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

Summary

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.

Abstract

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.

Introduction

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 cells^3,4. Clinically, as patients lose connectivity of the remaining upper and lower motor neurons, they present with features of neuronal hyper- and hypoexcitability throughout the disease^5,6,7,8,9, 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, FUS^11,12, C9orf72^13,14,15,16, and TDP43^17,18,19 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 synapses^20,24. 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 manner^25,26,27. 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 AM^30,31,32. 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 region^35,36. 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 moiety^35,36. 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 properties^37,38. 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 cell^28,39. 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 environment^41,42. Thus, it is an ideal tool for labeling recycling synaptic vesicles and subsequent tracking of these vesicles as they are later released following neuronal stimulation^41,42. 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 types^43,44. 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) promoter^37,46. 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 measured^37,46. 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 cells^{41,42,43,47,48}. 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 neurons^{48,49,50,51,52,53,54,55,56}. 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.

Figure 1: 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. Please click here to view a larger version of this figure.

Protocol

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 described^57,58. No strain bias appears to exist with the success of this culturing protocol. This method is described briefly below. The previous articles indicated should be referenced for complete details.

1. Euthanize pregnant female rats by CO₂ inhalation followed by secondary confirmation by cervical dislocation.
2. Harvest embryos and isolate brains in ice-cold 20 mM HEPES-buffered Hank's balanced salt solution (HBSS). From the outer cortex shell, separate and then discard striatum and hippocampi. Collect cortices.
3. Use fine forceps to remove meninges from cortices. To do this, hold the cortex in place with one pair of closed forceps using gentle pressure.
   NOTE: With the second pair of forceps in the second hand, pinch meninges. Peel away meninges using rolling motion with pinched pair. Reposition with the closed forceps and repeat as necessary until meninges is entirely removed.
4. Incubate cortices with 10 µg/mL of papain in HBSS for 4 min at 37 °C.
5. Wash three times in HBSS, and then triturate gently 5-10 times with a fire-polished glass Pasteur pipette to obtain a homogenous cell suspension.
   ​NOTE: Avoid bubbles; maintain the pipette within the cell suspension.
6. Plate neurons on 100 µg/mL poly-D-lysine coated 35 mm glass-bottom dishes at a density of 75,000 cells/dish in neurobasal medium with B27 supplement (2%), and penicillin-streptomycin.

2. Primary culture of motor neurons from embryonic rat spinal cord

NOTE: Primary motor neurons are prepared from E13.5 rat embryos as previously described, with few modifications^59,60. No strain bias appears to exist with the success of this culturing protocol. This method is described briefly below. The preceding articles indicated should be referenced for complete details.

1. Euthanize pregnant female rats as in step 1.1.
2. Dissect 10-20 spinal cords from embryos and break into small fragments mechanically using two pairs of forceps to pinch and pull, respectively.
3. Incubate in 0.025% trypsin for 8 min, followed by addition of DNase at 1 mg/mL.
4. Centrifuge through a 4% w/v BSA cushion at 470 x g for 5 min at 4 °C with no break.
5. Centrifuge pelleted cells for 55 min at 830 x g at 4 °C without brake through a 10.4% (v/v) density gradient medium (see Table of Materials) cushion. Carry forward the motor neuron band at the visual interface.
   NOTE: To ensure capture of all cells, use phenol-free media for the density gradient step and phenol-containing media for all the other steps. The interface of the density gradient and dissociation media is easily identifiable based on the color difference.
6. Spin collected bands through another 4% w/v BSA cushion at 470 x g for 5 min at 4 °C with no break.
7. Resuspend purified motor neurons in complete neurobasal medium with B27 supplement (2%), glutamine (0.25%), 2-mercaptoethanol (0.1%), horse serum (2%), and penicillin-streptomycin.
8. Plate neurons on 100 µg/mL of poly-lysine and 3 µg/mL of laminin-coated coverslips at a density of 50,000 cells/35 mm glass-bottom dish.

3. Neuronal transfections

NOTE: This step is carried out for neurons that will undergo GCaMP calcium imaging and/or neurons in which an exogenous DNA plasmid or RNA of interest is introduced in advance of synaptic evaluation. In contrast, styryl dye is loaded just before the imaging session and is addressed in section 6. The summary below is the transfection protocol used for primary rodent neurons in the presented examples, but it can easily be adapted and optimized to the user's needs.

1. Transfections for cultured primary cortical neurons occur at day 12 in vitro (DIV 12), whereas motor neurons are transfected at day 7 in vitro (DIV 7).
   NOTE: All the following steps occur within an aseptic biosafety cabinet.
2. Transfect 500 ng of GCaMP6m using transfection reagent at a ratio of 1:2 by volume. For co-transfections, add a total of 1.25 µg of total DNA/dish, maintaining this DNA to reagent ratio.
   NOTE: In the experimental examples shown, 750 ng of ALS/FTD-related plasmid of interest has been transfected to express C9orf72-ALS linked dipeptides, mutant FUS protein, etc. Ensure that the fluorescent tag of the co-transfected plasmid will not conflict with the FITC channel of GCaMP imaging. Likewise, if doing styryl dye imaging, ensure that the co-transfected plasmid will not conflict with the TRITC channel of imaging. Usage of synaptophysin-GCaMP3 is equally effective in this protocol. Therefore, transfect the same amount of this plasmid and follow all the steps as usual in section 7.
3. Incubate the DNA-transfection reagent complex at room temperature for 10 min.
4. Gently add the incubated complex to neurons dropwise with swirling. Return the dish to CO₂ incubator at 37 °C for 45-60 min.
5. Remove the entire culture media containing the DNA-transfection reagent complex and replace it with a 50-50 by volume preparation of pre-conditioned and fresh neuronal media. Then, put cells back into the CO₂ incubator for 48 h until imaging.

4. Preparation of buffer solutions and styryl dye stock solution

1. Make low and high aCSF solutions fresh before imaging using the ingredients listed in Table 1. Filter both the buffer solutions before use.
   NOTE: CaCl₂ dihydrate can form Ca(OH)₂ upon storage. For maximum efficacy, always use a recently opened container. If this is not possible, to remove Ca(OH)₂ from the external surface and ensure maximum solubility, solutions may be carbonated with gaseous CO₂ for 5 min before CaCl₂ addition. If this step is taken, adjust the pH of the resulting solution to maintain a pH value of 7.4, as excessive carbonation will result in Ca(HCO₃)₂ formation.

Low KCL aCSF buffer (pH 7.40 to 7.45)
Reagent	Concentration
HEPES	10 mM
NaCl	140 mM
KCl	5 mM
Glucose	10 mM
CaCl2 2H2O	2 mM
MgCl2 4H2O	1 mM
High KCL aCSF buffer (pH 7.40 to 7.45)
Reagent	Concentration
HEPES	10 mM
NaCl	95 mM
KCl	50 mM
Glucose	10 mM
CaCl2 2H2O	2 mM
MgCl2 4H2O	1 mM

Table 1: Composition of artificial cerebrospinal fluid (aCSF) buffers. This table includes the ingredients for preparing low and high KCl artificial cerebrospinal fluid buffers used while imaging and stimulating neurons. See section 4 for preparation instructions.

2. Prepare styryl dye stock solution by taking a 100 µg vial of dye and reconstituting it to a stock concentration of 10 mM with distilled water or neurobasal medium.

5. Microscope and perfusion system setup

NOTE: For imaging glass-bottom Petri dishes, an inverted confocal fluorescence microscope is preferred due to the flexibility of perfusion and for the use of a high numerical aperture oil immersion objective. Refer to the Table of Materials for the confocal microscope, camera, and objectives used for imaging of examples detailed in the Representative Results section.

1. Perform all the experiments at 37 °C with constant 5% CO₂ levels using an incubator system-coupled stage mount.
2. Control image acquisition using confocal software. Optimize acquisition settings at the start of the imaging session to choose excitation power and gain to ensure optimal visualization of signals without photobleaching.
   1. Keep excitation power, exposure time, detector gain and frame rate constant across all samples. Carry out time-lapse imaging using an aspect ratio of 512 x 512 and frame rate of 2 images/s to minimize dye bleaching.
      NOTE: While puncta should be clearly visible, laser intensity should be set to the minimum possible to avoid bleaching and phototoxicity. Set the confocal aperture to the narrowest setting to achieve optimal resolution of fluorescent puncta within neurites. Exposure time is set to 200 ms or less, consistent across samples. The maximum imaging speed of the camera used was 2 images/s. If using a faster camera, more images/s can be taken. Temporal imaging settings should be chosen if possible.
3. Select the following fluorescence excitation/dichroic/emission filter combinations for imaging using confocal software: Gcamp6m/Gcamp3 with FITC and styryl dye with TRITC, respectively.
4. Use the perfect focus feature of the confocal imaging acquisition software during time-lapse imaging to avoid z-drift.
   NOTE: Due to the rapid speed of imaging, a single plane is imaged. Ensuring the lack of z-drift during the experiment is very important.
5. Select the Time tab in the image acquisition panel to set the recording periods and intervals.
   1. Set Phase #1 to Interval 500 ms, Duration 3 - 5 min.
   2. Set Phase #2 to Interval 500 ms, Duration 5 min.
      NOTE: Phase #1 corresponds to baseline recording, Phase #2 to the stimulation period, respectively.
6. Assemble gravity perfusion apparatus for aCSF using a valve control system and a channel manifold.
   1. Load high KCl (see Table 1) into a 50 mL syringe at the top of the apparatus, with tubing running through the system. Set the flow rate to 1 mL/min.
7. Load a 35 mm glass dish containing neurons onto the confocal imaging stage, with the end of the perfusion tubing placed at the dish edge. Choose the field for imaging.

6. Styryl dye imaging of synaptic vesicle release

1. Incubate cells in low KCl aCSF (see Table 1) for 10 min at 37 °C, 48 h post-transfection.
2. Load primary cortical or motor neurons on glass-bottom Petri dishes with styryl dye.
   1. Remove low KCl aCSF by aspiration.
   2. Use a pipette to load neurons in the dark with 10 µM of styryl dye in aCSF containing 50 mM KCl for 5 min.
   3. Remove the loading solution and bath neurons in low KCl aCSF for 10 min to eliminate non-specific dye loading.
3. Place the 35 mm glass-bottom dish onto the imaging stage, and then observe either under a 20x air objective or 40x oil immersion objective of an inverted confocal microscope.
   NOTE: Use GFP fluorescence to locate transfected cells in case of cells co-transfected with a GFP-tagged plasmid.
4. Excite styryl dye using a 546 nm laser, collect emission using a 630-730 nm (TRITC) bandpass filter.
5. Select the imaging field and engage perfect focus. Next, take a single still image with brightfield, TRITC, and fluorescence marker channels to mark neuronal boundaries.
6. Initiate Run Now in the acquisition software. Carry out the basal recording for 3-5 min to exclude variations in dye intensity, if any (Phase #1).
   NOTE: It is essential to ensure that any decrease in dye intensity is due to synaptic vesicle release and not passive dye diffusion or photobleaching. This 3-5 min pre-stimulation period should result in a steady and maintained dye intensity level. The final 30 s of this recording will be used to determine a mean baseline fluorescence value for each ROI. Before evaluating experimental conditions, an additional non-stimulation condition of approximately 10 min of continuous recording using the intended acquisition settings can also be performed to ensure steady fluorescence values using the laser settings for an extended period.
7. At the switch to Phase #2, trigger On the button for the perfusion system. Then, constantly perfuse 50 mM KCl to neurons to facilitate dye unloading (Phase #2).
   1. Carry out recordings for 300 s after KCl addition. After this, acquisition stops; trigger the Off switch for the perfusion system.
8. Save the experiment and analyze data using confocal software as described in section 8 below.
   ​NOTE: The experiment may be stopped at this point, and analysis can be performed later. In the case where cells do not require transfection for the expression of proteins of interest, this protocol may be performed at any timepoint in vitro when seeking to examine the functionality of synaptic unloading. Following a test of control cells to ensure proper synaptic unloading, the experimenter should be blinded to the genetic or pharmacologic conditions of each dish tested to minimize bias.

7. Fluorescence imaging of Gcamp6m calcium transients

1. Transfect primary rodent cortical neurons cultured on 35 mm glass-bottom dishes with 500 ng of Gcamp6m as indicated in section 3.
   NOTE: If desired, co-transfect neurons with a plasmid of interest containing a fluorescent tag in the red or far-red range.
2. Incubate neurons with low KCl aCSF for 15 min 48 h post-transfection and then mount the dish on the imaging platform.
3. Visualize GCaMP6m fluorescence using a FITC filter (488 nm) and a 20x or 40x objective.
4. Select the imaging field and engage perfect focus. Next, take a single still image with brightfield, FITC, and fluorescence marker channels to mark neuronal boundaries.
5. Initiate Run Now in the acquisition software. Carry out the baseline recording for 5 min, and then perfuse with aCSF containing 50 mM KCL in the same manner as described in section 6 for styryl dye experiments.
   NOTE: The goal of this imaging is to measure evoked calcium transients. Should a neuron have basal firing activity and calcium fluxes during the pre-stimulation period, it is not used in data analysis. Instead, only cells with stable background fluorescence are used. Post-stimulation periods can also be extended to 60 min for calcium transients, with or without additional continuous high KCl perfusion.
6. Save the experiment and analyze data using confocal software as described in section 8 below. The experiment may be stopped at this point, and the analysis can be performed later.
   ​NOTE: If the cells do not require transfection for expression of proteins of interest, this protocol may be performed at any timepoint in vitro when seeking to examine calcium transients. Following a test of control cells to ensure measurement of calcium transients, the experimenter should be blinded to the genetic or pharmacologic conditions of each dish tested to minimize bias.

8. Image analysis

1. Open time-lapse images with confocal software.
2. Align images in time-lapse series by the command series : Image | Processing | Align Current Document. Select Align to the First Frame.
3. Select regions of interest (ROIs) along neurites using the ROI selection tool, a bean-shaped icon on the right of the image frame (Figure 3B). Also, mark an ROI representing background fluorescence intensity.
   NOTE: ROIs are selected by choosing areas of distinctly separated puncta along neuronal tracks indicated by the brightfield still image. At least five ROIs per neuron are chosen for analysis. The background ROI is chosen in a region of the field of view that does not contain neurites.
4. Measure raw fluorescence over time for selected ROIs using the following command series: Measure | Time Measurement.
   1. Initiate the Measure function at the upper portion of the Time Measurement panel. A graphical representation of raw fluorescence over time (Figure 3C) and quantitative data are both generated.
      NOTE: Each data point represents the raw fluorescence for that ROI for the frame associated with that specific measured time point.
5. Export raw fluorescence intensities to the spreadsheet software.
6. Analyze each ROI independently. First, normalize data by subtracting the background ROI intensity from the ROI of interest intensity at each time point.
   1. Take the raw fluorescence value from the background ROI at each specific time point and subtract this from the ROI of interest raw intensity value at that time point.
      NOTE: This is done for the entire recording period, both basal and stimulation phases.
7. Determine the average ROI of interest intensity for the last 30 s of baseline.
   1. Average the normalized raw basal values generated in step 8.6 from the final 30 s of the 3-5 min basal recording period.
      NOTE: The entire period of basal recording could be used to generate this value. However, as this value stabilizes and is maintained, the 60-time points of the final 30 s are of sufficient sampling size to represent the whole.
8. Compare this baseline value to the normalized intensity value at each time point, generating a change in fluorescence (ΔF) value.
   1. Take the value from step 8.7 and subtract the average baseline fluorescent value. Do this and subsequent steps for the entire recording period, both basal and stimulation phases.
9. Calculate this change concerning the baseline fluorescence value, generating a change in fluorescence/baseline fluorescence (ΔF/F). To do this, take the value from step 8.8 and divide it by the average baseline fluorescent value.
10. Finally, set the starting point value to 1 so that increases or decreases can be easily visualized graphically over time.
    1. Take the value generated in step 8.9 and add 1.
       ​NOTE: When this is done correctly, the 30 s of baseline period values should hover near the value of 1. In cells effectively releasing synaptic content, this value should trend from 1 to 0 following the start of stimulation. An example of steps 6-9 is presented in Figure 3E.

9. Data analysis

NOTE: The experimenter can be unblinded to experimental conditions to pool data for analysis appropriately. Use a sample size of at least 10 neurons per condition from each of three independent experiments. Only consider neurons for inclusion if at least five ROI regions can be designated. This level of experimental replication was sufficient in published studies to demonstrate a profound loss of synaptic unloading in ALS-related poly-GA-containing cells versus GFP controls (see Representative Results). However, if a more subtle phenotype is observed, the number of biological and/or technical replicates may require optimization by the user.

1. Using all calculated ΔF/F values over time from ROIs of a given experimental condition, determine a mean ΔF/F value for each time point along with an SEM.
2. Plot data using a graphing and statistics software program in xy-format, where x is the time elapsed, and y is the ΔF/F value calculated in step 9.1. Present all the values as mean ± SEM.
3. To determine statistical significance, use the Student's t-test for comparing two groups and one-way analysis of variance (ANOVA) followed by Tukey's posthoc analysis for comparing three or more groups.

Results

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...

Discussion

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...

Materials

List of materials used in this article

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