W blocks tau's interaction with the membrane in an annexin A2-dependent manner. J Cell Biol. 2011;192:647–661. [PMC free article] [PubMed]
48. Kaya AI, Ugur O, Altuntas O, Sayar K, Onaran HO. Long and short distance movements of beta(2)-adrenoceptor in cell membrane assessed by photoconvertible fluorescent protein dendra2-beta(2)-adrenoceptor fusion. Biochim Biophys Acta. 2011;1813:1511–1524. [PubMed]
49. O'Connell KM, Tamkun MM. Targeting of voltage-gated potassium channel isoforms to distinct cell surface microdomains. J Cell Sci. 2005;118:2155–2166. [PubMed]
50. Saheki Y, Bargmann CI. Presynaptic CaV2 calcium channel traffic requires CALF-1 and the alpha(2)delta subunit UNC-36. Nat Neurosci. 2009;12:1257–1265. [PMC free article] [PubMed]
51. Luo L. Fly MARCM and mouse MADM: genetic methods of labeling and manipulating single neurons. Brain Res Rev. 2007;55:220–227. [PMC free article] [PubMed]
52. Stark DA, Kulesa PM. Photoactivatable green fluorescent protein as a single-cell marker in living embryos. Dev Dyn. 2005;233:983–992. [PubMed]
53. Stark DA, Kulesa PM. An in vivo comparison of photoactivatable fluorescent proteins in an avian embryo model. Dev Dyn. 2007;236:1583–1594. [PubMed]
54. Kasemeier-Kulesa JC, McLennan R, Romine MH, Kulesa PM, Lefcort F. CXCR4 controls ventral migration of sympathetic precursor cells. J Neurosci. 2010;30:13078–13088. [PubMed]
55. Kulesa PM, Lefcort F, Kasemeier-Kulesa JC. The migration of autonomic precursor cells in the embryo. Auton Neurosci. 2009;151:3–9. [PubMed]
56. Kulesa PM, Stark DA, Steen J, Lansford R, Kasemeier-Kulesa JC. Watching the assembly of an organ a single cell at a time using confocal multi-position photoactivation and multi-time acquisition. Organogenesis. 2009;5:238–247. [PMC free article] [PubMed]
57. Kulesa PM, Teddy JM, Stark DA, Smith SE, McLennan R. Neural crest invasion is a spatially-ordered progression into the head with higher cell proliferation at the migratory front as revealed by the photoactivatable protein, KikGR. Dev Biol. 2008;316:275–287. [PMC free article] [PubMed]
58. Hatta K, Tsujii H, Omura T. Cell tracking using a photoconvertible fluorescent protein. Nat Protoc. 2006;1:960–967. [PubMed]
59. Aramaki S, Hatta K. Visualizing neurons one-by-one in vivo: optical dissection and reconstruction of neural networks with reversible fluorescent proteins. Dev Dyn. 2006;235:2192–2199. [PubMed]
60. Caron SJ, Prober D, Choy M, Schier AF In vivo birthdating by BAPTISM reveals that trigeminal sensory neuron diversity depends on early neurogenesis. Development. 2008;135:3259–3269. [PubMed]
. To determine the differential fates of early and late born neurons, zebrafish were generated with a huc::kaede transgene and trigeminal sensory neurons were photoconverted at 24 hours post-fertilization. In conjunction with expression of EGFP under the control of different subpopulation markers, neuronal birthdate and fate was determined based on color: green, yellow, or red. This method used two sequential photoconversion pulses to distinguish early and late born neurons. A late photoconversion pulse induced either yellow cells (produced by the red signal from the new photoconverted Kaede plus the EGFP subpopulation marker) or just red signal (if the neurons did not express the green subpopulation marker). This approach was used to show that the early born trigeminal sensory neurons in zebrafish are dedicated to sensing irritating stimuli, with later born neurons devoted to touch and nocioception.
61. McLean DL, Fetcho JR Spinal interneurons differentiate sequentially from those driving the fastest swimming movements in larval zebrafish to those driving the slowest ones. J Neurosci. 2009;29:13566–13577. [PubMed]
. This paper used the alx:Kaede transgene to monitor CiD interneurons and the HuC:Kaede transgene to monitor MCoD interneurons in zebrafish. Photoconversion of the whole zebrafish on progressive days during differentiation was used to correlate the order of cell development with their final position. This work showed that the CiD interneurons responsible for high-frequency swimming motions (as demonstrated by behavioral and electrophysiological analysis) were located dorsally in the spinal cord and developed first, whereas the MCoD interneurons responsible for slower swimming movements were located ventrally and developed later.
62. Kimura Y, Okamura Y, Higashijima S. alx, a zebrafish homolog of Chx10, marks ipsilateral descending excitatory interneurons that participate in the regulation of spinal locomotor circuits. J Neurosci. 2006;26:5684–5697. [PubMed]
63. Bajpai R, Chen DA, Rada-Iglesias A, Zhang J, Xiong Y, Helms J, Chang CP, Zhao Y, Swigut T, Wysocka J. CHD7 cooperates with PBAF to control multipotent neural crest formation. Nature. 2010;463:958–962. [PMC free article] [PubMed]
64. Horie T, Shinki R, Ogura Y, Kusakabe TG, Satoh N, Sasakura Y Ependymal cells of chordate larvae are stem-like cells that form the adult nervous system. Nature. 2011;469:525–528. [PubMed]
. Here the development of ependymal cells of chordate larvae into adult nervous system was studied in tunicates using Kaede expressed from the cis regulatory element of the Ciona B-tubulin gene. The entire larval nervous system was photoconverted and followed until adulthood was reached at ~72 hours. Activation of ciona larval CNS subregions indicated that the sensory vesicle and posterior brain contribute to the adult CNS. By fusing Kaede to transcription regulatory elements specific for different cell types, such as VACHT (cholinergic neurons), VGLUT (glutamatergics), VGAT (GABA-ergic) and CRALBP (glial/ependymal cells), the authors could determine which specific neural tissues in the larva contribute to the mature animal; these experiments revealed a small number of labeled cholinergic projection neurons, a small number of glutamatergic neurons in the ciliated funnel, and many ependymal cells in the adult, but no GABA-ergic neurons were noted.
65. Mutoh T, Miyata T, Kashiwagi S, Miyawaki A, Ogawa M. Dynamic behavior of individual cells in developing organotypic brain slices revealed by the photoconvertable protein Kaede. Exp Neurol. 2006;200:430–437. [PubMed]
66. Rieger S, Senghaas N, Walch A, Koster RW. Cadherin-2 controls directional chain migration of cerebellar granule neurons. PLoS Biol. 2009;7:e1000240. [PMC free article] [PubMed]
67. Sato T, Takahoko M, Okamoto H. HuC:Kaede, a useful tool to label neural morphologies in networks in vivo. Genesis. 2006;44:136–142. [PubMed]
68. Benton R. Sensitivity and specificity in Drosophila pheromone perception. Trends Neurosci. 2007;30:512–519. [PubMed]
69. Datta SR, Vasconcelos ML, Ruta V, Luo S, Wong A, Demir E, Flores J, Balonze K, Dickson BJ, Axel R The Drosophila pheromone cVA activates a sexually dimorphic neural circuit. Nature. 2008;452:473–477. [PubMed]
. This paper is the first practical demonstration of PA-GFP to trace neurons that are part of a behaviorally-relevant neural circuit. The authors express PA-GFP in projection neurons dedicated to responding to the pheromone cVA, and through photoactivation of their dendrites identify axonal arbors in a target region in the higher fly brain. Characterizing this anatomy required combining de novo labeling of these neurons with PA-GFP with image registration techniques. In addition, the fluorescence induced by photoactivation of dendrites of these neurons revealed cell bodies that could be targeted for cell-attached electrophysiological recordings. These experiments showed that axonal projections to higher brain of this class of projection neuron are sexually dimorphic and under control of the genetic sex determination hierarchy, with male neurons elaborating a male-specific process within the lateral horn.
70. Ruta V, Datta SR, Vasconcelos ML, Freeland J, Looger LL, Axel R A dimorphic pheromone circuit in Drosophila from sensory input to descending output. Nature. 2010;468:686–690. [PubMed]
. A method to identify neurons connected in a circuit was developed that takes advantage of highly diffusible PA-GFP variants. In this approach, the anatomy of a neuron of interest is revealed via photoactivation, followed by specific photoactivation of the newly-labeled axonal arbor; because the dendrites of the downstream neurons spatially overlap with this axonal arbor, targeting of the axonal arbor photoactivates the PA-GFP within the target neuron, thereby revealing its anatomical features. The authors were able to verify that this method effectively identifies connected neurons by performing electrophyisological recordings of the putatively connected neurons. Using this method the authors were able to trace behaviorally-relevant information from neurons in the sensory periphery to a set of neurons that may be responsible for behavioral output from the fly brain.
71. Claridge-Chang A, Roorda RD, Vrontou E, Sjulson L, Li H, Hirsh J, Miesenbock G Writing memories with light-addressable reinforcement circuitry. Cell. 2009;139:405–415. [PubMed]
. Optogenetic activation of neurons labeled by a TH-GAL4 driver in the fly nervous system was found to mimic negative reinforcement signals during memory formation, although the driver used marked four disparate sets of neurons. Using the driver to express PA-GFP and to trace the innervation patterns of each of those four sets of neurons, these researchers were able to identify the specific subset of labeled neurons that targeted the Drosophila mushroom body, a center for learning and memory in the fly brain. The results suggested that the paired posterior lateral (PPL1) cluster of dopaminergic neurons in the fly brain is responsible for aversive olfactory learning.
72. Schoonheim PJ, Arrenberg AB, Del Bene F, Baier H Optogenetic localization and genetic perturbation of saccade-generating neurons in zebrafish. J Neurosci. 2010;30:7111–7120. [PubMed]
. Channelrhodopsin and Halorhodopsin were expressed ubiquitously in zebrafish to map circuits involved in the optokinetic response. Fibers were implanted at various positions to identify a locus in the hindbrain necessary and sufficient for the optokinetic response. By performing this experiment in an animal expressing the photoconvertible protein, Kaede, a posthoc map was developed of the illumination region used to trigger behavior.
73. Arrenberg AB, Del Bene F, Baier H Optical control of zebrafish behavior with halorhodopsin. Proc Natl Acad Sci U S A. 2009;106:17968–17973. [PubMed]
. Halorhodopsin and Channelrhodopsin were expressed in neurons to identify a locus necessary and sufficient for forward swimming. By co-expressing either Dendra or Kaede, the authors could identify posthoc the region that was functionally targeted by their optical fiber.
74. Wyart C, Del Bene F, Warp E, Scott EK, Trauner D, Baier H, Isacoff EY Optogenetic dissection of a behavioural module in the vertebrate spinal cord. Nature. 2009;461:407–410. [PubMed]
. This paper focuses on establishing the role of Kolmer-Agduhr cell in regulating forward motion in zebrafish; as part of functional studies using the optogenetic reagent LiGluR, UAS:Kaede was deployed to identify the cells whose optogenetic stimulation triggered different forms of swimming-related behavior.
75. Neumann B, Nguyen KC, Hall DH, Ben-Yakar A, Hilliard MA. Axonal regeneration proceeds through specific axonal fusion in transected C. elegans neurons. Dev Dyn. 2011;240:1365–1372. [PMC free article] [PubMed]
76. McKinney MC, Stark DA, Teddy J, Kulesa PM. Neural crest cell communication involves an exchange of cytoplasmic material through cellular bridges revealed by photoconversion of KikGR. Dev Dyn. 2011;240:1391–1401. [PMC free article] [PubMed]
77. Martin SM, O'Brien GS, Portera-Cailliau C, Sagasti A. Wallerian degeneration of zebrafish trigeminal axons in the skin is required for regeneration and developmental pruning. Development. 2010;137:3985–3994. [PubMed]