Name: optical control of a neuronal protein using a genetically encoded unnatural amino acid in neurons
Uploaded: 2016-03-28T14:00:00
Description: Watch this Scientific Journal Video about Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons at JoVE.com

We thank Dr. S. Szobota for helpful discussion on neuron culture and Ca-P transfection. L.W. acknowledges support from The Salk Innovation Grant, the California Institute for Regenerative Medicine (RN1-00577-1), and the National Institutes of Health (1DP2OD004744).

All procedures in the current study were performed using Institutional Animal Care and Use Committee (IACUC) approved protocols for animal handling at The Salk Institute for Biological Studies, La Jolla, CA.
1. Uaa Incorporation in Kir2.1 and Expression of the Resultant PIRK in the Primary Neuronal Culture
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	<li>DNA Construction
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		<li>Select a target site of Kir2.1 for Uaa incorporation. Exploit prior knowledge and information about structure and function of Kir2.1, so that the cho...

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To achieve effective photo-modulation, the important initial step is to decide where to incorporate the photoresponsive Uaa in the target protein. Structural and functional information of the target protein is very helpful to guide the selection of candidate sites. At the same time, the purpose of light regulation would determine which site is most suitable. After choosing candidate sites, we recommend to test the sites in mammalian cell lines such as the Human Embryonic Kidney (HEK) cells for easier culture and manipula...

Compared to conventional electric stimulation, photostimulation offers greater temporal and spatial resolution with minimum interference to the physiological system of specimens. Since the demonstration of using lasers to stimulate neurons in 19711, many creative ways have been invented to exogenously control neuronal activity with light. Optical release of photocaged agonists has long been used to study physiological response of the neuronal network to ligands2,3,4. This technique has limited specificity due to diffusion of caged agonists. Genetic specificity is achieved by ectopically expressing light-sensitive opsin channels and pumps5,6,7, and it has been successfully applied to modulate selected neuronal networks in diverse model organisms. However, it would be difficult to apply this method to optically control various other neuronal proteins, since grafting photoresponsiveness from the opsin proteins to other proteins would require intense engineering that may alter the natural characteristics of the protein under study. Chemically tethering an exogenous photosensitive ligand to a protein has demonstrated another way to control the functionality of channel proteins8,9,10. The ligand is presented or withdrawn from the binding site of the protein through the photoisomerization of the azobenzene moiety. Tethering chemistry limits the application mainly to the extracellular side of membrane proteins, excluding the intracellular side and intracellular proteins.
Photoresponsive Uaas, after being incorporated into proteins, provide a general strategy to manipulate proteins with light. In early efforts, tRNAs chemically acylated with photocaged Uaas were microinjected into Xenopus oocytes to incorporate the Uaas into membrane receptors and ion channels11, which have advanced the understanding of their structure-function relationships12,13,14. This microinjection approach is mainly limited to large oocytes. Genetic incorporation of a Uaa bypasses the technically challenging tRNA acylation and microinjection by using an orthogonal tRNA/synthetase pair, which incorporates the Uaa through endogenous protein translation in live cells15,16,17,18. Uaa incorporation into neuronal proteins has been demonstrated in primary neurons and neural stem cells19,20. More recently, photoresponsive Uaa has been genetically incorporated into a neuronal protein in the mammalian brain in vivo for the first time21. These advancements make it possible to study neuronal proteins with Uaas in their native cellular environment.
Inwardly rectifying potassium channel Kir2.1 is a strong rectifier that passes K+ currents more readily into than out of the cell, and it is essential in regulating physiological processes including cell excitability, vascular tone, heart rate, renal salt flow and insulin release22. Overexpression of Kir2.1 hyperpolarizes the membrane potential of the target neuron, which becomes less excitable23,24. To optically control Kir2.1 in its native cells, Kang et al. genetically incorporated a photo-responsive Uaa into Kir2.1 expressed in mammalian cells, neurons and embryonic mouse brains21. A brief pulse of light was able to convert the Uaa into a natural amino acid Cys, thus activating the target Kir2.1 protein. When this photo-inducible inwardly rectifying potassium (PIRK) channel protein was expressed in rat hippocampal primary neurons, it suppressed neuronal firing in response to light activation. In addition, the PIRK channel was expressed in the embryonic mouse neocortex, and the light-activated PIRK current in cortical neurons was measured. The successful implementation of the Uaa technology in vivo in the mammalian brain opens the door to optically control neuronal proteins in their native environment, which will enable optical dissection of neuronal processes and mechanisms at the molecular level.
In this protocol we describe procedures for genetic incorporation of Uaas into primary neurons in culture and in the embryonic mouse brain in vivo. The photoresponsive Uaa Cmn and Kir2.1 are used to illustrate the process. Methods to assess successful Uaa incorporation and optical control of neuronal protein activity are provided. This protocol provides a clear guide to genetically encoding Uaas in neurons and in vivo, and to optically regulating neuronal protein function via a photoresponsive Uaa. We expect this protocol to facilitate the adoption of the in vivo Uaa technology for neuroscience and optogenetic biological studies.

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optical control of a neuronal protein using a genetically encoded unnatural amino acid in neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

To genetically incorporate a Uaa into a protein in neurons, the first important step is to design appropriate gene constructs to deliver and express genes efficiently in neurons. There are three genetic components for Uaa incorporation: (1) the target gene with the TAG amber stop codon introduced at the chosen site for Uaa incorporation (2) an orthogonal tRNA to recognize the mutated TAG stop codon, and (3) an orthogonal aminoacyl-tRNA synthetase to charge the Uaa onto the orthogonal tRNA...

Watch this Scientific Journal Video about Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons at JoVE.com

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

The overall goal of this procedure is to optically control a neuronal protein's function using a genetically encoded, unnatural amino acid in neurons. This method can help answer key questions about the contribution of a specific neuronal protein to neuronal signalling in brain function with spatial temporal resolution. The main advantage of this technique is it has a potential to be applied to various neuronal proteins to achieve optical regulation of various neuronal processes.Begin this procedure by plating hippocampal neurons onto cover slips in a 24-well plate with 500 microlitres of growth media, after filtering through a 40 micrometre nylon mesh. After that, incubate the neuronal culture at 35 degrees Celcius in a five percent carbon dioxide, 95 percent air humidified incubator for 2 to 3 weeks. On the day of transfection, make fresh 2.5 molar calcium chloride solution in double distilled water, and sterile filter it with a 0.22 micrometre filter.Next, make fresh 2X BBS at pH 7, and sterile filter it with a 0.22 micrometre filter. After that, replace the culture growth media with 500 microlitres of fresh, pre-warmed transfection media, and do not discard the old growth media. Prepare the transfection solution immediately before adding it to the culture by combining calcium chloride and double-distilled water, while slowly agitating the tube.Continue agitating the tube and slowly add DNA into the solution. Next, add 2X BBS buffer dropwise. Preparation of 2X BBS buffer is a critical step in calcium phosophate transfection, and it is a good idea to calibrate optimal pH for 2X BBS buffer between 6.90 to 7.15 for new DNA constructs or preps.Then, immediately add 30 microlitres of the transfection solution to each cover slip of the neuronal culture. Rock the culture dish a few times to mix the solution, and incubate it at 35 degrees Celcius for 45 minutes to one hour. After about an hour, a layer of fine calcium phosphate precipitates should be observed covering the neurons.Next, replace the transfecting media with 500 microlitres of pre-warmed washing buffer, and incubate it at 35 degrees Celcius for 15 to 20 minutes. The calcium phosphate precipitates should disappear after the wash. Now, replace the washing buffer with 500 microlitres of fresh growth media.Afterward, replace the growth media with the saved original media. Then, add pre-mixed, unnatural amino acid Cmn in 50 microlitres of warm growth media to the culture, that results in a one millimolar final concentration. Incubate the transfected culture at 35 degrees Celcius for 12 to 48 hours before recording.In this procedure, prepare the extra-and intracellular solutions. In our setup, the LED with emission of 385 nanometres is installed next to the microscope to deliver light to one centimetre away from the focal point at a 45 degree angle. Then, pull some patch pipettes with a glass electrode using a micro pipette puller.Afterward, take a neuron cultures coverslip from the incubator and rinse it once in the extracellular solution. Profuse the rig with extracellular solution. Subsequently, place the coverslip on the electrophysiology microscope platform, using standard patch clamping techniques.Patch a neuron transfected with M-citrine fluorescents. Next, record the neuronal activity using the current clamp method. First, adjust the resting potential to around negative 72 millivolts.Then, inject a step current to induce continuous firing of action potentials at five to 15 Hertz. While recording, deliver single LED pulses to the neuron, and monitor the changes of its action potentials. Then, add 0.5 millimolar barium chloride to the bath, to recover the action potentials.In the setting of acute brain slice, patch a neuron with M-cherry GFP fluorescents from the neocortical region. Then, record PIRK activity using the voltage ramp method. Specifically, monitor the Kir2.1 specific inward currents at negative 100 millivolts.Then, deliver single LED pulses to the neuron while recording, and monitor if PIRK proteins are activated. Once PIRK is activated, inward currents at negative 100 millivolts would increase significantly. At the end, add 0.5 millimolar barium chloride to the bath, and verify if PIRK is inactivated.Shown here are exemplary pictures of healthy rat hippocampal neuronal cultures. And two healthy neurons exhibiting plump cell bodies and pronounced dendrites and axons. These are the DIC and fluorescence images of rat hippocampal neurons cultures in vitro, and transfected for PIRK expression without CMN.In the presence of one millimolar CMN, transfected neurons are capable of expressing full-length PIRK M-citrine proteins, thus showing green fluorescence. And here is an example of a single light pulse, suppressing the activity of a PIRK expressed hippocampal neuron. These fluroescence images of mouse embryonic cortical neurons show the successful incorporation of CMN into GFP and Kir2.1 proteins in vivo.In this graph, the light-dependent activation of PIRK is indicated in the currents recorded from the mouse neocortical neurons. The implications of this technique extends to therapy of brain diseases caused by malfunction of specific neuronal proteins, for example, mutated Kir2.1 in Andersens syndrome, because this technique helps uncover which brain circuitry is affected by Kir2.1 deficits. Once mastered, this technique can be used to control neuronal proteins function with light in high specificity in live neurons.After watching this video, you should have a good understanding of how to incorporate unnatural amino acids into proteins in neurons to achieve photo control over neuronal proteins of interest.

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Research

JoVE Journal

Neuroscience

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; Gan et al., 2000). Here we provide a detailed description of each step required together with exemplary images of good and bad outcomes that will help when setting up the technique. In our experience, a few steps proved critical for the successful application of DiOLISTICS. These considerations include the quality of the DiI-coated bullets, the extent of fixative exposure, and the concentration of detergent used in the incubation solutions. Tips and solutions for common problems are provided. This is a versatile labeling technique that can be applied to multiple animal species at a wide range of ages. Unlike other fluorescent labeling techniques that are limited to preparations from young animals or restricted to mice because they rely on the expression of a fluorescent transgene, DiOLISTIC labeling can be applied to animals of all ages, species and genotypes and it can be used in combination with immunostaining to identify a specific subpopulation of cells. Here we demonstrate the use of DiOLISTICS to label neurons in brain slices from adult mice and adult non-human primates with the purpose of quantifying dendrite branching and dendritic spine morphology.

We demonstrate the use of the gene gun to introduce fluorescent dyes, such as DiI, into neurons in brain slices from rodents and non-human primates of different ages. In this particular case, we use adult mice (3-6 months old) and adult cynomologus monkeys (9-15 years old). This technique, originally described by the laboratory of Dr. Lichtman (Gan et al., 2000), is well suited for the study of dendritic branching and dendritic spine morphology and can be combined with traditional immunostaining, if detergents are kept at a low concentration.

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; ...

Efficient Derivation of Human Neuronal Progenitors and Neurons from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system (CNS) structure and circuitry in today's healthcare industry. Cell-based therapies hold great promise to restore the lost nerve tissue and function for CNS disorders. However, cell therapies based on CNS-derived neural stem cells have encountered supply restriction and difficulty to use in the clinical setting due to their limited expansion ability in culture and failing plasticity after extensive passaging1-3. Despite some beneficial outcomes, the CNS-derived human neural stem cells (hNSCs) appear to exert their therapeutic effects primarily by their non-neuronal progenies through producing trophic and neuroprotective molecules to rescue the endogenous cells1-3. Alternatively, pluripotent human embryonic stem cells (hESCs) proffer cures for a wide range of neurological disorders by supplying the diversity of human neuronal cell types in the developing CNS for regeneration1,4-7. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity7-10. In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic11-13. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules14 (please see a schematic in Fig. 1). Retinoic acid (RA) does not induce neuronal differentiation of undifferentiated hESCs maintained on feeders1, 14. And unlike mouse ESCs, treating hESC-differentiated embryoid bodies (EBs) only slightly increases the low yield of neurons1, 14, 15. However, after screening a variety of small molecules and growth factors, we found that such defined conditions rendered retinoic acid (RA) sufficient to induce the specification of neuroectoderm direct from pluripotent hESCs that further progressed to neuroblasts that generated human neuronal progenitors and neurons in the developing CNS with high efficiency (Fig. 2). We defined conditions for induction of neuroblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human neuronal cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of neuroblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human neuronal progenitors and neuronal cell types in the developing CNS for neural repair.

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Super-resolution Imaging of Neuronal Dense-core Vesicles

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and medical applications. Strengths of fluorescence include its sensitivity, specificity, and compatibility with live imaging. Unfortunately, conventional forms of fluorescence microscopy suffer from one major weakness, diffraction-limited resolution in the imaging plane, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, this limitation has been overcome with the introduction of super-resolution fluorescence microscopy techniques, such as photoactivated localization microscopy (PALM). Unlike its conventional counterparts, PALM can produce images with a lateral resolution of tens of nanometers. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of cellular structure and organization.
Unfortunately, PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. In this article, we show how to implement single-color PALM studies of vesicular structures in fixed, cultured neurons. PALM is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm. Key steps in our approach include labeling neurons with photoconvertible (green to red) chimeras of vesicle cargo, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a high-resolution PALM image. We also demonstrate the efficacy of our approach by presenting exceptionally well-resolved images of dense-core vesicles (DCVs) in cultured hippocampal neurons, which refute the hypothesis that extrasynaptic trafficking of DCVs is mediated largely by DCV clusters.

We describe how to implement photoactivated localization microscopy (PALM)-based studies of vesicles in fixed, cultured neurons. Key components of our protocol include labeling vesicles with photoconvertible chimeras, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a super-resolution image.

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of pyramidal neurons and dendritic spines, a ballistic labeling technique can be utilized. In the present protocol, pyramidal neurons are labeled with DilC18(3) dye and analyzed using neuronal reconstruction software to assess neuronal morphology and dendritic spines. To investigate neuronal structure, dendritic branching analysis and Sholl analysis are performed, allowing researchers to draw inferences about dendritic branching complexity and neuronal arbor complexity, respectively. The evaluation of dendritic spines is conducted using an automatic assisted classification algorithm integral to the reconstruction software, which classifies spines into four categories (i.e., thin, mushroom, stubby, filopodia). Furthermore, an additional three parameters (i.e., length, head diameter, and volume) are also chosen to assess alterations in dendritic spine morphology. To validate the potential of wide application of the ballistic labeling technique, pyramidal neurons from in vitro cell culture were successfully labeled. Overall, the ballistic labeling method is unique and useful for visualizing neurons in different brain regions in rats, which in combination with sophisticated reconstruction software, allows researchers to elucidate the possible mechanisms underlying neurocognitive dysfunction.

We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of ...