The authors thank Dr. Mike Ehlers and his laboratory members for originally demonstrating the protocol for subcellular fractionation of synaptic plasma membranes. We also thank Wendy Horsfall for management of the mouse colonies. This work was supported by operating grants from CIHR (AJR and AS).

The following protocol conforms to the guidelines of the Canadian Council of Animal Care and has been approved by the Faculty of Medicine and Pharmacy Animal Care Committee at the University of Toronto.
1. Prepare Required Reagents as Described in Table 1
<ol>
	<li>Add protease and phosphatase (if necessary) inhibitors to all sucrose solutions, buffers and ddH2O at concentrations outlined in Table 1. IMPORTANT: Perform all of the steps at 4&#160;&#176;C and pre-cool all of the reagents and equipment prior to the start of the experiment. Label all tubes, including ultracentrifuge tubes, with permanent marker.</li>
</ol>
2. Dissection of the Appropriate Brain Region
<ol>
	<li>Sacrifice the animal by cervical dislocation or decapitation. House and euthanize animals in accordance with institutional and governmental policies on animal care.</li>
	<li>Rapidly remove the brain from the skull and place on an chilled dissection block.</li>
	<li>Dissect the appropriate brain region from fresh tissue and proceed to Step 3.</li>
</ol>
3. Tissue Homogenization with Motor Driven Glass-Teflon Homogenizer
<ol>
	<li>Place 50-100 mg (for SPM), or 100-200 mg (for PSD) of the dissected tissue in a labeled 13 ml polypropylene tube containing 4 ml of 0.32 M HEPES-buffered sucrose solution. Transfer the sample to a 15 ml tapered glass-Teflon tissue grinder/homogenizer, set the motor drive to 900 rpm (setting 7) and homogenize the sample with 12 strokes over a 30 sec period. Use a different glass-Teflon homogenizer between samples, or rinse the homogenizer with cold distilled water between samples and wipe dry with a Kimwipe. NOTE: Up to 4 g of tissue can be homogenized in 4 ml of 0.32 M HEPES-buffered sucrose solution, however it is advisable to use less tissue to ensure proper fractionation.</li>
	<li>Transfer the homogenized sample back to the same 13 ml polypropylene tube. Reserve 100 &#956;l of homogenate and store at -80 &#176;C for subsequent protein quantification and western blot analysis of the total protein fraction.</li>
</ol>
4. Low Speed Centrifugation to Remove the Nuclear Fraction (Yields Supernatant S1)
<ol>
	<li>Centrifuge the homogenate in a fixed angle rotor at 900 x g&#160;for 10 min at 4 &#176;C. Transfer the supernatant (S1) to a new, labeled 13 ml tube and resuspend the nuclear fraction pellet (P1) in 500 &#956;l of 0.32 M HEPES-buffered sucrose. NOTE: The (P1) fraction can be stored at -80 &#176;C and used for western blot analysis of the nuclear fraction.</li>
</ol>
5. Enrichment of the Crude Synaptosomal Fraction (Yields Pellet P2)
<ol>
	<li>Centrifuge the supernatant (S1) at 10,000 x g&#160;for 15 min at 4 &#176;C. Remove the supernatant (S2) and reserve 500 &#956;l in a microcentrifuge tube to store at -80 &#176;C for subsequent protein quantification and western blot analysis of the cytosolic/light membrane fraction. Resuspend the remaining crude synaptosomal fraction pellet (P2) in 1 ml of 0.32 M HEPES-buffered sucrose solution and then add another 3 ml of 0.32 M HEPES-buffered sucrose solution.</li>
	<li>Centrifuge at 10,000 x&#160;g for 15 min at 4 &#176;C. Remove the supernatant (S2&#8217;) and reserve 500 &#956;l in a microcentrifuge tube to store at -80 &#176;C for subsequent protein quantification and western blot analysis of the cytosolic/light membrane fraction. Save the washed crude synaptosomal pellet (P2&#8217;) in the polypropylene tube.</li>
</ol>
6. Lysing (Hypoosmotic Shock) of the Crude Synaptosomal Fraction
<ol>
	<li>Lyse the crude synaptosomal pellet (P2&#8217;) in the polypropylene tube by resuspending it in 1 ml of ddH2O. Add another 3 ml of ddH2O, transfer to a glass-Teflon tissue homogenizer, and homogenize by hand with 3 strokes. NOTE: It is important to perform this step as rapidly as possible and quickly proceed to step 6.2.</li>
	<li>Transfer samples from the homogenizer back into the same 13 ml polypropylene tube and rapidly adjust the sample back to 4 mM HEPES with 16 &#956;l&#160;of 1 M HEPES solution, invert to mix.</li>
	<li>Rotate samples at 4&#160;&#176;C for 30 min to ensure complete lysing.</li>
</ol>
7. Enrichment of the Synaptosomal Membrane Fraction (P3)
<ol>
	<li>Centrifuge at 25,000 x&#160;g for 20 min at 4 &#176;C. Remove the crude vesicular fraction supernatant (S3) and reserve it in a 5 ml tube to store at -80 &#176;C for subsequent protein quantification and western blot analysis. Resuspend the synaptosomal membrane fraction (P3) in 1 ml of 0.32 M HEPES-buffered sucrose solution.</li>
</ol>
8. Preparation of the Discontinuous Sucrose Gradient
<ol>
	<li>Pipet exactly 3.5 ml of 1.2 M HEPES-buffered sucrose solution and exactly 3.0 ml each of 1.0 M and 0.8 M HEPES-buffered sucrose solution into separate 6 ml snap cap tubes. NOTE: This is done to pre-measure the volumes of the buffers that are used to make the sucrose gradient. Exact measurement of the volumes is important to ensure that the resulting gradients will be balanced for ultracentrifugation.</li>
	<li>Using a glass Pasteur pipet and bulb, transfer 1.2 M HEPES-buffered sucrose solution to the 12 ml polyallomer ultracentrifuge tube. Carefully layer the 1.0 M HEPES-buffered sucrose solution on top of the 1.2 M HEPES-buffered sucrose solution. Finally, layer the 0.8 M HEPES-buffered sucrose solution on top of the 1.0 M HEPES-buffered sucrose solution. Use a fresh Pasteur pipet for each sucrose solution and take care not to disturb the sucrose gradient. Vibrations from a bench vortex or microcentrifuge will disturb the integrity of the gradient.</li>
	<li>Using a Pasteur pipet, layer the resuspended synaptosomal membrane fraction (P3) on top of the prepared discontinuous sucrose gradient. Ensure that the gradients are balanced by weighing them inside the bucket of the swinging bucket rotor. If the buckets are not balanced, use a P200 pipet to carefully add 0.32 M HEPES-buffered sucrose to the top of the gradient and balance the rotor buckets.</li>
</ol>
9. Fractionation of the Synaptic Plasma Membrane (SPM)
<ol>
	<li>Ultracentrifuge in a swinging bucket rotor at 150,000 x&#160;g for 2 hr at 4 &#176;C. Carefully remove the sucrose gradients from the ultracentrifuge buckets. Using an 18 G needle and a 1 ml syringe, puncture the tube at the bottom of the 1.0 M / 1.2 M HEPES-buffered sucrose solution interphase and withdraw the synaptic plasma membrane layer (SPM).</li>
	<li>Make note of the volume that each SPM sample occupies in the 1 ml syringe. Place the collected SPM layer in 3.5 ml thick-wall ultracentrifuge tubes, add exactly 2.5 volumes of 4 mM HEPES to adjust the sucrose concentration from 1.2 M to 0.32 M. Once each sample has been adjusted to 0.32 M sucrose, balance the tubes with 0.32 M HEPES-buffered sucrose solution.</li>
	<li>Ultracentrifuge in a fixed angle rotor at 200,000 x&#160;g for 30 min at 4 &#176;C. Remove and discard the supernatant. Resuspend the synaptic plasma membrane pellet (SPM) in 300 &#956;l of 50 mM HEPES / 2 mM EDTA solution. Store samples at -80 &#176;C until used for western blotting.</li>
</ol>
10. Preparation of the Postsynaptic Density Fraction (PSD)
<ol>
	<li>Thaw SPM samples on ice.</li>
	<li>Make a solution of 0.54% Triton X-100 in 50 mM HEPES&#160;/&#160;2 mM EDTA solution.</li>
	<li>In a 5 ml polystyrene tube, combine 2.7 ml of Triton X-100/HEPES/EDTA solution and 300 &#956;l SPM fraction and rotate the sample for 15 min at 4 &#176;C</li>
	<li>Transfer samples to 3.5 ml thick-wall ultracentrifuge tubes and centrifuge samples at 32,000 x&#160;g for 20 min at 4 &#176;C.</li>
	<li>Reserve the supernatant (Triton X-100 soluble fraction [TS]) and store at -80&#160;&#176;C. NOTE: At this stage, the pellet represents the &#8220;PSD-1T&#8221; fraction. A second treatment with detergent will produce a &#8220;PSD-2T&#8221; fraction. If a PSD-1T fraction is desired, proceed to step 10.6.
	<ol>
		<li>To produce a PSD-2T fraction, resuspend the pellet in 3 ml of 0.5% Triton X-100 in 50 mM HEPES&#160;/ 2 mM EDTA solution. Rotate samples for 15 min at 4 &#176;C. Centrifuge samples at 32,000 x&#160;g for 20 min at 4 &#176;C and proceed to step 10.6.</li>
	</ol>
	</li>
	<li>Discard the supernatant and resuspend the postsynaptic (PSD) pellet in 50 mM HEPES / 2 mM EDTA solution. NOTE: The resuspension volume may depend on the size of the pellet; a volume of 50-75 &#956;l is recommended. Because the PSD fraction is a detergent-insoluble fraction, it is often necessary to add 1-2 &#956;l of 0.5% SDS to enable complete resuspension of protein. Store samples at -80 &#176;C until used for western blotting.</li>
</ol>

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H., Fletcher, E. L., Garner, C. C., Gundelfinger, E. D., Wassle, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+expression+of+the+presynaptic+cytomatrix+protein+bassoon+among+ribbon+synapses+in+the+mammalian+retina.">Differential expression of the presynaptic cytomatrix protein bassoon among ribbon synapses in the mammalian retina.</a> Eur J Neurosci. 11, 3683-3693 (1999).</li></ol>

The authors have no conflicts of interest to disclose.

There are several steps in the procedure that are critical for a successful outcome. In step 3, it is important that a consistent degree of homogenization is achieved for each sample. When homogenizing tissue with the motor driven homogenizer, a constant speed is used not only for the rotation of the pestle, but also with the number of strokes. The incubation time during the osmotic shock should be precise, since extended homogenization or incubation in the hypotonic solution will lyse mitochondria and contaminate the SP...

Neurons communicate through synapses, and the quality of this communication is regulated to a large extent by alterations in the composition of proteins at the synapse. In particular, the proteins located in the post-synaptic density participate in neuronal communication by intimately scaffolding neurotransmitter receptors with their signal transduction systems1. Furthermore, lasting changes in the strength of synaptic efficacy are controlled by the addition or removal of receptors at the post-synaptic density1-6. Therefore, the isolation and quantification of synaptic proteins is a necessary and useful technique to gain insight into the ways that neurons respond to stimuli and alter synaptic efficacy7. This article describes a common technique to isolate synaptic proteins from rodent brain tissue by ultracentrifugation on discontinuous sucrose gradients. The synaptic plasma membrane fraction can be enriched and isolated based on its density in sucrose, which has been empirically determined to be similar to 1.2 M sucrose.
Depending on the biological question, subcellular fractions can be separated by continuous or discontinuous gradients of either sucrose or Percoll. Continuous gradients allow for the separation of proteins into multiple fractions; this can be particularly useful to demonstrate the co-localization of proteins within a given fraction8. However, the preparation of continuous gradients is more laborious and is unnecessary for many applications. Discontinuous gradients are comparatively easier to prepare and can be used to separate proteins into a few, generally-defined fractions. Discontinuous gradients that are composed of three sucrose layers of increasing molarity have been widely used to isolate proteins associated with the synaptic plasma membrane (SPM). This synaptic plasma membrane fraction can be further processed to the post-synaptic density fraction (PSD) by detergent treatment and isolation of the detergent-insoluble fraction.
When this process was first described in the 1960&#8217;s9,10, electron microscopy was used to demonstrate the organelles and membranes that roughly define the synaptic plasma membrane and post-synaptic density fractions9-14. These studies demonstrated the inclusion of pre- and post-synaptic membranes and synaptic vesicles in the SPM fraction; after detergent treatment primarily the electron-dense, post-synaptic densities were visible. In the procedure, a hypotonic shock is used to pinch off the synaptic processes from the cell body10. This step takes advantage of the fact that mitochondria are more resistant to osmotic shock and remain intact, and so they sediment at the bottom of the sucrose gradient (Figure 1).
Using this same enrichment technique, the SPM and PSD fractions were first biochemically defined by polyacrylamide gel electrophoresis and sequencing of the major protein components15-17. Subsequently western blot analysis has been used to detect and quantify the levels of synaptic proteins and further define these fractions (Figure 2). We have used this technique in our laboratories to quantify changes in the synaptic levels of the dopamine transporter that occur when the Slc6a3 locus is duplicated in mice18. We have also used this technique in NMDA receptor deficient mice to uncover synapse-specific reductions in proteins that are part of the DISC1 interactome19.
It is evident from western blot analysis that SPM fractions contain synaptic vesicle membrane proteins, endosome markers, mitochondrial proteins, membrane associated synthetic enzymes and signal transduction molecules, as well as integral components of the post-synaptic density and synaptic plasma membranes20-23. Even PSD fractions can have contamination with abundant mitochondrial proteins and it may be necessary to perform a second gradient sedimentation or additional purification steps to remove them13. Recently, quantitative mass spectrometry has provided a list of over 100 proteins in the post-synaptic density alone, as well as an indication of the relative abundance of these components24,25.

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preparation of synaptic plasma membrane and postsynaptic density proteins using a discontinuous sucrose gradient

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in the late 1960&#8217;s, proteins associated with the synaptic plasma membrane can be isolated by ultracentrifugation on a sucrose density gradient. Once synaptic membranes are isolated, the macromolecular complex known as the post-synaptic density can be subsequently isolated due to its detergent insolubility. The techniques used to isolate synaptic plasma membranes and post-synaptic density proteins remain essentially the same after 40 years, and are widely used in current neuroscience research. This article details the fractionation of proteins associated with the synaptic plasma membrane and post-synaptic density using a discontinuous sucrose gradient. Resulting protein preparations are suitable for western blotting or 2D DIGE analysis.

The preparation of the sucrose density gradient should result in a clear separation of the three molar solutions of sucrose (0.8, 1.0, and 1.2 M sucrose). See Figure 3A for an example of the gradient before the protein sample is added. If the gradient is prepared too much in advance, or if it is prepared on a bench surface with vibration from other equipment, the gradient will be compromised and proper separation will not be achieved. If a clear separation of the three solutions is not visible, it is adv...

Watch this Scientific Journal Video about Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient at JoVE.com

Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient

This article details the enrichment of proteins associated with the synaptic plasma membrane by ultracentrifugation on a discontinuous sucrose gradient. The subsequent preparation of post-synaptic density proteins is also described. Protein preparations are suitable for western blotting or 2D DIGE analysis.

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in ...

preparation-synaptic-plasma-membrane-postsynaptic-density-proteins

Research

JoVE Journal

Neuroscience

31.1K Views.  University of Toronto.  В данной статье подробно обогащение белков, ассоциированных с плазматической мембраны синаптической с помощью ультрацентрифугирования на разрывной градиенте сахарозы. Последующая подготовка постсинаптических белков плотности также описывается. Белковы?...

Video: Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Membrane Potentials, Synaptic Responses, Neuronal Circuitry, Neuromodulation and Muscle Histology Using the Crayfish: Student Laboratory Exercises

The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that influence a cell's membrane potential and which we address in these experiments are: (1) Ion concentration of K+ on the outside of the membrane, and (2) the permeability of the membrane to specific ions. The crayfish abdominal extensor muscles are in groupings with some being tonic (slow) and others phasic (fast) in their biochemical and physiological phenotypes, as well as in their structure; the motor neurons that innervate these muscles are correspondingly different in functional characteristics. We use these muscles as well as the superficial, tonic abdominal flexor muscle to demonstrate properties in synaptic transmission. In addition, we introduce a sensory-CNS-motor neuron-muscle circuit to demonstrate the effect of cuticular sensory stimulation as well as the influence of neuromodulators on certain aspects of the circuit. With the techniques obtained in this exercise, one can begin to answer many questions remaining in other experimental preparations as well as in physiological applications related to medicine and health. We have demonstrated the usefulness of model invertebrate preparations to address fundamental questions pertinent to all animals.

The experiments demonstrate an easy approach for students to gain experience in examining muscle structure, synaptic responses, the effects of ion gradients and permeability on membrane potentials. Also, a sensory-CNS-motor-muscle circuit is presented to show a means to test effects of compounds on a neuronal circuit.

The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that ...

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 ...

Preparation of Synaptoneurosomes from Mouse Cortex using a Discontinuous Percoll-Sucrose Density Gradient

Synaptoneurosomes (SNs) are obtained after homogenization and fractionation of mouse brain cortex. They are resealed vesicles or isolated terminals that break away from axon terminals when the cortical tissue is homogenized. The SNs retain pre- and postsynaptic characteristics, which makes them useful in the study of synaptic transmission. They retain the molecular machinery used in neuronal signaling and are capable of uptake, storage, and release of neurotransmitters.
The production and isolation of active SNs can be problematic using medias like Ficoll, which can be cytotoxic and require extended centrifugation due to high density, and filtration and centrifugation methods, which can result in low activity due to mechanical damage of the SNs. However, the use of discontinuous Percoll-sucrose density gradients to isolate SNs provides a rapid method to produce good yields of translationally active SNs. The Percoll-sucrose gradient method is quick and gentle as it employs isotonic conditions, has fewer and shorter centrifugation spins and avoids centrifugation steps that pellet SNs and cause mechanical damage.

A method to prepare translationally active, intact synaptoneurosomes (SNs) from mouse brain cortex is described. The method uses a discontinuous Percoll-sucrose density gradient allowing for the quick preparation of active SNs.

Synaptoneurosomes (SNs) are obtained after homogenization and fractionation of mouse brain cortex. They are resealed vesicles or isolated terminals that ...

A General Method for Evaluating Incubation of Sucrose Craving in Rats

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and relapse to food taking 1. Food craving conceptualized in this way is akin to drug craving in response to drug-paired cues. A rich literature has been developed around understanding the behavioral and neurobiological determinants of drug craving; we and others have been focusing recently on translating techniques from basic addiction research to better understand addiction-like behaviors related to food 2-4.
As done in previous studies of drug craving, we examine sucrose craving behavior by utilizing a rat model of relapse. In this model, rats self-administer either drug or food in sessions over several days. In a session, lever responding delivers the reward along with a tone+light stimulus. Craving behavior is then operationally defined as responding in a subsequent session where the reward is not available. Rats will reliably respond for the tone+light stimulus, likely due to its acquired conditioned reinforcing properties 5. This behavior is sometimes referred to as sucrose seeking or cue reactivity. In the present discussion we will use the term "sucrose craving" to subsume both of these constructs.
In the past decade, we have focused on how the length of time following reward self-administration influences reward craving. Interestingly, rats increase responding for the reward-paired cue over the course of several weeks of a period of forced-abstinence. This "incubation of craving" is observed in rats that have self-administered either food or drugs of abuse 4,6. This time-dependent increase in craving we have identified in the animal model may have great potential relevance to human drug and food addiction behaviors.
Here we present a protocol for assessing incubation of sucrose craving in rats. Variants of the procedure will be indicated where craving is assessed as responding for a discrete sucrose-paired cue following extinction of lever pressing within the sucrose self-administration context (Extinction without cues) or as responding for sucrose-paired cues in a general extinction context (Extinction with cues).

Responding for food or drug-paired cues increases over the course of a period of abstinence and this may relate to an increased susceptibility to relapse behaviors. Here we detail a procedure for evaluating this "incubation of craving" in rats that have self-administered sucrose.

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and ...

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 ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

The Neuromuscular Junction: Measuring Synapse Size, Fragmentation and Changes in Synaptic Protein Density Using Confocal Fluorescence Microscopy

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. Structural changes at the NMJ can result in neurotransmission failure, resulting in weakness, atrophy and even death of the muscle fiber. Many studies have investigated how genetic modifications or disease can alter the structure of the mouse NMJ. Unfortunately, it can be difficult to directly compare findings from these studies because they often employed different parameters and analytical methods. Three protocols are described here. The first uses maximum intensity projection confocal images to measure the area of acetylcholine receptor (AChR)-rich postsynaptic membrane domains at the endplate and the area of synaptic vesicle staining in the overlying presynaptic nerve terminal. The second protocol compares the relative intensities of immunostaining for synaptic proteins in the postsynaptic membrane. The third protocol uses Fluorescence Resonance Energy Transfer (FRET) to detect changes in the packing of postsynaptic AChRs at the endplate. The protocols have been developed and refined over a series of studies. Factors that influence the quality and consistency of results are discussed and normative data are provided for NMJs in healthy young adult mice.

The neuromuscular junction (NMJ) is altered in a variety of conditions that can sometimes culminate in synaptic failure. This report describes fluorescence microscope-based methods to quantify such structural changes.

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.