This work was supported by the Intramural Programs of the National Institute of Neurological Disorders and Stroke (NINDS) and National Institute on Deafness and Other Communication Disorders (NIDCD), of the National Institutes of Health (NIH). We thank the NINDS EM facility and the NIDCD advanced imaging core (code # ZIC DC 000081-03) for assistance.

Care and handling of animals were in accordance with NIH Animal Care and Use Committee Guidelines. Postnatal day (P) 15-21 Sprague-Dawley rats, injected with 1-1.2% CTB bilaterally through the superior colliculus, were maintained on a 12:12-hr light:dark cycle.
1. Retinal Tissue Fixation
<ol>
	<li>Assemble the following materials and tools: A dissecting microscope, 2 forceps with very fine tips, scissors, cellulose filter paper, plastic pipette and a microscope slide.</li>
	<li>Anesthetize the rat in a closed chamber with 2.0 ml halothane (an inhalant anesthetic). Determine adequate anesthetization by these methods: lack of withdrawal of rear paw after toe pinch, or lack of blink reflex. Then decapitate immediately with a guillotine. Remove the eyes with a pair of iris scissors and place in a glass dish containing 4% paraformaldehyde in 0.1 M phosphate buffer (PB) at pH 7.4.</li>
	<li>Using the dissecting microscope, remove the cornea by cutting off the front of the eyeball. Remove the lens and vitreous from the inner retinal surface with forceps.</li>
	<li>Peel the sclera with the two forceps until the retina is isolated from the eyecup.</li>
	<li>Cut the retina immediately into 100 - 200 &#181;m-thick strips with a razor, and subject to pH-shift fixation.</li>
	<li>Fix retina strips in 4% paraformaldehyde in 0.1 M PB at pH 6.0 for 20 - 30 min and then in 4% paraformaldehyde plus 0.01% glutaraldehyde at pH 10.5 for 10 - 20 min at&#160;RT.</li>
	<li>After several washes in PB with 0.15 mM CaCl2 (pH 7.4 at 4&#176;C), cryoprotect the retinal strips with glycerol (60 min each in 10%, 20%, 30%, then O/N in 30%) in 0.1 M PB prior to freeze substitution.</li>
</ol>
2. Freeze-substitution
NOTE: This freeze-substitution method is modified from an earlier published protocol 19,20. Also, it is crucial that the instruments are very cold (wear gloves); otherwise, the tissue may thaw partially when touched with the instruments. All of these steps are done within the AFS chamber and the instruments are never allowed to move above the rim of the chamber. Similarly, proper cooling of all chemicals used in the AFS is necessary.
<ol>
	<li>Plunge-freeze the retinal strips in liquid propane at -184&#176;C in an EM cryopreservation instrument (CPC). Using a fine brush, place the samples on small pieces of double-stick tape attached to the metal stubs that go on the end of the plunging rods (wick off extra buffer using the brush).</li>
	<li>Plunge the rods into the liquid propane and keep there for about 5 sec, and then transfer to the automatic freeze-substitution instrument (AFS) using a small transport chamber that is filled with liquid nitrogen (&#39;LN2 cooled cryo transfer container&#39;).</li>
	<li>After placing the frozen sample and instruments (forceps and scalpel) into the AFS, cool the instruments in the chamber for several minutes before touching the tissue, or cool them by placing the tips of the instruments for a few seconds into the small transport chamber (filled with liquid nitrogen (&#39;LN2&#160;cooled cryo transfer container&#39;)).&#160;&#160;</li>
	<li>Once in the AFS, remove the sample and tape from the stub using a fine scalpel. Also, keep the nitrogen gas flow control, TF (TF is described as a &#39;regulator control for LN2 vaporiser&#39;) open during these procedures.</li>
	<li>Prior to placing the frozen tissue into the flat-embedding holders (i.e., before setting up the AFS), cut a thin circle from a clear plastic sheet and place it into the bottom of the holder so as to line the bottom of each well. This allows relatively easier removal of the polymerized specimen blocks when finished. 
	NOTE: Previously, we used an alternative method to the flat embedding holders, and employing double Reichert+gelatin capsules (see Petralia and Wenthold, 1999) 20.</li>
	<li>Use the following automatic sequence in the AFS (using instrument terminology): T1 = -90&#176;C for 32 hr, S1 = increase temperature by 4&#176;C/hr&#160;(11.3 hr), T2 = -45&#176;C for 50 hr, S2 = increase temperature by 5&#176;C/hr&#160;(9 hr), T3 = 0&#176;C for 40 hr (total = 142.3 hr). It may take 2 - 4 hr to place samples in the AFS (at -90&#176;C) prior to starting the timed sequence.</li>
	<li>Immerse the frozen sections in 1.5% uranyl acetate in methanol at -90&#176;C for &#62;32 hr in the AFS. Place two pieces of tissues (typically) in each of the seven wells in the flat-embedding aluminum holder (three fit into the AFS).
	<ol>
		<li>Place the tissue+tape into the uranyl acetate/methanol in the wells and remove the tape later, just prior to beginning embedding medium (such as Lowicryl HM20) infiltration, if it is too difficult to remove the tissue from the tape.</li>
	</ol>
	</li>
	<li>Then increase the temperature stepwise to -45&#176;C (+4&#176;C per hr; in the automatic sequence).</li>
	<li>Wash the samples three times in precooled methanol, by using a fine-tipped plastic pipette to remove the old solution from each flat-embedding holder, and then using another standard plastic pipette to add the precooled fresh methanol.</li>
	<li>Then, using the same method, infiltrate the samples progressively with low temperature embedding resins such as embedding medium (HM20/methanol at 1:1 and 2:1, each for 2 hr, followed by pure resin for 2 hr and then change the resins and keep O/N).</li>
	<li>Change the resin again the next day, adjusting the level of the resin to reach just to the top edge of the wells.</li>
	<li>Polymerize the samples (-45&#176;C to 0&#176;C in automatic sequence; +5&#176;C per hr) with UV light for 40 hr.</li>
	<li>Then remove the samples from the AFS. Typically, sample blocks still show some pinkness in color. Place the samples close to the fluorescent light in the chemical fume hood, at RT O/N or longer until they appear completely clear.</li>
</ol>
3. Postembedding EM Immunogold Immunocytochemistry
NOTE: Postembedding immunocytochemistry is performed as described 23,24,25.
<ol>
	<li>Cut 1 &#181;m sections with ultramicrotome, stain sections with 1% toluidine-blue, and examine them for section orientation; orient the tissue block to achieve the optimal transverse plane of sectioning.</li>
	<li>Cut 70 nm thick ultrathin sections with ultramicrotome and collect them on Formvar-carbon-coated nickel slot grids.</li>
	<li>Wash grids one time in distilled H2O followed by a Tris-buffered saline three times (TBS, 0.05 M Tris buffer, 0.7% NaCl, pH 7.6) wash.</li>
	<li>Incubate grids in 20 &#181;l drops of 5% BSA in TBS for 30 min , and then in 20 &#181;l drops of a mixture containing goat anti-CTB (1:3,000) and an antibody to one NMDAR subunit (anti-rabbit GluN2A 1:50, GluN2B 1:30), or an anti-rabbit GluA2/3 (1:30) in TBS-Triton (TBST, 0.01% Triton X-100, pH 7.6) with 2% BSA and 0.02 M NaN3 O/N at RT.</li>
	<li>Wash grids on three separate drops (20 &#181;l) of TBS (pH 7.6) for 10, 10, and 20 min, followed by TBS (pH 8.2) for 5 min.</li>
	<li>Incubate grids for 2 hr on drops (20 &#181;l) of a mixture containing donkey anti-rabbit IgG (1:20) coupled to 10 nm gold particles and donkey anti-goat IgG (1:20) coupled to 18 nm gold particles in TBST (pH 8.2) with 2% BSA and 0.02 M NaN3.</li>
	<li>Wash grids in 20 &#181;l drops of TBS (pH 7.6) for 5, 5, and 10 min, then in ultrapure water and then dry them.</li>
	<li>Counterstain grids with 5% uranyl acetate and 0.3% lead citrate in distilled H2O for 8 and 5 min under dark conditions, respectively.</li>
	<li>For triple-labeling experiments, incubate grids O/N&#160;at RT with 20 &#181;l drops of a mixture of anti-goat CTB (1:3,000), anti-mouse PSD-95 (1:100), and anti-rabbit GluN2A (1:50). Then incubate grids for 2 hr on 20 &#181;l drops of a mixture of IgGs coupled to 18, 10, and 5 nm gold particles in TBST (pH 8.2) with 2% BSA and 0.02 M NaN3. Keep the same procedures as double labeling for washing and counterstaining between and after antibody incubation.</li>
	<li>Controls were performed in which the secondary antibodies were applied alone.</li>
	<li>View grids on an EM and digitalize images. Process final figures in Adobe Photoshop 6.0 only for brightness and contrast if it is necessary24.</li>
</ol>
4. Quantification
<ol>
	<li>Manually select areas of the inner plexiform layer (IPL) without holes or cracks at 8,000X magnification, then randomly photomontage the full depth of IPL at 25,000X magnification.</li>
	<li>Identify RGC dendrites at cone bipolar dyads when they a) contain the retrogradely transported CTB signal (large gold particles), b) exhibit well-defined membranes, clefts, and postsynaptic densities, c) contain at least two small gold particles within the PSD, or more than one small gold particle along the extrasynaptic plasma membrane.</li>
	<li>Collect 45 - 55 RGC dendrites, based on above criteria, for quantitative analysis.</li>
	<li>Measure the lengths of the PSD and the extrasynaptic plasma membrane of individual RGC dendrites with NIH ImageJ software. Count gold particles within 10 nm of the membrane as membrane-associated, based on an average thickness of 7 - 9 nm for the plasma membrane&#160;26.</li>
	<li>Calculate particle density as the number of gold particles per linear micrometer (gold/&#181;m). Measure the distance between the center of each gold particle and the middle of the PSD (for synaptic location) or the edge of the PSD (for extrasynaptic location).</li>
	<li>Perform statistical analysis by software. Use two-tailed t-tests to compare means. Conclude significance when P &#60;0.05. Unless indicated otherwise, report values as mean &#177; SEM18.</li>
</ol>

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The authors have no disclosures.

We have described four techniques for successful quantitative post-embedding immunogold EM: 1) short and weak fixation, 2) freeze-substitution, 3) post-embedding immunogold staining, and 4) quantification.
EM immunogold allows the detection of specific proteins in ultrathin tissue sections. Antibodies labeled with gold particles can be directly visualized using EM. While powerful in detecting the subsynaptic localization of a membrane receptor, EM immunogold can be technically challenging, and...

Glutamate is the major excitatory neurotransmitter in the retina1. Retinal ganglion cells (RGCs), receiving glutamatergic synaptic input from bipolar cells2, are the output neurons of the retina that send visual information to the brain. Physiological studies showed that synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs) 3,4,5. Although excitatory postsynaptic currents (EPSCs) in RGCs are mediated by AMPARs and NMDARs3,5,6,7,8 , spontaneous miniature EPSCs (mEPSCs) on RGCs exhibit only an AMPARs-mediated component 4,5,9. However, reducing glutamate uptake revealed an NMDAR component in spontaneous EPSCs5, suggesting that NMDARs on RGC dendrites may be located outside of excitatory synapses . Membrane-associated guanylate kinases (MAGUKs) such as PSD-95 that cluster neurotransmitter receptors, including glutamate receptors and ion channels at synaptic sites, also exhibit distinct subsynaptic expression patterns 10,11,12,13,14.
Over recent decades, confocal immunohistochemistry and pre-embedding electron microscope (EM) immunohistochemistry have been employed to study membrane receptor expression. Although confocal immunostaining reveals broad patterns of receptor expression, its lower resolution makes it impossible to use to distinguish subcellular location. Pre-embedding EM studies in mammalian retina indicate that NMDAR subunits are present in postsynaptic elements at cone bipolar cell ribbon synapses 15,16,17. This is in apparent contrast to physiological evidence. However, diffusion of reaction product is a well-known artifact in the pre-embedding immunoperoxidase method. Hence, this approach does not usually give statistically reliable data and may exclude distinction between localization to synaptic membrane versus extrasynaptic membrane 18,19,20,21. On the other hand, physiological and anatomical data are consistent with a synaptic localization of AMPARs on RGCs 3,5,7,9,22. Thus, glutamate receptors and MAGUKs at retinal ribbon synapse are localized not only to the postsynaptic but also to the perisynaptic or extrasynaptic membrane compartments. However, a high-resolution quantitative analysis of these membrane proteins in a retinal ribbon synapse is still needed.
Here, we developed a postembedding EM immunogold technique to examine the subsynaptic localization of NMDAR subunits, AMPAR subunits and PSD-95 followed by estimating the number, density and variability of these proteins at synapses onto rat RGCs labeled using cholera toxin subunit B (CTB) retrograde tracing methods.

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			<td height="21" style="height:21px;">Donkey anti-mouse IgG-10 nm gold particles</td>
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			<td>25814</td>
			<td>Dilution 1/20</td>
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			<td height="21" style="height:21px;">Donkey anti-mouse IgG-5 nm gold particles</td>
			<td>EMS</td>
			<td>25812</td>
			<td>Dilution 1/20</td>
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			<td height="21" style="height:21px;">Donkey anti-goat IgG-18 nm gold particles</td>
			<td>Jackson ImmunoResearch</td>
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			<td>Dilution 1/20</td>
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			<td height="21" style="height:21px;">Formvar-Carbon coated nickel-slot grids.</td>
			<td>EMS</td>
			<td>FCF2010-Ni</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Uranyl acetate</td>
			<td>EMS</td>
			<td>22400-1</td>
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			<td height="21" style="height:21px;">Methanol</td>
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			<td>67-56-1</td>
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			<td height="21" style="height:21px;">Lead citrate</td>
			<td>Leica</td>
			<td></td>
			<td></td>
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			<td height="21" style="height:21px;">Leica EM AFS</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Leica EM CPC</td>
			<td>Leica</td>
			<td></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Ultromicrotome</td>
			<td>Leica</td>
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			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">JEOL 1200 EM</td>
			<td>JEOL</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">liquid nitrogen&#160;</td>
			<td>Roberts Oxygen</td>
			<td></td>
			<td></td>
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			<td height="21" style="height:21px;">Propane</td>
			<td>Roberts Oxygen</td>
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			<td height="21" style="height:21px;">CTB</td>
			<td>List Biological Laboratories</td>
			<td>104</td>
			<td>1-1.2%</td>
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			<td height="21" style="height:21px;">Anti-CTB</td>
			<td>List Biological Laboratories</td>
			<td>703</td>
			<td>Dilution 1/4000</td>
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</table>

high-resolution quantitative immunogold analysis of membrane receptors at retinal ribbon synapses

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs). Physiological data have indicated that glutamate receptors at RGCs are expressed not only in postsynaptic but also in perisynaptic or extrasynaptic membrane compartments. However, precise anatomical locations for glutamate receptors at RGC synapses have not been determined. Although a high-resolution quantitative analysis of glutamate receptors at central synapses is widely employed, this approach has had only limited success in the retina. We developed a postembedding immunogold method for analysis of membrane receptors, making it possible to estimate the number, density and variability of these receptors at retinal ribbon synapses. Here we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of retinal fixation, 2) freeze-substitution, 3) postembedding immunogold electron microscope (EM) immunocytochemistry and, 4) quantitative visualization of glutamate receptors at ribbon synapses.

The results presented here demonstrate strikingly different subsynaptic localization patterns of GluA 2/3 and NMDARs on RGC dendrites in rat retina, as described previously 24,25. 77% of GluA 2/3 immunogold particles in RGC dendritic profiles were located within the PSD (Figure 1A), similar to most central synapses. However, NMDARs were located either synaptically or extrasynaptically. 83% of GluN2A immunogold particles were localized in the PSD (Figure...

Watch this Scientific Journal Video about High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses at JoVE.com

High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA ...

high-resolution-quantitative-immunogold-analysis-membrane-receptors

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Neuroscience

7.4K Views.  National Institutes of Health. The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Video: High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

Automated Sholl Analysis of Digitized Neuronal Morphology at Multiple Scales

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3. Specifically, it affects the ability of neurons to receive inputs from other cells2 and contributes to the propagation of action potentials4,5. The morphology of the neurites also affects how information is processed. The diversity of dendrite morphologies facilitate local and long range signaling and allow individual neurons or groups of neurons to carry out specialized functions within the neuronal network6,7. Alterations in dendrite morphology, including fragmentation of dendrites and changes in branching patterns, have been observed in a number of disease states, including Alzheimer's disease8, schizophrenia9,10, and mental retardation11. The ability to both understand the factors that shape dendrite morphologies and to identify changes in dendrite morphologies is essential in the understanding of nervous system function and dysfunction.
Neurite morphology is often analyzed by Sholl analysis and by counting the number of neurites and the number of branch tips. This analysis is generally applied to dendrites, but it can also be applied to axons. Performing this analysis by hand is both time consuming and inevitably introduces variability due to experimenter bias and inconsistency. The Bonfire program is a semi-automated approach to the analysis of dendrite and axon morphology that builds upon available open-source morphological analysis tools. Our program enables the detection of local changes in dendrite and axon branching behaviors by performing Sholl analysis on subregions of the neuritic arbor. For example, Sholl analysis is performed on both the neuron as a whole as well as on each subset of processes (primary, secondary, terminal, root, etc.) Dendrite and axon patterning is influenced by a number of intracellular and extracellular factors, many acting locally. Thus, the resulting arbor morphology is a result of specific processes acting on specific neurites, making it necessary to perform morphological analysis on a smaller scale in order to observe these local variations12.
The Bonfire program requires the use of two open-source analysis tools, the NeuronJ plugin to ImageJ and NeuronStudio. Neurons are traced in ImageJ, and NeuronStudio is used to define the connectivity between neurites. Bonfire contains a number of custom scripts written in MATLAB (MathWorks) that are used to convert the data into the appropriate format for further analysis, check for user errors, and ultimately perform Sholl analysis. Finally, data are exported into Excel for statistical analysis. A flow chart of the Bonfire program is shown in Figure 1.

We have developed a computer program to analyze neuronal morphology. In combination with two existing open source analysis tools, our program performs Sholl analysis and determines the number of neurites, branch points, and neurite tips. The analyses are performed so that local changes in neurite morphology can be observed.

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3.  Specifically, it affects the ability of neurons to ...

Postsynaptic Recordings at Afferent Dendrites Contacting Cochlear Inner Hair Cells: Monitoring Multivesicular Release at a Ribbon Synapse

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the postsynaptic activity of a single ribbon synapse 1-4. Ribbon synapses of sensory cells release neurotransmitter continuously, the rate of which is modulated in response to graded changes in IHC membrane potential 5. Ribbon synapses have been shown to operate by multivesicular release, where multiple vesicles can be released simultaneously to evoke excitatory postsynaptic currents (EPSCs) of varying amplitudes 1, 4, 6-11. Neither the role of the presynaptic ribbon, nor the mechanism underlying multivesicular release is currently well understood.
The IHC is innervated by 10-20 auditory nerve fibers, and every fiber contacts the IHC with a unmyelinated single ending to form a single ribbon synapse. The small size of the afferent boutons contacting IHCs (approximately 1 &mu;m in diameter) enables recordings with exceptional temporal resolution to be made. Furthermore, the technique can be adapted to record from both pre- and postsynaptic cells simultaneously, allowing the transfer function at the synapse to be studied directly 2. This method therefore provides a means by which fundamental aspects of neurotransmission can be studied, from multivesicular release to the elusive function of the ribbon in sensory cells.

Whole-cell patch-clamp recordings from auditory nerve fiber dendrites at the inner hair cell ribbon synapse in the mammalian cochlea.

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the ...

Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and cones). The RPE is a monolayer of pigmented cuboidal cells and associates closely with the neural retina just above it. This association makes the RPE of great interest to researchers studying retinal diseases. The RPE is also the site of an in vivo assay of homology-directed DNA repair, the pun assay. The mouse eye is particularly difficult to dissect due to its small size (about 3.5mm in diameter) and its spherical shape. This article demonstrates in detail a procedure for dissection of the eye resulting in a whole mount of the RPE. In this procedure, we show how to work with, rather than against, the spherical structure of the eye. Briefly, the connective tissue, muscle, and optic nerve are removed from the back of the eye. Then, the cornea and lens are removed. Next, strategic cuts are made that result in significant flattening of the remaining tissue. Finally, the neural retina is gently lifted off, revealing an intact RPE, which is still attached to the underlying choroid and sclera. This whole mount can be used to perform the pun assay or for immunohistochemistry or immunofluorescent assessment of the RPE tissue.

A formal demonstration of the dissection of a mouse eye, resulting in a whole mount of the retinal pigment epithelium.

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods 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 ...

High-resolution Functional Magnetic Resonance Imaging Methods for Human Midbrain

Functional MRI (fMRI) is a widely used tool for non-invasively measuring correlates of human brain activity. However, its use has mostly been focused upon measuring activity on the surface of cerebral cortex rather than in subcortical regions such as midbrain and brainstem. Subcortical fMRI must overcome two challenges: spatial resolution and physiological noise. Here we describe an optimized set of techniques developed to perform high-resolution fMRI in human SC, a structure on the dorsal surface of the midbrain; the methods can also be used to image other brainstem and subcortical structures.
High-resolution (1.2 mm voxels) fMRI of the SC requires a non-conventional approach. The desired spatial sampling is obtained using a multi-shot (interleaved) spiral acquisition1. Since, T2* of SC tissue is longer than in cortex, a correspondingly longer echo time (TE ~ 40 msec) is used to maximize functional contrast. To cover the full extent of the SC, 8-10 slices are obtained. For each session a structural anatomy with the same slice prescription as the fMRI is also obtained, which is used to align the functional data to a high-resolution reference volume.
In a separate session, for each subject, we create a high-resolution (0.7 mm sampling) reference volume using a T1-weighted sequence that gives good tissue contrast. In the reference volume, the midbrain region is segmented using the ITK-SNAP software application2. This segmentation is used to create a 3D surface representation of the midbrain that is both smooth and accurate3. The surface vertices and normals are used to create a map of depth from the midbrain surface within the tissue4.
Functional data is transformed into the coordinate system of the segmented reference volume. Depth associations of the voxels enable the averaging of fMRI time series data within specified depth ranges to improve signal quality. Data is rendered on the 3D surface for visualization.
In our lab we use this technique for measuring topographic maps of visual stimulation and covert and overt visual attention within the SC1. As an example, we demonstrate the topographic representation of polar angle to visual stimulation in SC.

This article describes techniques to perform high-resolution functional magnetic resonance imaging with 1.2 mm sampling in human midbrain and subcortical structures using a 3T scanner. Use of these techniques to resolve topographic maps of visual stimulation in the human superior colliculus (SC) is given as an example.

Functional MRI (fMRI) is a widely used tool for non-invasively measuring correlates of human brain activity. However, its use has mostly been focused upon ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

The process by which the anterior region of the neural plate gives rise to the vertebrate retina continues to be a major focus of both clinical and basic research. In addition to the obvious medical relevance for understanding and treating retinal disease, the development of the vertebrate retina continues to serve as an important and elegant model system for understanding neuronal cell type determination and differentiation1-16. The neural retina consists of six discrete cell types (ganglion, amacrine, horizontal, photoreceptors, bipolar cells, and M&uuml;ller glial cells) arranged in stereotypical layers, a pattern that is largely conserved among all vertebrates 12,14-18.
While studying the retina in the intact developing embryo is clearly required for understanding how this complex organ develops from a protrusion of the forebrain into a layered structure, there are many questions that benefit from employing approaches using primary cell culture of presumptive retinal cells 7,19-23. For example, analyzing cells from tissues removed and dissociated at different stages allows one to discern the state of specification of individual cells at different developmental stages, that is, the fate of the cells in the absence of interactions with neighboring tissues 8,19-22,24-33. Primary cell culture also allows the investigator to treat the culture with specific reagents and analyze the results on a single cell level 5,8,21,24,27-30,33-39. Xenopus laevis, a classic model system for the study of early neural development 19,27,29,31-32,40-42, serves as a particularly suitable system for retinal primary cell culture 10,38,43-45.
Presumptive retinal tissue is accessible from the earliest stages of development, immediately following neural induction 25,38,43. In addition, given that each cell in the embryo contains a supply of yolk, retinal cells can be cultured in a very simple defined media consisting of a buffered salt solution, thus removing the confounding effects of incubation or other sera-based products 10,24,44-45.
However, the isolation of the retinal tissue from surrounding tissues and the subsequent processing is challenging. Here, we present a method for the dissection and dissociation of retinal cells in Xenopus laevis that will be used to prepare primary cell cultures that will, in turn, be analyzed for calcium activity and gene expression at the resolution of single cells. While the topic presented in this paper is the analysis of spontaneous calcium transients, the technique is broadly applicable to a wide array of research questions and approaches (Figure 1).

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

Xenopus laevis provides an ideal model system for studying cell fate specification and physiological function of individual retinal cells in primary cell culture. Here we present a technique for dissecting retinal tissues and generating primary cell cultures that are imaged for calcium activity and analyzed by in situ hybridization.

The process by which  the anterior region of the neural plate gives rise to the vertebrate retina  continues to be a major focus of both clinical and ...

High-throughput Analysis of Mammalian Olfactory Receptors: Measurement of Receptor Activation via Luciferase Activity

Odorants create unique and overlapping patterns of olfactory receptor activation, allowing a family of approximately 1,000 murine and 400 human receptors to recognize thousands of odorants. Odorant ligands have been published for fewer than 6% of human receptors1-11. This lack of data is due in part to difficulties functionally expressing these receptors in heterologous systems. Here, we describe a method for expressing the majority of the olfactory receptor family in Hana3A cells, followed by high-throughput assessment of olfactory receptor activation using a luciferase reporter assay. This assay can be used to (1) screen panels of odorants against panels of olfactory receptors; (2) confirm odorant/receptor interaction via dose response curves; and (3) compare receptor activation levels among receptor variants. In our sample data, 328 olfactory receptors were screened against 26 odorants. Odorant/receptor pairs with varying response scores were selected and tested in dose response. These data indicate that a screen is an effective method to enrich for odorant/receptor pairs that will pass a dose response experiment, i.e. receptors that have a bona fide response to an odorant. Therefore, this high-throughput luciferase assay is an effective method to characterize olfactory receptors&#8212;an essential step toward a model of odor coding in the mammalian olfactory system.

Olfactory receptor activation patterns encode odor identity, but the lack of published data identifying odorant ligands for mammalian olfactory receptors hinders the development of a comprehensive model of odor coding. This protocol describes a method to facilitate high-throughput identification of olfactory receptor ligands and quantification of receptor activation.

Odorants create unique and overlapping patterns of olfactory receptor activation, allowing a family of approximately 1,000 murine and 400 human receptors ...

Metabolomic Analysis of Rat Brain by High Resolution Nuclear Magnetic Resonance Spectroscopy of Tissue Extracts

Studies of gene expression on the RNA and protein levels have long been used to explore biological processes underlying disease. More recently, genomics and proteomics have been complemented by comprehensive quantitative analysis of the metabolite pool present in biological systems. This strategy, termed metabolomics, strives to provide a global characterization of the small-molecule complement involved in metabolism. While the genome and the proteome define the tasks cells can perform, the metabolome is part of the actual phenotype. Among the methods currently used in metabolomics, spectroscopic techniques are of special interest because they allow one to simultaneously analyze a large number of metabolites without prior selection for specific biochemical pathways, thus enabling a broad unbiased approach. Here, an optimized experimental protocol for metabolomic analysis by high-resolution NMR spectroscopy is presented, which is the method of choice for efficient quantification of tissue metabolites. Important strengths of this method are (i) the use of crude extracts, without the need to purify the sample and/or separate metabolites; (ii) the intrinsically quantitative nature of NMR, permitting quantitation of all metabolites represented by an NMR spectrum with one reference compound only; and (iii) the nondestructive nature of NMR enabling repeated use of the same sample for multiple measurements. The dynamic range of metabolite concentrations that can be covered is considerable due to the linear response of NMR signals, although metabolites occurring at extremely low concentrations may be difficult to detect. For the least abundant compounds, the highly sensitive mass spectrometry method may be advantageous although this technique requires more intricate sample preparation and quantification procedures than NMR spectroscopy. We present here an NMR protocol adjusted to rat brain analysis; however, the same protocol can be applied to other tissues with minor modifications.

The neurochemistry of mammalian brain is changed in many neurological and systemic diseases. Characteristic profiles of cerebral metabolites can be efficiently obtained based on crude extracts of brain tissue. To this end, high-resolution NMR spectroscopy is employed, enabling detailed quantitative analysis of metabolite concentrations (metabolomics).

Studies of gene expression on the RNA and protein levels have long been used to explore biological processes underlying disease. More recently, genomics ...

Peering at Brain Polysomes with Atomic Force Microscopy

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, formed by ribosomes, mRNAs, several proteins and non-coding RNAs, represent integrated platforms where translational controls take place. However, while the ribosome has been widely studied, the organization of polysomes is still lacking comprehensive understanding. Thus much effort is required in order to elucidate polysome organization and any novel mechanism of translational control that may be embedded. Atomic force microscopy (AFM) is a type of scanning probe microscopy that allows the acquisition of 3D images at nanoscale resolution. Compared to electron microscopy (EM) techniques, one of the main advantages of AFM is that it can acquire thousands of images both in air and in solution, enabling the sample to be maintained under near physiological conditions without any need for staining and fixing procedures. Here, a detailed protocol for the accurate purification of polysomes from mouse brain and their deposition on mica substrates is described. This protocol enables polysome imaging in air and liquid with AFM and their reconstruction as three-dimensional objects. Complementary to cryo-electron microscopy (cryo-EM), the proposed method can be conveniently used for systematically analyzing polysomes and studying their organization.

While ribosome structure has been extensively characterized, the organization of polysomes is still understudied. To overcome this lack of knowledge, we present here a detailed preparation protocol for accurate imaging of mammalian polysomes by atomic force microscopy (AFM) in air and liquid.

The translational machinery, i.e., the polysome or polyribosome, is one of the biggest and most complex cytoplasmic machineries in cells. Polysomes, ...