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 pups23.</li>
	<li>Place dissected hippocampi in 10 mL of Dissection Media (Table 1) and let them deposit at the bottom of the tube.</li>
	<li>Using a sterile pipette, carefully remove the Dissection Media, leaving the hippocampi undisturbed at the bottom of the tube.</li>
	<li>Add 10 mL of Media 1 (Table 1) to the hippocampi and incubate for 30 minutes at 37 &#176;C.</li>
	<li>Using a sterile pipette, carefully remove Media 1, leaving the hippocampi undisturbed at the bottom of the tube.</li>
	<li>Add 10 mL of Media 2 (Table 1) and leave the (capped) centrifuge tube under the hood horizontally for 45 minutes.</li>
	<li>Let the centrifuge tube stand vertically to allow the tissue to settle at the bottom of the tube.</li>
	<li>Using a sterile pipette, carefully remove Media 2, leaving the hippocampi undisturbed at the bottom of the centrifuge tube.</li>
	<li>Add 2 mL of Media 3 (Table 1).</li>
	<li>Using a p1000 pipette with a filtered tip, mechanically dissociate cells from the tissue.</li>
	<li>Transfer the supernatant, in which are located the isolated neurons, to a 15 mL centrifuge tube.</li>
	<li>Centrifuge the cell suspension for 2 min at 300 x g at room temperature (RT).</li>
	<li>After centrifugation, cells are located at the bottom of the centrifuge tube. Using a sterile pipette discard the supernatant.</li>
	<li>Resuspend cells in 1 mL of Media 4.</li>
	<li>Use a 70 &#181;m filter to eliminate undissociated cells.</li>
	<li>Count viable cells in a B&#252;rker chamber by adding 1 &#181;L of 0.4 % Trypan blue solution to 19 &#181;L of the cell suspension.</li>
	<li>Plate cells at 70,000 cells/well in a volume of 200 &#181;L per well.</li>
	<li>Allow the cells to attach for 2 h in a humidified incubator at 37 &#176;C and 5% CO2.</li>
	<li>Take out the chambered coverslips from the incubator and carefully replace the medium with 200 &#181;L of Culture Media.</li>
	<li>Leave the chambered coverslips in a humidified incubator at 37 &#176;C and 5% CO2.</li>
	<li>Replace one third of the medium with fresh culture media every 5-7 days.</li>
	<li>Wait until hippocampal primary neurons are fully matured (12-14 days after plating) to perform co-localization studies.</li>
</ol>
2. Immunofluorescence staining
<ol>
	<li>Take the chambered coverslips from the incubator.</li>
	<li>Remove the medium.</li>
	<li>Quickly wash the wells with 200 &#181;L of PBS.</li>
	<li>Add 4% paraformaldehyde (PFA) in PBS (200 &#181;L/well) to neurons to fix them quickly.</li>
	<li>Incubate the cells for 15 min at RT.</li>
	<li>Remove the PFA solution.</li>
	<li>Permeabilize the cells by adding PBS with 0.2% Triton X-100 (200 &#181;L/well).</li>
	<li>Incubate for 1 min at RT.</li>
	<li>Remove the solution and incubate the samples with 1% bovine serum albumin (BSA) in PBS (200 &#181;L/well) for 1 h at RT 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.</li>
	<li>Remove the solution.</li>
	<li>Add the primary antibody of choice diluted in a PBS solution containing 1% BSA and 0.2% Triton X-100 (120-200 &#181;L/well, depending on the antibody dilution and availability). Incubate the samples for 2 h.
	<ol>
		<li>As a negative control, do not add any primary antibody to one of the wells. Multiple antibodies of different species against different targets may be used at the same time. Use an antibody against MAP2 (a neuronal marker) raised in chicken, an antibody against either PSD95 or synaptophysin raised in mouse, and an antibody against a target protein raised in rabbit. This allows three-color SIM analyses.</li>
	</ol>
	</li>
	<li>Quickly wash the wells three times with PBS (200 &#181;L/well).</li>
	<li>Add secondary antibodies (dyLight and Alexa secondary antibodies can both be used) diluted in a PBS solution containing 1% BSA and 0.2% Triton X-100 (200 &#181;L/well). Incubate the samples for 1 h at RT.</li>
	<li>Quickly wash the wells three times with PBS (200 &#181;L/well).</li>
	<li>Add Hoechst dye at a concentration of 1 &#956;g/mL diluted in PBS (200 &#181;L/well) to stain nuclei. Incubate the samples for 10 minutes at RT.</li>
	<li>Quickly wash the wells twice with PBS.</li>
	<li>Mount cells using a SIM-compatible mountant. Use 10 &#181;L/well of ProLong Glass Antifade Mountant.</li>
	<li>Cover and protect the cells with a coverglass (e.g., a round coverglass with a diameter of 8 mm). Square ones can also be used.</li>
	<li>Store the chambered coverslips at RT and wait at least 48 h before acquiring images. Diamond Glass requires at least two days of curing before super-resolution acquisitions.</li>
</ol>
3. Antibody specificity control
NOTE: Use two strategies to assure antibody specificity. The first strategy is to use at least two different antibodies targeting the same substrate. The second strategy is antibody neutralization by incubation with the purified protein target or the epitope used to raise the antibody.
<ol>
	<li>Incubate the antibody of choice with five times excess of the recombinant target or epitope for 1 h at RT in 1% BSA in PBS.</li>
	<li>After the incubation, use the neutralized antibody at the usual concentration for staining as described above from 2.11.</li>
</ol>
4. Microscope calibration
NOTE: We routinely use an N-SIM Super-Resolution Microscope System manufactured by Nikon for the super-resolution studies. However, several other companies also offer super-resolution microscopes in their catalogues. Although specific indications for Nikon&#8217;s N-SIM system are described, the instructions that follow can be generalized to other systems. Before the acquisition of SIM images, the system requires a proper calibration with specific sub-resolution size fluorescent beads. An example is the TetraSpeck microspheres. These beads are stained with different fluorescent dyes to allow the calibration of different lasers with one sample.
<ol>
	<li>In a water bath sonicate around 1.8 x 108 fluorescent microspheres for 10 minutes. Nikon&#8217;s N-SIM system requires a sparsely populated multicolor beads sample for the calibration. This could differ for other systems that require a dense single layer of sub-resolution size fluorescent beads. Adjust the number of fluorescent particles accordingly.</li>
	<li>Dilute the fluorescent microspheres 1:500 in double distilled water.</li>
	<li>Sonicate a second time for an additional 10 minutes.</li>
	<li>Pipet 15 &#181;L of the diluted beads into a well of a chambered coverslip.</li>
	<li>Let the solution dry for 5 minutes at RT.</li>
	<li>Add 10 &#181;L of the mounting solution and place an 8 mm coverslip on top.</li>
	<li>Wait at least 48 hours to allow proper curing.</li>
	<li>Turn on the microscope and lasers.</li>
	<li>Let the system warm up to reach the thermal equilibrium of all microscope components. N-SIM Super-Resolution Microscope System requires at least 3 hours.</li>
	<li>Select the 100x objective.</li>
	<li>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.</li>
	<li>Insert the chambered coverslip in the microscope for viewing. Set the system to the chambered coverslip thickness by adjusting the objective correction collar. NIS software, the proprietary software provided with N-SIM Super-Resolution Microscope systems, has an automatic function to regulate correction collars.</li>
	<li>Adjust grating block focus for each channel to ensure focused structured pattern illumination on the sample. NIS software provides an automatic function for this task.</li>
	<li>Next, acquire raw 3D-SIM images of the multicolor microspheres. Reconstruct the raw images to obtain a super-resolved image using the microscope software or the open-source software platform for the analysis of biological images ImageJ24 and the plugin fairSIM25.</li>
	<li>Calculate, for each separated wavelength, the Fourier transform of the super-resolved image obtained in 4.14. If the transformed image fails to obtain a correct flower-like pattern, restart calibration from 4.11 since super-resolution has not been achieved.</li>
	<li>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 nm laterally.</li>
	<li>Next, perform channel registration by overlaying a multichannel 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.</li>
	<li>Confirm the quality of calibration by using the functions &#8220;Illumination Phase Steps&#8221; and &#8220;Illumination Pattern Focus&#8221; of SIMcheck26, a suite of plugins for the open-source application ImageJ. To this end, prepare a chambered coverslip to obtain a dense single layer of TetraSpeck microspheres and acquire a 3D-SIM image of the sample. Analyze the image in ImageJ and, if aberrations are detected, restart microscope calibration from step 4.11.</li>
</ol>
5. Acquisition
<ol>
	<li>Start analyzing the sample using a 40x objective in confocal or widefield mode. This allows navigation to the sample, maintaining good details and a large field of view.</li>
	<li>Use MAP2 antibody signal to identify an area representing neuronal processes.</li>
	<li>Acquire images of the sample in confocal mode to determine the quality of the staining. Poor confocal quality will reflect in poor SIM quality, therefore requiring the samples to be discarded.</li>
	<li>If the area and the quality of the images are satisfactory, switch the objective to 100x.</li>
	<li>Apply oil to the 100x objective.</li>
	<li>Acquire a widefield or confocal image that will be used later to assess the quality of the super-resolved image (Figure 1A,B).</li>
	<li>Switch to 3D-SIM mode.</li>
	<li>Using dialog windows to set 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 1 MHz.</li>
	<li>Using histogram windows, set lasers 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 30,000-45,000 to maximize the dynamic range of the acquisition.</li>
	<li>Set laser power between 0.1% and 50% when imaging the samples and exposure times between 50 ms and 2 s. Laser powers above 50% may cause rapid photobleaching of the fluorophores in use.</li>
	<li>Start acquiring the images in 3D-SIM mode.</li>
	<li>Use SIMcheck, a suite of free plugins for ImageJ, to assess the quality of acquisition of the raw images.</li>
	<li>If SIMCheck does not detect any artifacts or quality issues, acquire a minimum of 10 images from 4 technical replicates to allow statistical analysis.</li>
</ol>
6. Post-production: image reconstruction
NOTE: 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.
<ol>
	<li>Process the raw images using the microscope reconstruction analysis software to obtain a super-resolved image (Figure 1C). Alternatively, use the freely available ImageJ plugin fairSIM to reconstruct raw images.</li>
	<li>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 the reconstruction parameters such as Wiener filtering, apodization and zero-order suppression27. In NIS software, using the preview to monitor how changing the parameters affects the final resolved image, modify the parameters i) Illumination Modulation Contrast, ii) High Resolution Noise Suppression and iii) Out of Focus Suppression.</li>
	<li>Next, analyze the reconstructed image to unbiasedly detect artefacts by using NanoJ-SQUIRREL28, an ImageJ-based plugin to assess the quality of super-resolved images.</li>
	<li>If NanoJ-SQUIRREL detects artifacts, restart from the raw images and reconstruct them by modifying the reconstruction parameters such as Wiener filtering, apodization and zero-order suppression. In NIS software, using the preview to monitor how changing the parameters affects the final resolved image, modify the parameters Illumination Modulation Contrast, High Resolution Noise Suppression and Out of Focus Suppression.</li>
	<li>Use the super-resolved images to calculate the co-localization profile and/or Pearson&#8217;s and Mander&#8217;s coefficients.</li>
</ol>
7. Co-localization with profile analysis
NOTE: 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.
<ol>
	<li>Identify a single locus on the super-resolved image.</li>
	<li>Obtain the intensity profiles of the fluorescent signals of the locus of interest.</li>
	<li>Export the data.</li>
	<li>Use GraphPad Prism, or a similar analysis software, to normalize all signal peaks and obtain comparable signal intensities for each channel with the final goal of determining locus specific co-localization.</li>
</ol>
8. Quantification of Pearson&#8217;s and Mander&#8217;s coefficients
NOTE: If profile analysis has suggested single locus co-localization, a more general analysis of the whole image can be carried out by calculating Pearson&#8217;s and Mander&#8217;s coefficients29,30.
<ol>
	<li>Use JACoP31, an ImageJ plug-in, to determine the two parameters of co-localization: Pearson&#8217;s and Mander&#8217;s.</li>
</ol>
9. Statistical analysis
<ol>
	<li>Use GraphPad Prism, or a similar analysis and graphing software, to process data collected with JACoP.</li>
	<li>Use at least 40 SIM images for each condition analyzed to obtain graphs and for statistical relevance.</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guided+tour+into+subcellular+colocalization+analysis+in+light+microscopy.">A guided tour into subcellular colocalization analysis in light microscopy.</a> Journal of Microscopy. 224, 213-232 (2006).</li><li>Bae, J. R., Kim, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synapses+in+neurodegenerative+diseases.">Synapses in neurodegenerative diseases.</a> BMB Reports. 50 (5), 237-246 (2017).</li><li>Godin, A. G., Lounis, B., Cognet, L. <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+Approaches+for+Live+Cell+Imaging.">Super-resolution Microscopy Approaches for Live Cell Imaging.</a> Biophysical Journal. 107 (8), 1777-1784 (2014).</li><li>Dempsey, G. T., Vaughan, J. C., Chen, K. H., Bates, M., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+fluorophores+for+optimal+performance+in+localization-based+super-resolution+imaging.">Evaluation of fluorophores for optimal performance in localization-based super-resolution imaging.</a> Nature Methods. 8 (12), 1027-1036 (2011).</li><li>Karras, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Successful+optimization+of+reconstruction+parameters+in+structured+illumination+microscopy+-+A+practical+guide.">Successful optimization of reconstruction parameters in structured illumination microscopy - A practical guide.</a> Optics Communications. 436, 69-75 (2019).</li><li>Bereczki, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+markers+of+cognitive+decline+in+neurodegenerative+diseases:+a+proteomic+approach.">Synaptic markers of cognitive decline in neurodegenerative diseases: a proteomic approach.</a> Brain: A Journal of Neurology. 141 (2), 582-595 (2018).</li><li>Gilestro, G. F., Tononi, G., Cirelli, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Widespread+Changes+in+Synaptic+Markers+as+a+Function+of+Sleep+and+Wakefulness+in+Drosophila.">Widespread Changes in Synaptic Markers as a Function of Sleep and Wakefulness in Drosophila.</a> Science. 324 (5923), 109-112 (2009).</li><li>Adler, J., Parmryd, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantifying+colocalization+by+correlation:+the+Pearson+correlation+coefficient+is+superior+to+the+Mander's+overlap+coefficient.">Quantifying colocalization by correlation: the Pearson correlation coefficient is superior to the Mander's overlap coefficient.</a> Cytometry. Part A: The Journal of the International Society for Analytical Cytology. 77 (8), 733-742 (2010).</li></ol>

The authors have nothing to disclose.

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.

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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. ...

super-resolution-imaging-to-study-co-localization-proteins-synaptic

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JoVE Journal

Neuroscience

5.7K Views.  Istituto di Ricerche Farmacologiche Mario Negri IRCCS. This protocol shows how to employ super-resolution microscopy to study protein co-localization in primary neuronal cultures.

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

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 ...

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 ...

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 ...

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 ...

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 ...

Use of Primary Cultured Hippocampal Neurons to Study the Assembly of Axon Initial Segments

Neuronal axon initial segments (AIS) are sites of initiation of action potentials and have been extensively studied for their molecular structure, assembly and activity-dependent plasticity. Giant ankyrin-G, the master organizer of AIS, directly associates with membrane-spanning voltage gated sodium (VSVG) and potassium channels (KCNQ2/3), as well as 186 kDa neurofascin, a L1CAM cell adhesion molecule. Giant ankyrin-G also binds to and recruits cytoplasmic AIS molecules including beta-4-spectrin, and the microtubule-binding proteins, EB1/EB3 and Ndel1. Giant ankyrin-G is sufficient to rescue AIS formation in ankyrin-G deficient neurons. Ankyrin-G also includes a smaller 190 kDa isoform located at dendritic spines instead of the AIS, which is incapable of targeting to the AIS or rescuing the AIS in ankyrin-G-deficient neurons. Here, we described a protocol using cultured hippocampal neurons from ANK3-E22/23-flox mice, which, when transfected with Cre-BFP exhibit loss of all isoform of ankyrin-G and impair the formation of AIS. Combined a modified Banker glia/neuron co-culture system, we developed a method to transfect ankyrin-G null neurons with a 480 kDa ankyrin-G-GFP plasmid, which is sufficient to rescue the formation of AIS. We further employ a quantification method, developed by Salzer and colleagues to deal with variation in AIS distance from the neuronal cell bodies that occurs in hippocampal neuron cultures. This protocol allows quantitative studies of the de novo assembly and dynamic behavior of AIS.

Here, we described a protocol to quantitatively study the assembly and structure of the axon initial segments (AIS) of hippocampal neurons that lack pre-assembled AIS due to the absence of a giant ankyrin-G.

Neuronal axon initial segments (AIS) are sites of initiation of action potentials and have been extensively studied for their molecular structure, ...

Characterization of Neuromuscular Junctions in Mice by Combined Confocal and Super-Resolution Microscopy

Neuromuscular junctions (NMJs) are highly specialized synapses between lower motor neurons and skeletal muscle fibers that play an essential role in the transmission of molecules from the nervous system to voluntary muscles, leading to contraction. They are affected in many human diseases, including inherited neuromuscular disorders such as Duchenne muscular dystrophy (DMD), congenital myasthenic syndromes (CMS), spinal muscular atrophy (SMA), and amyotrophic lateral sclerosis (ALS). Therefore, monitoring the morphology of neuromuscular junctions and their alterations in disease mouse models represents a valuable tool for pathological studies and preclinical assessment of therapeutic approaches. Here, methods for labeling and analyzing the three-dimensional (3D) morphology of the pre- and postsynaptic parts of motor endplates from murine teased muscle fibers are described. The procedures to prepare samples and measure NMJ volume, area, tortuosity and axon terminal morphology/occupancy by confocal imaging, and the distance between postsynaptic junctional folds and acetylcholine receptor (AChR) stripe width by super-resolution stimulated emission depletion (STED) microscopy are detailed. Alterations in these NMJ parameters are illustrated in mutant mice affected by SMA and CMS.

This protocol describes a method for morphometric analysis of neuromuscular junctions by combined confocal and STED microscopy that is used to quantify pathological changes in mouse models of SMA and ColQ-related CMS.

Neuromuscular junctions (NMJs) are highly specialized synapses between lower motor neurons and skeletal muscle fibers that play an essential role in the ...