This work was financed by a VIDI grant number H64.09.016 from The Netherlands Organization for Scientific Research (NWO) to CPF. CPF is grateful to Dr. Silvina A. Fratantoni for her critical comments and corrections on the final manuscript. GMRDL/EMMM are supported by the Dutch Technology Foundation STW (project 12151 and 11350), which is part of the NWO, and which is partly funded by the Ministry of Economic Affairs. We thank the Catherine Kitts and Peter Drent of Nikon Instruments Europe BV for assistance and support. HX was supported by the Royal Dutch Academy of Arts and Sciences (grant 11CDP10) and WT was supported by grant the Netherlands Organization for Scientific research (grant 820.02.006).

All experimental procedures involving animals were optimized to reduce animal suffering and were approved by the Commission for Animal Experimentation, University of Amsterdam, DEC protocol # DED204 and DED250.
1. Coverslip Preparation
<ol>
	<li>Cut down the coverslips to a size of 15 mm x 15 mm using a carbide or diamond scribe, so that they fit into the wells of a 12-well plate.</li>
	<li>While working in a fume hood, start coverslip coating by fully submerging coverslips in concentrated nitric acid solution (70% wt/wt) in a glass container. To ensure even distribution shake, then incubate for a minimum of 4 hr. It is possible to reuse the concentrated nitric acid solution for approximately 2 times, but it can loose color by exposure to light.</li>
	<li>Remove the concentrated nitric acid solution and wash the coverslips four times in distilled water for 30 min, carefully shaking after each wash.</li>
	<li>With caution remove the water. 
	Note: the next three steps are performed in a sterile laminar flow cabinet.</li>
	<li>Soak the coverslips in 96% EtOH. Remove the coverslips from the EtOH solution and let them dry.</li>
	<li>Flame the coverslips and put them in a glass container. Use fine forceps to handle the coverslips (Dumont style no.5 forceps for example).</li>
	<li>Cover the container with aluminum foil and bake in a dry-heat oven for 12-16 hr at 180 &#176;C. Baked coverslips can be stored at room temperature for up to 1 month if covered tightly.</li>
</ol>
2. Coverslip Coating
The coverslip coating procedure favors neuron attachment to the glass surface and dendritic arborization 14.
<ol>
	<li>In a sterile laminar hood, use sterile small forceps to place the individual coverslips in single wells of a 12-well plate.</li>
	<li>Add 500 &#956;l of Poly-L-lysine solution (or sufficient amounts to submerge the coverslips).</li>
	<li>Wrap the plate in aluminum foil to prevent evaporation and leave it overnight at room temperature.</li>
	<li>Before starting the culture, aspirate the Poly-L-lysine solution carefully in a sterile laminar hood.</li>
	<li>Wash each well with 1 ml of sterile water twice, whilst preventing them to dry out.</li>
	<li>Aspirate water completely, add 1 ml of plating medium and leave the coverslips in the tissue culture incubator until ready to plate the cells. It is recommended to plate the cells within 24 hr.</li>
</ol>
3. Removal of Brains from E16-E19 Rat Embryos
<ol>
	<li>Sterilize the surgical instruments by heating them in a dry sterilizer overnight or washing them with 70% EtOH. Dry thoroughly if EtOH is used.</li>
	<li>Prepare several 30 mm dishes with 1x HBSS buffer and keep them on ice.</li>
	<li>Euthanize the rat dam with an intraperitoneal injection of Euthasol (160 mg/kg Euthasol in a volume of &#177; 0.4 ml).</li>
	<li>Check for the absence of reflexes.</li>
	<li>Spray the dam's abdomen with 70% EtOH.</li>
	<li>Make an incision along the abdomen and remove the uterus.</li>
	<li>Remove the embryos from the uterus and place them in a 100 mm diameter petri dish.</li>
	<li>Remove the heads of the embryos with large scissors and place the heads in a new petri dish containing cold 1x HBSS buffer. 
	Note: from here to step 7.1 the procedure is performed under sterile conditions in a laminar flow cabinet.</li>
	<li>Hold down the heads along the sides with big forceps.</li>
	<li>With small scissors, make a sagittal cut in the skin on top of the head, then laterally peel the skin down with a large forceps.</li>
	<li>Use the same approach as in step 3.10 to remove the skull. Make a sagittal cut starting at the caudal end; gently opening the skull without damaging the brain tissue. Fold the two halves of the skull away laterally exposing the brain.</li>
	<li>With the blunt spatula, scoop out the brain and place it in fresh cold 1x HBSS.</li>
</ol>
4. Dissection of the Hippocampi
It is very important that the dissection is done as quickly as possible in sterile conditions to ensure cell viability. Keep the samples cold on ice.
<ol>
	<li>Remove and discard the cerebellum with the fine scissors.</li>
	<li>Separate the two hemispheres of the brain by making a sagittal cut along the midline.</li>
	<li>Take each hemisphere and place both in a new 30 mm dish containing fresh cold 1x HBSS.</li>
	<li>Place each hemisphere such that the temporal lobe faces the bottom of the dish. 
	NOTE: From here on, it is recommended to use a dissecting microscope.</li>
	<li>Gently hold the midbrain using a small forceps and remove the midbrain with another pair of forceps. Leave the remainder of the hemisphere intact containing the cortex and the hippocampus.</li>
	<li>Turn over the tissue, so that the hippocampus is now facing the bottom of the dish.</li>
	<li>Gently hold the hemisphere in place with a fine forceps. By using another fine forceps, carefully and gently remove the meninges. It is easier to start at the olfactory bulb. Use caution so as not to damage the hippocampus.</li>
	<li>Orient the tissue so that the hippocampus is now facing up. The hippocampus can now be seen by its characteristic C-shaped structure.</li>
	<li>By using fine forceps, dissect out the hippocampus. Collect it in a new 30 mm dish containing fresh cold 1x HBSS.</li>
</ol>
5. Cell Dissociation and Plating
<ol>
	<li>Count the total number of hippocampi, then cut them into small pieces.</li>
	<li>Collect the pieces in a 15 ml centrifuge tube containing 3 ml 1x HBSS.</li>
	<li>Centrifuge at 300 x g for 5 min and carefully remove the supernatant.</li>
	<li>Add 6 &#956;l Trypsin per hippocampus.</li>
	<li>Incubate for max 20 min at 37 &#176;C. Swirl after 3 min.</li>
	<li>Wash two times with 5 ml of fresh cold 1x HBSS and discard the supernatant.</li>
	<li>After the second wash, add 1.5 ml of plating medium, pre-warmed to 37 &#176;C. The serum in the plating medium will inactivate Trypsin activity.</li>
	<li>Slowly triturate 30x with a fire-polished Pasteur pipette until all pieces of tissue are homogenously dispersed into single cells. Avoid any bubbling.</li>
	<li>Add 5 ml of plating medium pre-warmed to 37 &#176;C.</li>
	<li>Count cells using a Trypan blue vital staining.</li>
	<li>Seed 50,000 cells per well of a 12-well plate in 1 ml plating medium, prepared in Section 2 (total volume 2 ml).</li>
	<li>Gently rock the plate to evenly distribute the cells.</li>
	<li>Incubate at 37 &#176;C, 5% CO2. After 2-3 days, replace half of the plating medium (0.5 ml) with culture medium containing 10 &#181;M FUDR. 
	Note: Dendritic spine imaging is performed 16-17 days after plating (16-17 days in vitro, DIV).</li>
</ol>
6. Rat Hippocampal Primary Neuron Transfection using Lipofectamine
On DIV 14-15 neurons are transfected using the following protocol:
<ol>
	<li>Prepare plasmid DNA expressing GFP and incubation medium (10 ml of Neurobasal medium with 100 &#956;l of glutamax).</li>
	<li>Pre-warm the incubation medium to 37 &#176;C.</li>
	<li>Prepare DNA mix (Tube A). For each coverslip add 1 &#956;g of DNA in 100 &#956;l of plain Neurobasal medium. Gently mix.</li>
	<li>Prepare Lipofectamine mix (Tube B). For each coverslip add 2 &#956;l Lipofectamine in 100 &#956;l of plain Neurobasal medium. Gently mix.</li>
	<li>Add the Lipofectamine mix to the DNA mix dropwise.</li>
	<li>Incubate in the laminar flow hood for 30 min at room temperature.</li>
	<li>Add 1 ml of pre-warmed incubation medium to plate 1.</li>
	<li>Store plate 2 for 5 min at 37 &#176;C, 5% CO2.</li>
	<li>Gently add dropwise 200 &#956;l of the DNA/Lipofectamine mix to each well and incubate for 45 min at 37 &#176;C, 5% CO2.</li>
	<li>With the use of small forceps lift the coverslips containing the neurons and rinse them by dipping them in a 3 cm dish containing fresh warm Neurobasal medium and move them to plate 2.</li>
	<li>Incubate the transfected neurons at 37 &#176;C, 5% CO2 for 48 hr.</li>
	<li>Check for transfection efficiency 24 hr after transfection.</li>
</ol>
7. Immunostaining and Mounting of Rat Hippocampal Primary Neurons
To improve fluorescence intensity in transfected cells, perform an immunostaining protocol to enhance GFP detection 48 hr after transfection.
<ol>
	<li>Prepare 4% PFA and 0.05 M TBS.</li>
	<li>Warm the 4% PFA solution to 37 &#176;C.</li>
	<li>Gently aspirate the medium from the wells containing the coverslips.</li>
	<li>Add 500 &#956;l of warm 4% PFA carefully to prevent damaging the dendrites.</li>
	<li>Incubate at room temperature for 15 min.</li>
	<li>Wash with 1x HBSS 3 times for 5 min. 
	NOTE: at this point samples could be stored up to 3 weeks at 4 &#176;C or immunostaining can be started immediately.</li>
	<li>If samples were stored, wash with 0.05 M TBS 3 times for 5 min.</li>
	<li>Block with TBS-BSA (1%) solution at room temperature for 30 min.</li>
	<li>Wash with 0.05 M TBS 3 times for 5 min.</li>
	<li>Add the primary antibody diluted in incubation mix.</li>
	<li>Incubate the plates for 1 hr at room temperature.</li>
	<li>Further, Incubate overnight at 4 &#176;C with gentle shaking.</li>
	<li>Remove the primary antibody.</li>
	<li>Wash with 0.05 M TBS 3x for 5 min. 
	NOTE: from this point on, keep the coverslips protected from light.</li>
	<li>Add the secondary fluorescent-conjugated antibody diluted in incubation mix.</li>
	<li>Incubate at room temperature for 2 hr.</li>
	<li>Remove the secondary antibody.</li>
	<li>Wash with 0.05 M TBS 3 times for 5 min.</li>
	<li>Wash with 1X TB 2 times for 5 min.</li>
	<li>Use fine tweezers to remove to coverslips from the wells.</li>
	<li>Dry any excess of TB with a tissue and mount the coverslips using mounting medium.</li>
	<li>Seal with nail polish to prevent evaporation of mounting medium.</li>
</ol>
8. Dendritic Spine Imaging using Structure Illumination Microscopy
Dendritic spine imaging using the SIM system described in the materials has a lateral resolution (XY) value of approximately 85-110 nm and an axial (Z) resolution value between 200 - 250 nm, providing a factor of 2 times improvement in resolution compared to wide-field microscopy.
NOTE: Dendritic spine imaging using SIM is done typically 2 days after step 7.22, but could be done up to 3 weeks later if samples are kept in the dark and under a controlled temperature of 22 - 23 &#176;C.
<ol>
	<li>Turn on the 488 nm laser, the mercury lamp, the stage controller, the piezo controller, the halogen lamp for transmitted light and the PC and start up the SIM software in the "ANDOR for N-SIM" mode.</li>
	<li>Clean the 100x TIRF objective with 95% ethanol three times, and if necessary with petroleum ether.</li>
	<li>Use the following filter settings: 520 LP with a 488 dichroic.</li>
	<li>Put a drop of immersion oil on the objective. Check that there are no air-bubbles in the oil-drop. Move the objective upwards until the oil touches the sample. 
	Note: Place a cover over the stage to protect the sample from ambient light and dim the lights in the room as much as possible.</li>
	<li>Set the correction collar of the objective to 37 &#176;C, 200 &#181;m, to obtain the best symmetry of PSF. In order to set the correct collar position, first position the objective ring at the optimal nominal position and then check a 100 nm bead sample. According to the best PSF's symmetry, slightly change the collar position around the nominal one.</li>
	<li>For illumination, use 3D-SIM grating (3D 1layer 100X/1.49 all wavelengths). To begin the grating alignment place the selected grating block into the SIM illuminator, with the 100X 1.49 objective in place. Use a 100 nm bead sample mounted in media, with a concentration that can allow isolating 10-15 beads for a field of view (FOV). After setting the objective correction collar to the desired position, select the 3D-SIM illumination and start the software-guided alignment procedure. It will run (5 phases) x (1 direction) x (100 z-planes) images, from which it will reconstruct the FOV with beads. After selecting a single bead via an appropriate ROI, covering the entire bead including the out-of-focus blurred light, the software will start an automatic PSF fitting and it will adjust the grating position according to the result.</li>
	<li>To check the performance of the microscope, repeat the grating alignment every 2 weeks, because of the possible misalignment caused by table movements and/or temperature drifting. Furthermore, also check the laser intensity and stability according to the manufacturer's suggestions.</li>
	<li>Clean the sample surface with 95% ethanol three times. 
	Note: For the next steps see figure 3 for an overview of the control panels and the correct settings within the SIM software.</li>
	<li>In the SIM software, select the Optical Configuration Eye FITC and select an empty filter block in Turret 1 for visual inspection with white light.</li>
	<li>Set the focusing speed and the travel speed of the XY table to "Fine".</li>
	<li>Open the shutter and quickly focus on the sample.</li>
	<li>Close the shutter again and set the Z-coordinate to zero.</li>
	<li>Select the green filter in Turret 1 for visual inspection using the green channel.</li>
	<li>Set the intensity of the mercury lamp to the lowest setting.</li>
	<li>Move to the border of the sample carefully, making sure that the objective does not touch the sealant.</li>
	<li>Open the shutter and quickly scan through the sample with the lowest possible intensity.</li>
	<li>Upon encountering a sufficiently bright dendrite of interest and centering a segment of interest in the field of view, close the shutter.</li>
	<li>Set the software to Optical Configuration 3D-SIM 488 and the camera settings to read-out mode EM, gain 1 MHz 16-bit, exposure time 100 msec, laser power 5% and EM gain 200. 
	NOTE: Check that the green filter in Turret 2 is selected and that Turret 1 is empty.</li>
	<li>Check that the grating is set to &#8216;Moving' and click Live to view the sample with laser light and through the camera.</li>
	<li>Activate the Look Up Table.</li>
	<li>Center the object of interest if necessary and focus with the focusing speed set to Extra Fine. 
	NOTE: in the read-out mode EM gain 1 MHz 16-bit, the target intensity for a good SIM image is between 30,000-45,000.</li>
	<li>Quickly adjust the camera settings to get an intensity value between 30,000-45,000 in the read-out mode EM gain 1 MHz 16-bit. Initially use:
	<ol>
		<li>Laser power: 0% - 20% (with samples prepared as described above, 5% or 2.6 mW is sufficient)</li>
		<li>Exposure time: 50 msec-2 sec</li>
		<li>Read-out mode: EM gain 1 MHz 16-bit</li>
		<li>EM gain: 0-300</li>
		<li>Conversion gain: 1x - 5.1x</li>
		<li>Format for Live: No binning</li>
		<li>Format for Capture: No binning 
		NOTE: with these settings 6.3% &#177; 1.3% bleaching is routinely achieved, within a 10% limit of acceptable maximum bleaching, which could significantly affect the image quality 15.</li>
	</ol>
	</li>
	<li>Click Stop to turn off Live view.</li>
	<li>Configure the settings of the 3D Z stack in the ND Sequence panel to:
	<ol>
		<li>Range: 2 &#181;m</li>
		<li>Set size: 120 nm</li>
		<li>Z plane</li>
		<li>Click Home position</li>
		<li>Select the Optical Configuration 3D-SIM 488 in the Lambda section</li>
	</ol>
	</li>
	<li>Select 3D-SIM as the acquisition mode in the N-SIM pad.</li>
	<li>Run the ND Sequence acquisition and save the raw data.</li>
	<li>Select the Optical Configuration Eye FITC again and repeat steps 8.15-8.27 until the entire sample has been imaged</li>
	<li>3D Image reconstruction: The data acquired in step 8.23 can either be reconstructed right away or later on. Start by reconstructing the Z stack with the default reconstruction settings in Reconstruct Slice or Reconstruct Stack mode and adjust if necessary. 
	NOTE: for best results, Z stacks should be made with the indicated step size and reconstructed in Reconstruct Stack mode. Always check the validity of the reconstructed image by comparing it to the raw data or, preferably, a wide-field image. The parameters that can be adjusted for the reconstruction process are the contrast and high frequency noise suppression. Both of them influence how different raw image properties are taken into account during the reconstruction process. In case of low modulation depth raw data, the contrast parameter plays an important role. If the user has datasets with a low signal-to-noise ratio, then the high frequency noise suppression parameter can influence the reconstruction quality severely.</li>
	<li>When imaging in finished, center the XY stage and move the objective all the way down to its resting position.</li>
	<li>Unmount the sample and clean both the sample and objective with 95% ethanol.</li>
	<li>Shut down the software and the PC and switch off all other devices.</li>
	<li>3d reconstruction and spine classification of the acquired images can be carried out after converting the files to TIFF as described before using NeuronStudio software (see Figure 4) 16.</li>
	<li>Use the following parameters for reconstruction in the NeuronStudios software:
	<ol>
		<li>Volume: Voxel dimensions: X: 0.03 &#956;m; Y: 0.03 &#956;m; Z: 0.120 &#956;m.</li>
		<li>Dendrite detection: Attach ratio: 1.3; Minimum length: 5 &#956;m; Discretization Ratio: 1; Realign junctions: Yes.</li>
		<li>Spine detection: Minimum height: 0.2 &#956;m; Maximum height: 5.001 &#956;m; Maximum width: 3 &#956;m; Minimum stubby size: 10 voxels; Minimum non-stubby size: 5 voxels.</li>
		<li>Spine Classifier: Neck ratio (head-neck ratio): 1.1; Thin ratio: 2.5; Mushroom size: 0.35 &#956;m;</li>
	</ol>
	</li>
	<li>NeuronStudio parameters used for rendering: Neurite vertex shape: solid eclipse; Neurite vertex color: by type; Neurite edge shape: line; Neurite edge color: single color; Spine shape: solid eclipse; Spine color: by type. 
	Note: After 3D reconstruction is it also possible to set the volume render. The default threshold was set to 20 with the 'Regenerate volume rendering' option active. Opacity was set by 'Automatic Intensity' checked. 'Surface only' and 'Use Point Pre-Rendering' options were also active</li>
</ol>

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The authors have nothing to disclose.

In this article a working protocol to image dendritic spines from rat primary hippocampal neurons cultured in vitro using SIM is described. The primary hippocampal neuron culture method is an adaptation of the original method described by Kaech and Banker 18. The main differences are the use of Neurobasal/B27 culture medium, which eliminates the requirement of astroglial feeder cultures, and the addition of the mitotic inhibitor FUDR on day 3 which promotes neuronal survival while suppressing glial pr...

A dendritic spine is a small protrusion of the neuron membrane. This characteristic structure is specialized to typically receive input from a single synapse and represents the physical contact area between two neurons. Most functionally mature dendritic spines consist of a globular tip, termed head, and a thin neck that connects the head to the dendritic shaft. However, spines are not static and actively move and change their morphology continuously even in the adult brain 2. Within a 2 week period of time, rat primary hippocampal neuron cultures derived from late embryonic or early postnatal time develop complex dendritic arbors with numerous membrane protrusions that evolve from early filipodia to spine-like structures 3. Based on this dynamic behavior and other characteristics, dendritic spines are thought to provide an anatomical substrate for memory storage and synaptic transmission 4,5.
Given the critical role that dendritic spine size and shape have in synaptic function, it is important to measure their dimensions accurately. Spines vary from around 200 to 2,000 nanometers in length and can be readily identified by confocal laser-scanning fluorescence microscopy. However, spine dimensions other than length are usually below the conventional optical systems' resolution, theoretically fixed by diffraction around 200 nanometers 6. These resolving powers are insufficient for imaging finer details, such as the width of spine necks and heads. Much work has been dedicated to solve this problem and many relatively new super-resolution microscopy techniques have provided substantial progress. In particular, it is possible to achieve resolution beyond the classical limit without discarding any emission light by using laterally structured illumination microscopy (SIM) in a wide-field, non-confocal microscope 7-10. Using this technique in combination with non-linear microscopy techniques, it is theoretically possible to improve the lateral resolution of the optical microscope by an unlimited factor 11. However, in most experimental circumstances, SIM allows to surpass the resolution limit by a factor of two 1. Other super-resolution optical microscopy techniques such as Stimulated emission depletion (STED) microscopy 12 and photo-activation localization microscopy (PALM) 12 have been applied to imaging of dendritic spines. Localization-based methods such as PALM require very large numbers of raw images to achieve super-resolution and are therefore limited in speed. On the other hand, STED can achieve high imaging speed, although at relatively low photon counts and small fields of view, which may not be the case for SIM 13.
In this article the aim is to provide a working protocol to image dendritic spines from rat primary hippocampal neurons cultured in vitro using SIM. The protocol consists of two distinguishable phases: an initial one consisting of establishment, development, transfection and immunohistochemistry of rat primary hippocampal neuron cultures and a late phase dedicated to sample imaging.

<table>
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			<td>Fine forceps</td>
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			<td>Big forceps</td>
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			<td>Fine scissor</td>
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			<td>Big scissor</td>
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			<td>Blunt spatula</td>
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			<td>Dissecting microscope with illumination</td>
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			<td>Light microscope</td>
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			<td>37 &#176;C water bath</td>
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			<td>Laminar flow cell culture hood</td>
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			<td>High-temperature dry-oven</td>
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			<td>Bunsen burner</td>
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			<td>Cell culture incubator (5% CO2, 37 &#176;C)</td>
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			<td>Microcentrifuge</td>
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			<td>Orbital shaker</td>
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			<td>Nikon structured illumination microscope setup consisting of:</td>
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			<td>Nikon Eclipse Ti research inverted microscope with Perfect Focus System</td>
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			<td>Nikon CFI Apo TIRF 100x oil objective lens (N.A. 1.49)</td>
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			<td>4 Coherent Sapphire Lasers (458, 488, 514 and 561 nm exitation wavelength)</td>
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			<td>SIM Illuminator</td>
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			<td>Nikon Stage Controller</td>
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			<td>MCL Nano-Drive piezo controller</td>
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			<td>Nikon Intensilight C-HGFIE mercury lamp</td>
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			<td>SIM Microscope Enclosure temperature control</td>
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			<td>Andor EM-CCD Camera iXon DU897</td>
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			<td>PC with Microsoft Windows 7 Home Edition</td>
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			<td>Nikon&#8217;s NiS Elements 6.14 SIM software package</td>
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			<td>Nikon type A immersion oil</td>
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	</tbody>
</table>

imaging dendritic spines of rat primary hippocampal neurons using structured illumination microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

Described here is a standardized working protocol for imaging dendritic spines from rat primary hippocampal neurons in vitro using SIM. The protocol workflow and its crucial steps are shown in Figure 1. Overall, the protocol takes approximately 2 weeks of experimental work separated in a first phase of sample preparation, including culture, development and transfection of rat primary hippocampal neurons and immunohistochemistry, and second phase of sample imaging using SIM. The rat primary hippo...

Watch this Scientific Journal Video about Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy at JoVE.com

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

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15.7K Views.  University of Amsterdam. This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Video: Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

Generation of Neurospheres from Mixed Primary Hippocampal and Cortical Neurons Isolated from E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have a robust in vitro model that can be exploited for various neurobiology studies. Though primary neuron cultures (i.e., long-term hippocampal cultures) have provided scientists with models, it does not yet represent the complexity of brain network completely. In the wake of these limitations, a new model has emerged using neurospheres, which bears a closer resemblance to the brain tissue. The present protocol describes the plating of high and low densities of mixed cortical and hippocampal neurons isolated from the embryo of embryonic day 14-16 Sprague Dawley rats. This allows for the generation of neurospheres and long-term primary neuron culture as two independent platforms to conduct further studies. This process is extremely simple and cost-effective, as it minimizes several steps and reagents previously deemed essential for neuron culture. This is a robust protocol with minimal requirements that can be performed with achievable results and further used for a diversity of studies related to neuroscience.

Presented here is a protocol for the spontaneous generation of neurospheres enriched in neural progenitor cells from high density plated neurons. During the same experiment, when neurons are plated at a lower density, the protocol also results in prolonged primary rat neuron cultures.

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have ...

3D Modeling of Dendritic Spines with Synaptic Plasticity

Computational modeling of diffusion and reaction of chemical species in a three-dimensional (3D) geometry is a fundamental method to understand the mechanisms of synaptic plasticity in dendritic spines. In this protocol, the detailed 3D structure of the dendrites and dendritic spines is modeled with meshes on the software Blender with CellBlender. The synaptic and extrasynaptic regions are defined on the mesh. Next, the synaptic receptor and synaptic anchor molecules are defined with their diffusion constants. Finally, the chemical reactions between synaptic receptors and synaptic anchors are included and the computational model is solved numerically with the software MCell. This method describes the spatiotemporal path of every single molecule in a 3D geometrical structure. Thus, it is very useful to study the trafficking of synaptic receptors in and out of the dendritic spines during the occurrence of synaptic plasticity. A limitation of this method is that the high number of molecules slows the speed of the simulations. Modeling of dendritic spines with this method allows the study of homosynaptic potentiation and depression within single spines and heterosynaptic plasticity between neighbor dendritic spines.

The protocol develops a three-dimensional (3D) model of a dendritic segment with dendritic spines for modeling synaptic plasticity. The constructed mesh can be used for computational modeling of AMPA receptor trafficking in the long-term synaptic plasticity using the software program Blender with CellBlender and MCell.

Computational modeling of diffusion and reaction of chemical species in a three-dimensional (3D) geometry is a fundamental method to understand the ...

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic spines have been described for DD GABAergic neurons as the input sites from presynaptic cholinergic neurons in the motor circuit of Caenorhabditis elegans. This synaptic circuit can now serve as a powerful new in vivo model of spine morphogenesis and function that exploits the facile genetics and ready accessibility of C. elegans to live-cell imaging. 
This protocol describes experimental strategies for assessing DD spine structure and function. In this approach, a super-resolution imaging strategy is used to visualize the intricate shapes of actin-rich dendritic spines. To evaluate the DD spine function, the light-activated opsin, Chrimson, stimulates the presynaptic cholinergic neurons, and the calcium indicator, GCaMP, reports the evoked calcium transients in postsynaptic DD spines. Together, these methods comprise powerful approaches for identifying genetic determinants of dendritic spines in C. elegans that could also direct spine morphogenesis and function in the brain.

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are important cellular features of the nervous system. Here live imaging methods are described for assessing the structure and function of dendritic spines in C. elegans.&#160;These approaches support the development of mutant screens for genes that define dendritic spine shape or function.

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic ...

Serial Block-Face Scanning Electron Microscopy (SBEM) for the Study of Dendritic Spines

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In addition, some features of dendritic spines, such as volume of the spine and post-synaptic density (PSD) (representing post-synaptic part of the synapse), presence of presynaptic terminal, and smooth endoplasmic reticulum or atypical form of PSD (e.g., multi-innervated spines), can be observed only with 3D EM. By employing serial block-face scanning electron microscopy (SBEM) it is possible to obtain 3D EM data easier and in a more reproducible manner than when performing traditional serial sectioning. Here we show how to prepare mouse hippocampal samples for SBEM analysis and how this protocol can be combined with immunofluorescence study of dendritic spines. Mild fixation perfusion allows us to perform immunofluorescence studies with light microscopy on one half of the brain, while the other half was prepared for SBEM. This approach reduces the number of animals to be used for the study.

Serial Block-Face Scanning Electron Microscopy SBEM for the Study of Dendritic Spines

Serial Block-Face Scanning Electron Microscopy (SBEM) is applied to image and analyze dendritic spines in the murine hippocampus.

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In ...