We thank Dr. Gary Banker for suggestion on neuronal culture protocol. This work is supported by the Howard Hughes Medical Institute, a grant from NIH, and a George Barth Geller endowed professorship (V.B.).

NOTE: This culture method of hippocampal neurons from postnatal 0-day ANK3-E22/23f/f mice is adapted from Gary Banker&#8217;s glia/neuron co-culture system. Therefore, it is critical to perform all steps after dissection in a clean hood using sterilized tools. This protocol takes up to 1 month. The workflow is displayed in Figure 3. The protocol follows the animal guidelines of Duke University.
1. Preparing of coverslips and neuronal plating dish...

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The assembly of AIS is organized by 480 kDa ankyrin-G. However, ankyrin-G has shorter isoforms that can target to the AIS of wildtype neurons, which may lead to difficulty in interpretation of structure-function analyses of AIS assembly. Here we present a method using neurons from ANK3-E22/23-flox mice that allows study of de novo assembly of the AIS. By transfecting with Cre-BFP at 3 div, we eliminate the all endogenous isoforms of ankyrin-G. We could also co-transfect 480 kDa ankyrin-G to rescue the f...

The axon initial segment is located at the proximal axon in most vertebrate neurons. Functionally, AIS is where action potentials are initiated due to the high-density of voltage-gated sodium channels in this region. AIS of some excitatory neurons are also targeted by inhibitory interneurons through forming GABAergic synapses1,2,3. Therefore, AIS is a critical site to integrate cell signaling and modulate the excitability of neurons. AIS is normally 20-60 &#956;m in length and located within 20 &#956;m of the cell body. The length and position of AIS varies in neurons across brain regions, as well as in different developmental stages of the same neuron4,5. Accumulated evidence suggested that the composition and position of AIS are dynamic in responding to the change of neuronal activity4,5,6,7.
480 kDa ankyrin-G is the master organizer of AIS. 480 kDa ankyrin-G is a membrane associated adaptor protein that directly binds to voltage gated sodium channels as well as other major AIS proteins including beta4-spectrin, KCNQ2/3 channels that modulate sodium channel activity8,9, and 186 kDa neurofascin, a L1CAM that directs GABAergic synapses to the AIS2,10. 480 kDa ankyrin-G shares canonical ankyrin domains found in the short 190 kDa ankyrin-G isoform (ANK repeats, spectrin binding domain, regulatory domain), but are distinguished by a giant exon that is found only in vertebrates and is specifically expressed in neurons (Figure 1A)11,12. The 480 kDa ankyrin-G neuron specific domain (NSD) is required for AIS formation12. The 190 kDa ankyrin-G does not promote AIS assembly or target AIS in ankyrin-G-null neurons12. However, 190 kDa ankyrin-G is concentrated at the AIS containing 480 kDa ankyrin-G12. This ability of the 190 kDa ankyrin-G to target pre-assembled AIS of wildtype neurons has been a source of confusion in the literature and has slowed appreciation of the critical specialized functions of the 480 kDa ankyrin-G in AIS assembly. Therefore, it is critical to study AIS assembly in ankyrin-G-null neurons that lack a pre-assembled AIS.
Here, we present a method to study the assembly and structure of the AIS using cultured hippocampal neurons from ANK3-E22/23-flox mice that eliminates all isoforms of ankyrin-G13 (Figure 1B). By transfecting neurons with a Cre-BFP construct before AIS is assembled, we generated ankyrin-G-deficient neurons completely lacking an AIS (Figure 1B, Figure 2). The assembly of AIS is fully rescued following co-transfection of 480 kDa ankyrin-G-GFP plasmid with a Cre-BFP plasmid. This method provides a way to study the AIS assembly in a non-pre-assembled AIS environment. We also modified the glia-neuron co-culture system from Gary Banker without using antibiotics, previously designed for embryonic day 18 neurons, for application to postnatal mouse neurons and adapted a AIS quantitation method to average AIS measurements from multiple neurons to normalize the variation of AIS14,15.

<table>
	<tbody>
		<tr>
			<td>10xHBSS</td>
			<td>Thermo Fisher Scientific</td>
			<td>14065-056</td>
			<td></td>
		</tr>
		<tr>
			<td>18mm coverglass (1.5D)</td>
			<td>Fisher Scientific</td>
			<td>12-545-84-1D</td>
			<td></td>
		</tr>
		<tr>
			<td>190kDa ankyrin-G-GFP</td>
			<td>Addgene</td>
			<td>#31059</td>
			<td></td>
		</tr>
		<tr>
			<td>2.5% Tripsin without phenol red</td>
			<td>Thermo Fisher Scientific</td>
			<td>14065-056</td>
			<td></td>
		</tr>
		<tr>
			<td>480kDa ankyrin-G-GFP</td>
			<td>lab made</td>
			<td>Provide upon request</td>
			<td></td>
		</tr>
		<tr>
			<td>ANK3-E22/23f/f mice</td>
			<td>JAX</td>
			<td>Stock No: 029797</td>
			<td>B6.129-Ank3tm2.1Bnt/J;</td>
		</tr>
		<tr>
			<td>B27 serum-free supplement</td>
			<td>Thermo Fisher Scientific</td>
			<td>A3582801</td>
			<td></td>
		</tr>
		<tr>
			<td>Boric acid</td>
			<td>Sigma-Aldrich</td>
			<td>B6768</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer with 70-mm mesh</td>
			<td>BD Biosciences</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>Ceramic coverslip-staining rack</td>
			<td>Thomas Scientific</td>
			<td>8542E40</td>
			<td></td>
		</tr>
		<tr>
			<td>Cre-BFP</td>
			<td>Addgene</td>
			<td>#128174</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G7021</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Thermo Fisher Scientific</td>
			<td>11995073</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutaMAX-I supplement</td>
			<td>Thermo Fisher Scientific</td>
			<td>A1286001</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>Thermo Fisher Scientific</td>
			<td>11668030</td>
			<td></td>
		</tr>
		<tr>
			<td>MEM with Earle&#8217;s salts and L-glutamine</td>
			<td>Thermo Fisher Scientific</td>
			<td>11095-080</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurobasal Medium</td>
			<td>Thermo Fisher Scientific</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>Nitric acid 70%</td>
			<td>Sigma-Aldrich</td>
			<td>225711</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM I Reduced Serum Medium</td>
			<td>Thermo Fisher Scientific</td>
			<td>31985062</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>P6148</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-streptomycin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-lysine hydrochloride</td>
			<td>Sigma-Aldrich</td>
			<td>26124-78-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>1310-58-3</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

A complete set of experiment should include Cre-BFP only transfection as negative control, Cre-BFP plus 480 kDa ankyrin-G co-transfection as positive control and a non-transfected condition as technique control. In Cre-BFP only control, transfected neurons lack the accumulation of AIS markers, including ankyrin-G (ankG), beta4-spectrin (&#946;4), neurofascin (Nf) and voltage gated sodium channels (VSVG) (Figure 4A)16. In contrast, Cre and 480 kDa ankyrin-G co-transfec...

Watch this Scientific Journal Video about Use of Primary Cultured Hippocampal Neurons to Study the Assembly of Axon Initial Segments at JoVE.com

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

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, ...

This protocol is used to quantitatively study the assembly and structure of the axon initial segments of Hippocampal Neurons that lack pre-assembled AIS due to the absence of a giant ankyrin-G. To begin, dissect six to eight hippocampi from postnatal, one day old pups with HBSS medium in a Petri dish. Chop the hippocampi with dissection scissors into smaller pieces, and transfer them from the dish to a 15 milliliter tube.Wash the hippocampi twice with five milliliters of HBSS, and leave them in 4.5 milliliters of HBSS after the wash. Add 0.5 milliliters of 2.5%trypsin into 4.5 milliliters of HBSS, and incubate in a 37 degree Celsius water bath for 15 minutes, inverting the tube every five minutes. Well digested hippocampi should become sticky and form a cluster.If needed, extend the digestion for five more minutes. Wash hippocampi with HBSS three times for five minutes per wash. After the wash, add two milliliters of HBSS and pipette the hippocampi up and down with a Pasteur pipette 15 times.Triturate the tissue with a fire polished Pasteur pipette 10 times, and rest the tube for five minutes until all chunks set to the bottom. Repeat the trituration with the remaining chunks until most of them have disappeared. Then, use a one milliliter pipette tip to gently transfer the supernatant containing the dissociated neurons to plating dishes.Add it directly to the pre incubated plating medium and shake the plate gently. Two to four hours after seeding, check the plating dishes with a light microscope. The majority of the neurons should have attached to the cover slip.Using a fine tip forceps, flip the cover slips to the glial cell feeder dishes containing preconditioned neuronal culture medium with the wax dots side facing downwards. Neurons can grow in the glial cell feeder dishes for up to one month. Feed neurons every seven days with one milliliter of fresh neuronal culture medium.On the third day in vitro, flip the cover slips with the wax dot side facing up into a glial cell feeder dish with conditioned neuronal culture medium. Mix 0.25 micrograms of CRE BFP DNA with 0.5 micrograms of ankyrin-G GFP DNA in a 1.7 milliliter tube to transfect four cover slips. Add 100 microliters optimum.Then mix the tube and rested on a rack. If only CRE BFP is transfected, the GFP plasmic backbone is used to match the total amount of DNA. Mix three microliters of transfection reagent with 100 microliters of optimum in a new 1.7 milliliter tube and incubate the tube for five minutes at room temperature.Then, mix 100 microliters of the previously mixed DNA with 100 microliters of transfection reagent mix, and leave it for five to 10 minutes on a rack. Add 50 microliters of the DNA mix on top of each cover slip by inserting the tip just below the medium without touching the cover slips. Pipette slowly to avoid spreading the DNA mix.Slowly bring the dish back to the incubator and leave it there for 30 to 45 minutes. Flip the cover slips back to the home glial feeder dish with the wax dot side facing down and put the dish back in the incubator. For AIS quantification, open the Z series images in Fiji and generate a maximum projection of images.After subtracting the empty cover slip background signal from the image, draw a line along the AIS, making sure that the width of the line fully covers the AIS. Start the line before the AIS signal is raised above the background and stop after it drops to the background. Measure the mean pixel intensity along the line, and export it to a spreadsheet.Then, run a MATLAB script to generate the final figure. The experiment should include Cre-BFP only transfection as negative control, Cre-BFP plus 480 kilodalton ankyrin-G cotransfection as positive control, and a non-transfected condition as technique control. In the Cre-BFP only control, transfected neurons lack the accumulation of AIS markers, including ankyrin-G, beta 4 spectrin, neurofascin, and voltage gated sodium channels.In contrast, Cre and 480 kilodalton ankyrin-G co-transfected neurons have fully assembled AIS, which is demonstrated by the presence of AIS markers. It is important to confirm the quality of culture via comparison with the non-transfected dishes. Unhealthy neurons tend to show abnormal AIS structure, like discontinued or ectopic AIS.To evaluate how an ankyrin-G human neurodevelopmental disorder mutation affects AIS assembly, the average AIS intensity was plotted from the soma to the distal axon. AIS enriched proteins normally show a fast increase of signal from the proximal axon and a slow decrease of signal to the distal axon. When aligned with the non-transfected AIS, the mutant curve is wider and the peak of the curve is lower, suggesting a structure change of AIS.The wild type ankyrin-G assembled AIS closely aligned with the non-transfected one.

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4.9K Views.  Xi'an Jiaotong University. This protocol is used to quantitatively study the assembly and structure of the axon initial segments of Hippocampal Neurons that lack pre-assembled AIS due to the absence of a giant ankyrin-G. To begin, dissect six to eight hippocampi from postnatal, one day old pups with HBSS medium in a Petri dish. Chop the hippocampi with dissection scissors into smaller pieces, and transfer them from the dish to a 15 milliliter tube.Wash the hippocampi twice with five milliliters of HBSS, and leav...

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

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...

a technique to culture mouse hippocampal neurons

The video demonstrates a method for culturing neurons from murine hippocampi. Initially, the hippocampi undergo enzymatic digestion to break down the extracellular tissue matrix. Subsequently, the digested tissue is mechanically disrupted to isolate individual neurons. Finally, these neurons are cultured on glial feeder...

A Technique to Culture Mouse Hippocampal Neurons

live imaging of dense-core vesicles in primary cultured hippocampal neurons

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking.  Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and...

genetic study of axon regeneration with cultured adult dorsal root ganglion neurons

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein...

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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...

measuring synaptic vesicle endocytosis in cultured hippocampal neurons

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

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

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Primary Culture of Hippocampal Neurons from P0 Newborn Rats

The dissection and growth of cells from an individual brain area facilitates investigation of cellular and physiological parameters. We describe a method for primary cell culturing that produces neuron-enriched cultures in a serum-free...

stripe assay to study the attractive or repulsive activity of a protein substrate using dissociated hippocampal neurons

Stripe Assay to Study the Attractive or Repulsive Activity of a Protein Substrate Using Dissociated Hippocampal Neurons

Axon guidance molecules regulate neuronal migration and targeted growth-cone navigation. We present a powerful method, the stripe assay, to assess the ability of guidance molecules to attract or repulse neurons. In this protocol, we demonstrate the stripe assay by showing FLRT2's ability to repel cultured hippocampal neurons.

nucleofection and primary culture of embryonic mouse hippocampal and cortical neurons

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

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...

using live&#45;cell imaging to measure the effects of pathological proteins on axonal transport in primary hippocampal neurons

Author Spotlight: Understanding the Impact of Pathological Proteins on Axonal Transport in Neurodegenerative Diseases

Here, we demonstrate how to combine transfection of primary hippocampal rodent neurons with live-cell confocal imaging to analyze pathological protein-induced effects on axonal transport and identify mechanistic pathways mediating these...

Using Live&#45;Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

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

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...

Visualizing Proteins in Primary Hippocampal Neurons by Live&amp;#45;Cell Imaging

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imaging phluorin-tagged receptor insertion to the plasma membrane in primary cultured mouse neurons

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

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

measuring properties of the membrane periodic skeleton of the axon initial segment using 3d-structured illumination microscopy (3d-sim)

Measuring Properties of the Membrane Periodic Skeleton of the Axon Initial Segment using 3D-Structured Illumination Microscopy 3D-SIM

The present protocol describes a method to visualize and measure actin rings and other components of the membrane periodic skeleton of the axon initial segment using cultured rat hippocampal neurons and 3D-structured illumination microscopy...

Measuring Properties of the Membrane Periodic Skeleton of the Axon Initial Segment using 3D-Structured Illumination Microscopy (3D-SIM)

use of pre-assembled plastic microfluidic chips for compartmentalizing primary murine neurons

Use of Pre-Assembled Plastic Microfluidic Chips for Compartmentalizing Primary Murine Neurons

This protocol describes the use of plastic chips to culture and compartmentalize primary murine neurons. These chips are preassembled, user-friendly, and compatible with high-resolution, live, and fluorescence imaging. This protocol describes how to plate rat hippocampal neurons within these chips and perform fluidic isolation, axotomy and immunostaining.

assessing the expression of major histocompatibility complex class i on primary murine hippocampal neurons by flow cytometry

Assessing the Expression of Major Histocompatibility Complex Class I on Primary Murine Hippocampal Neurons by Flow Cytometry

This protocol describes a method for culturing primary hippocampal neurons from embryonic mouse brain. The expression of major histocompatibility complex class I on the extracellular surface of cultured neurons is then assessed by flow cytometric...

harvesting of primary hippocampal neurons for live&#45;cell imaging

Harvesting of Primary Hippocampal Neurons for Live&#45;Cell Imaging

measuring axonal cargo transport in mouse primary cortical cultured neurons

Measuring Axonal Cargo Transport in Mouse Primary Cortical Cultured Neurons

The present protocol describes the whole procedure of analysis for axonal transport. In particular, the calculation of transport velocity, not including the pausing, and the visualization method using open-access software "KYMOMAKER" are shown...

easy and reproducible low-density primary culture using frozen stock of embryonic hippocampal neurons

Easy and Reproducible Low-Density Primary Culture using Frozen Stock of Embryonic Hippocampal Neurons

A ready-to-use frozen stock of neurons is a powerful tool for evaluating synaptic functions. Here, we introduce an easy low-density primary culture from frozen stock using a 96-well...

an in vitro technique to study glutamate receptor trafficking in hippocampal neurons

This video demonstrates an antibody-feeding approach to study the trafficking of glutamate receptors in primary hippocampal neuronal cultures. This approach allows the observation of receptor populations at both the plasma and internal membranes. Discrimination between these subsets is achieved by labeling the receptors before and after membrane permeabilization, using the same primary antibody but secondary antibodies conjugated to different...

An In Vitro Technique to Study Glutamate Receptor Trafficking in Hippocampal 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 ...

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

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

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

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

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

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 ...

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.

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. ...

Growing axons develop a highly motile structure at their tip, termed the growth cone. The growth cone contacts extracellular environmental cues to navigate axonal growth. Netrin, slit, semaphorin, and ephrins are known guidance molecules that can attract or repel axons upon binding to receptors and co-receptors on the axon. The activated receptors initiate various signaling molecules in the growth cone that alter the structure and movement of the neuron. Here, we describe the detailed protocol for a stripe assay to assess the ability of a guidance molecule to attract or repel neurons. In this method, dissociated hippocampal neurons from E15.5 mice are cultured on laminin-coated dishes processed with alternating stripes of ectodomain of fibronectin and leucine-rich transmembrane protein-2 (FLRT2) and control immunoglobulin G (IgG) fragment crystallizable region (Fc) protein. Both axons and cell bodies were strongly repelled from the FLRT2-coated stripe regions after 24 h of culture. Immunostaining with tau1 showed that ~90% of the neurons were distributed on the Fc-coated stripes compared to the FLRT2-Fc-coated stripes (~10%). This result indicates that FLRT2 has a strong repulsive effect on these neurons. This powerful method is applicable not only for primary cultured neurons but also for a variety of other cells, such as neuroblasts.

Axon guidance molecules regulate neuronal migration and targeted growth-cone navigation. We present a powerful method, the stripe assay, to assess the ability of guidance molecules to attract or repulse neurons. In this protocol, we demonstrate the stripe assay by showing FLRT2's ability to repel cultured hippocampal neurons.

Growing axons develop a highly motile structure at their tip, termed the growth cone. The growth cone contacts extracellular environmental cues to ...

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 ...

Fluorescence-Activated Nuclei Negative Sorting of Neurons Combined with Single Nuclei RNA Sequencing to Study the Hippocampal Neurogenic Niche

Adult Hippocampal Neurogenesis (AHN), which consists of a lifelong maintenance of proliferative and quiescent neural stem cells (NSCs) within the sub-granular zone (SGZ) of the dentate gyrus (DG) and their differentiation from newly born neurons into granule cells in the granule cell layer, is well validated across numerous studies. Using genetically modified animals, particularly rodents, is a valuable tool to investigate signaling pathways regulating AHN and to study the role of each cell type that compose the hippocampal neurogenic niche. To address the latter, methods combining single nuclei isolation with next generation sequencing have had a significant impact in the field of AHN to identify gene signatures for each cell population. Further refinement of these techniques is however needed to phenotypically profile rarer cell populations within the DG. Here, we present a method that utilizes Fluorescence Activated Nuclei Sorting (FANS) to exclude most neuronal populations from a single nuclei suspension isolated from freshly dissected DG, by selecting unstained nuclei for the NeuN antigen, in order to perform single nuclei RNA sequencing (snRNA-seq). This method is a potential steppingstone to further investigate intercellular regulation of the AHN and to uncover novel cellular markers and mechanisms across species.

Presented here is a method to sequence single nuclei isolated from the mouse dentate gyrus that excludes most neurons through fluorescence-activated nuclei (FAN)-sorting. This approach generates high-quality expression profiles and facilitates the study of most other cell types represented in the niche, including scarce populations such as neural stem cells.