Name: real-time fluorescent measurement of synaptic functions in models of amyotrophic lateral sclerosis
Uploaded: 2021-07-16T14:30:00.000+00:00
Duration: 8 min 59 s
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).

The authors declare that they have no conflicts of interest.

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><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 CO&lt;sub&gt;2&lt;/sub&gt; 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&amp;nbsp; 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&#039;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&#039;s L-15 Medium</td><td>Thermo Scientific</td><td>11415064</td><td>For preparing neuronal cultures</td></tr><tr><td>Leibovitz&#039;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&amp;nbsp; Filter Units, 0.2 &amp;micro;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 ...

real-time-fluorescent-measurement-synaptic-functions-models

Research

JoVE Journal

Neuroscience

2.5K Views.  Thomas Jefferson University. 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.

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

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Amyloid fibrils are associated with a number of neurodegenerative diseases such as Huntington&#39;s, Parkinson&#39;s, or Alzheimer&#39;s disease. These amyloid fibrils can sequester endogenous metastable proteins as well as components of the proteostasis network (PN) and thereby exacerbate protein misfolding in the cell. There are a limited number of tools available to assess the aggregation process of amyloid proteins within an animal. We present a protocol for fluorescence lifetime microscopy (FLIM) that allows monitoring as well as quantification of the amyloid fibrilization in specific cells, such as neurons, in a noninvasive manner and with the progression of aging and upon perturbation of the PN. FLIM is independent of the expression levels of the fluorophore and enables an analysis of the aggregation process without any further staining or bleaching. Fluorophores are quenched when they are in close vicinity of amyloid structures, which results in a decrease of the fluorescence lifetime. The quenching directly correlates with the aggregation of the amyloid protein. FLIM is a versatile technique that can be applied to compare the fibrilization process of different amyloid proteins, environmental stimuli, or genetic backgrounds in vivo in a non-invasive manner.

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Fluorescence lifetime imaging monitors, quantifies and distinguishes the aggregation tendencies of proteins in living, aging, and stressed C. elegans disease models.

Amyloid fibrils are associated with a number of neurodegenerative diseases such as Huntington&#39;s, Parkinson&#39;s, or Alzheimer&#39;s disease. These ...

Biochemistry

DetectSyn: A Rapid, Unbiased Fluorescent Method to Detect Changes in Synapse Density

Synapses are the site of communication between neurons. Neuronal circuit strength is related to synaptic density, and the breakdown of synapses is characteristic of disease states like major depressive disorder (MDD) and Alzheimer's disease. Traditional techniques to investigate synapse numbers include genetic expression of fluorescent markers (e.g., green fluorescent protein (GFP)), dyes that fill a neuron (e.g., carbocyanine dye, DiI), and immunofluorescent detection of spine markers (e.g., postsynaptic density 95 (PSD95)). A major caveat to these proxy techniques is that they only identify postsynaptic changes. Yet, a synapse is a connection between a presynaptic terminal and a postsynaptic spine. The gold standard for measuring synapse formation/elimination requires time-consuming electron microscopy or array tomography techniques. These techniques require specialized training and costly equipment. Further, only a limited number of neurons can be assessed and are used to represent changes to an entire brain region. DetectSyn is a rapid fluorescent technique that identifies changes to synapse formation or elimination due to a disease state or drug activity. DetectSyn utilizes a rapid proximity ligation assay to detect juxtaposed pre- and postsynaptic proteins and standard fluorescent microscopy, a technique readily available to most laboratories. Fluorescent detection of the resulting puncta allows for quick and unbiased analysis of experiments. DetectSyn provides more representative results than electron microscopy because larger areas can be analyzed than a limited number of fluorescent neurons. Moreover, DetectSyn works for in vitro cultured neurons and fixed tissue slices. Finally, a method is provided to analyze the data collected from this technique. Overall, DetectSyn offers a procedure for detecting relative changes in synapse density across treatments or disease states and is more accessible than traditional techniques.

DetectSyn is an unbiased, rapid fluorescent assay that measures changes in relative synapse (pre- and postsynaptic engagement) number across treatments or disease states. This technique utilizes a proximity ligation technique that can be used both in cultured neurons and fixed tissue.

Synapses are the site of communication between neurons. Neuronal circuit strength is related to synaptic density, and the breakdown of synapses is ...

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

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

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

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

Using Enzyme-based Biosensors to Measure Tonic and Phasic Glutamate in Alzheimer's Mouse Models

Neurotransmitter disruption is often a key component of diseases of the central nervous system (CNS), playing a role in the pathology underlying Alzheimer's disease, Parkinson's disease, depression, and anxiety. Traditionally, microdialysis has been the most common (lauded) technique to examine neurotransmitter changes that occur in these disorders. But because microdialysis has the ability to measure slow 1-20 minute changes across large areas of tissue, it has the disadvantage of invasiveness, potentially destroying intrinsic connections within the brain and a slow sampling capability. A relatively newer technique, the microelectrode array (MEA), has numerous advantages for measuring specific neurotransmitter changes within discrete brain regions as they occur, making for a spatially and temporally precise approach. In addition, using MEAs is minimally invasive, allowing for measurement of neurotransmitter alterations in vivo. In our laboratory, we have been specifically interested in changes in the neurotransmitter, glutamate, related to Alzheimer's disease pathology. As such, the method described here has been used to assess potential hippocampal disruptions in glutamate in a transgenic mouse model of Alzheimer's disease. Briefly, the method used involves coating a multi-site microelectrode with an enzyme very selective for the neurotransmitter of interest and using self-referencing sites to subtract out background noise and interferents. After plating and calibration, the MEA can be constructed with a micropipette and lowered into the brain region of interest using a stereotaxic device. Here, the method described involves anesthetizing rTg(TauP301L)4510 mice and using a stereotaxic device to precisely target sub-regions (DG, CA1, and CA3) of the hippocampus.

Here, we describe the setup, software navigation, and data analysis for a spatially and temporally precise method of measuring tonic and phasic extracellular glutamate changes in vivo using enzyme-linked microelectrode arrays (MEA).

Neurotransmitter disruption is often a key component of diseases of the central nervous system (CNS), playing a role in the pathology underlying ...

Assay to Measure Nucleocytoplasmic Transport in Real Time within Motor Neuron-like NSC-34 Cells

Nucleocytoplasmic transport refers to the import and export of large molecules from the cell nucleus. Recently, a number of studies have shown a connection between amyotrophic lateral sclerosis (ALS) and impairments in the nucleocytoplasmic pathway. ALS is a neurodegenerative disease affecting the motor neurons and resulting in paralysis and ultimately in death, within 2-5 years on average. Most cases of ALS are sporadic, lacking any apparent genetic linkage, but 10% are inherited in a dominant manner. Recently, hexanucleotide repeat expansions (HREs) in the chromosome 9 open reading frame 72 (C9orf72) gene were identified as a genetic cause of ALS and frontotemporal dementia (FTD). Importantly, different groups have recently proposed that these mutants affect nucleocytoplasmic transport. These studies have mostly shown the final outcome and manifestations caused by HREs on nucleocytoplasmic transport, but they do not demonstrate nuclear transport dysfunction in real time. As a result, only severe nucleocytoplasmic transport deficiency can be determined, mostly due to high overexpression or exogenous protein insertion. 
This protocol describes a new and very sensitive assay to evaluate and quantify nucleocytoplasmic transport dysfunction in real time. The rate of import of a NLS-NES-GFP protein (shuttle-GFP) can be quantified in real time using fluorescent microscopy. This is performed by using an exportin inhibitor, thus allowing the shuttle GFP only to enter the nucleus. To validate the assay, the C9orf72 HRE translated dipeptide repeats, poly(GR) and poly(PR), which have been previously shown to disrupt nucleocytoplasmic transport, were used. Using the described assay, a 50% decrease in the nuclear import rate was observed compared to the control. Using this system, minute changes in nucleocytoplasmic transport can be examined and the ability of different factors to rescue (even partially) a nucleocytoplasmic transport defect can be determined.

This protocol describes a method to sensitively measure the nucleocytoplasmic transport rate within motor neuron-like NSC-34 cells by quantifying the real-time change in the nuclear import of a NLS-NES-GFP protein.

Nucleocytoplasmic transport refers to the import and export of large molecules from the cell nucleus. Recently, a number of studies have shown a ...

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.

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other ...

Measuring Glucose Uptake in Drosophila Models of TDP-43 Proteinopathy

Amyotrophic lateral sclerosis is a neurodegenerative disorder causing progressive muscle weakness and death within 2-5 years following diagnosis. Clinical manifestations include weight loss, dyslipidemia, and hypermetabolism; however, it remains unclear how these relate to motor neuron degeneration. Using a Drosophila model of TDP-43 proteinopathy that recapitulates several features of ALS including cytoplasmic inclusions, locomotor dysfunction, and reduced lifespan, we recently identified broad ranging metabolic deficits. Among these, glycolysis was found to be upregulated and genetic interaction experiments provided evidence for a compensatory neuroprotective mechanism. Indeed, despite upregulation of phosphofructokinase, the rate limiting enzyme in glycolysis, an increase in glycolysis using dietary and genetic manipulations was shown to mitigate locomotor dysfunction and increased lifespan in fly models of TDP-43 proteinopathy. To further investigate the effect on TDP-43 proteinopathy on glycolytic flux in motor neurons, a previously reported genetically encoded, FRET-based sensor, FLII12Pglu-700&#181;&#948;6, was used. This sensor is comprised of a bacterial glucose-sensing domain and cyan and yellow fluorescent proteins as the FRET pair. Upon glucose binding, the sensor undergoes a conformational change allowing FRET to occur. Using FLII12Pglu-700&#181;&#948;6, glucose uptake was found to be significantly increased in motor neurons expressing TDP-43G298S, an ALS causing variant. Here, we show how to measure glucose uptake, ex vivo, in larval ventral nerve cord preparations expressing the glucose sensor FLII12Pglu-700&#181;&#948;6 in the context of TDP-43 proteinopathy. This approach can be used to measure glucose uptake and assess glycolytic flux in different cell types or in the context of various mutations causing ALS and related neurodegenerative disorders.

Measuring Glucose Uptake in Drosophila Models of TDP-43 Proteinopathy

Glucose uptake is increased in Drosophila motor neurons affected by TAR DNA binding protein (TDP-43) proteinopathy, as indicated by a FRET-based, genetically encoded glucose sensor.

Amyotrophic lateral sclerosis is a neurodegenerative disorder causing progressive muscle weakness and death within 2-5 years following diagnosis. Clinical ...

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal ...

Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to use these cells for the purpose of studying the pathophysiology of amyotrophic lateral sclerosis (ALS) in the rat hSOD1G93A model. First, the protocol shows how to isolate and culture astrocytes and microglia from postnatal rat cortices, and then how to characterize and test these cultures for purity by immunocytochemistry using the glial fibrillary acidic protein (GFAP) marker of astrocytes and the ionized calcium-binding adaptor molecule 1 (Iba1) microglial marker. In the next stage, methods are described for dye-loading (calcium-sensitive Fluo 4-AM) of cultured cells and the recordings of Ca2+ changes in video imaging experiments on live cells.
The examples of video recordings consist of: (1) cases of Ca2+ imaging of cultured astrocytes acutely exposed to immunoglobulin G (IgG) isolated from ALS patients, showing a characteristic and specific response compared to the response to ATP as demonstrated in the same experiment. Examples also show a more pronounced transient rise in intracellular calcium concentration evoked by ALS IgG in hSOD1G93A astrocytes compared to non-transgenic controls; (2) Ca2+ imaging of cultured astrocytes during a depletion of calcium stores by thapsigargin (Thg), a non-competitive inhibitor of the endoplasmic reticulum Ca2+ ATPase, followed by store-operated calcium entry elicited by the addition of calcium in the recording solution, which demonstrates the difference between Ca2+ store operation in hSOD1G93A and in non-transgenic astrocytes; (3) Ca2+ imaging of the cultured microglia showing predominantly a lack of response to ALS IgG, whereas ATP application elicited a Ca2+ change. This paper also emphasizes possible caveats and cautions regarding critical cell density and purity of cultures, choosing the correct concentration of the Ca2+ dye and dye-loading techniques.

We present here a protocol on how to prepare primary cultures of glial cells, astrocytes, and microglia from rat cortices for time-lapse video imaging of intracellular Ca2+ for research on pathophysiology of amyotrophic lateral sclerosis in the hSOD1G93A rat model.

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to ...

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

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

Measuring Glucose Uptake in Drosophila Models of TDP-43 Proteinopathy

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis