Name: real-time fluorescent measurement of synaptic functions in models of amyotrophic lateral sclerosis
Uploaded: 2021-07-16T14:30:00
Description: Watch this Scientific Journal Video about Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis at JoVE.com

We would like to acknowledge the present and former members of the Jefferson Weinberg ALS Center for critical feedback and suggestions for optimizing these techniques and their analyses. This work was supported by funding from the NIH (RF1-AG057882-01 and R21-NS0103118 to D.T), the NINDS (R56-NS092572 and R01-NS109150 to P.P), the Muscular Dystrophy Association (D.T.), the Robert Packard Center for ALS Research (D.T.), the Family Strong 4 ALS foundation and the Farber Family Foundation (B.K.J., K.K, and P.P).

All animal procedures performed in this study were approved by the Institutional Animal Care and Use Committee of Jefferson University.
1. Primary culture of neurons from embryonic rat cortex
NOTE: Primary cortical neurons are isolated from E17.5 rat embryos as previously described57,58. No strain bias appears to exist with the success of this culturing protocol. This method is described briefly below. The pre...

Three steps common to both methods described are of crucial importance for experimental success and quantifiable outcomes. First, preparation of fresh aCSF before each round of experiments is essential, following the attached instructions. Failure to do so may prevent appropriate neuronal depolarization. A sample of untreated control neurons should constantly be tested before stimulation of any experimental groups to ensure proper cellular depolarization and provide a benchmark for positive results obtained in that imagi...

Modeling amyotrophic lateral sclerosis (ALS) in the laboratory is made uniquely challenging due to the overwhelmingly sporadic nature of over 80% of cases1, coupled with the vast number of genetic mutations known to be disease-causative2. Despite this, all cases of ALS share the unifying feature that before outright neuronal degeneration, there is dysfunctional communication between presynaptic motor neurons and postsynaptic muscle cells3,4. Clinically, as patients lose connectivity of the remaining upper and lower motor neurons, they present with features of neuronal hyper- and hypoexcitability throughout the disease5,6,7,8,9, reflecting complex underlying molecular changes to these synapses, which we, as ALS researchers, seek to understand.
Multiple transgenic models have illustrated that deterioration and disorganization of the neuromuscular junction occur with the expression of ALS-causative genetic mutations, including SOD110, FUS11,12, C9orf7213,14,15,16, and TDP4317,18,19 through morphological assessments, including evaluation of synaptic boutons, spine densities, and pre/postsynaptic organization. Mechanistically, since the landmark papers of Cole, Hodgkin, and Huxley in the 1930s, it has also been possible to evaluate synaptic responses through electrophysiological techniques in either in vitro cell culture or tissue slice preparations20. Through these strategies, many models of ALS have demonstrated synaptic transmission deficits. For example, a mutant variant of TDP43 causes enhanced firing frequency and decreases action potential threshold in NSC-34 (spinal cord x neuroblastoma hybrid cell line 34) motor-neuron-like cells21. This same variant also causes dysfunctional synaptic transmission at the neuromuscular junction (NMJ) before the onset of behavioral motor deficits in a mouse model22. It was previously showed that mutant FUS expression results in reduced synaptic transmission at the NMJ in a drosophila model of FUS-ALS before locomotor defects11. A recent report using induced pluripotent stem cells derived from C9orf72-expansion carriers revealed a reduction in the readily releasable pool of synaptic vesicles23. Altogether, these studies and others highlight the importance of building a more comprehensive understanding of the mechanisms underlying synaptic signaling in disease-relevant models of ALS. This will be pivotal in understanding the pathobiology of ALS and developing potential therapeutic targets for patients.
Methods of current and voltage clamping cells have been invaluable in determining membrane properties such as conductance, resting membrane potential, and quantal content of individual synapses20,24. However, one of the significant limitations of electrophysiology is that it is technically challenging and only provides insights from a single neuron at a time. Live-cell confocal microscopy, coupled with specific fluorescent probes, offers the opportunity to investigate the synaptic transmission of neurons in a spatiotemporal manner25,26,27. Although not a direct measure of neuronal excitability, this fluorescence approach can provide a relative measurement of two molecular correlations of synaptic function: synaptic vesicle release and calcium transients at synaptic terminals.
When an action potential reaches the presynaptic terminal region of neurons, calcium transients are triggered, facilitating the transition from an electrical signal to the process of neurotransmitter release28. Voltage-gated calcium channels localized to these areas tightly regulate calcium signaling to modulate the kinetics of neurotransmitter release29. The first reported fluorescence-based recordings of calcium transients were performed using either the dual-wavelength indicator Fura-2 AM or the single wavelength dye Fluo-3 AM30,31,32. While these dyes offered great new insight at the time, they suffer from several limitations such as non-specific compartmentalization within cells, active or passive dye loss from labeled cells, photobleaching, and toxicity if imaged over extended periods of time33. In the past decade, genetically encoded calcium indicators have become the workhorses for imaging various forms of neuronal activity. These indicators combine a modified fluorescent protein with a calcium chelator protein that rapidly switches fluorescence intensity after the binding of Ca2+ ions34. The application of these new indicators is vast, allowing for much easier visualization of intracellular calcium transients both in in vitro and in vivo settings. One family of these genetically encoded reporters, known as GCaMP, are now broadly utilized. These indicators contain a C-terminal calmodulin domain, followed by green fluorescent protein (GFP), and are capped by an N-terminal calmodulin-binding region35,36. Calcium-binding to the calmodulin domain triggers an interaction with the calmodulin-binding region, resulting in a conformational change in the overall protein structure and a substantial increase in the fluorescence of the GFP moiety35,36. Over the years, this family of reporters has undergone several evolutions to enable distinct readouts for particular calcium transients with specific kinetics (slow, medium, and fast), each with slightly different properties37,38. Here, the usage of the reporter GcaMP6 has been highlighted, which has been previously shown to detect single action potentials and dendritic calcium transients in neurons both in vivo and in vitro37.
Calcium transients in the presynaptic region trigger synaptic vesicle fusion events, causing neurotransmitter release into the synapse and initiation of signaling events in the postsynaptic cell28,39. Synaptic vesicles are both rapidly released and recycled, as the cell homeostatically maintains a stable cell membrane surface area and readily releasable pool of fusion capable membrane-bound vesicles40. The styryl dye used here has an affinity toward lipid membranes and specifically changes its emission properties based on the ordering of the surrounding lipid environment41,42. Thus, it is an ideal tool for labeling recycling synaptic vesicles and subsequent tracking of these vesicles as they are later released following neuronal stimulation41,42. The protocol that has been generated and optimized is an adaptation of the concepts described initially by Gaffield and colleagues, which allows us to visualize styryl dye-labeled synaptic vesicle puncta over time continuously41.
Here, two related fluorescence-based methodologies are described, reliably reporting specific cellular events involved in synaptic transmission. Protocols have been defined to probe the dynamics of depolarization-mediated presynaptic terminal calcium influx and synaptic vesicle exocytosis in cultured neurons. Here, methods and representative results are focused on using primary rodent cortical or motor neurons as the in vitro model system, as there are published studies using these cell types43,44. However, these methods are also applicable to differentiated human i3 cortical-like neurons45, as we have also had success with both protocols in presently ongoing experimentation in our laboratory. The general protocol is outlined in a stepwise linear format, shown in Figure 1. In brief, to study calcium dynamics in neurites, mature neurons are transfected with plasmid DNA to express the fluorescent reporter GCaMP6m under a Cytomegalovirus (CMV) promoter37,46. Transfected cells have a low level of basal green fluorescence, which increases in the presence of calcium. Regions of interest are specified to monitor fluorescence changes throughout our manipulation. This allows for highly spatially and temporally localized fluctuations in calcium to be measured37,46. For evaluating synaptic vesicle fusion and release, mature neurons are loaded with styryl dye incorporated into synaptic vesicle membranes as they are recycled, reformed, and reloaded with neurotransmitters in presynaptic cells41,42,43,47,48. The current dyes used for this purpose label synaptic vesicles along neurites and are used as a proxy for these regions in live-imaging experiments, as was shown by co-staining of styryl dye and synaptotagmin by Kraszewski and colleagues49. Included here are representative images of similar staining that have also been performed (Figure 2A). Previous investigators have extensively used such dyes to report synaptic vesicle dynamics at the neuromuscular junction and hippocampal neurons48,49,50,51,52,53,54,55,56. By selecting punctate regions of dye-loaded vesicles and by monitoring decreases in fluorescence intensity following vesicle release, functional synaptic transmission capacity and temporal dynamics of release can be studied following stimulation43. For both methods, a medium containing a high concentration of potassium chloride is employed to depolarize cells to mimic neuronal activity. Imaging parameters are specified to capture sub-second intervals spanning a baseline normalization followed by our stimulation capture period. Fluorescence measurements at each time point are determined, normalized to the background, and quantified over the experimental time period. Calcium-influx mediated GCaMP6m fluorescence increase or effective synaptic vesicle exocytosis styryl dye release fluorescence decrease can be detected through this strategy. Detailed methodological setup and parameters for these two protocols and a discussion on their advantages and limitations are described below.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62813/62813fig01.jpg" /> 
Figure 1:&#160;Visual rendering of overall general protocol process. (1) Isolate and culture primary rodent neurons in vitro to chosen maturation timepoint. (2) Introduce GCaMP DNA or styryl dye as reporters of synaptic activity. (3) Setup imaging paradigm using live-imaging equipped confocal microscope and associated software. Begin baseline recording period. (4) While cells are still undergoing live-image capture, stimulate neurons via high KCl bath perfusion. (5) Assess fluorescence intensity measurements over time to measure calcium transients or synaptic vesicle fusion. <a href="https://www.jove.com/files/ftp_upload/62813/62813fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>20x air objective</td>
			<td>Nikon</td>
			<td></td>
			<td>For imaging</td>
		</tr>
		<tr>
			<td>40x oil immersion objective</td>
			<td>Nikon</td>
			<td></td>
			<td>For imaging</td>
		</tr>
		<tr>
			<td>B27 supplement</td>
			<td>Thermo Scientific</td>
			<td>17504044</td>
			<td>Neuronal growth supplement</td>
		</tr>
		<tr>
			<td>BD Syringes without Needle, 50 mL</td>
			<td>Thermo Scientific</td>
			<td>13-689-8</td>
			<td>Part of gravity perfusion assembly</td>
		</tr>
		<tr>
			<td>Biosafety cell culture hood</td>
			<td>Baker</td>
			<td>SterilGARD III SG403A</td>
			<td>Asceptic cell culturing, transfection, and dye loading</td>
		</tr>
		<tr>
			<td>b-Mercaptoethanol</td>
			<td>Millipore Sigma</td>
			<td>M3148</td>
			<td>For culturing and maintenance of neuronal cultures</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Millipore Sigma</td>
			<td>A9418</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Millipore Sigma</td>
			<td>223506</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>Cell culture CO2 incubator</td>
			<td>Thermo Scientific</td>
			<td>13-998-123</td>
			<td>For culturing and maintenance of neurons</td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Eppendorf</td>
			<td>5810R</td>
			<td>For neuronal culture preparation</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Nikon</td>
			<td>Eclipse Ti +A1R core</td>
			<td>For fluorescence imaging</td>
		</tr>
		<tr>
			<td>CoolSNAP ES2&#160; CCD camera</td>
			<td>Photometrics</td>
			<td></td>
			<td>For image acquisition</td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>Millipore Sigma</td>
			<td>G8270</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>DNase</td>
			<td>Millipore Sigma</td>
			<td>D5025</td>
			<td>For neuronal culture preparation</td>
		</tr>
		<tr>
			<td>Female, timed-pregnancy Sprague Dawley rats</td>
			<td>Charles river</td>
			<td>400SASSD</td>
			<td>For preparing embryonic cortical and spinal motor neuron cultures</td>
		</tr>
		<tr>
			<td>FITC Filter cube</td>
			<td>Nikon</td>
			<td>77032509</td>
			<td>For imaging Gcamp calcium transients</td>
		</tr>
		<tr>
			<td>FM4-64 styryl dye</td>
			<td>Invitrogen</td>
			<td>T13320</td>
			<td>For imaging synaptic vesicle release</td>
		</tr>
		<tr>
			<td>Glass bottom petri dishes (Thickness #1.5)</td>
			<td>CellVis</td>
			<td>D35-10-1.5-N</td>
			<td>For growth of neurons on imaging-compatible culture dish</td>
		</tr>
		<tr>
			<td>Glass Pasteur pipette</td>
			<td>Grainger</td>
			<td>52NK56</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Hank&#39;s Balanced Salt Solution (HBSS)</td>
			<td>Millipore Sigma</td>
			<td>H6648</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Millipore Sigma</td>
			<td>H3375</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>High KCl artifical cerebrospinal fluid (aCSF)</td>
			<td></td>
			<td></td>
			<td>For imaging. Please see recipes*</td>
		</tr>
		<tr>
			<td>horse serum</td>
			<td>Millipore Sigma</td>
			<td>H1138</td>
			<td>For culturing and maintenance of neurons</td>
		</tr>
		<tr>
			<td>Laminar flow dissection hood</td>
			<td>NUAIRE</td>
			<td>NU-301-630</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Laminin</td>
			<td>Thermo Scientific</td>
			<td>23017015</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 Medium</td>
			<td>Thermo Scientific</td>
			<td>11415064</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 Medium, no phenol red</td>
			<td>Thermo Scientific</td>
			<td>21083027</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>L-Glutamine (200 mM)</td>
			<td>Thermo Scientific</td>
			<td>25030149</td>
			<td>Neuronal culture supplement</td>
		</tr>
		<tr>
			<td>Lipofectamine 2000 Transfection Reagent</td>
			<td>Thermo Scientific</td>
			<td>11668019</td>
			<td>For neuronal transfections</td>
		</tr>
		<tr>
			<td>Low KCl artifical cerebrospinal fluid (aCSF)</td>
			<td></td>
			<td></td>
			<td>For imaging. Please see recipes*</td>
		</tr>
		<tr>
			<td>Magnesium chloride</td>
			<td>Millipore Sigma</td>
			<td>208337</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>Microsoft Excel</td>
			<td>Microsoft</td>
			<td></td>
			<td>Software for data analysis/normalization</td>
		</tr>
		<tr>
			<td>Nalgene&#160; Filter Units, 0.2 &#181;m PES</td>
			<td>Thermo Scientific</td>
			<td>565-0020</td>
			<td>Filter unit for aCSF solution</td>
		</tr>
		<tr>
			<td>Neurobasal medium</td>
			<td>Thermo Scientific</td>
			<td>21103049</td>
			<td>For culturing and maintenance of neuronal cultures</td>
		</tr>
		<tr>
			<td>NIS-Elements Advanced Research</td>
			<td>Nikon</td>
			<td></td>
			<td>Software for image capture and analysis</td>
		</tr>
		<tr>
			<td>Nunc 15 mL Conical tubes</td>
			<td>Thermo Scientific</td>
			<td>339650</td>
			<td>For preparing neuronal culture and buffer solutions</td>
		</tr>
		<tr>
			<td>Nunc 50 mL conical tubes</td>
			<td>Thermo Scientific</td>
			<td>339652</td>
			<td>For preparing neuronal culture and buffer solutions</td>
		</tr>
		<tr>
			<td>Optiprep</td>
			<td>Millipore Sigma</td>
			<td>D1556</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Millipore Sigma</td>
			<td>P4762</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000 U/mL)</td>
			<td>Thermo Scientific</td>
			<td>15140122</td>
			<td>To prevent bacterial contamination of neuronal cultures</td>
		</tr>
		<tr>
			<td>Perfusion system</td>
			<td>Warner Instruments</td>
			<td>SF-77B</td>
			<td>For exchange of aCSF</td>
		</tr>
		<tr>
			<td>Perfusion tubing</td>
			<td>Cole-Parmer</td>
			<td>UX-30526-14</td>
			<td>Part of gravity perfusion assembly</td>
		</tr>
		<tr>
			<td>pGP-CMV-Gcamp6m plasmid</td>
			<td>Addgene</td>
			<td>40754</td>
			<td>For imaging calcium transients</td>
		</tr>
		<tr>
			<td>Poly-D-lysine hydrobromide</td>
			<td>Millipore Sigma</td>
			<td>P7886</td>
			<td>Coating agent for glass bottom petri dishes</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Millipore Sigma</td>
			<td>P3911</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Millipore Sigma</td>
			<td>S5761</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Millipore Sigma</td>
			<td>S9888</td>
			<td>Component of aCSF solutions</td>
		</tr>
		<tr>
			<td>Stage Top Incubator</td>
			<td>Tokai Hit</td>
			<td></td>
			<td>For incubation of live neurons during imaging period</td>
		</tr>
		<tr>
			<td>TRITC Filter cube</td>
			<td>Nikon</td>
			<td>77032809</td>
			<td>For imaging FM4-64</td>
		</tr>
		<tr>
			<td>Trypsin Inhibitor</td>
			<td>Millipore Sigma</td>
			<td>T6414</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.25%), phenol red</td>
			<td>Thermo Scientific</td>
			<td>25200056</td>
			<td>For preparing neuronal cultures</td>
		</tr>
		<tr>
			<td>Vibration Isolation table</td>
			<td>New Port</td>
			<td>VIP320X2430-135520</td>
			<td>Table/stand for microscope</td>
		</tr>
	</tbody>
</table>

real-time fluorescent measurement of synaptic functions in models of amyotrophic lateral sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Following the successful implementation of the above protocol, representative results are shown for a typical styryl dye synaptic vesicle release experiment. Cultured rat primary cortical neurons were loaded with dye using the method described in section 6. The specificity of dye loading was determined by co-labeling with synaptic vesicle marker synaptophysin. A majority of styryl dye positive puncta are co-positive for this marker (Figure 2A). To determine whether the settings used for styr...

Watch this Scientific Journal Video about Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis at JoVE.com

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

These methods enable quick assessment as to whether an experimental manipulation, disease causative protein, or RNA can impact synaptic processes, and whether therapeutic compounds can restore function. These techniques provide a reliable readout of synaptic function from a group of neurons in a relatively short period of time compared to electrophysiology. This protocol can be applied to any disease of neuronal development or degeneration.This works well for primary rodent neuronal cultures, and for neurons derived from induced pluripotent stem cells. To begin, optimize the image acquisition settings using confocal imaging acquisition software. Keep excitation power exposure time, detector gain and frame rate constant across all samples.For time lapse imaging, select an aspect ratio of 512 by 512, and a frame rate of two images per second to minimize dye bleaching. Then select the fluorescence excitation, dichroic and emission filter combinations for GCaMP six M/GCaMP three with Fitzy and stearoyl dye with Trixi. To avoid Z drift during time lapse imaging, use the perfect focus feature of the confocal imaging acquisition software.Next, select the time tab in the image acquisition panel. For phase one, set interval 500 milliseconds and duration three to five minutes. And for phase two, set interval 500 milliseconds and duration, five minutes.Then to assemble the gravity perfusion apparatus for artificial cerebrospinal fluid or ACSF, load the high potassium chloride ACSF buffer into a 50 milliliter syringe at the top of the apparatus and set the flow rate to one milliliter per minute. Then, load a 35 milliliter glass dish containing neurons onto the confocal imaging stage with the end of the perfusion tubing placed at the dish edge and choose the field for imaging. 48 hours post transfection, incubate the primary cortical or motor neurons in low potassium chloride ACSF buffer for 10 minutes at 37 degrees Celsius.Then remove the buffer by aspiration and using a pipette, load the neurons in the dark on a glass bottom Petri dish filled with ACSF containing 50 millimolar potassium chloride and 10 Micro molar styryl dye. After five minutes, remove the loading solution and bath the neurons in low potassium chloride ACSF buffer at 37 degrees Celsius for 10 minutes to eliminate the nonspecific dye loading. Then place the dish onto the imaging stage of the inverted confocal microscope and observe the cells under a 20 times air objective or 40 times oil immersion objective.Excite the steel dye using a 546 nanometer laser and collect emission using a 570 to 620 nanometer bandpass filter. After selecting the imaging field and engaging perfect focus, take a single still image with bright field, Trixi and fluorescence marker channels to mark neuronal boundaries. Then initiate Run Now in the acquisition software and carry out the basal recording for three to five minutes to exclude variations in dye intensity.Right at the switch to phase two, trigger the on button for the perfusion system and constantly perfuse 50 millimolar potassium chloride to the neurons to facilitate dye unloading. Carry out the recordings for five minutes then trigger the off switch for the perfusion system. Save the experiment for data analysis later using the confocal software.48 hours post transfection with GCaMP six M, incubate the primary rodent cortical neurons with low potassium chloride ACSF for 15 minutes. Then mount the dish on the imaging platform and visualized GCaMP six M fluorescence using a Fitzy filter and a 20 times or 40 times objective. After selecting the imaging field and engaging perfect focus, take a single still image with brightfield, Fitzy, and fluorescence marker channels to mark neuronal boundaries.Next, initiate Run Now in the acquisition software and carry out the basal recording for three to five minutes. Then perfuse ACSF containing 50 millimolar potassium chloride to the neurons as demonstrated previously and record for five minutes. When the image acquisition stops, save the experiment and proceed to data analysis using the confocal software.For image analysis, open the time lapse images in the confocal software and align the images by clicking on Image, followed by processing, followed by align current document, then select Align to the first frame. Select regions of interest along the neurites using the ROI selection tool and also mark an ROI representing background fluorescence intensity. Next, initiate the measure function from the time measurement panel to measure raw fluorescence over time for the selected ROIs.After measurement export the raw fluorescence intensities to the spreadsheet software. Cultured rat primary cortical neurons loaded with styryl dye are shown here. The specificity of dye loading was determined by co labeling with synaptic vesicle marker synaptophysin and the majority of sterile dye positive puncta are co positive for this marker.The analysis of the raw intensity values over the entire imaging period reveals that synaptophysin and intensity remains constant while styryl dye intensity decreases following stimulation. For green fluorescent protein transfected control neurons, successful synaptic vesicle release resulted in the striking loss of dye fluorescence upon high potassium chloride depolarization. This representative video shows a neuron selectively unloading styryl dye in a new right region following stimulation.In contrast, in neurons transfected with a C nine ORF 72 linked to dye peptide repeat construct to GA 50, impaired synaptic transmission is represented by retained dye fluorescence even after high potassium chloride depolarization. Representative fluorescence images of cortical neurons transfected with GCaMP six M before and after potassium chloride depolarization are shown here. Increased fluorescence values and cortical neurons transfected with GCaMP six M indicate calcium entry into neurites following potassium chloride inducted depolarization.At the end of the baseline recording period, the neurons expressed GCaMP six M with a low fluorescence. A dramatic fluorescence increase is then observed at the start of stimulation. Make sure fluorescence baseline recordings for styryl dye and GCaMP are stable.Always use perfect focus to avoid image drift, which would make post image data processing challenging. Further culturing of neurons is not recommended, as dishes are exposed to air during profusion. However, immuno staining or extraction of protein or RNA can assess potential causes of synaptic alterations.

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JoVE Journal

Neuroscience

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 ...

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

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

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

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

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

Real-time Imaging of Axonal Transport of Quantum Dot-labeled BDNF in Primary Neurons

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are increasingly viewed as an early feature of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD) and Huntington&#8217;s disease (HD). As yet unclear is the mechanism(s) by which axonal injury is induced. We reported the development of a novel technique to produce biologically active, monobiotinylated BDNF (mBtBDNF) that can be used to trace axonal transport of BDNF. Quantum dot-labeled BDNF (QD-BDNF) was produced by conjugating quantum dot 655 to mBtBDNF. A microfluidic device was used to isolate axons from neuron cell bodies. Addition of QD-BDNF to the axonal compartment allowed live imaging of BDNF transport in axons. We demonstrated that QD-BDNF moved essentially exclusively retrogradely, with very few pauses, at a moving velocity of around 1.06 &#956;m/sec. This system can be used to investigate mechanisms of disrupted axonal function in AD or HD, as well as other degenerative disorders.

Axonal transport of BDNF, a neurotrophic factor, is critical for the survival and function of several neuronal populations. Some degenerative disorders are marked by disruption of axonal structure and function. We demonstrated the techniques used to examine live trafficking of QD-BDNF in microfluidic chambers using primary neurons.

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are ...

A Protocol for the Administration of Real-Time fMRI Neurofeedback Training

Neurologic disorders are characterized by abnormal cellular-, molecular-, and circuit-level functions in the brain. New methods to induce and control neuroplastic processes and correct abnormal function, or even shift functions from damaged tissue to physiologically healthy brain regions, hold the potential to dramatically improve overall health. Of the current neuroplastic interventions in development, neurofeedback training (NFT) from functional Magnetic Resonance Imaging (fMRI) has the advantages of being completely non-invasive, non-pharmacologic, and spatially localized to target brain regions, as well as having no known side effects. Furthermore, NFT techniques, initially developed using fMRI, can often be translated to exercises that can be performed outside of the scanner without the aid of medical professionals or sophisticated medical equipment. In fMRI NFT, the fMRI signal is measured from specific regions of the brain, processed, and presented to the participant in real-time. Through training, self-directed mental processing techniques, that regulate this signal and its underlying neurophysiologic correlates, are developed. FMRI NFT has been used to train volitional control over a wide range of brain regions with implications for several different cognitive, behavioral, and motor systems. Additionally, fMRI NFT has shown promise in a broad range of applications such as the treatment of neurologic disorders and the augmentation of baseline human performance. In this article, we present an fMRI NFT protocol developed at our institution for modulation of both healthy and abnormal brain function, as well as examples of using the method to target both cognitive and auditory regions of the brain.

The ability to induce and/or control neural plasticity may be critical in future treatments for neurologic disorders and the recovery from brain injury. In this paper, we present a protocol on the use of neurofeedback training with functional magnetic resonance imaging to modulate human brain function.

Neurologic disorders are characterized by abnormal cellular-, molecular-, and circuit-level functions in the brain. New methods to induce and control ...

Live-cell Measurement of Odorant Receptor Activation Using a Real-time cAMP Assay

The enormous sizes of the mammalian odorant receptor (OR) families present difficulties to find their cognate ligands among numerous volatile chemicals. To efficiently and accurately deorphanize ORs, we combine the use of a heterologous cell line to express mammalian ORs and a genetically modified biosensor plasmid to measure cAMP production downstream of OR activation in real time. This assay can be used to screen odorants against ORs and vice versa. Positive odorant-receptor interactions from the screens can be subsequently confirmed by testing against various odor concentrations, generating concentration-response curves. Here we used this method to perform a high-throughput screening of an odorous compound against a human OR library expressed in Hana3A cells and confirmed that the positively-responding receptor is the cognate receptor for the compound of interest. We found this high-throughput detection method to be efficient and reliable in assessing OR activation and our data provide an example of its potential use in OR functional studies.

Characterizing the function of odorant receptors serves an indispensable part in the deorphanization process. We describe a method to measure the activation of odorant receptors in real time using a cAMP assay.

The enormous sizes of the mammalian odorant receptor (OR) families present difficulties to find their cognate ligands among numerous volatile chemicals. ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Real-time In Vitro Monitoring of Odorant Receptor Activation by an Odorant in the Vapor Phase

Olfactory perception begins with the interaction of odorants with odorant receptors (OR) expressed by olfactory sensory neurons (OSN). Odor recognition follows a combinatorial coding scheme, where one OR can be activated by a set of odorants and one odorant can activate a combination of ORs. Through such combinatorial coding, organisms can detect and discriminate between a myriad of volatile odor molecules. Thus, an odor at a given concentration can be described by an activation pattern of ORs, which is specific to each odor. In that sense, cracking the mechanisms that the brain uses to perceive odor requires the understanding odorant-OR interactions. This is why the olfaction community is committed to &#34;de-orphanize&#34;&#160;these receptors. Conventional in vitro systems used to identify odorant-OR interactions have utilized incubating cell media with odorant, which is distinct from the natural detection of odors via vapor odorants dissolution into nasal mucosa before interacting with ORs. Here, we describe a new method that allows for real-time monitoring of OR activation via vapor-phase odorants. Our method relies on measuring cAMP release by luminescence using the Glosensor assay. It bridges current gaps between in vivo and in vitro approaches and provides a basis for a biomimetic volatile chemical sensor.

Physiologically, odorant receptors are activated by odorant molecules inhaled in the vapor phase. However, most in vitro systems utilize liquid phase odorant stimulation. Here, we present a method that allows real-time in vitro monitoring of odorant receptor activation upon odorant stimulation in vapor phase.

Olfactory perception begins with the interaction of odorants with odorant receptors (OR) expressed by olfactory sensory neurons (OSN). Odor recognition ...

Comprehensive Autopsy Program for Individuals with Multiple Sclerosis

We describe a rapid tissue donation program for individuals with multiple sclerosis (MS) that requires scientists and technicians to be on-call 24/7, 365 days a year. Participants consent to donate their brain and spinal cord. Most patients were followed by neurologists at the Cleveland Clinic Mellen Center for MS Treatment and Research. Their clinical courses and neurological disabilities are well-characterized. Soon after death, the body is transported to the MS Imaging Center, where the brain is scanned in situ by 3 T magnetic resonance imaging (MRI). The body is then transferred to the autopsy room, where the brain and spinal cord are removed. The brain is divided into two hemispheres. One hemisphere is immediately placed in a slicing box and alternate 1 cm-thick slices are either fixed in 4% paraformaldehyde for two days or rapidly frozen in dry ice and 2-methylbutane. The short-fixed brain slices are stored in a cryopreservation solution and used for histological analyses and immunocytochemical detection of sensitive antigens. Frozen slices are stored at -80 &#176;C and used for molecular, immunocytochemical, and in situ hybridization/RNA scope studies. The other hemisphere is placed in 4% paraformaldehyde for several months, placed in the slicing box, re-scanned in the 3 T magnetic resonance (MR) scanner and sliced into centimeter-thick slices. Postmortem in situ MR images (MRIs) are co-registered with 1 cm-thick brain slices to facilitate MRI-pathology correlations. All brain slices are photographed and brain white-matter lesions are identified. The spinal cord is cut into 2 cm segments. Alternate segments are fixed in 4% paraformaldehyde or rapidly frozen. The rapid procurement of postmortem MS tissues allows pathological and molecular analyses of MS brains and spinal cords and pathological correlations of brain MRI abnormalities. The quality of these rapidly-processed postmortem tissues (usually within 6 h of death) is of great value to MS research and has resulted in many high-impact discoveries.

Multiple sclerosis is an inflammatory demyelinating disease with no cure. Analysis of brain tissue provides important clues to understanding the pathogenesis of the disease. Here we discuss the methodology and downstream analysis of MS brain tissue collected through a unique rapid autopsy program in operation at the Cleveland Clinic.

We describe a rapid tissue donation program for individuals with multiple sclerosis (MS) that requires scientists and technicians to be on-call 24/7, 365 ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.