This work was supported by the National Institute on Deafness and Other Communication Disorders (NIDCD) Intramural Program.

1. Neuron transfection
<ol>
	<li>Culture embryonic day 18 (E18) rat hippocampal neurons on poly-d-lysine-coated MatTek 35-mm glass-bottom dishes1. On 16-18 days in vitro(DIV), transfect neurons using the Clontech CalPhos Mammalian Transfection Kit. First, replace the culture medium with 1.5 ml <a href="http://www.invitrogen.com/site/us/en/home/Products-and-Services/Applications/Cell-Culture/Mammalian-Cell-Culture/Classical_Media/dmem.html">Dulbecco's Modified Eagle Medium</a> (DMEM) per 35-mm dish 0.5 hour prior to transfection. Save the original culture medium in a sterile 15 ml tube for later (step 1.6) use.</li>
	<li>Mix 10 &mu;g pEGFP-N1 plasmid DNA with sterile H2O (Clontech) and 12.4 &mu;l 2 M calcium solution (Clontech) to a total volume of 100 &mu;l.</li>
	<li>Add the mixture from step 1.2) to 100 &mu;l 2&times;HBS dropwise while vortexing 2&times;HBS at medium speed.</li>
	<li>Let the mixture sit at room temperature for 20 minutes and then add the final mixture from step 1.3) into DMEM-incubated neurons.</li>
	<li>Put the neurons back into the 37 &deg;C incubator for 1-1.5 hours.</li>
	<li>Remove the calcium phosphate-containing medium, then wash cells with DMEM three times. Before returning the culture dish to the incubator, exchange the DMEM medium with the original culture medium.</li>
</ol>
2. FRAP experiment on a spine
<ol>
	<li>Neurons are used for the FRAP experiment two to four days after transfection.</li>
	<li>Replace the culture medium from the 35-mm glass-bottom dish, by immediately adding pre-warmed Tyrode Solution, which contains (in mM) NaCl 145, KCl 5, HEPES 10, Glucose 10, Glycine 0.005, CaCl2 2.6, and MgCl2 1.3 (pH adjusted to 7.4 with NaOH).</li>
	<li>A Zeiss LSM 710 confocal microscope is used for the FRAP experiment. The Zeiss TempModule system is used to control the temperature (37&deg;C), the humidity and the CO2 (5%) of the working system. Make sure that the CO2 tank is connected and the water bottle, which is used for balancing humidity, is filled with water.</li>
	<li>Find a transfected mature dendrite with the 100&times;objective (&alpha;plan-APOCHROMAT 100&times;/1.46 oil). If the transfected cells in the dish are sparse, search for a desired cell with the 40&times; objective (plan-NEOFLUAR 40&times;/1.3 oil), and then switch to the 100&times; objective to capture images.</li>
	<li>Use 5&times; optical zoom and a 256&times;256 pixel resolution to image a short piece of dendrite with several spines. To capture images, use nominal speed 9 (pixel dwell time 3.15 &mu;sec) which takes 0.5 second to finish a scan. The pinhole is set to 2&mu;m to obtain strong fluorescence. When taking images, try to use low laser transmission, for example 1-5 %, to avoid photobleaching the entire image.</li>
	<li>Select the spine of interest. In our experiment, we choose mushroom spines with their spine head diameters of ~1 &mu;m.</li>
	<li>To do a FRAP experiment, take 5 control images before bleaching, then bleach the spine of interest 10 times at nominal 100% laser transmission [The argon laser power is 30 mW, with 50% (15 mW) in the 488 laser line, and approximately 3-4 mW reaches the microscope through the 488 laser line], and then capture a series of images immediately after bleaching. For this experiment, images are captured every 1 second for 15 seconds after bleaching. The interval of time should be adjusted according to different targeting proteins and different experimental designs (Fig. 1).</li>
	<li>Save images.</li>
</ol>
3. Data analysis
<ol>
	<li>Open images with ImageJ software.</li>
	<li>Align the stack of images using the align tool ('plugins' &rarr;'stacks - shuffling' &rarr;'align slices in stack' &rarr;'translation' and/or 'rigid body') in ImageJ, so that the spine of interest does not float, in other words - it remains in the same position on the image.</li>
	<li>Measure relative fluorescence intensity of the spine of interest (Fs), a transfected but unbleached region (control), and a background region (Fb) in time lapse images, using the 'Intensity v Time Monitor' tool of ImageJ ('plugins' &rarr;'stacks - shuffling' &rarr;'Intensity v Time Monitor'). The control region could be a piece of well-focused distal dendrite. The background region is a non-fluorescent region.</li>
	<li>Calculate the photobleaching rate (r) by comparing the fluorescence of the control region before (Fc0) and after (Fc) photobleaching. 
	r = Fc/Fc0.</li>
	<li>Normalize the fluorescence intensity of the target spine (F) as follows: 
	F = (Fs - Fb) / r.</li>
	<li>Curve fit the fluorescence intensity of the target spine with a one-phase exponential equation of Graphpad Prism software (Fig. 2) or other similar software.</li>
	<li>Calculate the mobile fraction (fm) and the immobile fraction (fi) by the following equations: 
	fm = F&infin;/ F0, 
	where F&infin; is the fluorescence intensity after full recovery, and F0 is the fluorescence intensity before photobleaching. 
	fi = 1 - fm. </li>
</ol>
4. Representative Results:
In this study, we perform a FRAP experiment on mature hippocampal neurons. At 18-22 DIV, mushroom spines are already formed. Using our method, the dynamic changes of the fluorescence intensity in a small region, such as a spine, can be recorded.
To analyze the fluorescence recovery process of EGFP, we take 5 images as controls before bleaching and then 1 image every 1 second immediately after bleaching for 15 seconds. The resolution of the image is sufficent for quantitative analysis. The fluorescence recovery profiles of tagged fluorescence proteins are highly reproducible.
We also briefly show how to define the mobile and immobile fractions of a fluorescence protein, using ImageJ and Graphpad Prism software. The FRAP method and analysis we show here can be broadly used in neuroscience, cell biology and other studies.
<img src="/files/ftp_upload/2568/2568fig1.jpg" alt="Figure 1" /> 
Figure 1. FRAP measurements of EGFP fluorescence in a spine from a cultured hippocampal neuron. The red arrowheads indicate the time of photobleaching. Photographs represent the same area before (Pre) and at 0, 1, 5, 10, 15 seconds after photobleaching. The region of the spine, control and background are marked with letters S, C and B, respectively. Neurons were maintained at 37&deg;C during the experiment. Scale bar, 1 &mu;m.
<img src="/files/ftp_upload/2568/2568fig2.jpg" alt="Figure 2" /> 
Figure 2. FRAP curves of EGFP fluorescence over a 15-second period. The green line shows the original curve; the red line shows the normalized curve. The dots on curves show the FRAP every 1 second. The curves were fitted by one-phase exponential equations. The average fluorescence before photobleaching was counted as 100%. In this experiment, the mobile fraction (MF) is 94% and the immobile fraction (IF) is 6%.

<ol>
	<li>Zheng, C. Y., Petralia, R. S., Wang, Y. X., Kachar, B., Wenthold, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SAP102+is+a+highly+mobile+MAGUK+in+spines.">SAP102 is a highly mobile MAGUK in spines.</a> J Neurosci. 30, 4757-4766 (2010).</li><li>Grati, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+turnover+of+stereocilia+membrane+proteins:+evidence+from+the+trafficking+and+mobility+of+plasma+membrane+Ca(2+)-ATPase+2.">Rapid turnover of stereocilia membrane proteins: evidence from the trafficking and mobility of plasma membrane Ca(2+)-ATPase 2.</a> J Neurosci. 26, 6386-6395 (2006).</li><li>Liu, L. N., Aartsma, T. J., Thomas, J. C., Zhou, B. C., Zhang, Y. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FRAP+Analysis+on+Red+Alga+Reveals+the+Fluorescence+Recovery+Is+Ascribed+to+Intrinsic+Photoprocesses+of+Phycobilisomes+than+Large-Scale+Diffusion.">FRAP Analysis on Red Alga Reveals the Fluorescence Recovery Is Ascribed to Intrinsic Photoprocesses of Phycobilisomes than Large-Scale Diffusion.</a> PLoS ONE. 4, e5295-e5295 (2009).</li><li>Wiedenmann, J., Nienhaus, G. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+imaging+with+EosFP+and+other+photoactivatable+marker+proteins+of+the+GFP+family.">Live-cell imaging with EosFP and other photoactivatable marker proteins of the GFP family.</a> Expert Rev Proteomics. 3, 361-374 (2006).</li><li>Shaner, N. C., Patterson, G. H., Davidson, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+fluorescent+protein+technology.">Advances in fluorescent protein technology.</a> J Cell Sci. 120, 4247-4260 (2007).</li><li>Sprague, B. L., McNally, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FRAP+analysis+of+binding:+proper+and+fitting.">FRAP analysis of binding: proper and fitting.</a> Trends Cell Biol. 15, 84-91 (2005).</li></ol>

No conflicts of interest declared.

FRAP analysis has been broadly used in vivo and in vitro1-2studies. This technique commonly utilizes GFP <a href="http://en.wikipedia.org/wiki/Fusion_protein" title="Fusion 
protein">fusion proteins</a>, although it could also use red alga fusion proteins3. This analysis is sensitive and can be used to characterize the mobility of GFP-tagged proteins.
To produce meaningful FRAP analysis, it is important to avoid unnecessary photobleaching before and during the FRAP experiment. There are two ways to achieve this. First, the process to search and observe the experimental neuron should be fast. Especially, observation of neurons with a 100X objective for a long time significantly bleaches the fluorescence. Second, high laser power and frequent scanning often increase the possibility of photobleaching. Thus, it will be necessary to calculate the photobleaching rate in a control region and then normalize the FRAP curve in the experimental region. The normalizing method has been described in the protocol (see step 3.3-3.5 for details).
The photobleaching step is also critical for ensuring good FRAP results. In this experiment, we bleach the spine of interest 10 times at 100% laser transmission. This condition is sufficient to bleach the fluorescence of a spine to background level in a fixed preparation. Thus, we set the fluorescence intensity to the same number as background at 0 seconds after photobleaching. Depending on the speed of the first scan, a significant amount of fluorescence recovery might already be detected when the protein of interest is highly mobile.
Many fluorescent protein probes have been developed to study protein dynamics with FRAP. A complementary approach is, for example, to use photoactivatable GFP (PA-GFP), or photoconvertible variants. Together with FRAP technique, these tools are becoming indispensible for live cell imaging studies4-6.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>35 mm glass-bottom dishes </td>
<td>&nbsp;</td>
<td>MatTek</td>
<td>P35GC-1.0-14-C</td>
<td>&nbsp;</td>
<tr>
<td>CalPhos Mammalian Transfection Kit</td>
<td>&nbsp;</td>
<td>Clontech</td>
<td>631312</td>
<td>&nbsp;</td>
<tr>
<td>pEGFP-N1 plasmid</td>
<td>&nbsp;</td>
<td>Clontech</td>
<td>6085-1</td>
<td>&nbsp;</td>
<tr>
<td>Zeiss confocal microscope</td>
<td>&nbsp;</td>
<td>Zeiss</td>
<td>LSM 710</td>
<td>&nbsp;</td>
<tr>
<td>Zeiss TempModule system</td>
<td>&nbsp;</td>
<td>Zeiss</td>
<td>N.A.</td>
<td>&nbsp;</td>
<tr>
<td>ImageJ software</td>
<td>&nbsp;</td>
<td>NIH</td>
<td>N.A.</td>
<td>&nbsp;</td>
<tr>
<td>Graphpad Prism software</td>
<td>&nbsp;</td>
<td>Graphpad software</td>
<td>N.A.</td>
<td>&nbsp;</td>		</tbody>
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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.

Watch this Scientific Journal Video about Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons at JoVE.com

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.

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 ...

fluorescence-recovery-after-photobleaching-frap-fluorescence-tagged

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Neuroscience

54.1K Views.  National Institutes of Health, Bethesda. 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. 

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

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

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 ...

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 ...

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

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

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

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

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 ...

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

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.

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. ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

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

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