This work is supported by startup funds from The Jackson Laboratory.

1. Preparing Mouse Glia Culture for Neuronal Culture Conditioning
<ol>
	<li>T75 flasks are coated with collagen solution (1:3 dilution of Purecol in ddH2O). The flasks are set upright and left to dry overnight in a tissue culture hood.</li>
	<li>Dissection buffer (aCSF: 119 mM NaCl, 5 mM KCl, 1 mM MgCl2, 30 mM dextrose, 25 mM HEPES, pH 7.4, without calcium) and glia media (DMEM supplemented with 10% FBS, 10 units/ml penicillin, 10 &mu;g/ml streptomycin, and Glutamax) are prepared and stored at 4 &deg;C until ready for use.</li>
	<li>On the day of glia culture, the glia media is warmed for at least 30 min in a 37 &deg;C water bath before dissection of mouse brains. Embryonic or neonatal mice are euthanized by rapid decapitation. Cerebral cortexes are dissected out under a dissection microscope.</li>
	<li>Dissected brain tissues are digested in papain solution (100 ml papain and 20 &mu;l 1% DNase I in 2 ml dissection buffer) at 37 &deg;C for 20 min.</li>
	<li>Following papain digestion, brain tissues are rinsed with 10 ml pre-warmed glia media. Fresh glia media (2 ml) is then added to the digested brain tissues, and the tissues are triturated with a fire-polished glass Pasteur pipette.</li>
	<li>Fresh glia media (8 ml) is added to the dissociated tissues. A 70 &mu;m cell-strainer is used to filter the dissociated tissue suspension. Cells are then seeded into T75 flasks (usually 2-3 embryos can seed one T75 flask) and placed into a 37 &deg;C cell culture incubator.</li>
	<li>The next day, the media from the T75 flasks is aspirated. The flasks are rinsed with 10 ml PBS. Fresh glia media (10-15 ml) is then added to each flask.</li>
	<li>Within 10-12 days, glia in the T75 flasks should be confluent. Glial cells are rinsed with PBS, and 5 ml of 0.05% trypsin is added to each T75 flask.</li>
	<li>After trypsin incubation at 37 &deg;C for 5 min, glial cells are dissociated and glial cells from each T75 flask are split into two different collagen-coated T75 flasks.</li>
	<li>Culture inserts (Millipore PICM03050) are prepared by placing them into 6-well plates, coating them with collagen, and air-drying overnight. Glia media (2 ml each) is placed underneath each culture insert.</li>
	<li>Glia from confluent T75 flasks are trypsinized and seeded onto the culture inserts (one T75 flask per 6-well plate, 2 ml each insert). These glia culture inserts will be used to condition the neuronal cultures.</li>
</ol>
2. Neuronal Culture
<ol>
	<li>The glass coverslips for neuronal culture are cleaned by soaking in 70% HNO3 at least overnight. Coverslips are then washed with ddH2O at least 3 times, and dried in an Isotemp Oven (&gt;200 &deg;C) overnight.</li>
	<li>Cleaned coverslips are placed in a 6-well plate, one coverslip per well. Coverslips are coated with poly-L-Lysine solution (2 ml, 0.1% in 100 mM boric acid, pH = 8.5) overnight in a 37 &deg;C incubator.</li>
	<li>The poly-L-Lysine solution is removed from the coverslips and coverslips are washed 3 times with autoclaved ddH2O. Coverslips are then incubated in a 37 &deg;C incubator overnight in plating media (Neurobasal supplemented with 5% heat-inactivated horse serum, 10 units/ml penicillin, 10 &mu;g/ml streptomycin, Glutamax; 2% B27 supplement is added to the plating medium on the day it is used).</li>
	<li>On the following day, the plating media is removed and 2 ml of fresh plating media with B-27 is added to each well that has a glass coverslip.</li>
	<li>Pregnant mice (E15-18) are euthanized with CO2, and embryonic mice are euthanized by decapitation. The hippocampus or striatum is dissected from the brain tissue and placed into a 15 ml conical tube with ice-cold dissection buffer, for culture of hippocampal neurons or striatal medium spiny neurons, respectively.</li>
	<li>Dissected brain tissues are dissociated with papain as described in steps 1.4) and 1.5).</li>
	<li>Following dissociation, the number of cells in each neuronal suspension is counted with a hemocytometer. Neurons are then seeded on each coverslip at a density of 0.4 x 106 cells per well of a 6-well plate, and cultured in a 37 &deg;C incubator overnight (neurons are DIV 0).</li>
	<li>To prepare glia culture inserts for conditioning the neuronal cultures, the glia media is removed from the glia culture inserts and fresh plating medium is added (2 ml in the insert and 2 ml underneath the insert).</li>
	<li>The next day (DIV 1), coverslips with neurons are placed underneath the glia culture inserts, one coverslip in each well.</li>
	<li>Neurons are fed with half the volume of fresh pre-warmed feeding media (with B-27) every 3-4 days until they are ready for transfection and imaging experiments.</li>
</ol>
3. Transfection
<ol>
	<li>Neuronal transfection is performed using Lipofectamine 2000. For each neuronal culture (one culture per coverslip), 2 &mu;l Lipofectamine 2000 and 1 &mu;g of DNA are used. Fresh Neurobasal medium with no supplement is used to dilute Lipofectamine and DNA.</li>
	<li>DNA and Lipofectamine are incubated for 20 min at room temperature after mixing.</li>
	<li>Save 1 ml of the media from underneath each glia culture insert, and 1 ml from inside each insert, and transfer the media to a conical tube. Add the same volume of feeding media + B27 to the saved medium and incubate at 37 &deg;C. This medium is used for neuronal culture following transfection.</li>
	<li>Following the 20-min Lipofectamine/DNA incubation, glia culture inserts are removed momentarily from the 6-well plates. The 200 &mu;l Lipofectamine/DNA mixture is added to each coverslip with neurons. The glia culture inserts are then returned to each well, above the neurons. Generation of bubbles under the membrane of the culture inserts should be avoided. Neurons are incubated with the Lipofectamine/DNA mixture at 37 &deg;C for 2-4 hr.</li>
	<li>After the 2-4 hr incubation period, glia inserts are removed again. The culture media with the Lipofectamine/DNA mixture is removed from each well. The media saved in step 3.3) is added to neurons (2 ml per well). Glia culture inserts are returned to each well, and the glia media from each insert is removed, and 2 ml of the media from step 3.3) is added into each insert.</li>
	<li>Neurons are cultured for an additional 24-72 hr to allow expression of transfected cDNA before TIRF imaging is performed on these neurons.</li>
</ol>
4. TIRF Imaging
<ol>
	<li>Artificial Cerebrospinal Fluid (aCSF: 119 mM NaCl, 2.5 mM KCl, 30 mM glucose, 2 mM CaCl2, 1 mM MgCl2, 25 mM HEPES, pH = 7.4) will be used for live-imaging experiments. Our TIRF imaging system is custom set-up with a 100-mW Cyan laser for excitation, and an Electron Multiplying Charge Coupled Device (EMCCD) camera for a detector.</li>
	<li>Prior to imaging experiments, the aCSF is warned in a 37 &deg;C water bath for at least 30 min. The heating insert for live imaging is placed on the microscope stage, and the heating insert, laser, and camera are warmed for at least 30 min before the start of the imaging experiments.</li>
	<li>A coverslip with transfected neurons is assembled into the live-imaging chamber. The pre-warmed aCSF is placed in the chamber after the chamber is assembled; this step should be completed as quickly as possible.</li>
	<li>Once the chamber is placed on the heating insert on the microscope stage, transfected neurons are identified visually under epi-fluorescent imaging mode.</li>
	<li>After healthy transfected neurons are identified (refer to Figures 1 and 2), we typically perform 1 min of photo-bleach to eliminate pre-existing pHluorin fluorescence on the plasma membrane, as pHluorin-receptors on the plasma membrane have very high fluorescence levels when TIRF imaging is first started, which interferes with visualization of individual insertion events.</li>
	<li>Following photobleach, the gain setting of the electron multiplier on the EMCCD camera is set to maximum, and recording is performed for 1 min at 10 Hz.</li>
	<li>Each recording will contain 600 images. Individual insertion events are identified by playing the recording back using ImageJ. Individual insertion events are analyzed manually using ImageJ.</li>
</ol>
5. Representative Results
Consistent neuronal culture is the key to successful live-imaging experiments. Figure 1 shows mouse hippocampal neurons cultured using our protocol from DIV 11 to DIV 18. It is clear from the images that the neurons have developed extensive processes and that the dendritic processes are covered densely with dendritic spines, indications that these neurons are healthy in culture. Our culture protocol could also be used for other types of neurons in the brain. For example, we recently used this protocol to culture striatal medium spiny neurons in a study to examine dopamine D2 receptor (DRD2) insertion in cultured mouse striatal medium spiny neurons 9. Figure 2 shows mouse striatal medium spiny neurons cultured using our protocol at DIV 8. We have also successfully performed TIRF imaging of superecliptic pHluorin-tagged DRD2s in this type of neuron. Figure 3 shows expression of pHluorin-tagged DRD2 (pH-DRD2) in a medium spiny neuron and visualization of pH-DRD2 insertion in this neuron.
<img alt="Figure 1" src="/files/ftp_upload/4450/4450fig1.jpg" /> 
Figure 1. Mouse hippocampal culture using the interface culture method. DIV: days in vitro, the day of culture is considered as DIV 0. From the images, it is evident that hippocampal neurons mature in this type of culture between DIV 11 and DIV 15. At DIV 11 and 13, neuronal dendrites are covered with filipodia and immature spines. At DIV 15 and 18, neuronal dendrites are covered with large spines, an indication of more mature neurons. Left panels: low-magnification images of mouse hippocampal neurons at different ages. Right panels: high-magnification images (zoom in on neuronal dendrites of the left panel) of neuronal dendritic processes with dendritic spines at different ages.
<img alt="Figure 2" src="/files/ftp_upload/4450/4450fig2.jpg" /> 
Figure 2. Mouse striatal medium spiny neuron culture using the interface culture method. Neurons in the images are at DIV 8.
<img alt="Figure 3" src="/files/ftp_upload/4450/4450fig3.jpg" /> 
Figure 3. A mouse striatal medium spiny neuron transfected with super ecliptic pHluorin-tagged dopamine D2 receptor (pH-DRD2). Image on the left is maximum intensity projection of a time-lapse recording (600 images, 100 msec per image). White arrowheads indicate individual vesicular insertion of pH-DRD2. This image retains insertion information in the x-y dimension but loses the time information. Image on the right shows y-t maximum intensity projection, in which the x-y-t stack is rotated 90 degrees around the y-axis, and the maximum pixel on each x-axis line is projected onto a single y-axis pixel. This image retains insertion information along the y-t axis but loses the x-axis information.

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No conflicts of interest declared.

For unknown reasons, mouse neurons are always more difficult to culture than rat neurons. In our experience, a mixed culture of neurons and glia works well for primary cultured mouse neurons. However, such a mixed culture is not suitable for TIRF imaging experiments, as in this type of culture neurons and their processes tend to grow on top of glia cells, situating the neuronal somata and dendritic processes beyond the reach of TIRF microscopy. Therefore, a lower-density neuronal culture with few glia on the covers...

<table border="1">
	<tbody>
 <tr>
 <td>Name of Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>purified bovine collagen solution (Purecol)</td>
 <td>Advanced Biomatrix</td>
 <td>5005-B</td>
 <td></td>
 </tr>
 <tr>
 <td>Hank's Balanced Salt Solution (HBSS)</td>
 <td>GIBCO</td>
 <td>14185-045</td>
 <td></td>
 </tr>
 <tr>
 <td>penicillin-streptomycin (Pen Strep)</td>
 <td>GIBCO</td>
 <td>15140-122</td>
 <td></td>
 </tr>
 <tr>
 <td>sodium pyruvate</td>
 <td>GIBCO</td>
 <td>11360-070</td>
 <td></td>
 </tr>
 <tr>
 <td>DMEM High Glucose</td>
 <td>GIBCO</td>
 <td>10313-021</td>
 <td></td>
 </tr>
 <tr>
 <td>fetal bovine serum (FBS)</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>GlutaMAX</td>
 <td>GIBCO</td>
 <td>35050-061</td>
 <td></td>
 </tr>
 <tr>
 <td>papain</td>
 <td>Worthington Biochemical Corp.</td>
 <td>LS003126</td>
 <td></td>
 </tr>
 <tr>
 <td>Deoxyribonuclease I from bovine pancreas (DNase I)</td>
 <td>SIGMA</td>
 <td>DN25-10MG</td>
 <td></td>
 </tr>
 <tr>
 <td>Dulbecco's Phosphate Buffered Saline (DPBS)</td>
 <td>GIBCO</td>
 <td>14190-144</td>
 <td></td>
 </tr>
 <tr>
 <td>0.05% trypsin</td>
 <td>GIBCO</td>
 <td>25300-054</td>
 <td></td>
 </tr>
 <tr>
 <td>poly-l-lysine hydrobromide</td>
 <td>SIGMA</td>
 <td>P2636-1G</td>
 <td></td>
 </tr>
 <tr>
 <td>boric acid</td>
 <td>Fisher-Scientific</td>
 <td>BP 168-500</td>
 <td></td>
 </tr>
 <tr>
 <td>Neurobasal Medium</td>
 <td>GIBCO</td>
 <td>21103-049</td>
 <td></td>
 </tr>
 <tr>
 <td>B-27 Serum-Free Supplement</td>
 <td>GIBCO</td>
 <td>17504-044</td>
 <td></td>
 </tr>
 <tr>
 <td>heat inactive horse serum</td>
 <td>GIBCO</td>
 <td>26050-070</td>
 <td></td>
 </tr>
 <tr>
 <td>Lipofectamine 2000</td>
 <td>Invitrogen</td>
 <td>11668 019</td>
 <td></td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Fisher-Scientific</td>
 <td>BP310-500</td>
 <td></td>
 </tr>
 <tr>
 <td>Culture Insert</td>
 <td>Millipore</td>
 <td>PICM03050</td>
 <td></td>
 </tr>
	 </tbody>
 </table>

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

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

Watch this Scientific Journal Video about Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons at JoVE.com

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 membrane.

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

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13.1K Views.  The Jackson Laboratory. 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.

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

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

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

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

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

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

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion channel, and transporter cell surface presentation on a minutes time scale.&#160;There is a broad diversity of mechanisms that control endocytic trafficking of individual proteins. Studies investigating the molecular underpinnings of trafficking have primarily relied upon surface biotinylation to quantitatively measure changes in membrane protein surface expression in response to exogenous stimuli and gene manipulation.&#160;However, this approach has been mainly limited to cultured cells, which may not faithfully reflect the physiologically relevant mechanisms at play in adult neurons.&#160;Moreover, cultured cell approaches may underestimate region-specific differences in trafficking mechanisms.&#160;Here, we describe an approach that extends cell surface biotinylation to the acute brain slice preparation.&#160;We demonstrate that this method provides a high-fidelity approach to measure rapid changes in membrane protein surface levels in adult neurons.&#160;This approach is likely to have broad utility in the field of neuronal endocytic trafficking.

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission.&#160;To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons.&#160;Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion ...

Primary Culture of Mouse Dopaminergic Neurons

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions such as motor control, motivation, and working memory. Nigrostriatal dopaminergic neurons selectively degenerate in Parkinson&#39;s disease (PD). This progressive neuronal loss is unequivocally associated with the motors symptoms of the pathology (bradykinesia, resting tremor, and muscular rigidity). The main agent responsible of dopaminergic neuron degeneration is still unknown. However, these neurons appear to be extremely vulnerable in diverse conditions. Primary cultures constitute one of the most relevant models to investigate properties and characteristics of dopaminergic neurons. These cultures can be submitted to various stress agents that mimic PD pathology and to neuroprotective compounds in order to stop or slow down neuronal degeneration. The numerous transgenic mouse models of PD that have been generated during the last decade further increased the interest of researchers for dopaminergic neuron cultures. Here, the video protocol focuses on the delicate dissection of embryonic mouse brains. Precise excision of ventral mesencephalon is crucial to obtain neuronal cultures sufficiently rich in dopaminergic cells to allow subsequent studies. This protocol can be realized with embryonic transgenic mice and is suitable for immunofluorescence staining, quantitative PCR, second messenger quantification, or neuronal death/survival assessment.

Dopaminergic neurons play a vital regulatory role in the brain. Their loss is associated with Parkinson's disease. In this video, we show how to generate primary cultures of central dopaminergic neurons from embryonic mouse mesencephalon. Such cultures are useful to study the extreme vulnerability of these neurons to various stresses.

Dopaminergic neurons represent less than 1% of the total number of neurons in the brain. This low amount of neurons regulates important brain functions ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

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 ...

Primary Culture of Neurons Isolated from Embryonic Mouse Cerebellum

The use of primary cell cultures has become one of the major tools to study the nervous system in vitro. The ultimate goal of using this simplified model system is to provide a controlled microenvironment and maintain the high survival rate and the natural features of dissociated neuronal and nonneuronal cells as much as possible under in vitro conditions. In this article, we demonstrate a method of isolating primary neurons from the developing mouse cerebellum, placing them in an in vitro environment, establishing their growth, and monitoring their viability and differentiation for several weeks. This method is applicable to embryonic neurons dissociated from cerebellum between embryonic days 12&ndash;18.

Conducting in vitro experiments to reflect in vivo conditions as adequately as possible is not an easy task. The use of primary cell cultures is an important step toward understanding cell biology in a whole organism. The provided protocol outlines how to successfully grow and culture embryonic mouse cerebellar neurons.

The use of primary cell cultures has become one of the major tools to study the nervous system in vitro. The ultimate goal of using this simplified model ...