We thank Dr. Yong-Ling Zhu for providing synaptophysin-pHluorin2x construct, and Dr. James E. Rothman for providing VAMP2-phluorin. We thank Dr. Susan Cheng and Virginia Crocker of NINDS Electron Microscopy Facility for their technical support and help. This work was supported by the National Institute of Neurological Disorders and Stroke Intramural Research Program in USA and a grant from the KRIBB Research Initiative Program (Korean Biomedical Scientist Fellowship Program), Korea Research Institute of Bioscience and Biotechnology, Republic of Korea.

NOTE: The following protocol describes the pHluorin imaging methods and EM methods used in cultured hippocampal neurons. pHluorin monitors synaptic vesicle protein uptake in living cells and EM detects uptake of synaptic vesicle and ultrastructural changes.
Animal care and procedure followed NIH guidelines and were approved by the NIH Animal Care and Use Committee.
1. pHluorin Imaging
<ol>
	<li>Hippocampal neuron culture
	<ol>
		<li>Prepare Hippocampus Buffer (HB) by combining 4 mM NaHCO3 and 5 mM HEPES and adjust to pH 7.3 with 5 M NaOH. Make the culture medium by mixing neurobasal medium, 2% B27, 0.5 mM L-glutamine and 1% penicillin-streptomycin. Additionally, prepare a mixture of HB buffer with 20% Fetal Bovine Serum (HB/20% FBS). 
		NOTE: This culturing protocol is based on Sankaranarayanan, et al.29 and Wu, et al.24</li>
		<li>Decapitate mouse pups between postnatal day 0 to day 2 into culture medium, and extract the brain into 4 &#176;C HB/20% FBS. Remove the brainstem and thalamus to expose the hippocampi. Dissect out the hippocampi, after exposing it by removal of the brainstem and thalamus, and transfer to fresh 4 &#176;C HB/20% FBS. 
		NOTE: Typically, yield is around 4 x 105 cells/mL for one pup (two hippocampi). Clean off the adhering membranes using tweezers or scissors, then transfer to fresh 4 &#176;C HB/20% FBS. Unroll the dentate gyrus and isolate the subiculum by cutting with scissors, then transfer to fresh 4 &#176;C HB/20% FBS.</li>
		<li>Divide each hippocampus, by cutting from end to end, into about 10 slices and transfer them to a 15 mL polypropylene conical tube.</li>
		<li>After allowing the tissue to settle, wash with 10 mL of HB/20% FBS, and then wash three times with 10 mL of HB.</li>
		<li>Prepare digestion solution of 137 mM NaCl, 5 mM KCl, 7 mM Na2HPO4, and 25 mM HEPES, and adjust to pH 7.2 with 5 M NaOH. Remove the supernatant from the hippocampi. Add 10 mg of trypsin and 1 mg of DNase to 2 mL of digestion solution, and filter through a sterile 0.22 &#181;m membrane directly onto the sample pellet.</li>
		<li>Incubate the hippocampi for 5 min at 37 &#176;C, then wash twice with 10 mL of HB/20% FBS, and then wash once with 10 mL of HB.</li>
		<li>Dissociate the cells with 6 mg of MgSO4&#8729;7H2O and 1 mg of DNase to 2 mL HB and sterile filter onto the hippocampi pellet through a sterile 0.22 &#181;m filter. Gently dissociate the cells by careful triturating, while taking care to avoid introducing air bubbles. Allow tissue particulates to settle for 2 min, and then slowly transfer the supernatant to another tube.</li>
		<li>Add 3 mL HB/20% FBS to the cell suspension, and centrifuge or 10 min at 4 &#176;C and 1,000 rpm. Discard the supernatant and resuspend in culture medium.</li>
		<li>Plate 60,000 cells suspended in 150 &#181;L culture medium on a Poly-D-Lysine coated 25-mm diameter coverslip, without spill off the coverslip. Add 2 mL of pre-warmed culture medium 2 h after plating. Coverslips are maintained in 6-well culturing plates or sterile Petri dishes.</li>
		<li>Maintain cells at 37 &#176;C in a 5% CO2 humidified incubator in culture medium for 14-21 days prior to recording. During the culture growth, change the top half of the medium twice a week.</li>
	</ol>
	</li>
	<li>Transfection
	<ol>
		<li>6-7 days after plating, transfect synaptophysin-pHluorin 2X (SpH) or VAMP2-pHluorin into hippocampal neurons. For SpH, use a Cytomegalovirus (CMV) promoter, inserted into a pcDNA3 vector30. For VAMP2-pHluorin, use a CMV promoter inserted into a pCI vector31.</li>
		<li>Transfect either vector by lipid carrier into target cells, using 1 &#181;g of plasmid. Use culture medium from protocol step 1.1.1, which lacks serum, for the transfection. Change the medium 2 h after transfection to reduce toxicity. 
		NOTE: In case of low expression of SpH in the boutons, increase the DNA concentration to 2 &#181;g or the incubation time with the lipid carrier, unless the cells are unhealthy. Typically, 4-10 cells (0.006-0.008%) of neurons were transfected.</li>
	</ol>
	</li>
	<li>Light microscopy
	<ol>
		<li>Prepare normal saline solution composed of 119 mM NaCl, 2 mM CaCl2, 2.5 mM KCl, 25 mM HEPES (pH 7.4), 30 mM glucose, 2 mM MgCl2, 0.01 mM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), and 0.05 mM DL-2-amino-5-phosphonovaleric acid (AP5). Take a coverslip from the culture medium plate and place on an imaging chamber that allows field stimulation, using lubricant and sealant to avoid leaks. Avoid letting the glass dry during the transfer of a coverslip from the plate to the chamber by immediately adding 750 &#181;L normal saline solution. 
		NOTE: CNQX and AP5 were used to block postsynaptic activity, which has the potential for recurrent activity. Before placing a chamber on an inverted fluorescence microscope, use cleaning tissue to confirm that the chamber does not leak. Slow leaking causes focus changes during the recording by the mixing of normal saline and immersion oil, which results in changes in the refractive index.</li>
		<li>Stimulation and recording
		<ol>
			<li>Image on an inverted widefield microscope with a 60X (1.4 numerical aperture) oil immersion lens with a metal halide lamp. Visualize pHluorin with a filter set for an excitation peak of 480 nm, a 490 nm long pass mirror, a 500-550 nm emission filter, and a manual flip shutter. Capture images every 100 ms, with 2 x 2 binning, using an Electron Multiplying Charge Coupled Device (EMCCD) camera. 
			NOTE: Several features should be considered to achieve the proper conditions to avoid photobleaching, including the image capture interval, and filter and binning, which are dependent on equipment. In the case of confocal imaging, laser power and exposure time should be considered to avoid photobleaching. The setup was determined after recording at least 3 min without stimulus and the recording was performed for at least 10 s without stimulus to check for photobleaching.</li>
			<li>Choose an area with a high density of boutons for ease of analysis in each experiment. Identify transfected cells by their green fluorescence signal, with emphasis on round or oval expression patterns in the bouton and continuous expression between boutons.</li>
			<li>Induce endocytosis by field stimulation using a 1 ms pulse, 20 mA action potential (AP) supplied from a pulse stimulator and delivered through a platinum electrode within the stimulus isolation unit. Image fluorescence activity over the course of the stimulation and during cell recovery. 
			NOTE: In the case of dead cells, expression was stronger than living boutons and showed a patchy pattern between boutons.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Image analysis
	<ol>
		<li>For analyzing the fluorescence intensity in a single bouton, set up a Region of Interest (ROI) as a 1.5 x 1.5 &#181;m square; the size of a bouton is within about 1.5 &#181;m. Use traces prior to stimulation to check for photobleaching and subtract as background.</li>
		<li>Normalize fluorescence change (&#916;F) with the equation: <img alt="Equation" height="11" src="/files/ftp_upload/55862/55862eq1.jpg" width="172" /> 
		where Fmax and F0 refer to maximal increase after stimulation and baseline fluorescence, respectively. Measure endocytosis rates as the rate of decay during the first 4-10 s after the maximum point of pHluorin fluorescence. Obtain the time constant (&#964;) of endocytosis by fitting the pHluorin fluorescence decay from the peak increase to the baseline with a mono-exponential function.</li>
	</ol>
	</li>
</ol>
2. Electron Microscopy
<ol>
	<li>Prepare a Poly-D-Lysine coated 6-well plate by applying 1.5 mL of 0.01% sterile-filtered Poly-D-Lysine solution to each well for 1 h at room temperature, then washing three times with sterilized water. Dissect, culture, and maintain hippocampal neurons as in steps 1.1.1, 1.1.2, and 1.1.3, respectively.</li>
	<li>Prepare a high K+ stimulation solution with HRP as 31.5 mM NaCl, 2 mM CaCl2, 90 mM KCl, 25 mM HEPES (pH 7.4), 30 mM glucose, 2 mM MgCl2, 0.01 mM CNQX, 0.05 mM AP5, and 5 mg/mL HRP, then adjust to pH 7.4 with 5 M NaOH.
	<ol>
		<li>Stimulate the hippocampal neuron culture with 1.5 mL high K+ stimulation solution at room temperature (referred to as K+) by addition of 1.5 mL to each well for 90 s. In the resting condition (referred to as R), apply the same concentration of 5 mg/mL HRP for 90 s, but with normal saline solution. For the recovery sample, apply high K+ stimulation solution as with the K+ sample, then rapidly wash and replace with normal saline and incubate for 10 min.</li>
	</ol>
	</li>
	<li>Fixation and staining
	<ol>
		<li>Prepare 0.1 M Na cacodylate buffer using 21.4 g/L Na cacodylate at pH 7.4. Fix cells with 4% glutaraldehyde in 0.1 M Na cacodylate buffer for at least 1 h at room temperature. Wash three times with 0.1 M Na cacodylate buffer for 7 min each.</li>
		<li>Prepare Diaminobenzidine (DAB) solution, comprised of 0.5 mg/mL of DAB with 0.3% H2O2 in ddH2O, and filter with a 0.22 &#181;m filter. Apply 1.5 mL DAB solution for 30 min at 37 &#176;C. Wash three times with 0.1 M Na cacodylate buffer for 7 min each. 
		CAUTION: DAB is a toxic and suspected carcinogen. Please use gloves and lab coats. 
		NOTE: Labeling with DAB occurs due to its oxidation by H2O2, as catalyzed by HRP. Small increases in the components result in an increased signal in the sample. Increasing the concentration of HRP speeds up the effect of the catalyst. Sufficiently large concentrations of H2O2 allow for debilitating side reactions with HRP, inhibiting the effect of labeling32. In this work, the concentrations of this labeling system were chosen based on currently available research33,34.</li>
		<li>Incubate the neurons with 1.5 mL of 1% OsO4 in 0.1 M Na cacodylate buffer for 1 h at 4 &#176;C as post fixation. Wash three times with 1.5 mL of 0.1 M Na cacodylate buffer for 7 min each. 
		CAUTION: Due to the toxicity and reactivity of OsO4, keeping the sample on ice in a chemical hood is preferable in many cases to using a fridge for the incubation.</li>
		<li>Prepare 0.1 M sodium acetate buffer with 13.61 g/L sodium acetate and 11.43 mL/L glacial acetic acid at pH 5.0. Wash three times with 1.5 mL 0.1 M acetate buffer at pH 5.0 for 7 min each and incubate with 1.5 mL 1% uranyl acetate in 0.1 M acetate buffer at pH 5.0 for 1 h at 4 &#176;C. Wash three times with 1.5 mL 0.1 M acetate buffer for 7 min each.</li>
	</ol>
	</li>
	<li>Epoxy embedding
	<ol>
		<li>Dehydrate the neuron culture with single 1.5 mL washes of 50%, 70%, and 90% ethanol, for 7 min in each and then 3 washes of 1.5 mL 100% ethanol for 7 min each in a fume hood.</li>
		<li>Mix 485 mL/L bisphenol-A-(epichlorhydrin) epoxy resin, 160 mL/L dodecenyl succinic anhydride (DDSA), 340 mL/L Methyl-5-Norbornene-2,3-Dicarboxylic Anhydride (NMA), and 15 mL/L 2,4,6-tris(dimethylaminomethyl)phenol (DMP-30) to create the epoxy resin. Mix thoroughly, then store under vacuum to remove air bubbles. 
		NOTE: It is critical to remove air bubbles in the resin, especially those smaller than visible to the naked eye, because they can cause cavities in the resin during sectioning.</li>
		<li>Infiltrate the sample by replacing the ethanol with 50% epoxy resin in ethanol for 30 min at room temperature on a shaker, then 70% epoxy resin in ethanol for 30 min at room temperature on a shaker.</li>
		<li>Switch the epoxy resin solution with 100% epoxy resin and incubate for 10 min at 50 &#176;C. Perform two exchanges of fresh 100% epoxy resin with incubations for 1 h at 50 &#176;C. Add fresh 100% epoxy resin and allow to harden at 50 &#176;C overnight and then at 60 &#176;C for over 36 h to harden.</li>
	</ol>
	</li>
	<li>Remove each sample from the muti-well plate with a jeweler&#39;s handsaw. Select regions of interest, dense concentrations of cells, using an inverted light microscope, and then cut 70 to 80 nm blocks for sectioning by microtome. Mount the cut region in the microtome chuck and load the microtome. Position the chuck in the microtome and mount a diamond knife with the edge parallel to the surface of the block. Collect sections of 70 to 80 nm thickness directly onto individual grids.</li>
	<li>Dissolve uranyl acetate into water for a 1% solution by weight, and separately dissolve lead citrate into water for a 3% solution by weight. Counterstain the sections by submersion with 1% aqueous uranyl acetate for 15 min and then 3% aqueous lead citrate for 5 min to improve the contrast of the samples.</li>
	<li>EM imaging
	<ol>
		<li>Examine the sections with a transmission electron microscope and record images with a CCD digital camera at a primary magnification of 10,000-20,000X3.</li>
	</ol>
	</li>
	<li>Statistics
	<ol>
		<li>Perform a t-test to identify significant differences by comparing the mean and standard error of measurement (s.e.m.) between the control and experimental samples.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here we demonstrate two methods for monitoring synaptic vesicle endocytosis. In the first method, we monitored pHluorin fused with a synaptic vesicle protein in transfected neurons and subsequently electrically stimulated. Secondly, we used EM imaging of HRP uptake as induced by KCl. We used different stimuli for two reasons. First, high potassium application induces depolarization of all neurons in the culture. This facilitates EM examination, given that our EM methodology could not distinguish between non-stimulated and stimulated neurons. Second, ultra-structural morphological changes were more reliably observed after intense stimulation, such as the high potassium stimulation, whereas pHluorin fluorescence changes can be observed after a brief train of action potentials or even a single action potential.
Light microscopy imaging using pHluorin shows synaptic vesicle protein fusion and uptake with high temporal resolution in living cells. It can be used to study mechanisms that regulate the endocytosis time course and the size of synaptic vesicle pools. pHluorin imaging has not only been used for monitoring synaptic protein endocytosis, but also for monitoring evoked or spontaneous single vesicle release, short-term synaptic plasticity, and for the measurement of the release probability35. Using pHluorin has shown that the synaptic vesicular protein pools being released are not the same as those being retrieved36,37. Recently, pHluorin fused with synaptic proteins was used to detect the fate of newly fused vesicles38, and bulk endocytosis39. Therefore, pHluorin fused with a synaptic protein is a valuable tool for studying exocytosis and endocytosis in living synapses.
The limitation of this technique is that the decay of the fluorescence intensity reflects not only endosomes, but also vesicular re-acidification. These two components could potentially be separated by treatment with bafilomycin, an inhibitor of V-ATPase10,40. Moreover, the fluorescence increase during electrical stimulation reflects the net outcome of exocytosis and endocytosis. Although pHluorin imaging could not detect membrane invagination, this limitation can be overcome by other techniques, particularly EM.
In EM, strongly stimulated cells showed increased HRP-positive synaptic vesicles and bulk endosomes. These bulk endosomes gradually disappeared by then regenerating into synaptic vesicles24. This technique shows synaptic vesicle uptake by revealing the ultrastructure in the presynaptic terminal in both control and stimulated cells. A drawback to this technique is assessing synaptic vesicle uptake in fixed cells rather than living cells. Additionally, the possibility of a fixative induced artifact cannot be excluded. Known artifacts affecting the membrane include blebbing, membrane erosion, and protein changes41. These differences from the naturally occurring state of a cell are an inherent concern to all methods of visualizing cells, and particular attention was paid to the EM data presented to assess the possibility of such artifacts41,42,43. While this technique alone cannot show functional endocytosis of synaptic proteins, combining it with light microscopy, which can provide live images of the endocytosis process, creates a more complete view of this cellular mechanism.

Neurotransmitters are stored in synaptic vesicles and released by exocytosis. The synaptic vesicle membrane and protein are then internalized by endocytosis, and reused in the next round of exocytosis. Endocytosis of synaptic vesicles is important for maintaining synaptic vesicle pools and removes protruding vesicles from the plasma membrane. The pH-sensitive green fluorescent protein pHluorin, which is quenched in acidic circumstances and dequenched in neutral pH, has been used to measure endocytosis time courses in live cells1,2,3. The pHluorin protein is typically attached to the lumenal side of synaptic vesicle proteins, such as synaptophysin or VAMP2/synaptobrevin. At rest, pHluorin is quenched in the 5.5 pH lumen of synaptic vesicles. Vesicle fusion to the plasma membrane exposes the vesicular lumen to the extracellular solution where the pH is ~7.3, resulting in an increase in pHluorin fluorescence. After exocytosis, the increased fluorescence decays, due to endocytosis of synaptic vesicle proteins followed by vesicle re-acidification within those recovered vesicles. Although the decay reflects both endocytosis and vesicular re-acidification, it mostly reflects endocytosis, because re-acidification is faster than endocytosis in most conditions1,4. The time constant of re-acidification is 3-4 s or less5,6, which is generally faster than the 10 s or more required for vesicle endocytosis4,5. If experiments are needed to distinguish endocytosis from re-acidification, acid quenching experiments using the 4-Morpholineethanesulfonic acid (MES) solution (25 mM) with a pH of 5.5 can be used to determine whether synaptic vesicle proteins are retrieved from the plasma membrane via endocytosis1,3,4. Thus, the pHluorin fluorescence intensity increase reflects a balance of exo- and endocytosis, and the decrease after nerve stimulation specifically reflects endocytosis.
pHluorin imaging may be used not only to measure the time course of endocytosis, but also the size of synaptic vesicle pools7,8, and the probability of evoked release and spontaneous release9. Many factors and proteins involved in regulating endocytosis, such as calcium, soluble NSF-attachment protein receptor (SNARE) proteins, brain-derived neurotrophic factor(BDNF), and calcineurin have been identified using pHluorin imaging1,2,10,11,12,13,14,15,16. Moreover, release of neurotransmitter could be detected in not only primary neurons but in neuroblastoma cells with TIRFM17. Recently, pHluorin variants, dsRed, mOrange and pHTomato were developed for monitoring simultaneous recordings of multiple factors in a single synapse18,19. For example, pHTomato has been fused with synaptophysin and used with a genetically encoded calcium indicator (GCaMP5K) to monitor presynaptic vesicle fusion and Ca2+ influx in the postsynaptic compartment20. Therefore, pHluorin attached to synaptic proteins provides a useful method to analyze the relationship between endocytosis and exocytosis.
EM is another method commonly used to study endocytosis, due to the high spatial resolution that shows ultrastructural changes during endocytosis. Two general areas are the ability to visualize pathological changes within neuronal cells21 and track vesicle proteins22. In particular, the observation of synaptic vesicle uptake, membrane curvature coated by clathrin in the periactive zone, and endosomal structures are possible with EM3,23,24,25,26,27,28. While EM involves potential artifacts, such as fixative-induced malformations, that may affect endocytosis, and data analysis is labor intensive, the resolution provides an attractive opportunity to visualize cellular structure. Potential fixative problems and the limitation in EM temporal resolution can be overcome by high pressure freezing, providing a fast and non-chemical method of stabilizing the delicate structures present during endocytosis27.

<table>
	<tbody>
		<tr>
			<td>Lipofectamine LTX with Plus</td>
			<td>Thermo Fisher</td>
			<td>15338-100</td>
			<td>Transfection of plasmid DNA including synaptophysin or VAMP2-pHluorin</td>
		</tr>
		<tr>
			<td>neurobasal medium</td>
			<td>Thermo Fisher</td>
			<td>21103-049</td>
			<td>Growth medium for neuron, Warm up to 37&#176;C before use</td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Thermo Fisher</td>
			<td>17504-044</td>
			<td>Gradient for neuronal differentiation</td>
		</tr>
		<tr>
			<td>Glutamax</td>
			<td>Thermo Fisher</td>
			<td>35050-061</td>
			<td>Gradient for neuronal culture</td>
		</tr>
		<tr>
			<td>Poly-D-Lysine coated coverslip</td>
			<td>Neuvitro</td>
			<td>GG-25-pdl</td>
			<td>Substrate for neuronal growth and imaging of pHluorin</td>
		</tr>
		<tr>
			<td>Trypsin XI from bovine pancrease</td>
			<td>Sigma</td>
			<td>T1005</td>
			<td>Neuronal culture-digest hippocampal tissues</td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancreas</td>
			<td>Sigma</td>
			<td>D5025</td>
			<td>Neuronal culture-inhibits viscous cell suspension</td>
		</tr>
		<tr>
			<td>pulse stimulator</td>
			<td>A-M systems</td>
			<td>model 2100</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>Slotted bath with field stimulation</td>
			<td>Warner Instruments</td>
			<td>RC-21BRFS</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>stimulus isolation unit</td>
			<td>Warner Instruments</td>
			<td>SIU102</td>
			<td>Apply electrical stimulation</td>
		</tr>
		<tr>
			<td>lubricant</td>
			<td>Dow corning</td>
			<td>111</td>
			<td>pHluorin imaging-seal with coverslip and imaging chamber, avoid leak from chamber</td>
		</tr>
		<tr>
			<td>AP5</td>
			<td>Tocris</td>
			<td>3693</td>
			<td>Gradient for normal saline, selective NMDA receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td>
		</tr>
		<tr>
			<td>CNQX</td>
			<td>Tocris</td>
			<td>190</td>
			<td>Gradient for normal saline, competitive AMPA/kainate receptor antagonist, inhibit postsynaptic activity which have potential for recurrent activity</td>
		</tr>
		<tr>
			<td>Illuminator</td>
			<td>Nikon</td>
			<td>C-HGFI</td>
			<td>Metal halide light source for pHluorin</td>
		</tr>
		<tr>
			<td>EMCCD camera</td>
			<td>Andor</td>
			<td>iXon3</td>
			<td>pHluorin imaging, detect pHluorin fluorescence intensity</td>
		</tr>
		<tr>
			<td>Inverted microscopy</td>
			<td>Nikon</td>
			<td>Ti-E</td>
			<td>Imaging for synaptophysin or VAMP2 pHluorin transfected cells</td>
		</tr>
		<tr>
			<td>NIS-Elements AR</td>
			<td>Nikon</td>
			<td>NIS-Elements Advanced Research</td>
			<td>Software for imaging acquisition and analysis</td>
		</tr>
		<tr>
			<td>Igor Pro</td>
			<td>WaveMetrics</td>
			<td>Igor pro</td>
			<td>Software for imaging analysis and data presentation</td>
		</tr>
		<tr>
			<td>imaging chamber</td>
			<td>Warner Instruments</td>
			<td>RC21B</td>
			<td>pHluorin imaging, apply field stimulation on living cells</td>
		</tr>
		<tr>
			<td>poly-l-lysine</td>
			<td>Sigma</td>
			<td>P4832</td>
			<td>Electron microscopy, substrate for neuronal growth, apply on multiwell plate for 1 h at room temperature then wash with sterilized water 3 times</td>
		</tr>
		<tr>
			<td>Horseradish peroxidase(HRP)</td>
			<td>Sigma</td>
			<td>P6782</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles by catalyzing DAB in presence hydrogen peroxide, final concentration is 5 mg/mL in normal saline, make fresh before use</td>
		</tr>
		<tr>
			<td>Na cacodylate</td>
			<td>Electron Microscopy Sciences</td>
			<td>12300</td>
			<td>Electron microscopy, buffer for fixatives and washing, final concentration is 0.1 N</td>
		</tr>
		<tr>
			<td>3,3&#8242;-Diaminobenzidine(DAB)</td>
			<td>Sigma</td>
			<td>D8001</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles, substrate for HRP, final concentration is 0.5 mg/mL in DDW and filtered, make fresh before use</td>
		</tr>
		<tr>
			<td>Hydrogen peroxide solution</td>
			<td>Sigma</td>
			<td>H1009</td>
			<td>Electron microscopy, labeling of endocytosed synaptic vesicles by inducing HRP-DAB reaction, final concentration is 0.3% in DDW, make fresh before use</td>
		</tr>
		<tr>
			<td>glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16365</td>
			<td>Electron microscopy, fixatives, final concentration is 4% in Na-cacodylate buffer, make fresh before use, shake well before to use</td>
		</tr>
		<tr>
			<td>TEM</td>
			<td>JEOL</td>
			<td>200CX</td>
			<td>Electron microscopy, imaging of endocytosed vesicles and ultrastructural changes</td>
		</tr>
		<tr>
			<td>CCD digital camera</td>
			<td>AMT</td>
			<td>XR-100</td>
			<td>Electron microscopy, capturing images</td>
		</tr>
		<tr>
			<td>Lead citrate</td>
			<td>Leica microsystems</td>
			<td>16707235</td>
			<td>Electron microscopy, grid staining</td>
		</tr>
	</tbody>
</table>

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.

Using the lipid carrier method, SpH was expressed in hippocampal neurons, allowing for the identification of boutons (Figure 1a). Electrical stimulation of the cells induced exocytosis, and a corresponding increase in fluorescence intensity. The increase in fluorescence (&#916;F) was stopped by ending the stimulus (Figure 1b). The increased fluorescence was followed by a slow decrease, due to endocytosis. In the case of VAMP2-pHluorin, VAMP2 diffuses along the axon from a bouton after stimulus4. The raw data used arbitrary units, and was normalized to the baseline to obtain the rate of decay and &#964; (Figure 2a-c). Since the measurements were made from a normalized trace, the rate of decay reflects the initial decay of fluorescence in the percentage of the peak &#916;F per second (&#916;F/s). In our experimental conditions, endocytosis (usually longer than 10 s) was much slower than reacidification (~3-4 s)4,5. Thus, the fluorescence decay of pHluorin primarily reflects endocytosis4,5. In presynaptic boutons, &#916;F is larger than the axon except bouton and the beginning time-of-increase was matched with initiation of electrical stimulus. In axon except bouton, &#916;F was lower in comparison to regions with boutons and beginning time-of-increase was delayed compared to the initiation of electrical stimulus (Figure 2b). &#916;F induced in bouton and axon except bouton was 82.9 &#177; 9.0% (n = 5 experiments) and 23.2 &#177; 4.0% (n = 5 experiments), respectively (Figure 2b). &#916;F of SpH decayed mono-exponentially with a &#964; of 17.7 &#177; 0.3 (n = 5 experiments) and 20.7 &#177; 0.2 (n = 4 experiments) in 50 AP and 200 AP, respectively (Figure 1b). In 200 AP, &#964; was not significantly different before or after normalizing with the baseline. In the case of VAMP2-pHluorin transfected cells, the expression level of VAMP2-pHluorin was higher than SpH (Figure 3a-c). Traces of VAMP2-pHluorin were also obtained by normalizing to the baseline (Figure 3b).
EM was performed in cultured neurons and the clathrin coated vesicles were examined in comparison to control samples, which were incubated with 5 mg/mL of HRP for 90 s in absence of the stimulus (Figure 4a-b). The clathrin coated pits were observed at a probability of 0.05 per bouton in the absence of stimulus. Stimulation with high K+ induced bulk endosome and synaptic vesicles uptake, which were identified by HRP labeling (Figure 5a). Synaptic vesicles were defined as vesicles with a diameter less than 50 nm and bulk endosomes were defined as having a diameter over 80 nm or with a cross sectional area of more than an 80 nm vesicle. Induced bulk endosome uptake was decreased by recovery with normal saline (Figure 5b).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55862/55862fig1.jpg" /> 
Figure 1: Images of SpH transfected neurons and traces of SpH transfected cell responses to electrical stimulation. (a) A sample image of cultured hippocampal neurons transfected with SpH by liposome delivery. SpH was highly expressed in boutons, presenting circle or oval patterns, with relatively lower expression along the axon. White boxes indicate ROI containing boutons (1.5 x 1.5 &#181;m). Scale bar: 20 &#181;m. (b) Responses of SpH transfected cells to an electrical stimulus of 50 AP at 20 Hz (n = 5 experiments) or 200 AP at 20 Hz (n = 4 experiments). Each experiment was based on 20-60 boutons. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55862/55862fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55862/55862fig2.jpg" /> 
Figure 2: Rate of decay and tau value for endocytosis. (a) Responses to 50 AP at 20 Hz were converted from arbitrary units to normalized traces (n = 5). Fluorescence change was normalized to the baseline. This figure has been modified from Wu et al., 20163. (b) Normalized responses to 200 AP at 20 Hz in boutons (black, n = 4 experiments, left panel) and axon except bouton (red, n = 4 experiments, left panel). Fluorescence change was normalized to the baseline. Dashed box is magnified in the right panel. (c) Using normalized baseline, the rate of decay or &#964; was calculated in 50 AP (n = 5 experiments) and 200 AP (n = 4 experiments) at 20 Hz. Before calculation, &#916;F was normalized to 100%. &#964; of endocytosis was obtained from normalized trace between the time of maximum fluorescence and the end of experiment. Rate of decay (mean + s.e.m., upper panel) and &#964; (mean + s.e.m., lower panel) induced by 50 AP and 200 AP.&#160; <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55862/55862fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/55862/55862fig3.jpg" /> 
Figure 3: Images of VAMP2-pHluorin transfected neurons and examples of traces. (a) A sample image of cultured hippocampal neurons transfected with VAMP2-pHluorin by liposome delivery. White boxes indicate the ROI containing boutons (1.5 x 1.5 &#181;m). Scale bar: 20 &#181;m. (b) Traces for VAMP2-pHluorin responses to 200 AP at 20 Hz (n = 4 experiments). Fluorescence change was normalized to the baseline. (c) Fbase (a.u.) (mean + s.e.m.) in VAMP2-pHluorin (n = 4 experiments) and SpH (n = 7 experiments) transfected boutons. **, p &#60;0.01. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55862/55862fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/55862/55862fig4.jpg" /> 
Figure 4: Images of EM in presynaptic terminal. (a) Synaptic vesicles were shown in cultured hippocampal neurons using EM. This figure has been modified from Wu et al., 20163. Scale bar = 200 nm. (b) Magnified image showing clathrin coated pits. Scale bar: 50 nm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55862/55862fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/55862/55862fig5.jpg" /> 
Figure 5: Images of presynaptic terminal in stimulated neurons with high potassium. (a) EM images of neurons. The cells were fixed without stimulus (R), immediately after stimulation with 90 mM KCl for 90 s containing soluble HRP (K+), or after stimulation and then incubated for 10 min in normal saline. This figure has been modified from Wu et al., 20163. Scale bar = 200 nm. (b) Compared to control, HRP (-) synaptic vesicles were decreased and HRP (+) vesicles were increased by KCl. Bulk endosome uptake was induced by KCl and gradually decreased by recovery (mean + s.e.m., each group was from 40-100 synaptic profiles). *p &#60;0.1; **p &#60;0.01, significant to resting; ##p &#60;0.01, significant to K+. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55862/55862fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

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

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

measuring-synaptic-vesicle-endocytosis-in-cultured-hippocampal-neurons

Nanyang Technological University

Research

JoVE Journal

Neuroscience

9.8K Views.  National Institute of Neurological Disorders and Stroke. Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.

Video: Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

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 ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

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

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

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 ...

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

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

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

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

Inducing Dendritic Growth in Cultured Sympathetic Neurons

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

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

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

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

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

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

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

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 ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

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 ...

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...