Name: an enhanced green fluorescence protein-based assay for studying neurite outgrowth in primary neurons
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This work was supported by funds from the Research Grants Council Hong Kong, Health and Medical Research Fund (Hong Kong), CUHK direct grant scheme, the United College endowment fund and the TUYF Charitable Trust.

All procedures followed were in accordance with the ethical standards of the animal experimentation ethics committee of the Chinese University of Hong Kong.
1. Preparation of Coverslips
<ol>
	<li>Place a sterile 18 mm circular coverslip into each well of a 12-well tissue culture plate.</li>
	<li>Coat the coverslip with 5 &#181;g/mL poly-D-lysine solution in a humidified 37 &#176;C incubator for at least 1 h.</li>
	<li>Aspirate the poly-D-lysine solution from the tissue culture plate and rinse the coated coverslips once with sterile water.</li>
</ol>
2. Rat Embryonic Neuron Dissection
<ol>
	<li>Sacrifice a timed-pregnant Sprague-Dawley rat at a gestational age of 18 days (E18) by either cervical dislocation or CO2 asphyxiation. 
	NOTE: Please check local regulations for the sacrifice of pregnant rats.</li>
	<li>Open the abdominal cavity of the pregnant rat with dissecting scissors and transfer the uterus to a 10 cm Petri dish.</li>
	<li>Open the uterus and the amniotic sac carefully with small dissecting scissors and remove the placenta from the rat embryo using small dissecting scissors. Transfer the whole embryo to a 10 cm Petri dish with pre-chilled phosphate buffered saline supplemented with glucose (PBS-glucose, 10 mM sodium phosphates, 2.68 mM potassium chloride, 140 mM sodium chloride and 3 g/L glucose) using a pair of small forceps.</li>
	<li>Cut along the sagittal suture of the skull and open it carefully with a pair of small dissecting scissors. Transfer the embryonic brain with a small flat spatula to a 10 cm Petri dish with ice-cold PBS-glucose.</li>
	<li>Separate the two cerebral hemispheres from the cerebellum and brain stem using two #5 tweezers under a dissection microscope. 
	NOTE: Please see reference12 for the structure of the rat brain.</li>
	<li>Remove the meninges using the #5 tweezers.</li>
	<li>Isolate the cortex from the cerebral hemispheres with two straight #5 tweezers.</li>
	<li>Transfer the isolated cortex to ice-cold PBS-glucose in a 15 mL centrifuge tube.</li>
</ol>
3. Primary Cortical Neuron Culture
NOTE: All procedures in steps 3 and 4 are performed inside a Class II Biosafety cabinet.
<ol>
	<li>Settle the isolated cortex by gravity at 4 &#176;C for 5 min and aspirate the PBS-glucose.</li>
	<li>Add 1 mL of 0.05% Trypsin-EDTA to the isolated cortex and mix gently by tapping and incubate the tissue in a 37 &#176;C water bath for 10 min to allow enzymatic digestion. Tap the tube gently to mixing every 2 min.</li>
	<li>Add 4 mL of maintenance medium (e.g., Neurobasal Medium) to the tissue/trypsin mixture. 
	NOTE: All the maintenance medium used in this protocol is supplemented with Penicillin-Streptomycin and B-27 supplement13.</li>
	<li>Dissociate the tissue gently by trituration using a 1 mL pipette.</li>
	<li>Pellet the dissociated cells by centrifugation at 200 x g for 5 min. Aspirate the supernatant.</li>
	<li>Repeat steps 3.5 to 3.7 twice.</li>
	<li>Resuspend the cell pellet in 1 mL of maintenance medium.</li>
	<li>Add 10 &#181;L of 0.4% Trypan Blue solution to 10 &#181;L of cell suspension for counting of viable cells by a hemocytometer.</li>
	<li>Plate the neurons at a density of 65,000/cm2 (viable cells) in 1 mL of maintenance medium per well in a 12-well plate.</li>
</ol>
4. Cell Transfection and Fixation
<ol>
	<li>At 2 days in vitro (DIV2), transfect 0.5 &#181;g of EGFP construct (pEGFP-C1) to neurons together either with or without of 0.5 &#181;g of POI by using 1 &#181;L of transfection reagent (e.g., Lipofectamine 2000). Use manufacturer&#39;s instructions. 
	NOTE: Mammalian expression constructs were prepared by using an endotoxin free plasmid preparation kit. Treatment with chemicals/molecules (in this manuscript cytochalasin D (Cyto D) and nerve growth factor (NGF) were used) can be done at 6 h after transfection.</li>
	<li>Aspirate the culture medium 24 h post-transfection and wash the transfected cells once with 37 &#176;C PBS (10 mM sodium phosphates, 2.68 mM potassium chloride, 140 mM sodium chloride).</li>
	<li>Fix the cells with 4% paraformaldehyde in PBS for 10 min in the dark at room temperature.</li>
	<li>Wash the fixed cells three times with PBS.</li>
	<li>Add a minimal amount of fluorescence mounting medium on a microscope glass slide. Carefully transfer the coverslip from the 12-well plate onto the mounting medium with the sample facing the glass slide. 
	NOTE: Seal the edge of the coverslip with nail polish if an aqueous mounting medium is used.</li>
</ol>
5. Measurement of Neurite Outgrowth
<ol>
	<li>Use a 40x objective for capturing images using an epi-fluorescent microscope.</li>
	<li>Capture images from at least 40 intact neurons with EGFP signal per transfection.</li>
	<li>Open the captured image in ImageJ software with the NeuronJ plugin11 to measure the length of the longest neurite, from the cell body to the tip of the growth cone, of each imaged neuron.</li>
	<li>Analyze the data obtained with the software to determine the effect of the targeted proteins in neurite outgrowth.</li>
</ol>

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The authors declare that they have no conflicts of interest with the contents of this article.

As stated before, PC12 and its subclones are widely employed for studying neurite extension because they have excellent transfection efficiency2,3. In contrast, primary neurons have a low transfection rate, which is a major obstacle for studying neurite outgrowth regulators by transfection7. Here, we describe a convenient protocol for quantifying neurite outgrowth in primary neurons. Despite the low overall transfection efficiency, more than 80% of the transfected neurons were successfully co-transfected with the two proteins: FE65 and EGFP (Figure 2A). Therefore, by analyzing the EGFP labelled neurons, the effect of FE65 on neurite outgrowth could be precisely determined (Figure 2B).
Another advantage of the EGFP-based approach described is that immunofluorescence staining can be omitted. Immunofluorescence staining of Tuj 1, a neuron-specific class III &#946;-tubulin, is widely employed as a morphological marker when studying neurite elongation18,19,20,21. In addition to Tuj 1, staining of the POI is required for the identification of transfected neurons19,22,23. It is known that the consistency of immunofluorescence staining, which is crucial for neurite outgrowth measurement, is affected by many factors including sample preparation and antibodies24. Hence, the overall simplicity of the EGFP-based method could improve the accuracy of the assay.
In our protocol, neurons are fixed for neurite measurement 24 h post-transfection. Thus, neurons are exposed to the transfection reagent for 24 h. It is long known that transfection reagents are toxic to cells25. If the effect of the POI on neurite extension needs to be determined beyond DIV3, fresh medium replenishment may help to minimize the toxicity. Moreover, alternative gene delivery methods may be considered such as Ca2+ phosphate co-precipitation and nucleofection, both of which are shown to have a less toxic effect to cells26. Additionally, we show here that the highest effect of FE65 on neurite outgrowth is observed in the neurons transfected with that 0.5 &#181;g of FE65 and 0.5 &#181;g of EGFP plasmids by using 1 &#181;L of transfection reagent. For other POIs, the optimal amounts of plasmid DNA and transfection reagents should be determined as proteins have widely different turnover rates.
In addition to gene delivery method, cell density is another critical parameter to be considered. If the neuron density is too high, it becomes difficult to identify the origins of neurites as they would overlap with each other. Although plating cells at a low density may resolve the issue, the survival of primary neurons would be significantly reduced at a low cell density27. Primary rat hippocampal neurons have been shown to grow healthily at the density between 40,000 to 100,000/cm2&#160;28. In this protocol, a density of 65,000/cm2 of rat cortical neurons is used. Nevertheless, it is important to determine the appropriate cell density for different types of neurons under different experimental conditions.
The neurite measurement of developing rat primary neurons (i.e.&#160;DIV3 neurons) is described here. However, more mature neuronal phenotypes can be observed in neurons beyond DIV329. As the effects of POIs or drugs on neurite extension in mature neurons may be different from the developing neurons, investigation by using neurons from different stages would provide a more comprehensive perspective. It is worth noting that we were still able to observe EGFP expression in the neurons 7 d post-transfection. This may facilitate the study of POIs or drugs on neurite outgrowth in mature neurons.
Human induced pluripotent stem cell (hiPSC)-derived neurons are valuable tools in identification of novel therapeutic approaches. For instance, investigation of neurite outgrowth in these cells could reveal novel targets in neuroregeneration as the use of hiPSC-derived neurons avoids the species differences. Similar to primary rodent neurons, hiPSC-derived neurons are difficult to transfect30, which may reduce the co-transfection efficiency of EGFP and the POI. Hence, the use of polycistronic mammalian expression vectors could ensure that all the transfected cells express the entire set of transfected genes including EGFP and POIs. Additionally, alternative gene delivery methods, such as electroporation, may improve transfection efficiency. Again, cell viability is an issue which needs to be considered when using such gene delivery approaches.
It is known that the rat embryonic brain cortex contains different types of neurons including spiny stellate neurons, bipolar neurons and long-projecting neurons31. While this protocol stated here can reveal the general effect of POIs on neurite outgrowth, it is possible that the same POI may exhibit different responses in different types of neurons. For example, myostatin increases the number of sensory axons but not that of motor axons32. In such a scenario, modification of the protocol is necessary, such as prior isolation of specific types of neurons by flow cytometry. Alternatively, specific neuronal marker antibodies may be used for the identification of the required types of neurons during imaging.

Neurites, including both axons and dendrites, are the projections from neurons involved in the establishment of the neural networks. The dynamic outgrowth of neurites is essential for neurodevelopment. However, the underlying regulatory mechanisms underneath remain unclear. In particular, neurite damage is often observed in various neurodegenerative diseases and after brain injuries1. Therefore, investigation of the roles of putative molecules in various neurite outgrowth regulatory pathways would improve our understanding of the process. Moreover, it may reveal novel therapeutic targets for various neurological disorders. Neuronal cell lines are valuable models for studying neuronal processes including neurite outgrowth as they are easy to manipulate and transfect2,3. However, genetic drift has been reported to occur in some commonly used cell lines, which could lead to variations in their physiological responses4. Moreover, differential protein expression has been shown between neuronal cell lines and primary neurons. For instance, PC12, a neuronal cell line derived from rat adrenal gland that is widely used for studying neurite outgrowth2,3, does not express NMDA receptors5. Furthermore, it has been proposed that the reduced responsiveness of the mouse neuroblastoma line neuro-2a to neurotoxins in comparison to primary neurons is due to the lack of expression of certain membrane receptors and ion channels6. Therefore, primary neurons are a more desirable and representative model for the investigation of neurite outgrowth. However, the use of primary neurons is hindered by their low transfection efficiency7.
Here, we describe a method that involves the co-transfection of the protein of interest (POI) and EGFP to primary rat cortical neurons. The EGFP acts as a morphological marker for the identification of successfully transfected neurons and permits the measurement of neurites. We validated this method by using compounds/molecules that have been reported to modulate neurite outgrowth. Moreover, FE65, a neuronal adaptor protein that has been shown to stimulate neurite outgrowth, was used to illustrate this approach8,9. This protocol involves (1) the isolation of primary cortical neurons from embryonic day 18 (E18) rat embryos, (2) the co-transfection of neurons with EGFP and the POI (FE65 in this study) and (3) the imaging and analysis of the neurons by using the image processing software ImageJ with the NeuronJ plugin10,11.

<table><tbody><tr><td>#5 tweezers</td><td>Regine</td><td>5-COB</td><td></td></tr><tr><td>18 mm Circle Cover Slips</td><td>Thermo Scientific</td><td>CB00180RA</td><td>Sterilize before use</td></tr><tr><td>B27 Supplement</td><td>Gibco</td><td>17504044</td><td></td></tr><tr><td>Cytochalasin D</td><td>Invitrogen</td><td>PHZ1063</td><td>Dissolved in DMSO</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma-Aldrich</td><td>G8270</td><td></td></tr><tr><td>Dimethyl Sulfoxide</td><td>Sigma-Aldrich</td><td>D2650</td><td></td></tr><tr><td>Dissecting Scissors, 10 cm</td><td>World Precision Instruments</td><td>14393</td><td></td></tr><tr><td>Dissecting Scissors, 12.5 cm</td><td>World Precision Instruments</td><td>15922</td><td></td></tr><tr><td>EndoFree Plasmid Maxi Kit</td><td>QIAGEN</td><td>12362</td><td></td></tr><tr><td>Fluorescence Mounting Medium</td><td>Dako</td><td>S302380</td><td></td></tr><tr><td>Lipofectamine 2000 Transfection Reagent</td><td>Invitrogen</td><td>11668019</td><td></td></tr><tr><td>Neurobasal Medium</td><td>Gibco</td><td>21103049</td><td></td></tr><tr><td>NGF 2.5S Native Mouse Protein</td><td>Gibco</td><td>13257019</td><td></td></tr><tr><td>Nugent Utility Forceps, 10 mm, Straight Tip</td><td>World Precision Instruments</td><td>504489</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>P6148</td><td></td></tr><tr><td>pEGFP-C1</td><td>Clontech</td><td>#6084-1</td><td></td></tr><tr><td>pCI FE65</td><td></td><td></td><td>Please see references 8 and 15</td></tr><tr><td>PBS Tablets</td><td>Gibco</td><td>18912014</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>Poly-D-lysine hydrobromide</td><td>Sigma-Aldrich</td><td>P7280</td><td></td></tr><tr><td>Spatula</td><td>Sigma-Aldrich</td><td>S4147</td><td></td></tr><tr><td>Trypsin-EDTA (0.05%), phenol red</td><td>Gibco</td><td>25300062</td><td></td></tr><tr><td>Trypan Blue Solution, 0.4%</td><td>Gibco</td><td>15250061</td><td></td></tr></tbody></table>

an enhanced green fluorescence protein-based assay for studying neurite outgrowth in primary neurons

Neurite outgrowth is a fundamental event in the formation of the neural circuits during nervous system development. Severe neurite damage and synaptic dysfunction occur in various neurodegenerative diseases and age-related degeneration. Investigation of the mechanisms that regulate neurite outgrowth would not only shed valuable light on brain developmental processes but also on such neurological disorders. Due to the low transfection efficiency, it is currently challenging to study the effect of a specific protein on neurite outgrowth in primary mammalian neurons. Here, we describe a simple method for the investigation of neurite outgrowth by the co-transfection of primary rat cortical neurons with EGFP and a protein of interest (POI). This method allows the identification of POI transfected neurons through the EGFP signal, and thus the effect of the POI on neurite outgrowth can be determined precisely. This EGFP-based assay provides a convenient approach for the investigation of pathways regulating neurite outgrowth.

To test this methodology, we used Cyto D and nerve growth factor NGF, which have been shown to inhibit and stimulate neurite outgrowth respectively14,15,16. The neurite length of neurons transfected with EGFP were measured after treatment with Cyto D or NGF. The transfection efficiency of EGFP to the neurons was 2.7% (1,068 neurons counted). As shown in Figure 1A, Cyto D suppressed neurite extension in a dose-dependent manner. Conversely, neurite outgrowth was potentiated in the neurons treated with NGF (Figure 1B).
Next, we investigated the utility of this system by transfecting the neuronal adaptor FE65, which has been shown to promote neurite outgrowth. Primary rat cortical neurons were co-transfected with FE65 and EGFP. Despite the low transfection efficiency, immuno-fluorescence analysis revealed that over 80% of the neurons were successfully co-transfected with EGFP and FE65 (Figure 2A). Similar to previous reports8,9, FE65 significantly stimulated the neurite outgrowth by 2x (Figure 2B). We also analyzed the expression of EGFP and FE65 at different time points by Western blot analysis. As shown in Figure 2C, EGFP and FE65 were detected 6 h and 12 h post-transfection, respectively. Similar expression levels of the proteins were observed in 1 d to 7 d post-transfection neurons. This indicates that the analysis of neurite outgrowth could be done as early as 6 h post-transfection or in more mature neurons. Together, this data suggest that the mentioned protocol is suitable for determining the role of putative neurite outgrowth regulatory proteins by classical transfection.
We also monitored the effect of gene dosage on neurite outgrowth by transfecting primary rat cortical neurons with different amounts of FE65 plasmid DNA. As shown in Figure 2D, a dose-dependent increase in neurite extension was observed from 0 - 0.5 &#181;g of FE65 plasmid DNA. However, there was no significant difference between neurons transfected with either 0.5 &#181;g or 1 &#181;g of plasmid DNA (Figure 2D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60031/60031fig1d.jpg" /> 
Figure 1: Neurite outgrowth is modulated by Cyto D and NGF. E18 rat cortical neurons were transfected on DIV2 with an EGFP expression vector. 6 h post-transfection, the cells were treated with (A) 0-0.5 &#181;g/mL Cyto D or (B) 0-100 ng/mL NGF for 24 h. Then the neurons were fixed and imaged accordingly. Images were captured with 40x objective using an epi-fluorescence microscope and the length of the longest neurite from the cell body to the tip of the growth cone was measured by using ImageJ with the NeuronJ plugin. Three independent experiments were performed and at least 40 neurons were measured in each group. The bar chart showed the fold change in mean neurite length. Unpaired t-test was adopted for the statistical analysis. *p &#60; 0.001, **p &#60; 0.05. Error bars = S.E.M. <a href="https://www.jove.com/files/ftp_upload/60031/60031fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60031/60031fig2k.jpg" /> 
Figure 2: FE65 stimulates neurite outgrowth. E18 rat cortical neurons were transfected on DIV2 with either empty vector control (EV) or FE65 together with an EGFP expression vector. Cells were fixed and imaged 24 h after transfection. (A) The transfected neurons were counterstained with anti-FE65 antibody and the number of cells with EGFP or FE65 singly transfected and EGFP + FE65 co-transfected were counted. Three independent experiments were performed and at least 100 cells were counted in each experiment. Data were expressed as the percentage of cells with EGFP and EGFP + FE65 signals. *p &#60; 0.001. Error bars = S.D. (B) The length of the longest neurite from the cell body to the tip of the growth cone was measured by using ImageJ with the NeuronJ plugin. Representative neuron images were shown in the right panel. FE65 was stained by a goat anti-FE65 antibody as described8,17. Three independent experiments were performed and at least 40 neurons were measured per transfection. The bar chart showed the fold change in mean neurite length. The data were analyzed by unpaired t-test. *p &#60; 0.001. Error bars = S.E.M. Scale bar = 10 &#181;m. (C) Western blot analysis of the expression of levels of EGFP and FE65 at the post-transfection time points as indicated. EGFP and FE65 were detected by mouse anti-GFP and anti-myc (to FE65 C-terminal myc tag), respectively. (D) The average neurite length of neurons transfected with various amounts of FE65 plasmid DNA as indicated. Statistical analyses were performed using one-way ANOVA tests with Bonferroni post-hoc test. *p &#60; 0.001, **p &#60; 0.05. Error bars = S.E.M. Scale bar = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/60031/60031fig2elarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons at JoVE.com

An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons

In this report, we describe a simple protocol for studying neurite outgrowth in embryonic rat cortical neurons by co-transfecting with EGFP and the protein of interest.

Neurite outgrowth is a fundamental event in the formation of the neural circuits during nervous system development. Severe neurite damage and synaptic ...

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Nanyang Technological University

Research

JoVE Journal

Developmental Biology

5.9K Views.  The Chinese University of Hong Kong, Hong Kong Special Administrative Region. In this report, we describe a simple protocol for studying neurite outgrowth in embryonic rat cortical neurons by co-transfecting with EGFP and the protein of interest.

Video: An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Biology

A Neuronal and Astrocyte Co-Culture Assay for High Content Analysis of Neurotoxicity

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing measurement of multiple cellular phenotypes within a single assay. In this study, we utilized HCA to develop a novel assay for neurotoxicity. Neurotoxicity assessment represents an important part of drug safety evaluation, as well as being a significant focus of environmental protection efforts. Additionally, neurotoxicity is also a well-accepted in vitro marker of the development of neurodegenerative diseases such as Alzheimer's and Parkinson's diseases. Recently, the application of HCA to neuronal screening has been reported. By labeling neuronal cells with &beta;III-tubulin, HCA assays can provide high-throughput, non-subjective, quantitative measurements of parameters such as neuronal number, neurite count and neurite length, all of which can indicate neurotoxic effects. However, the role of astrocytes remains unexplored in these models. Astrocytes have an integral role in the maintenance of central nervous system (CNS) homeostasis, and are associated with both neuroprotection and neurodegradation when they are activated in response to toxic substances or disease states. GFAP is an intermediate filament protein expressed predominantly in the astrocytes of the CNS. Astrocytic activation (gliosis) leads to the upregulation of GFAP, commonly accompanied by astrocyte proliferation and hypertrophy. This process of reactive gliosis has been proposed as an early marker of damage to the nervous system. The traditional method for GFAP quantitation is by immunoassay. This approach is limited by an inability to provide information on cellular localization, morphology and cell number. We determined that HCA could be used to overcome these limitations and to simultaneously measure multiple features associated with gliosis - changes in GFAP expression, astrocyte hypertrophy, and astrocyte proliferation - within a single assay. In co-culture studies, astrocytes have been shown to protect neurons against several types of toxic insult and to critically influence neuronal survival. Recent studies have suggested that the use of astrocytes in an in vitro neurotoxicity test system may prove more relevant to human CNS structure and function than neuronal cells alone. Accordingly, we have developed an HCA assay for co-culture of neurons and astrocytes, comprised of protocols and validated, target-specific detection reagents for profiling &beta;III-tubulin and glial fibrillary acidic protein (GFAP). This assay enables simultaneous analysis of neurotoxicity, neurite outgrowth, gliosis, neuronal and astrocytic morphology and neuronal and astrocytic development in a wide variety of cellular models, representing a novel, non-subjective, high-throughput assay for neurotoxicity assessment. The assay holds great potential for enhanced detection of neurotoxicity and improved productivity in neuroscience research and drug discovery.

This article describes a novel protocol and reagent set designed for sensitive measurement of neurotoxic effects of compounds and treatments on co-cultures of neurons and astrocytes using high content analysis. Results demonstrate that high content analysis represents an exciting novel technology for neurotoxicity assessment.

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing ...

A Neurite Outgrowth Assay and Neurotoxicity Assessment with Human Neural Progenitor Cell-Derived Neurons

Neurite outgrowth assay and neurotoxicity assessment are two major studies that can be performed using the presented method herein. This protocol provides reliable analysis of neuronal morphology together with quantitative measurements of modifications on neurite length and synaptic protein localization and abundance upon treatment with small molecule compounds. In addition to the application of the presented method in neurite outgrowth studies, neurotoxicity assessment can be performed to assess, distinguish and rank commercial chemical compounds based on their potential developmental neurotoxicity effect.
Even though cell lines are nowadays widely used in compound screening assays in neuroscience, they often differ genetically and phenotypically from their tissue origin. Primary cells, on the other hand, maintain important markers and functions observed in vivo. Therefore, due to the translation potential and physiological relevance that these cells could offer neurite outgrowth assay and neurotoxicity assessment can considerably benefit from using human neural progenitor cells (hNPCs) as the primary human cell model.
The presented method herein can be utilized to screen for the ability of compounds to induce neurite outgrowth and neurotoxicity by taking advantage of the human neural progenitor cell-derived neurons, a cell model closely representing human biology.&#34;

The presented protocol describes a method for a neurite outgrowth assay and neurotoxicity assessment of small molecule compounds.

Neurite outgrowth assay and neurotoxicity assessment are two major studies that can be performed using the presented method herein. This protocol provides ...

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane proteins become exposed at the cell surface. One of these proteins is Synaptotagmin-1 (Syt1). An antibody directed against the luminal domain of Syt1, once added to the culture medium, is taken up during the exo-endocytotic cycle. This uptake is proportional to the amount of SV recycling and can be quantified through immunofluorescence. Here, we combine Syt1 antibody uptake with double transfection of cultured hippocampal neurons. This allows us to (1) localize presynaptic sites based on expression of recombinant presynaptic marker Synaptophysin, (2) determine their functionality using Syt1 uptake, and (3) characterize the targeting and effects of a protein of interest, GFP-Rogdi.

We describe an optical assay for synaptic vesicle (SV) recycling in cultured neurons. Combining this protocol with double transfection to express a presynaptic marker and protein of interest allows us to locate presynaptic sites, their synaptic vesicle recycling capacity, and determine the role of the protein of interest.

At active presynaptic nerve terminals, synaptic vesicles undergo cycles of exo- and endocytosis. During recycling, the luminal domains of SV transmembrane ...

Neuroscience

Primary Cell Culture of Purified GABAergic or Glutamatergic Neurons Established through Fluorescence-activated Cell Sorting

The overall goal of this protocol is to generate purified neuronal cultures derived from either GABAergic or glutamatergic neurons. Purified neurons can be cultured in defined media for 16 days in vitro and are amenable to any analyses typically performed on dissociated cultures, including electrophysiological, morphological and survival analyses. The major advantage of these cultures is that specific cell types can be selectively studied in the absence of complex external influences, such as those arising from glial cells or other neuron types. When planning experiments with purified cells, however, it is important to note that neurons strongly depend on glia-conditioned media for their growth and survival. In addition, glutamatergic neurons further depend on glia-secreted factors for the establishment of synaptic transmission. We therefore also describe a method for co-culturing neurons and glial cells in a non-contact arrangement. Using these methods, we have identified major differences between the development of GABAergic and glutamatergic neuronal networks. Thus, studying cultures of purified neurons has great potential for furthering our understanding of how the nervous system develops and functions. Moreover, purified cultures may be useful for investigating the direct action of pharmacological agents, growth factors or for exploring the consequences of genetic manipulations on specific cell types. As more and more transgenic animals become available, labeling additional specific cell types of interest, we expect that the protocols described here will grow in their applicability and potential.

This protocol describes a cell sorting based method for the purification and culture of fluorescent GABAergic or glutamatergic neurons from the neocortex and hippocampus of postnatal mice or rats.

The overall goal of this protocol is to generate purified neuronal cultures derived from either GABAergic or glutamatergic neurons. Purified neurons can ...

ROS Live Cell Imaging During Neuronal Development

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the different ROS, hydrogen peroxide (H2O2) is best characterized with respect to roles in cellular signaling. H2O2 has been implicated during the development in several species. For example, a transient increase in H2O2 has been detected in zebrafish embryos during the first days following fertilization. Furthermore, depleting an important cellular H2O2 source, NADPH oxidase (NOX), impairs nervous system development such as the differentiation, axonal growth, and guidance of retinal ganglion cells (RGCs) both in&#160;vivo and in vitro. Here, we describe a method for imaging intracellular H2O2 levels in cultured zebrafish neurons and whole larvae during development using the genetically encoded H2O2-specific biosensor, roGFP2-Orp1. This probe can be transiently or stably expressed in zebrafish larvae. Furthermore, the ratiometric readout diminishes the probability of detecting artifacts due to differential gene expression or volume effects. First, we demonstrate how to isolate and culture RGCs derived from zebrafish embryos that transiently express roGFP2-Orp1. Then, we use whole larvae to monitor H2O2 levels at the tissue level. The sensor has been validated by the addition of H2O2. Additionally, this methodology could be used to measure H2O2 levels in specific cell types and tissues by generating transgenic animals with tissue-specific biosensor expression. As zebrafish facilitate genetic and developmental manipulations, the approach demonstrated here could serve as a pipeline to test the role of H2O2 during neuronal and general embryonic development in vertebrates.

This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the ...

A Scalable Method to Study Neuronal Survival in Primary Neuronal Culture with Single-cell and Real-Time Resolution

Neuronal loss is at the core of many neuropathologies, including stroke, Alzheimer's disease, and Parkinson's disease. Different methods were developed to study the process of neuronal survival upon cytotoxic stress. Most methods are based on biochemical approaches that do not allow single-cell resolution or involve complex and costly methodologies. Presented here is a versatile, inexpensive, and effective experimental paradigm to study neuronal survival. This method takes advantage of sparse fluorescent labeling of the neurons followed by live imaging and automated quantification. To this aim, the neurons are electroporated to express fluorescent markers and co-cultured with non-electroporated neurons to easily regulate cell density and increase survival. 
Sparse labeling by electroporation allows a simple and robust automated quantification. In addition, fluorescent labeling can be combined with the co-expression of a gene of interest to study specific molecular pathways. Here, we present a model of stroke as a neurotoxic model, namely, the oxygen-glucose deprivation (OGD) assay, which was performed in an affordable and robust homemade hypoxic chamber. Finally, two different workflows are described using IN Cell Analyzer 2200 or the open-source ImageJ for image analysis for semi-automatic data processing. This workflow can be easily adapted to different experimental models of toxicity and scaled up for high-throughput screening. In conclusion, the described protocol provides an approachable, affordable, and effective in vitro model of neurotoxicity, which can be suitable for testing the roles of specific genes and pathways in live imaging and for high-throughput drug screening.

Here, we present a simple, potent, and versatile methodology to investigate neuronal survival upon cytotoxic stress in primary cortical neurons with cellular resolution in real time or in fixed material.

Neuronal loss is at the core of many neuropathologies, including stroke, Alzheimer's disease, and Parkinson's disease. Different methods were developed to ...

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.

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 ...

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 ...

Immunofluorescence Assay to Evaluate Effect of Target Proteins on Neurite Outgrowth

Source: Chan, W., W., R. et al. <a href="https://app.jove.com/v/60031">An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons.</a> J. Vis. Exp. (2019).
In this video, we demonstrate an EGFP-based immunofluorescence assay to identify the efficacy of target proteins on the neurite outgrowth of primary cortical neurons. The primary neurons are transfected with a fluorescent protein, EGFP, and a target protein, and then an immunoassay is performed to investigate the effect of the target protein on neurite outgrowth.

Source: Chan, W., W., R. et al. An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons. J. Vis. Exp. 
...

An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons

Summary

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Chapters in this video

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

A Neuronal and Astrocyte Co-Culture Assay for High Content Analysis of Neurotoxicity

A Neurite Outgrowth Assay and Neurotoxicity Assessment with Human Neural Progenitor Cell-Derived Neurons

An Optical Assay for Synaptic Vesicle Recycling in Cultured Neurons Overexpressing Presynaptic Proteins

Primary Cell Culture of Purified GABAergic or Glutamatergic Neurons Established through Fluorescence-activated Cell Sorting

ROS Live Cell Imaging During Neuronal Development

A Scalable Method to Study Neuronal Survival in Primary Neuronal Culture with Single-cell and Real-Time Resolution

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

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Immunofluorescence Assay to Evaluate Effect of Target Proteins on Neurite Outgrowth