The authors would like to thank the Swedish Research Council (349-2007-8680) for funding and Dalsj&#246;fors K&#246;tt AB (Dalsj&#246;fors, Sweden) for donation of bovine adrenal glands.

1. Cell Culture of Bovine Chromaffin Cells Isolated by Enzymatic Digestion from Adrenal Glands
<ol>
	<li>To isolate chromaffin cells collected from bovine adrenal glands, sterilize cells with 70% ethanol solution. Trim away fat and connective tissue using a scalpel. To remove blood, rinse the adrenal veins using Locke&#39;s solution (NaCl 154 mM, KCl 5.6 mM, NaHCO3 3.6 mM, hydroxyethyl piperazineethanesulfonic acid (HEPES) 5.6 mM at pH 7.4) that has been thermostated to 37 &#176;C.</li>
	<li>To digest the tissue, inflate each gland by injecting approximately 2.5 mL of warm sterile-filtered 0.2% collagenase P solution through the adrenal vein using a syringe and incubate the tissue for 20 min at 37 &#176;C. Examine the glands to ensure the digestion process of the medulla is completed. The tissue should feel soft and not tense.</li>
	<li>Collect the digested medulla by snipping off the glands in the longitudinal direction. Mince the medulla tissue using a scalpel. Filter the tissue suspension over a steel sieve and dilute with Locke&#39;s solution to approximately a 50 mL volume to reduce the activity of collagenase P.</li>
	<li>Pellet the cells at 300 x g in a centrifuge for 10 min at room temperature and collect into 50 mL sterile test tubes. Re-suspend the obtained pellet in 20 mL Locke&#39;s solution and filter the solution over a sterile 100-&#181;m nylon mesh into tubes.</li>
	<li>Mix the chromaffin cell suspension with sterile Percoll (1:1) and centrifuge the cell solution at 18,600 x g for 20 min at room temperature. Collect the top layer of the density gradient and filter the solution over a 100-&#181;m nylon mesh into tubes.</li>
	<li>To exclude Percoll, dilute the cell suspension with Locke&#39;s solution and centrifuge at 300 x g for 10 min at room temperature before re-suspending the pellet in Locke&#39;s solution. The estimated cell density of the isolated cell suspension is approximately 4 million cells/mL.</li>
	<li>For amperometric recording of exocytosis and intracellular cytometry measurements, plate chromaffin cells in collagen IV coated 60 mm plastic dishes at a density of about 17.5 &#215; 103 cells/cm2 and incubate the cells at 37 &#176;C in a 5% CO2 environment. Perform the electrochemical experiments within 1-3 days of cell culture.</li>
	<li>For the TEM imaging experiments, plate chromaffin cells in 75 cm2 cell culture flasks at a density of 7-8 million cells per flask and incubate at 37 &#176;C in a 5% CO2 environment for 1 day.</li>
</ol>
2. Single Cell Exocytosis Amperometry Experiments24
<ol>
	<li>To fabricate carbon fiber microelectrodes for these experiments, take a glass capillary with an outer diameter suitable for the electrode holder at the head stage that will be used in these experiments. Use a borosilicate glass capillary with an outer diameter of 1.2 mm and an inner diameter of 0.69 mm.
	<ol>
		<li>Connect one of the capillary ends to a water aspiration tube. Place 5 &#181;m diameter carbon fibers on something, such as a piece of white paper, to enhance visualization.</li>
		<li>Identify a single carbon fiber and hold the fiber down on one end with a finger to keep the carbon fiber in place while positioning the open end of the capillary in proximity to the free end of the carbon fiber. Gently aspirate the carbon fiber into the glass capillary so that the carbon fiber sticks out through both ends of the capillary. Move away the aspiration force.</li>
	</ol>
	</li>
	<li>Place the capillary with the carbon fiber into a micropipette puller and pull the glass capillary into two separate glass pipette tips. To disconnect the carbon fiber that is connected between the two glass tips, use a pair of scissors to cut the carbon fiber and gain two carbon fiber microelectrodes.</li>
	<li>Under a microscope, place the carbon fiber on top of a thicker microscope slide that allows manual cutting of the carbon fiber at the edge of where the fiber is extending from the glass capillary coating using a scalpel.</li>
	<li>After cutting the carbon fiber, leaving a glass coated carbon fiber tip, insert a few mm of the electrode tip into an Epoxy solution for 10 min to pull up epoxy and seal any possible open space between the carbon fiber and the surrounding glass. Lift the electrodes slowly out of the epoxy solution to prevent bulky glue drops from forming at the tip of the electrode.</li>
	<li>Place the electrodes on a holder (e.g., a wooden flat stick) with two-side sticky heat resistant tape to which electrodes can be attached. Bake the epoxy treated electrodes overnight in an oven at 100 &#176;C. The electrode tips easily break if they come into contact with a surface, so ensure the electrodes always are safely stored using a holder where the electrodes are fixed in place and tips do not risk being touched.</li>
	<li>To obtain a flat disc electrode surface, place the electrode in the holder of a microgrinder and bevel each carbon fiber electrode at an angel of 45&#176;. After beveling, mark the capillary on its topside using a permanent marker to later know how to locate the disc electrode surface at a 45&#176;&#160;angle when placing the electrode near cells for the exocytosis measurement. 
	NOTE: This step is important as in these experiments, the oval disc electrode needs to be placed flat on top of cells with its surface parallel to the surface of the Petri dish. Also, beveling electrodes are done the same day as experiments to ensure a fresh and clean electrode surface.</li>
	<li>Before use, place each carbon fiber microelectrode in a test solution (e.g., 0.1 mM dopamine in PBS buffer (pH 7.4)) to monitor the steady state current using cyclic voltammetry. For a cyclic voltammetry scan, apply a triangle potential waveform from -0.2 V to +0.8 V vs a Ag/AgCl reference electrode at 100 mV/s to ensure good reaction kinetics data are obtained that are in agreement with theoretically calculated values for a 5 &#181;m diameter disc carbon fiber microelectrode25.</li>
	<li>For amperometry recording of exocytosis, place the Petri dish with cultured chromaffin cells onto an inverted microscope. It is important to shield the microscope set up with a Faraday cage to eliminate electronic noise during the amperometry recording, due to the very small currents being measured. Use a microscope heating stage to maintain a temperature of 37 &#176;C during the cell experiments .</li>
	<li>Use a low-noise patch clamp instrument to apply a constant potential of +700 mV at the working electrode versus a Ag/AgCl reference electrode that is also placed in the Petri dish away from the working electrode. For recording exocytosis of chromaffin cells, digitalize the signal at 10 kHz and apply an internal low pass Bessel filter at 2 kHz to filter the recorded signal.</li>
	<li>To perform amperometric recording at single cells, mount the freshly beveled and tested carbon fiber disc microelectrode in the electrode holder of the head stage that is used with the potentiostat.
	<ol>
		<li>Gently place the electrode with the flat oval disc shape electrode surface facing down towards the apical surface of the cell and place the electrode in contact with the cell membrane using a micromanipulator.</li>
		<li>Adjust the distance from the electrode to the cell by monitoring the cell deformation caused by the electrode upon placement on top of the cell membrane and then carefully retract the electrode to a distance where the cell regains a shape close to its original cell shape. The ideal for kinetic and quantitative recording is to create a thin liquid film of a hundred nanometers to separate the electrode and cell surface, which are similar conditions for postsynaptic detection of chemical release in a synapse.</li>
	</ol>
	</li>
	<li>To stimulate the cells to exocytosis, position a glass micropipette with a tip size of 2-3 &#181;m diameter, filled with 5 mM BaCl2 solution at a distance of at least 20 &#181;m away from the cell, to prevent any solution leaking from the pipette tip to influence the experiment and apply a 5 s injection pulse of 5 mM BaCl2 solution at the cell surface to stimulate the cell to exocytosis.</li>
	<li>To study the reversible effects of osmotic pressure on the exocytosis process, prepare an isotonic buffer solution (150 mM NaCl, 5 mM KCl, 1.2 mM MgCl2, 5 mM glucose, 10 mM HEPES, pH 7.4) with 310 mOsm/kg iso-osmotic pressure and a hypertonic buffer made by adjusting the NaCl concentration of the isotonic buffer solution with an osmolality corresponding to 730 mOsm/kg.</li>
	<li>Place the cells in the isotonic buffer and stimulate the cell to exocytosis by applying a barium injection pulse while recording the amperometric current transients during the initiated exocytosis activity for approximately 3 min.
	<ol>
		<li>To compare the exocytosis responses in isotonic conditions to hypertonic conditions, incubate the cells for 10 min in hypertonic buffer solution and thereafter apply a barium injection pulse to stimulate exocytosis and perform 3 min of amperometric recording.</li>
		<li>For the reversible response of cells, incubate cells again for 10 min in isotonic buffer solution and stimulate cells by applying a 5s barium injection pulse and perform 3 min of recording the exocytosis response from the cell.</li>
	</ol>
	</li>
	<li>As control experiments to determine the effect on exocytosis activity by multiple barium stimulations, perform an amperometric recording of exocytosis release from three consecutive BaCl2 stimulations on the same cell placed in isotonic buffer and using the same time protocol for the consecutive cell incubation experiments with different osmolarity.</li>
</ol>
3. Intracellular Electrochemical Cytometry24,26
<ol>
	<li>To fabricate electrodes for vesicle quantal size measurements, use the same materials and start preparing a 5 &#181;m in diameter carbon fiber electrode according to the descriptions in sections 2.1 and 2.2 of microelectrode fabrication for amperometric exocytosis measurements.</li>
	<li>Under a microscope, use a scalpel to cut the carbon fiber extending out of the glass tip so that a carbon fiber with a length ranging from 30 to 100 &#181;m is left sticking out from the glass tip.</li>
	<li>To prepare a flame etched tip of the carbon fiber electrode, use a butane flame. To achieve an evenly etched, cylindrical shaped electrode tip, hold the cylindrical shaped carbon fiber electrode while rotating it and place the carbon fiber extending from the glass into the blue edge of the butane flame until the tip of the carbon develops a red color. This often takes less than 2 s. If successful, this results in an etched electrode tip size of 50-100 nm in diameter (a SEM image of such tip is shown in Figure 5B). After flame etching, place the electrode under a microscope to evaluate the electrode tip.</li>
	<li>Insert the cylindrical nanotip microelectrode in epoxy solution for 3 min followed by a 15 s dipping of the electrode tip into an acetone solution. This allows epoxy to seal the potential gap space between the carbon fiber and the insulating glass capillary wall, while the acetone clears the epoxy off the etched carbon fiber electrode surface. To cure the epoxy, bake the electrodes in an oven overnight at 100 &#176;C.</li>
	<li>Before use, test the steady state current of each carbon fiber microelectrode, as explained in section 2.7 using cyclic voltammetry. For experiments, only use electrodes that display a plateau current around 1.5-2.5 nA27.</li>
	<li>For intercellular amperometry measurements, place the cells at the microscope as described in 2.8 and use the same experimental settings of the potentiostat as described in 2.9.</li>
	<li>To perform intracellular amperometric measurement, and to prevent significant physical damage to the cell, insert the nanotip cylindrical microelectrode into the cell by adding a gentle mechanical force, just enough to push the electrode through the cell plasma membrane and into the cell cytoplasm using a micromanipulator.</li>
	<li>After insertion, and with the cell membrane sealed around the cylindrical electrode, start the amperometric recording in situ at the live cell. At the oxidation potential that is applied to the electrode, vesicles will adsorb to the electrode surface and stochastically rupture.&#160; Hence, there is no need for any kind of stimulus to initiate this process.</li>
	<li>To determine the osmotic effect on vesicle quantal size, collect the intracellular cytometry measurements from a group of cells that have been incubated in isotonic and in hypertonic buffer using the experimental condition as described in section 2.13.</li>
</ol>
4. Data Analysis of Amperometry Recordings
<ol>
	<li>To analyze the recorded amperometry data from exocytosis and intracellular vesicle quantal size analysis, use a software program that allows analysis of current transients in the recorded current versus time trace so that kinetic peak parameters and the integrated total charge for individual current peaks can be determined. For data analysis, we have used software developed in the Sulzer lab that was written for the data analysis program Igor28.</li>
	<li>When analyzing the amperometric spikes, select a threshold limit for peaks of three times the root-mean square (RMS) standard deviation of the noise for each recording.</li>
	<li>Collect the amperometric spikes from each cell recording manually to prevent false spikes that do not follow the Gaussian shape, double spikes which can be the result of merged exocytosis events, and only accept Gaussian shaped spikes with a threshold of three times higher than the RMS noise.</li>
	<li>Collect the data for total charge per single amperometric peak and use Faradays law (N=QnF) to calculate the molar amount of catecholamine neurotransmitter detected per exocytosis or intracellular single vesicle rupture event, where N is the moles of neurotransmitters going through a redox reaction at the electrode surface, Q=the total charge detected under each spike, n=number of electrons transferred in the redox reaction (n=2 for catecholamines), and the Faraday&#180;s constant F=96485 C/mol.</li>
	<li>Perform statistical analysis of the data collected by first calculating the average charge detected per event for each cell. Then for variance between cells, calculate the average charge between cells and use a student&#39;s t-test between cells assuming unequal variance. This study used the MATLAB program for statistical analysis.</li>
</ol>
5. TEM Imaging for Vesicle Size Analysis
<ol>
	<li>To perform TEM imaging of cells at the different osmotic conditions, first incubate the cells in isotonic or hypertonic buffer for 10 min at 37 &#176;C in a 5% CO2 environment before chemical fixation of cells.</li>
	<li>Perform cell fixation using the Karnovsky fixative method29. Using this method, incubate the cells with a solution containing 0.01% sodium azide, 1% formaldehyde, and 1.25% glutaraldehyde in which the sample can be stored at 4 &#176;C.</li>
	<li>After fixation, wash the cells with 0.15 M sodium cacodylate buffer.</li>
	<li>To stain the cell samples post fixation, and in preparation for TEM imaging, incubate the cells with a 1% osmium tetroxide solution for 2 h at 4 &#176;C, followed by 1 h incubation in 0.5% uranyl acetate solution at room temperature in the dark.</li>
	<li>In a final fixation step, dehydrate the cells first by rinsing the cells in 100% ethanol and then rinse cells with acetone.</li>
	<li>Embed the cell samples in epoxyresin and thereafter perform centrifugation at 4,000 x g for 30-40 min and allow the cell sample to polymerize at 40 &#176;C for 15 h followed by 48 h incubation at 60 &#176;C.</li>
	<li>In preparation for imaging, section the cell sample embedded in Agar 100 resin into slices with thickness of 60 nm using an ultramicrotome.</li>
	<li>Subject the cells to a 4% uranyl acetate/25% ethanol solution for 4 min followed by 20 s of rinsing with water. Wash the cells with Reynolds lead citrate for 3 min, followed by 20 s of rinsing with water.</li>
	<li>Perform transmission electron microscopy imaging of cell samples. In our experiments, we have used a transmission electron microscope that has been operated at 120 kV.</li>
	<li>From each recorded image of a cell section illustrating the cell ultrastructure that displays a population of the vesicles in the cell cytoplasm, first calibrate the image pixel size by relating pixels to the recorded scale bar in each TEM image. Use imaging software to determine the vesicle sizes at each image of cells exposed to isotonic or hypertonic conditions. In our experiments, we used the software ImageJ for image analysis. It is worth noting that for image analysis of cellular ultrastructure from TEM images of cells, size adjustment of these measurements based on thickness of cell sections needs to be considered and have been previously described by Almers&#180;s lab30.</li>
</ol>

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V., Sulzer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+exocytotic+events+recorded+by+amperometry.">Analysis of exocytotic events recorded by amperometry.</a> Nat Methods. 2 (9), 651-658 (2005).</li><li>Morris, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+formaldehyde+glutaraldehyde+fixative+of+high+osmolality+for+use+in+electron+microscopy.">A formaldehyde glutaraldehyde fixative of high osmolality for use in electron microscopy.</a> J Cell Biol. 27, 137-139 (1965).</li><li>Parsons, T. D., Coorssen, J., Horstmann, H., Almers, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Docked+granules,+the+exocytic+burst,+and+the+need+for+ATP+hydrolysis+in+endocrine+cells.">Docked granules, the exocytic burst, and the need for ATP hydrolysis in endocrine cells.</a> Neuron. 15 (5), 1085-1096 (1995).</li></ol>

The authors have nothing to disclose.

We here present a protocol and the advantages of combining three complementary analytical methods to analyze secretory vesicles and the exocytosis process to gain a better understanding of how a physical force such as osmotic pressure can affect secretory vesicles and the exocytosis process in secretory cells. These methods include carbon fiber microelectrode amperometry, which is an established method for recording exocytosis activity, intracellular electrochemical cytometry, which is used to determine quantal size of v...

Chromaffin cells in adrenal glands are neuroendocrine cells that release neurotransmitter molecules into the blood stream. This occurs through a cellular process that involves the docking and fusion of neurotransmitter-filled vesicles, resulting in content release from vesicles to the extracellular space in a process called exocytosis. The neurotransmitters (adrenaline and noradrenaline) in chromaffin cells are actively transported by membrane proteins into large dense core vesicles (LDCVs) and stored at high concentrations (~0.5-1 M)1,2. Accumulation of neurotransmitters inside the LDCVs is accomplished by the affinity of catecholamine molecules to the intravesicular dense core protein matrix comprised of chromogranin proteins (~169 mg/mL)3,4,5,6, and an intravesicular cocktail solution containing key components for catecholamine loading and storage into the vesicle such as ATP (125-300 mM)7, Ca2+ (50-100 &#181;M in the solution and ~40 mM bound to the protein matrix)8, Mg2+ (5 mM)9, ascorbate (10-30 mM)10, and a pH of ~5.511,12. The LDCVs maintain an iso-osmotic condition with the cell cytoplasm (310 mOsm/kg)13, even though the theoretical solute concentration inside the vesicles sum up to more than 750 mM. The composition of the intravesicular components is not only essential for the loading and storage of catecholamines, but also for aggregation of solutes to the dense core protein matrix. This significantly reduces the osmolarity of the vesicles is significantly reduced and can affect the amount catecholamine that is released during exocytosis5,6.
Studies on the effect of extracellular osmotic pressure on the exocytosis process by amperometric recording have reported that high extracellular osmotic pressure inhibits exocytosis activity and reduces the number of neurotransmitters secreted from single vesicle compartments4,14,15,16,17,18,19. The explanations of these observations have speculated on the possible enhancement of macromolecular crowding in the cell cytoplasm inhibiting vesicle fusion events, and an alteration in membrane biophysical properties affecting the number of neurotransmitters released during exocytosis. These thoughts assumed that high extracellular osmotic stress does not affect the vesicle quantal size, which defines the number of neurotransmitter molecules stored in a vesicle compartment at prior stage of triggered exocytosis14,15,17,19,20,21. In amperometric measurements of exocytosis release in single cells, a carbon fiber disc microelectrode is placed in close contact with the cell surface, creating an experimental set up mimicking the synapse configuration, where the amperometric electrode serves as a postsynaptic detector (Figure 1)22,23. By stimulating a cell to exocytosis, one can induce neurotransmitter-filled vesicles to fuse with the cell plasma membrane and release part or the full vesicle content into the extracellular space. These neurotransmitter molecules released at the surface of the electrode can be detected electrochemically if the neurotransmitters are electroactive (e.g., catecholamines) by applying a redox potential of +700 mV vs a Ag/AgCl reference electrode. Consequently, a series of current spikes mark the detection of individual exocytosis events. From the current versus time trace in an amperometric recording, the area under each single amperometric spike represents the charge detected per exocytosis event and can be converted to the mole of neurotransmitters released, using the Faraday equation. Hence, the amperometric recordings provide quantitative information on the amount neurotransmitters expelled from single exocytosis events and report on the frequency of exocytosis events, but do not present quantitative information on the secretory vesicles content before vesicle fusion and neurotransmitter release has been initiated.
Therefore, to get a better understanding of how secretory vesicles in the cell cytoplasm respond to extracellular osmotic stress before the cell is triggered to undergo exocytosis, other complementary analytical methods can be used to enrich this information. For instance, to investigate if osmotic stress alters vesicle volume, transmission electron microscopy (TEM) imaging analysis can be used to measure the vesicle size of cells after chemical fixation. To examine if osmotic stress affects vesicle quantal size, a recently developed amperometric technique called intracellular electrochemical cytometry, can be applied for quantification of vesicle neurotransmitter content at secretory vesicles in their native state when still residing in the cytoplasm of live cells26. In the intracellular electrochemical cytometry technique, a nanotip cylindrical carbon fiber electrode is gently inserted into the cytoplasm of live cells and, by applying a +700 mV potential to this electrode (vs a Ag/AgCl reference electrode), the catecholamine content in vesicles can be quantified by the detection of a redox current spike from single vesicles colliding, adsorbing, and subsequently stochastically rupturing at the electrode surface and thereby releasing their contents against the electrode surface26. Hence, in the amperometric current versus time trace, each single vesicle rupturing event can result in a current transient and, by integrating the area of each current spike, the vesicle quantal size can be calculated using Faraday&#180;s law.
Therefore, by linking the quantitative information gained from vesicle size measurements using TEM imaging together with vesicle quantal size analysis, as recorded by intracellular electrochemical cytometry, vesicle neurotransmitter concentration can also be determined. This allows vesicle characterization when cells are exposed to different osmotic conditions and provides a better view on how vesicles may respond to extracellular osmotic stress at the stage prior to exocytosis. The results from combining these methods have shown that in the presence of extracellular high osmotic pressure, vesicles shrink and adjust their quantal size and comparing the quantitative information on the relative changes from these measurements shows that while shrinking, vesicles adjust their contents and size to maintain a constant neurotransmitter concentration24. Thus, this understanding is valuable in connecting to the observations made on neurotransmitter release in cells exposed to osmotic stress. In these protocols, we describe the use of these three complementary methodologies that allow the characterization of how secretory vesicles in their native environment respond to extracellular osmolality and the effects of this response on the exocytosis process. In addition to our previous observations regarding the effect of high osmotic pressure on exocytosis24, we present additional experiments that describe cell recovery after osmotic shock and the effect of multiple barium stimulations in chromaffin cells.

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monitoring the effect of osmotic stress on secretory vesicles and exocytosis

Amperometry recording of cells subjected to osmotic shock show that secretory cells respond to this physical stress by reducing the exocytosis activity and the amount of neurotransmitter released from vesicles in single exocytosis events. It has been suggested that the reduction in neurotransmitters expelled is due to alterations in membrane biophysical properties when cells shrink in response to osmotic stress and with assumptions made that secretory vesicles in the cell cytoplasm are not affected by extracellular osmotic stress. Amperometry recording of exocytosis monitors what is released from cells the moment a vesicle fuses with the plasma membrane, but does not provide information on the vesicle content before the vesicle fusion is triggered. Therefore, by combining amperometry recording with other complementary analytical methods that are capable of characterizing the secretory vesicles before exocytosis at cells is triggered offers a broader overview for examining how secretory vesicles and the exocytosis process are affected by osmotic shock. We here describe how complementing amperometry recording with intracellular electrochemical cytometry and transmission electron microscopy (TEM) imaging can be used to characterize alterations in secretory vesicles size and neurotransmitter content at chromaffin cells in relation to exocytosis activity before and after exposure to osmotic stress. By linking the quantitative information gained from experiments using all three analytical methods, conclusions were previously made that secretory vesicles respond to extracellular osmotic stress by shrinking in size and reducing the vesicle quantal size to maintain a constant vesicle neurotransmitter concentration. Hence, this gives some clarification regarding why vesicles, in response to osmotic stress, reduce the amount neurotransmitters released during exocytosis release. The amperometric recordings here indicate this is a reversible process and that vesicle after osmotic shock are refilled with neurotransmitters when placed cells are reverted into an isotonic environment.

We here describe the protocol for how combining TEM imaging together with two electrochemical methodologies, carbon fiber amperometry and intracellular electrochemical cytometry, can provide information that gains a broader view alluding to the effect of extracellular osmotic pressure on secretory vesicles and the exocytosis process. By comparing representative amperometric recordings of exocytosis release at single chromaffin cells using the experimental set up (shown in Figure 1), a signif...

Watch this Scientific Journal Video about Monitoring the Effect of Osmotic Stress on Secretory Vesicles and Exocytosis at JoVE.com

Monitoring the Effect of Osmotic Stress on Secretory Vesicles and Exocytosis

Osmotic stress affects exocytosis and the amount of neurotransmitter released during this process. We demonstrate how combining electrochemical methods together with transmission electron microscopy can be used to study the effect of extracellular osmotic pressure on exocytosis activity, vesicle quantal size, and the amount of neurotransmitter released during exocytosis.

Amperometry recording of cells subjected to osmotic shock show that secretory cells respond to this physical stress by reducing the exocytosis activity ...

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8.4K Views.  Chalmers University of Technology. Osmotic stress affects exocytosis and the amount of neurotransmitter released during this process. We demonstrate how combining electrochemical methods together with transmission electron microscopy can be used to study the effect of extracellular osmotic pressure on exocytosis activity, vesicle quantal size, and the amount of neurotransmitter released during exocytosis.

Video: Monitoring the Effect of Osmotic Stress on Secretory Vesicles and Exocytosis

Low-stress Route Learning Using the Lashley III Maze in Mice

Many behavior tests designed to assess learning and memory in rodents, particularly mice, rely on visual cues, food and/or water deprivation, or other aversive stimuli to motivate task acquisition. As animals age, sensory modalities deteriorate. For example, many strains of mice develop hearing deficits or cataracts. Changes in the sensory systems required to guide mice during task acquisition present potential confounds in interpreting learning changes in aging animals. Moreover, the use of aversive stimuli to motivate animals to learn tasks is potentially confounding when comparing mice with differential sensitivities to stress. To minimize these types of confounding effects, we have implemented a modified version of the Lashley III maze. This maze relies on route learning, whereby mice learn to navigate a maze via repeated exposure under low stress conditions, e.g. dark phase, no food/water deprivation, until they navigate a path from the start location to a pseudo-home cage with 0 or 1 error(s) on two consecutive trials. We classify this as a low-stress behavior test because it does not rely on aversive stimuli to encourage exploration of the maze and learning of the task. The apparatus consists of a modular start box, a 4-arm maze body, and a goal box. At the end of the goal box is a pseudo-home cage that contains bedding similar to that found in the animal&#x2019;s home cage and is specific to each animal for the duration of maze testing. It has been demonstrated previously that this pseudo-home cage provides sufficient reward to motivate mice to learn to navigate the maze1. Here, we present the visualization of the Lashley III maze procedure in the context of evaluating age-related differences in learning and memory in mice along with a comparison of learning behavior in two different background strains of mice. We hope that other investigators interested in evaluating the effects of aging or stress vulnerability in mice will consider this maze an attractive alternative to behavioral tests that involve more stressful learning tasks and/or visual cues.

The Lashley III maze is a route-learning task that does not rely on aversive stimuli or visual cues. It is thus a highly attractive option for evaluating learning and memory, especially in aging mice or otherwise where stress is a consideration.

Many behavior tests designed to assess learning and memory in rodents, particularly mice, rely on visual cues, food and/or water deprivation, or other ...

Brain Imaging Investigation of the Impairing Effect of Emotion on Cognition

Emotions can impact cognition by exerting both enhancing (e.g., better memory for emotional events) and impairing (e.g., increased emotional distractibility) effects (reviewed in 1). Complementing our recent protocol 2 describing a method that allows investigation of the neural correlates of the memory-enhancing effect of emotion (see also 1, 3-5), here we present a protocol that allows investigation of the neural correlates of the detrimental impact of emotion on cognition. The main feature of this method is that it allows identification of reciprocal modulations between activity in a ventral neural system, involved in 'hot' emotion processing (HotEmo system), and a dorsal system, involved in higher-level 'cold' cognitive/executive processing (ColdEx system), which are linked to cognitive performance and to individual variations in behavior (reviewed in 1). Since its initial introduction 6, this design has proven particularly versatile and influential in the elucidation of various aspects concerning the neural correlates of the detrimental impact of emotional distraction on cognition, with a focus on working memory (WM), and of coping with such distraction 7,11, in both healthy 8-11 and clinical participants 12-14.

We present a protocol that allows investigation of the neural mechanisms mediating the detrimental impact of emotion on cognition, using functional magnetic resonance imaging. This protocol can be used with both healthy and clinical participants.

Emotions can impact cognition by exerting both enhancing (e.g., better memory for emotional events) and impairing (e.g., increased emotional ...

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 ...

Modeling Neural Immune Signaling of Episodic and Chronic Migraine Using Spreading Depression In Vitro

Migraine and its transformation to chronic migraine are healthcare burdens in need of improved treatment options. We seek to define how neural immune signaling modulates the susceptibility to migraine, modeled in vitro using spreading depression (SD), as a means to develop novel therapeutic targets for episodic and chronic migraine. SD is the likely cause of migraine aura and migraine pain. It is a paroxysmal loss of neuronal function triggered by initially increased neuronal activity, which slowly propagates within susceptible brain regions. Normal brain function is exquisitely sensitive to, and relies on, coincident low-level immune signaling. Thus, neural immune signaling likely affects electrical activity of SD, and therefore migraine. Pain perception studies of SD in whole animals are fraught with difficulties, but whole animals are well suited to examine systems biology aspects of migraine since SD activates trigeminal nociceptive pathways. However, whole animal studies alone cannot be used to decipher the cellular and neural circuit mechanisms of SD. Instead, in vitro preparations where environmental conditions can be controlled are necessary. Here, it is important to recognize limitations of acute slices and distinct advantages of hippocampal slice cultures. Acute brain slices cannot reveal subtle changes in immune signaling since preparing the slices alone triggers: pro-inflammatory changes that last days, epileptiform behavior due to high levels of oxygen tension needed to vitalize the slices, and irreversible cell injury at anoxic slice centers. 
In contrast, we examine immune signaling in mature hippocampal slice cultures since the cultures closely parallel their in vivo counterpart with mature trisynaptic function; show quiescent astrocytes, microglia, and cytokine levels; and SD is easily induced in an unanesthetized preparation. Furthermore, the slices are long-lived and SD can be induced on consecutive days without injury, making this preparation the sole means to-date capable of modeling the neuroimmune consequences of chronic SD, and thus perhaps chronic migraine. We use electrophysiological techniques and non-invasive imaging to measure neuronal cell and circuit functions coincident with SD. Neural immune gene expression variables are measured with qPCR screening, qPCR arrays, and, importantly, use of cDNA preamplification for detection of ultra-low level targets such as interferon-gamma using whole, regional, or specific cell enhanced (via laser dissection microscopy) sampling. Cytokine cascade signaling is further assessed with multiplexed phosphoprotein related targets with gene expression and phosphoprotein changes confirmed via cell-specific immunostaining. Pharmacological and siRNA strategies are used to mimic and modulate SD immune signaling.

Modeling Neural Immune Signaling of Episodic and Chronic Migraine Using Spreading Depression In Vitro

Migraine and its transformation to chronic migraine are immense healthcare burdens in need of improved treatment options. We seek to define how neural immune signaling modulates the susceptibility to migraine, modeled in vitro using spreading depression in hippocampal slice cultures, as a means to develop novel therapeutic targets.

Migraine and its transformation to chronic migraine are healthcare burdens in need of improved treatment options.  We seek to define how neural immune ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Super-resolution Imaging of Neuronal Dense-core Vesicles

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and medical applications. Strengths of fluorescence include its sensitivity, specificity, and compatibility with live imaging. Unfortunately, conventional forms of fluorescence microscopy suffer from one major weakness, diffraction-limited resolution in the imaging plane, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, this limitation has been overcome with the introduction of super-resolution fluorescence microscopy techniques, such as photoactivated localization microscopy (PALM). Unlike its conventional counterparts, PALM can produce images with a lateral resolution of tens of nanometers. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of cellular structure and organization.
Unfortunately, PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. In this article, we show how to implement single-color PALM studies of vesicular structures in fixed, cultured neurons. PALM is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm. Key steps in our approach include labeling neurons with photoconvertible (green to red) chimeras of vesicle cargo, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a high-resolution PALM image. We also demonstrate the efficacy of our approach by presenting exceptionally well-resolved images of dense-core vesicles (DCVs) in cultured hippocampal neurons, which refute the hypothesis that extrasynaptic trafficking of DCVs is mediated largely by DCV clusters.

We describe how to implement photoactivated localization microscopy (PALM)-based studies of vesicles in fixed, cultured neurons. Key components of our protocol include labeling vesicles with photoconvertible chimeras, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a super-resolution image.

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and ...

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the over-production of amyloid-beta and the deposition of amyloid-beta plaques in brain regions of the afflicted individuals. Throughout the years scientists have generated numerous Alzheimer&#8217;s disease mouse models that attempt to replicate the amyloid-beta pathology. Unfortunately, the mouse models only selectively mimic the disease features. Neuronal death, a prominent effect in the brains of Alzheimer&#8217;s disease patients, is noticeably lacking in these mice. Hence, we and others have employed a method of directly infusing soluble oligomeric species of amyloid-beta - forms of amyloid-beta that have been proven to be most toxic to neurons - stereotaxically into the brain. In this report we utilize male C57BL/6J mice to document this surgical technique of increasing amyloid-beta levels in a select brain region. The infusion target is the dentate gyrus of the hippocampus because this brain structure, along with the basal forebrain that is connected by the cholinergic circuit, represents one of the areas of degeneration in the disease. The results of elevating amyloid-beta in the dentate gyrus via stereotaxic infusion reveal increases in neuron loss in the dentate gyrus within 1 week, while there is a concomitant increase in cell death and cholinergic neuron loss in the vertical limb of the diagonal band of Broca of the basal forebrain. These effects are observed up to 2 weeks. Our data suggests that the current amyloid-beta infusion model provides an alternative mouse model to address region specific neuron death in a short-term basis. The advantage of this model is that amyloid-beta can be elevated in a spatial and temporal manner.

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the ...

Characterization of Immune Cell-derived Extracellular Vesicles and Studying Functional Impact on Cell Environment

The neuroinflammatory state of the central nervous system (CNS) plays a key role in physiological and pathological conditions. Microglia, the resident immune cells in the brain, and sometimes the infiltrating bone marrow-derived macrophages (BMDMs), regulate the inflammatory profile of their microenvironment in the CNS. It is now accepted that the extracellular vesicle (EV) populations from immune cells act as immune mediators. Thus, their collection and isolation are important to identify their contents but also evaluate their biological effects on recipient cells. The present data highlight chronological requirements for EV isolation from microglia cells or blood macrophages including the ultracentrifugation and size-exclusion chromatography (SEC) steps. A non-targeted proteomic analysis permitted the validation of protein signatures as EV markers and characterized the biologically active EV contents. Microglia-derived EVs were also functionally used on primary culture of neurons to assess their importance as immune mediators in the neurite outgrowth. The results showed that microglia-derived EVs contribute to facilitate the neurite outgrowth in vitro. In parallel, blood macrophage-derived EVs were functionally used as immune mediators in spheroid cultures of C6 glioma cells, the results showing that these EVs control the glioma cell invasion in vitro. This report highlights the possibility to evaluate the EV-mediated immune cell functions but also understand the molecular bases of such a communication. This deciphering could promote the use of natural vesicles and/or the in vitro preparation of therapeutic vesicles in order to mimic immune properties in the microenvironment of CNS pathologies.

The present report highlights chronological requirements for extracellular vesicle (EV) isolation from microglia or blood macrophages. Microglia-derived EVs were evaluated as regulators of the neurite outgrowth while blood macrophage-derived EVs were studied in the control of C6 glioma cell invasion in in vitro assays. The goal is to better understand these EV functions as immune mediators in specific microenvironments.

The neuroinflammatory state of the central nervous system (CNS) plays a key role in physiological and pathological conditions. Microglia, the resident ...

Induction of Diffuse Axonal Brain Injury in Rats Based on Rotational Acceleration

Traumatic brain injury (TBI) is a major cause of death and disability. Diffuse axonal injury (DAI) is the predominant mechanism of injury in a large percentage of TBI patients requiring hospitalization. DAI involves widespread axonal damage from shaking, rotation or blast injury, leading to rapid axonal stretch injury and secondary axonal changes that are associated with a long-lasting impact on functional recovery. Historically, experimental models of DAI without focal injury have been difficult to design. Here we validate a simple, reproducible and reliable rodent model of DAI that causes widespread white matter damage without skull fractures or contusions.

This protocol validates a reliable, easy-to-perform and reproducible rodent model of brain diffuse axonal injury (DAI) that induces widespread white matter damage without skull fractures or contusions.

Traumatic brain injury (TBI) is a major cause of death and disability. Diffuse axonal injury (DAI) is the predominant mechanism of injury in a large ...

Implantation of Osmotic Pumps and Induction of Stress to Establish a Symptomatic, Pharmacological Mouse Model for DYT/PARK-ATP1A3 Dystonia

Genetically modified mouse models face limitations, especially when studying movement disorders, where most of the available transgenic rodent models do not present a motor phenotype resembling the clinical aspects of the human disease. Pharmacological mouse models allow for a more direct study of the pathomechanisms and their effect on the behavioral phenotype. Osmotic pumps connected to brain cannulas open up the possibility of creating pharmacological mouse models via local and chronic drug delivery. For the hereditary movement disorder of rapid-onset dystonia-parkinsonism, the loss-of-function mutation in the &#945;3-subunit of the Na+/K+-ATPase can be simulated by a highly specific blockade via the glycoside ouabain. In order to locally block the &#945;3-subunit in the basal ganglia and the cerebellum, which are the two brain structures believed to be heavily involved in the pathogenesis of rapid-onset dystonia-parkinsonism, a bilateral cannula is stereotaxically implanted into the striatum and an additional single cannula is introduced into the cerebellum. The cannulas are connected via vinyl tubing to two osmotic pumps, which are subcutaneously implanted on the back of the animals and allow for the chronic and precise delivery of ouabain. The pharmacological mouse model for rapid-onset dystonia-parkinsonism carries the additional advantage of recapitulating the clinical and pathological features of asymptomatic and symptomatic mutation carriers. Just like mutation carriers of rapid-onset dystonia parkinsonism, the ouabain-perfused mice develop dystonia-like movements only after additional exposure to stress. We demonstrate a mild stress paradigm and introduce two modified scoring systems for the assessment of a motor phenotype.

We provide a protocol to generate a pharmacological DYT/PARK-ATP1A3 dystonia mouse model via implantation of cannulas into basal ganglia and cerebellum connected to osmotic pumps. We describe the induction of dystonia-like movements via application of a motor challenge and the characterization of the phenotype via behavioral scoring systems.

Genetically modified mouse models face limitations, especially when studying movement disorders, where most of the available transgenic rodent models do ...