We thank Steve Finkbeiner and members of the Finkbeiner lab for pioneering robotic microscopy. We also thank Dan Peisach for building the initial software required for image processing and automated survival analysis. This work was funded by the National Institute for Neurological Disorders and Stroke (NINDS) R01-NS097542, the University of Michigan Protein Folding Disease Initiative, and Ann Arbor Active Against ALS.

All vertebrate animal work was approved by the Committee on the Use and Care of Animals at the University of Michigan (protocol # PRO00007096). Experiments are carefully planned to minimize the number of animals sacrificed. Pregnant female wild-type (WT), non-transgenic Long Evans rats (Rattus norvegicus) are housed singly in chambers equipped with environmental enrichment, and cared for by the Unit for Laboratory Animal Medicine (ULAM) at the University of Michigan, in accordance with the NIH-supported Guide for the Care and Use of Laboratory Animals. All rats were kept in routine housing for as little time as possible prior to euthanasia, consistent with the recommendations of the Guidelines on Euthanasia of the American Veterinary Medical Association and the University of Michigan Methods of Euthanasia by Species Guidelines.
1. Material Preparation
<ol>
	<li>Dissect cortical neurons from embryonic day 19&#8211;20 rat pups and culture rat cortical neurons at 0.5 x 106 cells per milliliter on poly-D-lysine coated plates for 4 days in vitro, as described previously13,14,15,16,17,18,19.</li>
	<li>Prepare the plasmid DNA of interest following the steps outlined by an endotoxin-free plasmid DNA isolation kit (see Table of Materials). Quantify the resultant DNA using a spectrophotometer.</li>
	<li>On in vitro day 4 (DIV4), aliquot, filter sterilize, and incubate the following media at 37 &#176;C: 6 mL reduced serum media (RSM; e.g., OptiMEM), 25 mL neuronal basal media (NBM), 40 mL NBKY (NBM + 1 mM kynurenic acid + 10 mM MgCl2, adjusted to a pH of 7.4), 10 mL NBC (NBM + 1x neuronal cell culture supplement + 1x L-glutamine supplement + 1x Pen Strep). 
	NOTE: Volumes listed are sufficient for transfecting one 96-well plate. Refer to the Table of Materials for specific reagents.</li>
</ol>
2. Transfection of Rat Cortical Neurons
<ol>
	<li>Modify the provided Example transfection sheet (see Supplemental File 1) by adjusting the plate type, plate map, number of DNAs, DNA concentration, and number of wells (green boxes). 
	NOTE: The total DNA sums to 0.2 &#181;g per well, regardless of whether one (e.g., DNA A) or multiple (e.g., DNA B and C) DNA constructs are added to each well.</li>
	<li>Working from the spreadsheet, combine the appropriate amount of RSM and DNA in one tube. Combine the appropriate amount of RSM and transfection reagent (e.g., Lipofectamine) in a separate tube.</li>
	<li>Incubate at room temperature (RT) for 5 min.</li>
	<li>Combine the DNA and transfection reagent RSM mixtures and incubate at RT for 20 min.</li>
	<li>During this incubation step, use a multichannel pipette and sterile plastic troughs to wash cells 2x with 100 &#181;L per well of NBM. Reserve the conditioned media (CM) and store at 37 &#176;C. For this and following steps, take care to minimize the amount of time neurons are exposed to air.</li>
	<li>Remove the NBM media and replace with 100 &#181;L per well of NBKY.</li>
	<li>After 20 min have passed, add 50 &#181;L of the transfection reagent/DNA mixture dropwise to each well.</li>
	<li>Incubate cells with the transfection reagent/DNA complexes for 20 min at 37 &#176;C.</li>
	<li>Rinse 2x with NBKY and replace with 100 &#181;L of CM and 100 &#181;L of NBC per well.</li>
	<li>Successfully transfected cells should be visible by fluorescence microscopy within 16&#8211;24 h of transfection. To gauge efficiency, use a fluorescent microscope to check the transfection after overnight incubation at 37 &#176;C. 
	NOTE: This technique results in an overall transfection efficiency of 5 to 10%.</li>
</ol>
3. Imaging
<ol>
	<li>Place the plate on a fluorescent microscope with a motorized stage, and establish a fiduciary (e.g., a mark on the bottom of the plate) that will allow the user to align the plate each time it is imaged. Save an image of this fiduciary for reference.</li>
	<li>Navigate to a field of interest and note the x-y coordinates relative to the fiduciary.</li>
	<li>Focus on transfected cells expressing a fluorescent label.</li>
	<li>Take fluorescent images in the appropriate channel or channels (e.g., red fluorescent protein [RFP], green fluorescent protein [GFP], 4&#8242;,6-diamidino-2-phenylindole [DAPI]), either manually or in an automated manner. By taking several images at regularly-spaced intervals, a montage of the well can be assembled during image processing (see step 4). 
	NOTE: The spacing depends on several factors, including magnification, the optics of the microscope, and the detector size. In general, the optical spacing between adjacent images will be between 90&#8211;95% of the size of each individual image, to allow for a small degree of image overlap and feature alignment.</li>
	<li>Repeat this process as often as required, aligning to the original fiduciary each time. For survival analysis, imaging takes place every 6&#8211;24 h, depending on the cell type and the purpose of the experiment.</li>
</ol>
4. Image Processing
NOTE: Following image acquisition, a series of processing steps are required prior to image analysis. These include, but are not limited to, stitching, stacking, and background subtraction (Figure 1). The goal of these steps is to produce an image stack, or time series, in which cells are clearly discernible from their background and easy to follow over multiple time points. A dedicated FIJI macro (Image_Processing.ijm, see Supplemental File 2), performs basic stitching, stacking, and background subtraction. An explanation of each step and the parameters to consider when performing image processing is provided in the discussion section.
<ol>
	<li>Adjust the raw data or input directory to match the formatting shown in Figure 2.</li>
	<li>If time points are not contiguous (i.e., T1, T2, T3), rename these folders so that they are. This step is critical to ensure that the Image_Processing macro does not crash during stacking.</li>
	<li>Double-click on the Fiji icon to open the program, then click and drag the Image_Processing macro onto the Fiji bar. This will open the macro within Fiji.</li>
	<li>Adjust lines 2-7 of the Image_Processing macro to specify the input directory containing images, the desired output directory for stitched and stacked images, the number of imaging timepoints, number of fluorescent channels and plate format.</li>
	<li>Determine the order in which the images were acquired. To test this, manually stitch a montage of images in FIJI by maneuvering to the Plugins drop down menu | Stitching | Grid/Collection stitching. Adjust the settings within the dropdown menus Type and Order until an accurately stitched image is produced.</li>
	<li>Adjust the GRID_TYPE and STITCH_ORDER variables in lines 8 and 9 of the Image_Processing macro to match these selections.</li>
	<li>Specify the number of images per well by adjusting line 10 in the Image_Processing macro. 
	NOTE: For a 2 x 2 montage of images, this line would read DIM = 2.</li>
	<li>If background subtraction is required, adjust line 14 in the Image_Processing macro to BGSUB = true.</li>
	<li>Set the rolling ball radius by adjusting line 15 in the Image_Processing macro. 
	NOTE: For optimal results, set the radius to at least the diameter of the largest foreground object in the image.</li>
	<li>Click Run to start the Image_Processing macro. Once started, Image_Processing will automatically advance through stitching, stacking, and background subtraction.</li>
</ol>
5. Scoring Cell Death
NOTE: See the Discussion section for more information on scoring cell death and censoring data.
<ol>
	<li>Locate the image stacks produced by the Image_Processing script. Open these in FIJI.</li>
	<li>Use the point tool within FIJI to individually label each cell with a number. Pressing t after each point will add the cell identifier to the ROI (region-of-interest) Manager. 
	NOTE: The identifiers can be visualized by clicking the labels and show all checkboxes in the ROI Manager.</li>
	<li>Progress through the timepoints in each image stack and record the timepoint when each cell either dies or needs to be censored in the file Survival_spreadsheet.csv (see Supplemental File 3).
	<ol>
		<li>Each cell occupies a single row in the spreadsheet, where a unique identifier (ID) for each cell consists of its corresponding well and ROI number within that well. tp_death is the last time point a cell is observed to be alive, while time_death represents the actual time of death in hours. For each cell, input these data. It is critical to maintain this structure for subsequent analysis using survival.R (see Supplemental File 4). 
		NOTE: The criteria for determining cell death are crucial and may vary depending on cell type. Three main criteria are used in the identification of dead neurons11,20 (Figure 3): loss of fluorescence intensity (e.g., Neuron 1 at 69 h), rounding of the cell body (e.g., Neuron 2 at 188 h), and the loss of neurite integrity or blebbing (e.g., Neuron 2 at 188 h).</li>
	</ol>
	</li>
	<li>Record the censor status of the cell in the last column. 
	NOTE: Here, due to the peculiar way censoring is handled by R, censored cells are marked by 0, while uncensored cells are marked by 1. Note that all cells that live to the last time point are censored, and therefore marked as 0.</li>
</ol>
6. Performing Cox Proportional Hazards Analysis and Visualizing Results
<ol>
	<li>If necessary, download R studio at https://cran.r-project.org/mirrors.html.</li>
	<li>Open R studio and double-click the icon for the survival.R script.</li>
	<li>Place the cursor on line 2 of survival.R and click the run button in the main R studio window in order to load the survival library.</li>
	<li>Change line 5 of the survival.R script to match the location of the file Survival_spreadsheet.csv. Click on run to load the survival data as a dataframe.</li>
	<li>Highlight lines 8 and 9 and click the run button in the R studio console window in order to perform Cox proportional hazards analysis. Results and output statistics appear in the console window of R studio.</li>
	<li>Highlight lines 12-16 of survival.R and hit the run button in order to produce a cumulative risk of death plot, which will appear in the plots tab in R studio. This file can be saved by clicking on the export button above the plot.</li>
	<li>If it is desirable to plot the survival data as a Kaplan-Meier curve, highlight lines 19-24 of survival.R and hit the run button.</li>
</ol>

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Flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein labelled Annexin V.</a> Journal of Immunological Methods. 184, 39-51 (1995).</li><li>Chien, K. R., Abrams, J., Serroni, A., Martin, J. T., Farber, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Accelerated+phospholipid+degradation+and+associated+membrane+dysfunction+in+irreversible,+ischemic+liver+cell+injury.">Accelerated phospholipid degradation and associated membrane dysfunction in irreversible, ischemic liver cell injury.</a> Journal of Biological Chemistry. 253, 4809-4817 (1978).</li><li>Zwaal, R. F. A., Comfurius, P., Bevers, E. 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B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+Vitro+Modulation+of+TrkB+Receptor+Signaling+upon+Sequential+Delivery+of+Curcumin-DHA+Loaded+Carriers+Towards+Promoting+Neuronal+Survival.">In Vitro Modulation of TrkB Receptor Signaling upon Sequential Delivery of Curcumin-DHA Loaded Carriers Towards Promoting Neuronal Survival.</a> Pharmaceutical Research. 34 (2), 492-505 (2017).</li><li>Arrasate, M., Finkbeiner, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+microscope+system+for+determining+factors+that+predict+neuronal+fate.">Automated microscope system for determining factors that predict neuronal fate.</a> Proceedings of the National Academy of Science of the United States of America. 102, 3840-3845 (2005).</li><li>Christensen, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multivariate+survival+analysis+using+Cox&#8217;s+regression+model.">Multivariate survival analysis using Cox&#8217;s regression model.</a> Hepatology. 7, 1346-1358 (1987).</li><li>Barmada, S. 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The authors have nothing to disclose.

Here, methodology to directly monitor neuronal survival on a single-cell basis is presented. In contrast to traditional assays for cell death that are limited to discrete time points and entire populations of cells, this method allows for the continuous assessment of a variety of factors such as cellular morphology, protein expression, or localization, and can determine how each factor influences cellular survival in a prospective manner.
This methodology can be modified to fit a wide array of...

Abnormal cell death is a driving factor in many diseases, including cancer, neurodegeneration, and stroke1. Robust and sensitive assays for cell death are essential to the characterization of these disorders, as well as the development of therapeutic strategies for extending or reducing cellular survival. There are currently dozens of techniques for measuring cell death, either directly or through surrogate markers2. For example, cell death can be assessed visually with the help of vital dyes that selectively stain dead or living cells3, or by monitoring the appearance of specific phospholipids on the plasma membrane4,5,6. Measurements of intracellular components or cellular metabolites released into the media upon cellular dissolution can also be used as proxies for cell death7,8. Alternatively, cellular viability can be approximated by assessing metabolic activity9,10. Though these methods provide rapid means of assessing cell survival, they are not without caveats. Each technique observes the culture as a single population, rendering it impossible to distinguish between individual cells and their unique rates of survival. Furthermore, such population-based assays are unable to measure factors that may be important for cell death, including cellular morphology, protein expression, or localization. In many cases, these assays are limited to discrete time points, and do not allow for the continuous observation of cells over time.
&#160;In contrast, longitudinal fluorescence microscopy is a highly flexible methodology that directly and continuously monitors the risk of death on a single-cell basis11. In brief, longitudinal fluorescence microscopy enables thousands of individual cells to be tracked at regular intervals for extended periods of time, allowing precise determinations of cell death and the factors that enhance or suppress cell death. At its base, the method involves the transient transfection or transduction of cells with vectors encoding fluorescent proteins. A unique fiduciary is then established, and the position of each transfected cell in relation to this landmark allows the user to image and track individual cells over the course of hours, days, or weeks. When these images are viewed sequentially, cell death is marked by characteristic changes in fluorescence, morphology, and fragmentation of the cell body, enabling the assignment of a time of death for each cell. The calculated rate of death, determined by the hazard function, can then be quantitatively compared between conditions, or related to select cellular characteristics using univariate or multivariate Cox proportional hazards analysis12. Together, these approaches enable the accurate and objective discrimination of rates of cell death among cellular populations, and the identification of variables that significantly predict cell death and/or survival (Figure 1).
Although this method can be used to monitor survival in any post-mitotic cell type in a variety of plating formats, this protocol will describe conditions for transfecting and imaging rat cortical neurons cultured in a 96-well plate.

<table>
	<tbody>
		<tr>
			<td>Neurobasal Medium</td>
			<td>GIBCO</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM</td>
			<td>GIBCO</td>
			<td>31985-070</td>
			<td></td>
		</tr>
		<tr>
			<td>CompactPrep Plasmid Maxi Kit</td>
			<td>Qiagen</td>
			<td>12863</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride Hexahydrate</td>
			<td>Sigma</td>
			<td>M9272</td>
			<td></td>
		</tr>
		<tr>
			<td>Kynurenic Acid Hydrate</td>
			<td>TCI</td>
			<td>H0303</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly D-Lysine</td>
			<td>Millipore</td>
			<td>A-003-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamax</td>
			<td>GIBCO</td>
			<td>35050-061</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>Invitrogen</td>
			<td>52887</td>
			<td></td>
		</tr>
		<tr>
			<td>B27 supplement</td>
			<td>Thermo Fisher</td>
			<td>A3582801</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin Streptomycin</td>
			<td>GIBCO</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well plates</td>
			<td>TPP</td>
			<td>0876</td>
			<td></td>
		</tr>
	</tbody>
</table>

monitoring neuronal survival via longitudinal fluorescence microscopy

Standard cytotoxicity assays, which require the collection of lysates or fixed cells at multiple time points, have limited sensitivity and capacity to assess factors that influence neuronal fate. These assays require the observation of separate populations of cells at discrete time points. As a result, individual cells cannot be followed prospectively over time, severely limiting the ability to discriminate whether subcellular events, such as puncta formation or protein mislocalization, are pathogenic drivers of disease, homeostatic responses, or merely coincidental phenomena. Single-cell longitudinal microscopy overcomes these limitations, allowing the researcher to determine differences in survival between populations and draw causal relationships with enhanced sensitivity. This video guide will outline a representative workflow for experiments measuring single-cell survival of rat primary cortical neurons expressing a fluorescent protein marker. The viewer will learn how to achieve high-efficiency transfections, collect and process images enabling the prospective tracking of individual cells, and compare the relative survival of neuronal populations using Cox proportional hazards analysis.

Using the transfection procedure described here, DIV4 rat cortical neurons were transfected with a plasmid encoding the fluorescent protein mApple. Beginning 24 h post-transfection, cells were imaged by fluorescence microscopy every 24 h for 10 consecutive days. The resultant images were organized as indicated in Figure 2, then stitched, stacked, and scored for cell death (Figure 1). Figure 3 shows a...

Watch this Scientific Journal Video about Monitoring Neuronal Survival via Longitudinal Fluorescence Microscopy at JoVE.com

Monitoring Neuronal Survival via Longitudinal Fluorescence Microscopy

Here, we present a protocol to monitor survival on a single-cell basis and identify variables that significantly predict cell death.

Standard cytotoxicity assays, which require the collection of lysates or fixed cells at multiple time points, have limited sensitivity and capacity to ...

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6.0K Views.  University of Michigan School of Medicine. Here, we present a protocol to monitor survival on a single-cell basis and identify variables that significantly predict cell death.

Video: Monitoring Neuronal Survival via Longitudinal Fluorescence Microscopy

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

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. ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a closed-loop event-triggered manner. Although many studies are focusing on the possible benefits and side-effects of TES in healthy and pathologic brains, there are still many fundamental open questions regarding the mechanism of action of the stimulation. Therefore, there is a clear need for a robust and reproducible method to test the acute and the chronic effects of TES in rodents. TES can be combined with regular behavioral, electrophysiological, and imaging techniques to investigate neuronal networks in vivo. The implantation of transcranial stimulation electrodes does not impose extra constraints on the experimental design while it offers a versatile, flexible tool to manipulate brain activity. Here we provide a detailed, step-by-step protocol to fabricate and implant transcranial stimulation electrodes to influence brain activity in a temporally constrained manner for months.

This detailed protocol describes transcranial stimulation electrode placement on the temporal bone in order to investigate the short- and long-term effects of transcranial electrical stimulation in freely moving rats.

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a ...

In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

This technique combines optical fiber mediated fluorescence recordings with the precise delivery of recombinant adeno-associated virus based gene reporters. This new and easy to use in vivo fluorescence monitoring system was developed to record the transcriptional rhythm of the clock gene, Cry1, in the suprachiasmatic nucleus (SCN) of freely moving mice. To do so, a Cry1 transcription fluorescence reporter was designed and packaged into Adeno-associated virus. Purified, concentrated virus was injected into the mouse SCN followed by the insertion of an optic fiber, which was then fixed onto the surface of the brain. The animals were returned to their home cages and allowed a 1-month post-operative recovery period to ensure sufficient reporter expression. Fluorescence was then recorded in freely moving mice via an in vivo monitoring system that was constructed at our institution. For the in vivo recording system, a 488 nm laser was coupled with a 1 &#215; 4 beam-splitter that divided the light into four laser excitation outputs of equal power. This setup enabled us to record from four animals simultaneously. Each of the emitted fluorescence signals was collected via a photomultiplier tube and a data acquisition card. In contrast to the previous bioluminescence in vivo circadian clock recording technique, this fluorescence in vivo recording system allowed the recording of circadian clock gene expression during the light cycle.

In Vivo Monitoring of Circadian Clock Gene Expression in the Mouse Suprachiasmatic Nucleus Using Fluorescence Reporters

This newly developed fluorescence-based technology enables long-term monitoring of the transcription of circadian clock genes in the suprachiasmatic nucleus (SCN) of freely moving mice in real-time and at a high temporal resolution.

This technique combines optical fiber mediated fluorescence recordings with the precise delivery of recombinant adeno-associated virus based gene ...

Brain Death Induction in Mice Using Intra-Arterial Blood Pressure Monitoring and Ventilation via Tracheostomy

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain death (BD) still represents the major source for solid transplants. Unfortunately, the irreversible loss of brain function is known to induce multiple pathophysiological changes, including hemodynamic as well as hormonal modifications, finally leading to a systemic inflammatory response. Models that allow a systematic investigation of these effects in vivo are scarce. We present a murine model of BD induction, which could aid investigations into the devastating effects of BD on allograft quality. After implementing intra-arterial blood pressure measurement via the common carotid artery and reliable ventilation via a tracheostomy, BD is induced by steadily increasing intracranial pressure using a balloon catheter. Four hours after BD induction, organs may be harvested for analysis or for further transplantation procedures. Our strategy enables the comprehensive analysis of donor BD in a murine model, therefore allowing an in-depth understanding of BD-related effects in solid organ transplantation and potentially paving the way to optimized organ preconditioning.

We present a murine model of brain death induction in order to evaluate the influence of its pathophysiological effects on organs as well as on consecutive grafts in the context of solid organ transplantation.

While both living donation and donation after circulatory death provide alternative opportunities for organ transplantation, donation after donor brain ...

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.