This work was supported by National Institutes of Health grants 2 R15 GM061539-02 (to B.A.S.), 2 R15 NS40425-03 (to J.E.L.), MH 66179 (to Dr. Gary Banker of Oregon Health &#160;&#38;&#160;Science&#160;University/OHSU), and P30 NS061800 (to Dr. Sue Aicher of OHSU). We thank Barbara Smoody for extensive support with the culture of hippocampal neurons, and Drs. Brian Long and James Abney for a critical reading of this manuscript.

1. Sample Preparation
<ol>
	<li>Prepare DNA encoding a photoconvertible or photoactivatable chimera targeted to DCVs, using standard molecular biology techniques. 
	Note: One possibility is DNA encoding a tissue plasminogen activator-Dendra2 chimera (tPA-Dendra2). Dendra2 is a photoconvertible protein that switches from green to red emission upon exposure to ultraviolet light12.</li>
	<li>Culture hippocampal neurons on high performance #1.5 cover slips for ~5-10 days, following standard protocols13.</li>
	<li>Transfect developing hippocampal neurons using a cationic lipid reagent. 
	Note: Viral-mediated infection is more effective for mature neurons. This alternative approach has been described in detail elsewhere14.
	<ol>
		<li>Prepare a 1.5 ml tube containing ~1 &#181;g of DNA and 250 &#181;l of minimal essential medium (MEM) and a second 1.5 ml tube containing 4 &#181;l of the cationic lipid reagent and 250 &#181;l of MEM.</li>
		<li>Incubate the solutions for 5 min.</li>
		<li>Combine the two solutions and incubate the resulting mixture for an additional 30 min.</li>
		<li>During the incubation, prepare a dish containing several ml&#160;of culture medium warmed to 37 &#176;C.</li>
		<li>Gently transfer cover slips containing neurons to the dish containing warm medium and add the transfection mixture.</li>
		<li>Tilt the dish gently and then place it in an incubator at 37 &#176;C for 90 min.</li>
		<li>Return neurons to their &#34;home dish&#34; and wait ~12-18 hr.</li>
	</ol>
	</li>
	<li>Fix cells for 45 min in 4% paraformaldehyde/4% sucrose in phosphate buffered saline (PBS, pH 7.4) warmed to 37 &#176;C. 
	Note: In PALM experiments, it is important to achieve good preservation of ultrastructure, and standard formaldehyde fixation protocols used for immunofluorescence staining typically are not sufficient15. One important reason for poor structural preservation is insufficient fixation time because formaldehyde fixation involves adduct formation and protein crosslinking, and the latter process is quite slow16. A 45 min fixation helps to mitigate this problem with no detectable overfixation-induced loss of fluorescence.</li>
	<li>Wash cover slips ~10x with filtered PBS. 
	Note: This reduces fluorescent contaminants. Image samples relatively quickly after fixation. In the interim, store cells at 4 &#176;C in filtered PBS in a light-tight container.</li>
	<li>Mount cover slips containing neurons in a chamber in filtered PBS. 
	Note: The sample must be mounted in an aqueous medium, like PBS, with refractive index lower than that of glass, to implement total internal reflection fluorescence (TIRF)-based illumination. TIRF-based illumination is very useful because only cover slip-proximal fluorophores are excited, and there is a large associated drop in background fluorescence17.</li>
</ol>
2. Image Acquisition
<ol>
	<li>Turn on the super-resolution imaging system.
	<ol>
		<li>Turn on the arc lamp.</li>
		<li>Flip the &#34;system/pc&#34; and &#34;component&#34; switches (on the power remote switch) to &#34;on.&#34;</li>
		<li>Turn on the computer (after the microscope control tablet/docking station comes on). 
		Note: The docking station can be used to control and monitor many attributes of the optics and light path, notably choice of objective and focus.</li>
		<li>Double click the software icon and choose the &#34;start system&#34; option in the login dialog.</li>
	</ol>
	</li>
	<li>Examine the sample.
	<ol>
		<li>Open the front and top access panels on the laser safety cabinet.</li>
		<li>Tilt the transmitted&#160;light arm to access the stage and objectives.</li>
		<li>Put oil on the alpha Plan-Apochromat 100X numerical aperture/NA = 1.46 (TIRF) objective. 
		Note: A 63X high-NA objective can be used in lieu of the 100X.</li>
		<li>Mount the sample on the stage, and raise the objective toward the sample.</li>
		<li>Monitor the lens position using the XYZ function of the docking station. Stop when the lens is in the ballpark of focus (e.g., z ~2.50 mm).</li>
		<li>Fully close the access panels. 
		Note: To engage the laser safety.</li>
		<li>Fine tune the focus using the oculars and buttons in the locate tab. 
		Note: The locate tab provides access to buttons that produce arc lamp-based illumination and facilitates direct observation of the sample with the oculars. Similarly, the acquisition tab provides access to tools that produce laser-based illumination and facilitates image capture by the camera.
		<ol>
			<li>Go to the locate tab, choose &#34;Trans On.&#34;</li>
			<li>Click the ocular expansion symbol. 
			Note: This brings up a schematic of the light path. Attributes of the light path can be altered by left clicking appropriate icons in the schematic or by using the touch screen on the docking station. For example, a filter appropriate for viewing emission from Dendra2 can be chosen using the reflector revolver icon.</li>
			<li>Look into the eyepieces, and bring the cells into focus using the docking station as a guide. 
			Note: The sample will be very sparsely populated with fluorescent cells because hippocampal neurons are difficult to transfect, and it thus can be a little tricky to focus using reflected illumination. If this is the case, use transmitted illumination and the z reading of the docking station as a guide.</li>
			<li>Toggle the light &#34;Off&#34; after focusing.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Identify a cell that is fairly bright, exhibits punctate fluorescence, and has a classic morphology that is indicative of good health.
	<ol>
		<li>Choose reflected light and the filter appropriate for viewing green emission from Dendra2.</li>
		<li>Scan the cover slip for cells expressing Dendra2 chimeras using low intensity excitation light.</li>
	</ol>
	</li>
	<li>Switch to the acquisition tab.
	<ol>
		<li>Use the &#34;experiment manager&#34; to load a PALM acquisition configuration that is appropriately designed or easily modified. 
		Note: If no such experiment exists, design a starting experimental configuration using the following settings as a guide: Laser power sliders: 405 nm (~0.01% for a 50 mW laser); 488 nm (~3% for a 100 mW laser); 561 nm (~40% for a 100 mW laser); Exposure time: ~50 msec; Camera gain: ~200; Multidimensional acquisition: time series (cycles = 30,000, interval = 0); Tracks: 488 nm with epi-illumination, and 561 nm with TIRF-based illumination (see step 2.5.1); Objective: 100X (NA = 1.46); TIRF angle slider: incidence angle ~67-72&#176;; Pixel size: 100 nm; Field of view: TIRF (the alternative choices yield higher laser intensities and are used for fluorophores that are more difficult to bleach).</li>
		<li>Click &#34;yes&#34; when a pop up menu appears with a query about switching on lasers.</li>
	</ol>
	</li>
	<li>Bring the image of the cell into focus at the camera plane.
	<ol>
		<li>Choose the 488 nm epi-illumination laser track. 
		Note: Tracks are beam configurations used during imaging. Here, track names are determined by the excitation wavelength that produces the emission that is passed to the camera. For example, the 561 nm track uses both the 405 and 561 nm laser lines, but the filter cube passes only emission from photoconverted Dendra2 excited by 561 nm light to the camera.</li>
		<li>Focus the image of the cell on the camera using the &#34;continuous&#34; acquisition button.</li>
	</ol>
	</li>
	<li>Collect a conventional widefield image.
	<ol>
		<li>Use the &#34;Snap&#34; button.</li>
		<li>Save the image by going to the &#34;File&#34; menu and choosing &#34;save/save as.&#34;</li>
	</ol>
	</li>
	<li>Set up the TIRF-based illumination.
	<ol>
		<li>Choose the 488 track and switch to &#34;TIRF&#34; using the EPI/TIRF button.</li>
		<li>Choose &#34;continuous&#34; acquisition.</li>
		<li>Adjust the TIRF-based illumination slider. 
		Note: TIRF is achieved when there is a sudden darkening of background AND the specimen can be focused in ONE plane18. If new features become apparent via focusing up into the sample, the incidence angle is too low because in TIRF there is only one plane of focus.</li>
		<li>Double check the TIRF angle using the 561 track with the 405 nm laser &#34;off.&#34;</li>
		<li>Turn the 405 nm laser back &#34;on&#34; before initiating a PALM experiment.</li>
	</ol>
	</li>
	<li>Collect PALM data.
	<ol>
		<li>Hit &#34;start experiment.&#34; 
		Note: If desired, open the &#34;online processing options&#34; tab and check &#34;online processing PALM&#34; and &#34;Fit Gauss2D&#34; (before starting the experiment) to monitor the reconstructed PALM image during raw data collection. This will facilitate assessment of the quality of the data and thus the value of continuing the relatively time-consuming data collection process.</li>
		<li>Visually monitor photoconversion.</li>
		<li>Increase the intensity of the 405 nm (photoconverting) laser when Dendra2 photoconversion diminishes (typically after acquisition of ~10,000 raw images).</li>
		<li>Continue the experiment if photoconversion increases. Otherwise, stop the experiment (without loss of data) by hitting the &#34;stop&#34; current step button.</li>
		<li>Save the images when data collection is complete (after ~15 min).</li>
	</ol>
	</li>
</ol>
3. Image Processing, Display, and Analysis
<ol>
	<li>In the absence of online processing, process raw images that were saved to disk.
	<ol>
		<li>Click on the processing tab, open the method tab, and select &#34;PALM.&#34;</li>
		<li>Choose &#34;PALM&#34; again from the four suboptions.</li>
		<li>Go to the file menu, open and view the file of interest, and hit &#34;select.&#34;</li>
		<li>Hit &#34;apply.&#34; 
		Note: This will initiate peak/fluorophore finding and peak localization with default options. If necessary, peaks also can be grouped using the &#34;PAL-Group&#34; tab. Grouping assigns peaks that appear in several consecutive frames to a single molecule if the peak coordinates are sufficiently similar. In addition, the data can be corrected for sample drift using the software&#8217;s model-based drift alignment option.</li>
	</ol>
	</li>
	<li>Filter out poorly localized and poorly sampled components in the peak-localized data.
	<ol>
		<li>Discard fluorophores with localization precisions, &#963;, &#62;35 nm using the &#34;PAL-Filter&#34; tool.</li>
		<li>Discard fluorophores in areas where the mean distance between peaks, d, is too large to resolve vesicles of diameter, D.
		<ol>
			<li>Extract the mean localization precision,&#160;&#963;, from the localization precision histogram.</li>
			<li>Calculate a maximum acceptable value for d based on &#963;, and the Shannon-Nyquist criteron19, as embodied in the resolution approximation20 R = D = [(2.35&#963;)2 + (2d)2]1/2.</li>
			<li>Set the threshold in the &#34;Remove PALM outliers&#34; tool to delete peaks surrounded by fewer than &#960;D2/(4d2) neighbors within a circle of radius D/2.</li>
		</ol>
		</li>
		<li>Convert the processed PALM data into a PALM image using the &#34;PAL-Rendering&#34; tool; see Figure 1.</li>
		<li>Quantify sizes and separations by choosing the &#34;Profile&#34; function.
		<ol>
			<li>Check the associated line and table options, and draw a line along the desired direction in the image.</li>
			<li>Quantify diameters by determining full widths at half maximal intensity.</li>
			<li>Quantify separations by determining peak-to-peak spacings of intensity profiles.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

PALM and related super-resolution fluorescence microscopy techniques have recently emerged as a valuable complement to better-established forms of optical microscopy and electron microscopy (EM)22-24. Positive attributes of PALM include relatively simple sample preparation and minimal sample perturbation. In principle, PALM also can be used to study living cells. Probably the main negative attribute of PALM is time-consuming data acquisition, which significantly hampers studies of faster dynamic processes in l...

A number of cellular processes depend on accurate and efficient vesicle-mediated trafficking of biomolecules to specific subcellular destinations. One prominent example is synaptic assembly, which is preceded by long-ranged, vesicle-mediated delivery of synaptic constituents from sites of biogenesis in the neuronal soma to potentially distal pre- and postsynaptic sites1.
Fluorescence microscopy is a powerful and popular method of studying vesicle trafficking. Strengths of the technique include its sensitivity, specificity, and compatibility with live imaging2. Unfortunately, until relatively recently, the technique has suffered from one major weakness, diffraction-limited resolution2, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, lateral resolution in fluorescence microscopy surpassed the diffraction barrier with the introduction of super-resolution fluorescence microscopy techniques, such as PALM3. The lateral resolution of PALM, tens of nanometers, is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm4. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of vesicles, including some aspects of their trafficking to specific subcellular sites.
PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. Here we describe how to implement PALM studies of vesicular structures, and we demonstrate the efficacy of our approach for the case of DCVs in hippocampal neurons. In particular, we use PALM to address the hypothesis that trafficking of DCVs to synapses in hippocampal neurons is mediated by DCV clusters5-8.
Cluster-mediated trafficking of vesicles to synapses in developing neurons is an intriguing possibility because it may facilitate rapid synaptic stabilization and assembly9,10. Proponents of clustered trafficking of DCVs cite the large apparent size of extrasynaptic fluorescent puncta harboring exogenous DCV cargo as evidence supporting clustering6. However, these puncta appear in images generated using diffraction-limited fluorescence microscopy techniques, which are not suited to distinguishing size effects arising from diffraction from those arising from clustering.
To resolve this issue, we collected conventional widefield fluorescence and PALM images of hippocampal neurons expressing chimeras targeted to DCVs. Analysis of these images revealed that &#62;92% of putative extrasynaptic DCV clusters in conventional images are resolved as 80 nm (individual DCV-sized)11 puncta in PALM images. Thus, these data largely invalidate the clustering hypothesis as applied to DCVs in developing hippocampal neurons.

<table border="0" cellpadding="0" cellspacing="0" width="1105">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Zeiss PALM</td>
			<td style="width:169px;">Carl Zeiss, Inc.</td>
			<td style="width:119px;">Elyra PS.1</td>
			<td style="width:572px;">With Zeiss Efficient Navigation (ZEN) software and fluorescence filter Set 77 HE GFP/mRFP/Alexa633&#160;</td>
		</tr>
		<tr height="28">
			<td height="28" style="height:28px;width:245px;">Lipofectamine 2000 Transfection Reagent</td>
			<td style="width:169px;">Life Technoligies</td>
			<td style="width:119px;">11668-019</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Minimum Essential Medium</td>
			<td style="width:169px;">Life Technologies</td>
			<td style="width:119px;">11095-080</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Phosphate-Buffered Saline</td>
			<td style="width:169px;">Life Technologies</td>
			<td style="width:119px;">10010049</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:245px;">Paraformaldehyde</td>
			<td style="width:169px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">19208</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">Sucrose</td>
			<td style="width:169px;">Sigma-Aldrich</td>
			<td style="width:119px;">S-8501</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:245px;">growth glass coverslips 18mm 1.5D</td>
			<td style="width:169px;">Fisher Scientific</td>
			<td style="width:119px;">NC0059095</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Figure 1 shows one end product of imaging and processing. In this PALM image, lateral coordinates of localized fluorophores are shown using the centroid display mode, and the super-resolution image of the associated DCV is shown using the Gaussian display mode.
Figure 2A shows analogous widefield and PALM images of the soma and proximal processes of an eight days in vitro hippocampal neuron expressing tPA-Dendra2. Important features of the images incl...

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Super-resolution Imaging of Neuronal Dense-core Vesicles

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 ...

super-resolution-imaging-of-neuronal-dense-core-vesicles

Research

JoVE Journal

Neuroscience

9.6K Views.  Lewis & Clark College. 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.

Video: Super-resolution Imaging of Neuronal Dense-core Vesicles

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

Prolonged Incubation of Acute Neuronal Tissue for Electrophysiology and Calcium-imaging

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the experimental time, and increases the number of animals that are utilized per study. This limitation specifically impacts protocols such as calcium imaging that require prolonged pre-incubation with bath-applied dyes. Exponential bacterial growth within 3 - 4 h after slicing is tightly correlated with a decrease in tissue health. This study describes a method for&#160;limiting the proliferation of bacteria in acute preparations to maintain viable neuronal tissue for prolonged periods of time (&#62;24 h) without the need for antibiotics, sterile procedures, or tissue culture media containing growth factors. By cycling the extracellular fluid through UV irradiation and keeping the tissue in a custom holding chamber at 15 - 16 &#176;C, the tissue shows no difference in electrophysiological properties, or calcium signaling through intracellular calcium dyes at &#62;24 h postdissection. These methods will not only extend experimental time for those using acute neuronal tissue, but will reduce the number of animals required to complete experimental goals, and will set a gold standard for acute neuronal tissue incubation.

Once removed from the body, neuronal tissue is greatly affected by environmental conditions, leading to eventual degradation of the tissue after 6 - 8 h. Using a unique incubation method, which closely monitors and regulates the extracellular environment of the tissue, tissue viability can be significantly extended for &#62;24 h.

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the ...

Analyzing Dendritic Morphology in Columns and Layers

In many regions of the central nervous systems, such as the fly optic lobes and the vertebrate cortex, synaptic circuits are organized in layers and columns to facilitate brain wiring during development and information processing in developed animals. Postsynaptic neurons elaborate dendrites in type-specific patterns in specific layers to synapse with appropriate presynaptic terminals. The fly medulla neuropil is composed of 10 layers and about 750 columns; each column is innervated by dendrites of over 38 types of medulla neurons, which match with the axonal terminals of some 7 types of afferents in a type-specific fashion. This report details the procedures to image and analyze dendrites of medulla neurons. The workflow includes three sections: (i) the dual-view imaging section combines two confocal image stacks collected at orthogonal orientations into a high-resolution 3D image of dendrites; (ii) the dendrite tracing and registration section traces dendritic arbors in 3D and registers dendritic traces to the reference column array; (iii) the dendritic analysis section analyzes dendritic patterns with respect to columns and layers, including layer-specific termination and planar projection direction of dendritic arbors, and derives estimates of dendritic branching and termination frequencies. The protocols utilize custom plugins built on the open-source MIPAV (Medical Imaging Processing, Analysis, and Visualization) platform and custom toolboxes in the matrix laboratory language. Together, these protocols provide a complete workflow to analyze the dendritic routing of Drosophila medulla neurons in layers and columns, to identify cell types, and to determine defects in mutants.

Here, we show how to analyze dendritic routing of Drosophila medulla neurons in columns and layers. The workflow includes a dual-view imaging technique to improve the image quality and computational tools for tracing, registering dendritic arbors to the reference column array and for analyzing the dendritic structures in 3D space.

In many regions of the central nervous systems, such as the fly optic lobes and the vertebrate cortex, synaptic circuits are organized in layers and ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

Toxicity of misfolded proteins and mitochondrial dysfunction are pivotal factors that promote age-associated functional neuronal decline and neurodegenerative disease across species. Although these neurotoxic challenges have long been considered to be cell-intrinsic, considerable evidence now supports that misfolded human disease proteins originating in one neuron can appear in neighboring cells, a phenomenon proposed to promote pathology spread in human neurodegenerative disease.
C. elegans adult neurons that express aggregating proteins can extrude large (~4 &#181;m) membrane-surrounded vesicles that can include the aggregated protein, mitochondria, and lysosomes. These large vesicles are called &#8220;exophers&#8221; and are distinct from exosomes (which are about 100x smaller and have different biogenesis). Throwing out cellular debris in exophers may occur by a conserved mechanism that constitutes a fundamental, but formerly unrecognized, branch of neuronal proteostasis and mitochondrial quality control, relevant to processes by which aggregates spread in human neurodegenerative diseases.
While exophers have been mostly studied in animals that express high copy transgenic mCherry within touch neurons, these protocols are equally useful in the study of exophergenesis using fluorescently tagged organelles or other proteins of interest in various classes of neurons.
Described here are the physical features of C. elegans exophers, strategies for their detection, identification criteria, optimal timing for quantitation, and animal growth protocols that control for stresses that can modulate exopher production levels. Together, details of protocols outlined here should serve to establish a standard for quantitative analysis of exophers across laboratories. This document seeks to serve as a resource in the field for laboratories seeking to elaborate molecular mechanisms by which exophers are produced and by which exophers are reacted to by neighboring and distant cells.

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

This protocol describes approaches for detection and quantitation of large aggregate and/or organelle extrusions (~4 &#181;m) produced by C. elegans cells in the form of membrane-bound exophers. We describe strains, growth conditions, scoring criteria, timing, and microscopy considerations needed to facilitate dissection of this debris expulsion mechanism.

Toxicity of misfolded proteins and mitochondrial dysfunction are pivotal factors that promote age-associated functional neuronal decline and ...

Super-Resolution Imaging to Study Co-Localization of Proteins and Synaptic Markers in Primary Neurons

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. Imaging studies are widely used to investigate their function and plasticity in physiological and pathological conditions. Because of their size and structure, localization studies of proteins require high-resolution imaging techniques. In this protocol, we describe a procedure to study in primary neurons the co-localization of target proteins with synaptic markers at a super-resolution level using structured illumination microscopy (SIM). SIM is a patterned-light illumination technique that doubles the spatial resolution of wide-field microscopy, reaching a detail of around 100 nm. The protocol indicates the required controls and settings for robust co-localization studies and an overview of the statistical methods to analyze the imaging data properly.

This protocol shows how to employ super-resolution microscopy to study protein co-localization in primary neuronal cultures.

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

ROS Live Cell Imaging During Neuronal Development

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the different ROS, hydrogen peroxide (H2O2) is best characterized with respect to roles in cellular signaling. H2O2 has been implicated during the development in several species. For example, a transient increase in H2O2 has been detected in zebrafish embryos during the first days following fertilization. Furthermore, depleting an important cellular H2O2 source, NADPH oxidase (NOX), impairs nervous system development such as the differentiation, axonal growth, and guidance of retinal ganglion cells (RGCs) both in&#160;vivo and in vitro. Here, we describe a method for imaging intracellular H2O2 levels in cultured zebrafish neurons and whole larvae during development using the genetically encoded H2O2-specific biosensor, roGFP2-Orp1. This probe can be transiently or stably expressed in zebrafish larvae. Furthermore, the ratiometric readout diminishes the probability of detecting artifacts due to differential gene expression or volume effects. First, we demonstrate how to isolate and culture RGCs derived from zebrafish embryos that transiently express roGFP2-Orp1. Then, we use whole larvae to monitor H2O2 levels at the tissue level. The sensor has been validated by the addition of H2O2. Additionally, this methodology could be used to measure H2O2 levels in specific cell types and tissues by generating transgenic animals with tissue-specific biosensor expression. As zebrafish facilitate genetic and developmental manipulations, the approach demonstrated here could serve as a pipeline to test the role of H2O2 during neuronal and general embryonic development in vertebrates.

This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.