Name: measuring synaptic vesicle endocytosis in cultured hippocampal neurons
Uploaded: 2017-09-04T12:30:00.000+00:00
Duration: 7 min 30 s
Description: Watch this Scientific Journal Video about Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons at JoVE.com

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&amp;deg;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&amp;prime;-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

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Neuroscience

9.8K Views.  National Institute of Neurological Disorders and Stroke. Synaptic vesikel endocytose is gedetecteerd door de lichte microscopie van pHluorin gefuseerd met synaptic vesikel eiwit en elektronenmicroscopie van vesikel opname.

Video: Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

Biology

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

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 ...

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 ...

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

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 ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...

Transmission Electron Microscopic Imaging of Synaptic Vesicle Endocytosis in Mouse Hippocampal Neurons 

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/55862/measuring-synaptic-vesicle-endocytosis-in-cultured-hippocampal-neurons'>Source: Villarreal, S., et. al., Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons. J. Vis. Exp. (2017)</a>This video demonstrates the analysis of synaptic vesicle endocytosis in cultured hippocampal neurons through horseradish peroxidase labeling and electron microscopy.

Source: Villarreal, S., et. al., Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons. J. Vis. Exp. (2017)This video demonstrates the ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

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