The authors would like to thank Edoardo Micotti for constructive criticism of the manuscript. This study was supported by BrightFocus A2019296F, by Fondo di Beneficenza - Gruppo Intesa Sanpaolo (LC), by Fondazione Regionale per la Ricerca Biomedica (Care4NeuroRare CP_20/2018) (CN), by the Marie Sk&#322;odowska-Curie Innovative Training Network (JK) and by Fondazione Telethon TCP15011, Alzheimer&#39;s Association AARG-17-505136 (LF).

1. Primary cultures
<ol>
	<li>Culture mouse hippocampal primary neurons in chambered coverslips (such as Ibidi &#181;-Slide 8 Well or Nunc Lab-Tek Chambered Coverglass) that match the objective requirement for #1.5 (0.17 mm) coverslip thickness.</li>
	<li>Coat chambered coverslips with 100 &#181;L of poly-L-lysine (100 &#181;g/mL).</li>
	<li>The next day, wash the chambered coverslips twice with sterile phosphate-buffered saline (PBS).</li>
	<li>To obtain mouse primary neurons, isolate hippocampi from P1-P4 pups<sup cl...

<ol>
	<li>Foster, M., Sherrington, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> A textbook of physiology, part three: The central nervous system (7th ed.). , (1897).</li><li>Choquet, D., Triller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Dynamic+Synapse.">The Dynamic Synapse.</a> Neuron. 80 (3), 691-703 (2013).</li><li>McAllister, A. 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G., Galbraith, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Super-resolution+microscopy+at+a+glance.">Super-resolution microscopy at a glance.</a> Journal of Cell Science. 124 (10), 1607-1611 (2011).</li><li>Gustafsson, M. G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surpassing+the+lateral+resolution+limit+by+a+factor+of+two+using+structured+illumination+microscopy.">Surpassing the lateral resolution limit by a factor of two using structured illumination microscopy.</a> Journal of Microscopy. 198 (2), 82-87 (2000).</li><li>Gustafsson, M. G. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptopathic+mechanisms+of+neurodegeneration+and+dementia:+Insights+from+Huntington's+disease.">Synaptopathic mechanisms of neurodegeneration and dementia: Insights from Huntington's disease.</a> Progress in Neurobiology. 153, 18-45 (2017).</li><li>Won, H., Mah, W., Kim, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autism+spectrum+disorder+causes,+mechanisms,+and+treatments:+focus+on+neuronal+synapses.">Autism spectrum disorder causes, mechanisms, and treatments: focus on neuronal synapses.</a> Frontiers in Molecular Neuroscience. 6, (2013).</li><li>Pfeiffer, B. E., Huber, K. 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Elucidating the structure and composition of the synapse is crucial for understanding the physiological and pathological processes that regulate memory and cognition. While in the normal state, synapses are the building blocks of memory, they also underlie complex neurological disorders such as Alzheimer&#8217;s disease32. The protocol described here serves to study the co-localization of neuronal proteins with a super-resolution microscopy technique called SIM. Using a particular pattern of illum...

The understanding and view of the synapse has changed enormously since its first description by Foster and Sherrington in 18971. Since then, our knowledge of neuronal communication and the molecular processes behind it has grown exponentially2. It has become clear that synapses can be thought of as a two-compartment system: a pre-synaptic compartment containing vesicles for the release of neurotransmitters and a post-synaptic compartment with receptors3. This simplistic view, in the past twenty years, has evolved into a complex network of the proteins required to transduce transmitter binding into signaling4.
The gains in the understanding are partially due to super-resolution techniques that overcame the diffraction limit of conventional light microscopy to suit the dimension of synapses better5,6,7,8,9,10. Due to the diffraction limit, an optical microscope cannot reach a resolution above 200 nm laterally11,12. To bypass this limit, super-resolution techniques were created, using different approaches and reaching different sub-diffraction limit resolutions: SIM, STED (Stimulated Emission Depletion Microscopy), PALM (PhotoActivated Localization Microscopy) and STORM (Stochastic Optical Reconstruction Microscopy)13,14. SIM doubles the spatial resolution of laser-based wide-field microscopy systems by inserting a diffraction grating into the excitation beam path15. The movable grating diffracts the laser beams to create a known illumination pattern, usually stripes. This purposely structured light pattern is superimposed to the unknown spatial distribution of the fluorescent dye (of the sample). The interference fringes formed by the two patterns encode for otherwise indistinguishable fine details with normal wide-field microscopy. The final super-resolved image is obtained by combining and decoding with mathematical methods several raw images of the same sample obtained by the translations and rotations of the diffraction grating. The resolution of the super-resolved images reaches 100 nm in the lateral and 500 nm in the axial directions for 2D-SIM15 or 100 nm in the lateral and 250 nm in the axial directions for 3D-SIM16.
The new understanding of the synapse is even more important in the light of the many neurological disorders where synaptic dysfunction plays a major role in onset and progression17,18. Alzheimer&#8217;s disease, Down syndrome, Parkinson&#8217;s disease, prion diseases, epilepsy, autism spectrum disorders and fragile X syndrome among others have been linked to abnormalities in synaptic composition, morphology and function19,20,21,22.
Recently, using a set of SUMO-specific antibodies, we used SIM to show co-localization in primary hippocampal neurons of the SUMO proteins with the pre- and post-synaptic markers synaptophysin and PSD95 at super-resolution level23. This enabled us to confirm biochemical and confocal microscopy evidence of SUMO localization in neurons.
Here, we describe a protocol to study the localization of proteins in mouse hippocampal primary neurons. At the same time, this protocol may be adapted to different types of primary neuronal cultures.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
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			<td >0.4% Trypan blue solution&#160;</td>
			<td >Thermo Fisher Scientific</td>
			<td >15250061</td>
			<td >Chemical</td>
		</tr>
		<tr >
			<td >70 &#181;m filter&#160;</td>
			<td>Corning</td>
			<td>352350</td>
			<td>Equiment</td>
		</tr>
		<tr >
			<td >Alexa</td>
			<td>Thermo Fisher Scientific</td>
			<td>-</td>
			<td>Antibody</td>
		</tr>
		<tr >
			<td >Antibody SENP1</td>
			<td >Santa Cruz</td>
			<td >sc-271360</td>
			<td>Antibody</td>
		</tr>
		<tr >
			<td >B27 Supplement</td>
			<td >Life Technologies</td>
			<td >17504044</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >Bovine serum albumin&#160;</td>
			<td>Merck</td>
			<td>5470</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >CaCl2</td>
			<td >Merck Life Science</td>
			<td >21115</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >Chambered coverslips</td>
			<td>Ibidi</td>
			<td>80826</td>
			<td>Equiment</td>
		</tr>
		<tr >
			<td >DyLight</td>
			<td>Thermo Fisher Scientific</td>
			<td>-</td>
			<td>Antibody</td>
		</tr>
		<tr >
			<td >FBS (Hyclone)</td>
			<td>GIBCO</td>
			<td>SH3007002 (CHA1111L)</td>
			<td>Serum</td>
		</tr>
		<tr >
			<td >FluoSpheres carboxylate-modified microspheres, 0.1 &#956;m, yellow&#8211;green fluorescent</td>
			<td>Thermo Fisher Scientific</td>
			<td>F8803</td>
			<td>Equiment</td>
		</tr>
		<tr >
			<td >Glucose</td>
			<td >Merck Life Science</td>
			<td >G8769</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >Glutamax</td>
			<td>GIBCO</td>
			<td>35050061</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >HEPES</td>
			<td >Merck Life Science</td>
			<td>H3537</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >L-Cystein&#160;</td>
			<td >Merck Life Science&#160;</td>
			<td>C6852-25g</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >MAP2</td>
			<td>Merck</td>
			<td>AB15452</td>
			<td>Antibody</td>
		</tr>
		<tr >
			<td >MEM&#160;</td>
			<td >Life Technologies</td>
			<td >21575022</td>
			<td>Medium</td>
		</tr>
		<tr >
			<td >MgCl</td>
			<td >Merck Life Science</td>
			<td >M8266</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >NaOH</td>
			<td >VWR International</td>
			<td>1,091,371,000</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >Neurobasal A</td>
			<td >Life Technologies</td>
			<td >10888022</td>
			<td>Medium</td>
		</tr>
		<tr >
			<td >N-SIM Super Resolution Microscope</td>
			<td>Nikon</td>
			<td>-</td>
			<td>Instrument</td>
		</tr>
		<tr >
			<td >Papain</td>
			<td >Merck Life Science&#160;</td>
			<td >P-3125</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >paraformaldehyde&#160;</td>
			<td>Thermo Fisher Scientific</td>
			<td>28908</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >Pen/Strep 10x</td>
			<td >Life Technologies</td>
			<td >15140122</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >phosphate-buffered saline&#160;</td>
			<td>Gibco</td>
			<td>10010023</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >Poly-L lysine</td>
			<td>Sigma</td>
			<td>P2636</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >ProLong Diamond Glass Antifade Mountant</td>
			<td>Thermo Fisher Scientific</td>
			<td>P36970</td>
			<td>Chemical</td>
		</tr>
		<tr >
			<td >PSD95&#160;</td>
			<td>NeuroMab</td>
			<td>K28/43</td>
			<td>Antibody</td>
		</tr>
		<tr >
			<td >Round coverglass</td>
			<td>Thermo</td>
			<td>12052712</td>
			<td>Equiment</td>
		</tr>
		<tr >
			<td >SUMO1</td>
			<td>Abcam</td>
			<td>ab32058</td>
			<td>Antibody</td>
		</tr>
		<tr >
			<td >Synaptophysin&#160;</td>
			<td>Merck</td>
			<td>S5768&#160;</td>
			<td>Antibody</td>
		</tr>
		<tr >
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			<td >Trypsin inhibitor&#160;</td>
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			<td>T9003-500MG</td>
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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.

We present here the standard workflow to study neuronal proteins co-localization. We first calibrated the microscope and next we performed SIM analysis of the samples. To calibrate the system, we used fluorescent microspheres of 0.1 &#956;m diameter. Upon obtaining super-resolved 3D-SIM images of the beads, the underlying image data are Fourier-transformed to re-convert them to a spatial frequency representation. In Figure 2A, the distinct flower pattern is presented as an indication of supe...

Watch this Scientific Journal Video about Super-Resolution Imaging to Study Co-Localization of Proteins and Synaptic Markers in Primary Neurons at JoVE.com

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

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. ...

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 is a patterned light illumination technique that doubles the spatial resolution of widefield microscopy, reaching a detail of around 100 nanometers. 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.The key to allowing the analysis at a super-resolution level is, however, the reagents used during acquisition, such as chambered coverslips and mounting solutions compatible with the diffraction index of the objective. To obtain a mouse hippocampal primary neuron, isolate hippocampi from P1 to P4 pups. Plate cells at 70, 000 cells per well in a volume of 200 microliters per well.Wait until hippocampal primary neurons are fully matured 12 to 14 days after plating to perform co-localization studies. Add 4%paraformaldehyde, PFA, in PBS 200 microliters per well to neurons to fix them quickly. Remove the solution and incubate the samples with 1%bovine serum albumin, BSA, in PBS at 200 microliters per well for one hour at room temperature to passively cover all free-binding surfaces of the plate with an irrelevant protein for the analysis.A BSA-based blocking buffer without Triton X-100 reduces the antibody background more efficiently than the same buffer with 0.2%Triton X-100. Add the primary antibody. As a negative control, do not add any primary antibody to one of the wells.Mount cells using a SIM-compatible mountant, e.g. ProLong Glass Antifade Mountant. Cover and protect the cells with a coverglass, for example, a round coverglass with a diameter of eight millimeters.Square ones can also be used. Store the chambered coverslips at room temperature and wait at least 48 hours before acquiring images. Diamond glass requires at least two days of curing before super-resolution acquisitions.Use two strategies to assure antibody specificity. The first one is the use of at least two different antibodies targeting the same substrate. The second strategy is antibody neutralization by incubation with the purified protein target, or the used to raise the antibody.We routinely use an N-SIM super-resolution microscope manufactured by Nikon for our analysis. The instructions that follow are to be intended for the acquisition of SIM images independently of the microscope of use. It is also important to maximize the performance of the microscope to perform an accurate calibration of the instruments, including correction ring, laser alignment, and grating block focusing.Before the acquisition of SIM images, the system requires a proper calibration with specific sub-resolution size fluorescent beads. An example are TetraSpeck microspheres. These beads are stained with different fluorescent dyes to allow the calibration of different lasers with one sample.In a water bath, sonicate around 1.8 times 10 to the 8th fluorescent microspheres for 10 minutes. Dilute the fluorescent microspheres one in 500 in double distilled water. Sonicate a second time for an additional 10 minutes.Pipette 15 microliters of the diluted beads into a well of a chambered coverslip. Let the solution dry for five minutes at room temperature. Add 10 microliters of the mounting solution, e.g.ProLong Glass, and place an eight millimeter coverslip on top. Wait at least 48 hours to allow a proper curing. Turn on the microscope and lasers.Let the system warm up to reach thermal equilibrium of all microscope components and the SIM super-resolution microscope system requires at least three hours. Select the 100X objective. Start the calibration by aligning the lasers to the center of the diffraction grating block.In the N-SIM system, a micrometer knob and a dedicated camera allow centering of the light beams to the target. Insert the chambered coverslip of the microscope for viewing. Set the system to the chambered coverslip thickness by adjusting the objective correction cover.NIS Software, the proprietary software provided with N-SIM super-resolution microscope systems, has an automatic function to regulate correction of colors. Adjust grating block focus for each channel to ensure focused structured pattern illumination on the sample. The software provides an automatic function for this task.Acquire raw 3D SIM images of the multicolor microspheres. Calculate for each separate wavelength the Fourier transform of the super-resolved image. If the transformed image fails to obtain a correct flower-like pattern, restart calibration since super-resolution has not been achieved.In the super-resolved image, select a single microsphere and calculate its intensity profile for each channel to measure the resolution achieved. It should now be close to 100 nanometers laterally. Perform channel registration by overlaying a multi-channel acquisition of the microspheres.The goal is to collimate all channel signals laterally and axially. This will eliminate chromatic aberrations due to the misalignment of the different channels and help the co-localization analysis. Confirm the quality of calibration by using the functions illumination phase step and illumination patterned focus of SIMcheck, a suite of plugins for the open source application ImageJ.Start analyzing the sample using a 40X objective in confocal or widefield mode. This allows navigation of the sample, maintaining good detail and a large field of view. Use MAP2 antibody signal to detect an area representing neuronal processes.Acquire images of the sample in confocal mode to determine the quality of the staining. Switch to the objective 100X. Apply oil to the 100X objective.Acquire a widefield or confocal image that will be used later to assess the quality of the super-resolved image. Switch to 3D SIM mode. Using dialogue windows to set the parameters for acquisition, select the highest bit depth setting available to maximize color information.Typically, 16 bit is the standard choice. Moreover, to improve signal-to-noise ratio, select a low-frequency value for acquisition, such as one megahertz. Using histogram windows, set laser power to obtain a linear response of signal to avoid loss of information, limit saturated pixels in the images.The N-SIM system uses an Andor iXon3 camera. When working at 16 bit, choose a target intensity of 16, 000 to ensure the linear response of the camera. Alternatively, choose a range between 30, 000 to 45, 000 to maximize the dynamic range of the acquisitions.Set the laser power between 0.1%and 50%when imaging the samples and the exposure times between 50 milliseconds and two seconds. Laser powers above 50%may cause rapid photobleaching of the fluorophores in use. Start acquiring the images in 3D SIM mode.Use SIMcheck, a suite of free plugins for ImageJ, to assess the quality of acquisitions of the raw images. If SIMcheck does not detect any artifacts or quality issues, acquire a minimum of 10 images from full technical replicates to allow statistical analysis. Acquired images are raw data that needs to be processed to obtain reconstructed SIM images that will be used for further analysis.In the protocol, we suggest to use synaptophysin and PSD-95 as pre and post synaptic markers. Additional markers can, however, be used. A huge body of literature supports the use of other proteins, such as drebrin and bassoon as synaptic markers.3D SIM acquired images are raw images that need to be processed to obtain reconstructed super-resolved images. Incorrect reconstruction of raw images can lead to artifacts that would affect the analysis of the samples. Great attention should therefore be paid to properly choosing reconstruction parameters.Process the raw images using the microscope reconstruction analysis software to obtain a super-resolved image. Alternatively, use a freely available ImageJ plugin fairSIM to reconstruct the raw images. Calculate the Fourier transform of the super-resolved images using the microscope reconstruction software or ImageJ plugin SIMcheck.A good reconstructed image should return, for each channel, a flower-like image. If the reconstructed images fail to recreate a flower-like shape, restart from the raw images and reconstruct them by modifying reconstruction parameters, such as linear filtering, apodization, and zero order suppression. In the NIS Software using the preview to monitor how changing the parameters affects the final resolved image, modify the parameter's illumination modulation contrast, high-resolution noise suppression, and out of focus suppression.Analyze the reconstructed image to unbiasedly detect artifacts by using NanoJ-SQUIRREL, an ImageJ-based plugin, to assess the quality of super-resolved images. In our analysis of co-localization, we used two approaches. The first one is a visual approach based on the profile analysis that shows single events of co-localization and that identifies the contribution of each channel.As a first step to study co-localization between synaptic markers and a protein of interest, take a super-resolved image and analyze a single locus to determine signal overlap. Identify a single locus on the super-resolved image. Obtain the intensity profiles of the fluorescent signals to the locus of interest.If profile analysis has suggested single locus co-localization, a more general analysis of the whole image can be carried out by calculating Pearson's and Mander's coefficients. Use JACoP, an ImageJ plugin, to determine two parameters of co-localization, Pearson's and Mander's. After having calibrated the system and assessed the quality of the reconstructed images, we next started analyzing the primary neuronal cultures stained with an antibody against MAP2, a neuronal marker, PSD-95, a postsynaptic marker, and our target protein SUMO1.We first analyzed the sample performing four-channel confocal microscopy with a 40X objective. Upon selecting an area representing neuronal processes, we switched to a 100X objective. We acquired both confocal and SIM images of the same area to assess quality of reconstruction with NanoJ-SQUIRREL and perform co-localization analysis.Elucidating the structure and composition of the synapse is crucial for the understanding of both physiological and pathological processes that regulate memory and cognition. While in normal state synapses are the building blocks of memory, they are also at the basis of complex neurological disorders, such as one of the most common forms of dementia Alzheimer's disease. The protocol described here allows the study of protein composition of synapses with a super-resolution microscopy technique called structured illumination microscopy.

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Research

JoVE Journal

Neuroscience

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

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 ...

Real-time Imaging of Axonal Transport of Quantum Dot-labeled BDNF in Primary Neurons

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are increasingly viewed as an early feature of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD) and Huntington&#8217;s disease (HD). As yet unclear is the mechanism(s) by which axonal injury is induced. We reported the development of a novel technique to produce biologically active, monobiotinylated BDNF (mBtBDNF) that can be used to trace axonal transport of BDNF. Quantum dot-labeled BDNF (QD-BDNF) was produced by conjugating quantum dot 655 to mBtBDNF. A microfluidic device was used to isolate axons from neuron cell bodies. Addition of QD-BDNF to the axonal compartment allowed live imaging of BDNF transport in axons. We demonstrated that QD-BDNF moved essentially exclusively retrogradely, with very few pauses, at a moving velocity of around 1.06 &#956;m/sec. This system can be used to investigate mechanisms of disrupted axonal function in AD or HD, as well as other degenerative disorders.

Axonal transport of BDNF, a neurotrophic factor, is critical for the survival and function of several neuronal populations. Some degenerative disorders are marked by disruption of axonal structure and function. We demonstrated the techniques used to examine live trafficking of QD-BDNF in microfluidic chambers using primary neurons.

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...