Primary rat hippocampal neurons were supplied by Dr. George Lykotrafitis and Shiju Gu of the Biomedical Engineering Department at the University of Connecticut (Storrs, CT, USA). The Abberior STED instrument housed in the Advanced Light Microscopy Facility in the Center for Open Research Resources and Equipment was acquired with NIH grant S10OD023618 awarded to Christopher O&#39;Connell. This research was funded by NIH grant R01AG065879 awarded to Nathan N. Alder.

Unveiling Mitochondrial Ultrastructure in Neurons Using STED Microscopy

The authors have nothing to disclose.

This protocol presents the use of human neuroblastoma cell line SH-SY5Y and primary rat hippocampal neurons with the novel IMM-targeting PKMO dye for live cell STED imaging. Due to the novelty of PKMO, there is currently little published using this dye for live STED imaging. Using these cell types for STED imaging poses challenges, specifically because neuronal cells have narrower mitochondria. One limitation of this protocol is the PKMO dye used, as it can be toxic to cells. Different cells and cell lines respond differently to the dye, thus, adjustments to dye concentration and incubation time to optimize results for strong signal without harming cells may be required. A suggested solution is to lower the concentration and increase the staining time63; however, this may result in poorer staining without increasing cell viability.
Similarly to PKMO, the commercial dye Live Orange mito (Table of Materials) also exhibits some cell toxicity. This dye was used for a variety of cultured cells but was unable to exhibit comparable staining in RA-differentiated SH-SY5Y cells successfully with the same parameters as their undifferentiated counterparts (our unpublished observations). However, amenable staining protocols may be optimized for this probe and chosen cell types. With this dye, detector gating times of 1-1.05 to 7-7.05 ns were used, with all other parameters in Table 1 remaining the same. Generally, staining cells with 200-250 nM Live Orange mito for 45 min yielded comparable results as the PKMO results shown. Higher concentration staining for less time or lower concentration staining for the same amount of time or slightly longer can yield different results and may be favorable to other cell types or growth conditions.
Imaging primary rat hippocampal neurons differs from immortalized cells due to the nature of the axon and dendrite projections as well as mitochondrial distribution at the time of imaging. One difficulty in this part of the protocol is that seeding density determines whether the primary cultures will be able to adhere and grow healthily, and at higher densities, the projections tend to overgrow by DIV 10. Therefore, the mitochondria imaged from these primary neurons will likely come from the cell body and not the projections; however, successful growth from a lower starting cell density yields better imaging results at later growth times. The key is to ensure low background and out-of-focus light to have the best contrast for STED. To address concerns regarding cell population, culturing primary hippocampal cells in B27-supplemented neuron growth media prevents the growth of glial cells, and the source reports that &#60;5% of cells are astrocytes and the absence of NbActiv1 supplement in the growth media reduces the number of astrocytes in cultures to &#60;2%87. For both cultured SH-SY5Y cells and primary rat hippocampal neurons, the PDL coating used for growth contributes to background haze in images. Sufficient signal-to-noise is accomplished with the settings reported in (Table 1) and deconvolution removes most of the background observed.
In addition to the imaging covered here, it is also possible to add treatments or stress to cells before or during imaging. For example, adding tert-butyl hydrogen peroxide (tBHP) induces oxidative stress, and it is possible to monitor changes in mitochondria over time after addition. The addition of amyloid-&#946; (A&#946;) with a fluorescent tag allows monitoring of the distribution of this peptide in relation to mitochondria as well as the mitochondrial structure over time. Mitochondrial health has been heavily implicated in AD and is widely supported to play a role in A&#946; toxicity43,71,72. Notably, the differentiation status of SH-SY5Y cells affects A&#946; protein precursor (A&#946;PP) localization85, and experiments using A&#946;PP should be carefully constructed.
As an example of how this protocol can be adapted, it is shown that the fluorescent variant A&#946;(1-42)-HiLyte 647 can be added to PKMO-stained cells 15 min before imaging (Supplementary Figure 1). The imaging parameters are similar (Supplementary Table 2), with the main difference being that a smaller pinhole is needed when imaging narrower mitochondria. Imaging A&#946;-HiLyte647 with STED requires less overall excitation (6%-8%) and STED depletion (10%-12%) laser power and fewer accumulations (six). Detector gating is also extended from 0.1 to 10 ns. Although STED resolution of A&#946; is not necessary, the overall signal-to-noise ratio and A&#946; particle size of raw STED were better than those of the confocal images, and subsequent deconvolution can also be performed. Collecting STED images and deconvolving raw STED z-stack projections of A&#946; appears particularly useful when merging with raw STED or deconvolved STED images of the PKMO stain (Supplementary Figure 1B,C). Both channels were collected in a single frame step. Measurements of time-dependent localization, similar to those listed in Figure 2 and shown in Figure 5, where applicable, and cristae architecture differences can be obtained following stress treatment or other additions.
Other possible methods for dual-labeling in live cell STED of mitochondria not reported here but have been reported by others include the use of SNAP-tagged proteins93, Halo-tagged proteins, and the use of other cell-permeable dyes with generic targets, such as mtDNA63. Notably, the labeling strategy of SNAP and Halo tagging influences the resulting fluorescence signal intensity and longevity when imaging94. Additionally, while this protocol presents several examples of analyses that can be applied to segmented mitochondria, there are many other analyses that software packages can perform on these images.

Mitochondria are eukaryotic organelles of endosymbiotic origin that are responsible for regulating several key cellular processes, including intermediary metabolism and ATP production, ion homeostasis, lipid biosynthesis, and programmed cell death (apoptosis). These organelles are topologically complex, containing a double membrane system that establishes multiple subcompartments1 (Figure 1A). The outer mitochondrial membrane (OMM) interfaces with the cytosol and establishes direct inter-organelle contacts2,3. The inner mitochondrial membrane (IMM) is an energy-conserving membrane that maintains ion gradients stored primarily as an electric membrane potential (&#916;&#936;m) to drive ATP synthesis and other energy-requiring processes4,5. The IMM is further subdivided into the inner boundary membrane (IBM), which is closely appressed to the OMM, and protruding structures called cristae that are bound by the cristae membrane (CM). This membrane delineates the innermost matrix compartment from the intracristal space (ICS) and the intermembrane space (IMS).
Mitochondria have a dynamic morphology based on continuous and balanced processes of fission and fusion that are governed by mechanoenzymes of the dynamin superfamily6. Fusion allows for increased connectivity and formation of reticular networks, whereas fission leads to mitochondrial fragmentation and enables the removal of damaged mitochondria by mitophagy7. Mitochondrial morphology varies by tissue type8 and developmental stage9 and is regulated to allow cells to adapt to factors including energetic needs10,11 and stressors12. Standard morphometric features of mitochondria, such as the extent of network formation (interconnected vs. fragmented), perimeter, area, volume, length (aspect ratio), roundness, and degree of branching, can be measured and quantified by standard optical microscopy because the sizes of these features are greater than the diffraction limit of light (~200 nm)13.
Cristae architecture defines the internal structure of mitochondria (Figure 1B). The diversity of cristae morphologies can be broadly categorized as flat (lamellar or discoidal) or tubular-vesicular14. All cristae attach at the IBM through tubular or slot-like structures termed cristae junctions (CJs) that can serve to compartmentalize the IMS from the ICS and the IBM from the CM15. Cristae morphology is regulated by key protein complexes of the IMM, including (1) the mitochondrial contact site and cristae organizing system (MICOS) that resides at CJs and stabilizes IMM-OMM contacts16, (2) the optic atrophy 1 (OPA1) GTPase that regulates cristae remodeling17,18,19, and (3) F1FO ATP synthase that forms stabilizing oligomeric assemblies at cristae tips (CTs)20,21. In addition, the IMM is enriched in nonbilayer phospholipids phosphatidylethanolamine and cardiolipin that stabilize the highly curved IMM22. Cristae are also dynamic, demonstrating morphological changes under various conditions, such as different metabolic states23,24, with different respiratory substrates25, under starvation and oxidative stress26,27, with apoptosis28,29, and with aging30. Recently it has been shown that cristae could undergo major remodeling events on a timescale of seconds, underscoring their dynamic nature31. Several features of cristae can be quantified, including dimensions of structures within individual cristae (e.g., CJ width, crista length, and width) and parameters that relate individual crista to other structures (e.g., intra-cristae spacing and cristae incident angle relative to the OMM)32. These quantifiable cristae parameters show a direct correlation with function. For instance, the extent of mitochondrial ATP production is positively related to the abundance of cristae, quantified as cristae density or cristae number normalized to another feature (e.g., cristae per OMM area)33,34,35. Because IMM morphology is defined by nanoscale features, it comprises mitochondrial ultrastructure, which requires imaging techniques that provide resolution greater than the light diffraction limit. As described below, such techniques include electron microscopy and super-resolution microscopy (nanoscopy).
The neural and glial cells of the central nervous system (CNS) are particularly reliant on mitochondrial function. On average, the brain constitutes only 2% of the total body weight, but utilizes 25% of the total body glucose and accounts for 20% of body oxygen consumption, making it vulnerable to impairments in energy metabolism36. Progressive neurodegenerative diseases (NDs), including Alzheimer&#39;s disease (AD), amyotrophic lateral sclerosis (ALS), Huntington&#39;s disease (HD), multiple sclerosis (MS), and Parkinson&#39;s disease (PD), are some of the most extensively studied pathologies to date, with research efforts ranging from understanding the molecular underpinnings of these diseases to seeking potential therapeutic prevention and interventions. NDs are associated with increased oxidative stress originating in part from reactive oxygen species (ROS) generated by the mitochondrial electron transport chain (ETC)37, as well as altered mitochondrial calcium handling38 and mitochondrial lipid metabolism39. These physiological alterations are accompanied by noted defects in mitochondrial morphology that are associated with AD40,41,42,43,44, ALS45,46, HD47,48,49, MS50, and PD51,52,53. These structural and functional defects can be coupled by complex cause-effect relationships. For example, given that cristae morphology stabilizes OXPHOS enzymes54, mitochondrial ROS are not only generated by the ETC, but they also act to damage the infrastructure in which the ETC resides, promoting a feed-forward ROS cycle that enhances susceptibility to oxidative damage. Furthermore, cristae disorganization has been shown to trigger processes such as mitochondrial DNA (mtDNA) release and inflammatory pathways connected to autoimmune, metabolic, and age-related disorders55. Therefore,&#160;analysis of&#160;mitochondrial structure is key to a full understanding of NDs and their molecular underpinnings.
Popular methods of viewing cristae, including transmission electron microscopy, electron tomography and cryo-electron tomography (cryo-ET), and X-ray tomography, in particular cryo-soft X-ray tomography, have revealed important findings and work with a variety of sample types56,57,58,59,60. Despite recent advancements toward better observation of organellar ultrastructure, these methods still come with the caveat of requiring sample fixation and, therefore, cannot capture real-time dynamics of cristae directly. Super-resolution fluorescence microscopy, particularly in the forms of structured illumination microscopy (SIM), stochastic optical reconstruction microscopy (STORM), photoactivated localization microscopy (PALM), expansion microscopy (ExM), and stimulated emission depletion (STED) microscopy, have become popular ways of viewing structures requiring resolution below the diffraction limit that constrains classical methods of optical microscopy. When ExM is used in conjunction with another super-resolution technique, the results are impressive, but the sample must be fixed and stained in a gel61. By comparison, SIM, PALM/STORM, and STED have all been successfully used with live samples, and new and promising dyes that generally stain the IMM provide a novel and easy approach for live imaging of mitochondria cristae dynamics62,63,64,65,66. Recent advancements in live dyes for STED imaging have improved dye brightness and photostability, and these dyes target the IMM at a higher degree of specificity than their predecessors. These developments allow the collection of long-term timelapse and z-stack experiments with super-resolution imaging, opening the door to better live cell analysis of mitochondrial ultrastructure and dynamics.
Herein, protocols for live cell imaging of undifferentiated and differentiated SH-SY5Y cells stained with the PKmito Orange (PKMO) dye using STED63 are provided. The SH-SY5Y cell line is a thrice subcloned derivative from the parental cell line, SK-N-SH, generated from a bone marrow biopsy of metastatic neuroblastoma67,68,69,70. This cell line is a commonly used in vitro model in ND research, particularly with diseases such as AD, HD, and PD, in which mitochondrial dysfunction is heavily implicated10,43,71,72,73. The ability to differentiate SH-SY5Y cells into cells with a neuron-like phenotype through manipulating culture media has proven a suitable model for neuroscience research without relying on primary neuronal cells10,74. In this protocol, retinoic acid (RA) was added to the cell culture medium to induce the differentiation of SH-SY5Y cells. RA is a vitamin A derivative and has been shown to regulate the cell cycle and promote the expression of transcription factors that regulate neuronal differentiation75. A protocol for culturing and live cell imaging of neurons isolated from the rat hippocampus is also provided. The hippocampus has been shown to be affected by mitochondrial degeneration and, along with the cortex, plays an important role in aging and ND76,77,78,79,80.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >0.05% Trypsin-EDTA</td>
			<td >Gibco</td>
			<td >25300054</td>
			<td ></td>
		</tr>
		<tr >
			<td >0.4% Trypan blue</td>
			<td >Invitrogen</td>
			<td >T10282</td>
			<td ></td>
		</tr>
		<tr >
			<td >0.5% Trypsin-EDTA, no phenol red</td>
			<td >Gibco</td>
			<td >15400054</td>
			<td ></td>
		</tr>
		<tr >
			<td >100 X antibiotic-antimycotic</td>
			<td >Gibco</td>
			<td >15240062</td>
			<td ></td>
		</tr>
		<tr >
			<td >100 X/1.40 UPlanSApo oil immersion lens</td>
			<td >Olympus</td>
			<td ></td>
			<td >Equipped in Olympus IX83 microscope for STED setup described in Section 4</td>
		</tr>
		<tr >
			<td >All-trans-retinoic acid</td>
			<td >Sigma</td>
			<td >R2625</td>
			<td ></td>
		</tr>
		<tr >
			<td >Amyloid-&#946; (1-42, HiLyte Fluor647, 0.1 mg)</td>
			<td >AnaSpec</td>
			<td >AS-64161</td>
			<td >Other fluorescent conjugates available</td>
		</tr>
		<tr >
			<td >B27 supplement (50 X), serum free</td>
			<td >Gibco</td>
			<td >17504044</td>
			<td ></td>
		</tr>
		<tr >
			<td >Cell Counter (Countess II FL)</td>
			<td >Life Technologies</td>
			<td >AMQAF1000</td>
			<td ></td>
		</tr>
		<tr >
			<td >Centrifuge</td>
			<td >Eppendorf</td>
			<td >5804-R</td>
			<td ></td>
		</tr>
		<tr >
			<td >Counter slides</td>
			<td >Invitrogen</td>
			<td >C10283</td>
			<td ></td>
		</tr>
		<tr >
			<td >Conical tubes (15 mL)</td>
			<td >Thermo Fisher Scientific</td>
			<td >339650</td>
			<td ></td>
		</tr>
		<tr >
			<td >Cuvettes (Quartz Cells)</td>
			<td >Starna Cells, Inc.</td>
			<td >9-Q-10</td>
			<td >Used with Spectrometer as described in Section 1.3</td>
		</tr>
		<tr >
			<td >DMEM (high glucose with sodium pyruvate)</td>
			<td >Gibco</td>
			<td >11995073</td>
			<td >Used for SH-SY5Y cell materials as described in Section 1</td>
		</tr>
		<tr >
			<td >DMEM (high glucose no sodium pyruvate)</td>
			<td >Gibco</td>
			<td >11965092</td>
			<td >Used for primary cell materials as described in Section 2</td>
		</tr>
		<tr >
			<td >DMEM (phenol red-free)</td>
			<td >Gibco</td>
			<td >31053028</td>
			<td >Used for imaging as described in Section 3</td>
		</tr>
		<tr >
			<td >DMSO</td>
			<td >Sigma</td>
			<td >D8418</td>
			<td ></td>
		</tr>
		<tr >
			<td >DNAase I from bovine pancreas</td>
			<td >Sigma</td>
			<td >DN25</td>
			<td >Used for primary cell materials as described in Section 2.2.1 and 2.2.2</td>
		</tr>
		<tr >
			<td >DPBS (no calcium, no magnesium)</td>
			<td >Gibco</td>
			<td >14190144</td>
			<td ></td>
		</tr>
		<tr >
			<td >E18 Rat Hippocampus</td>
			<td >Transnetyx Tissue</td>
			<td >SDEHP</td>
			<td ></td>
		</tr>
		<tr >
			<td >Ethanol (200 proof)</td>
			<td >Fisher Bioreagents</td>
			<td >BP28184</td>
			<td ></td>
		</tr>
		<tr >
			<td >Fetal bovine serum (FBS), not heat-inactivated</td>
			<td >Gibco</td>
			<td >26140079</td>
			<td >For cultured cells, in Section 1</td>
		</tr>
		<tr >
			<td >Fetal bovine serum (heat inactivated)</td>
			<td >Gibco</td>
			<td >10082147</td>
			<td >For primary cell culture, Section 2</td>
		</tr>
		<tr >
			<td >Filter sterilization unit (0.1 &#181;m, 500 mL)</td>
			<td >Thermo Fisher Scientific</td>
			<td >5660010</td>
			<td ></td>
		</tr>
		<tr >
			<td >FIJI (Is Just ImageJ) and Trainable Weka Segmentation (TWS) plug-in</td>
			<td >--</td>
			<td >--</td>
			<td >Free, open-source image analysis software that includes plug-ins including Trainable Weka Segmentation described in Section 5; TWS plug-in from ref. 90 of the main text</td>
		</tr>
		<tr >
			<td >GlutaMAX supplement (100 X)</td>
			<td >Gibco</td>
			<td >35050061</td>
			<td >Glutamine supplement used for primary cell materials described in Section 2.1.2</td>
		</tr>
		<tr >
			<td >Hausser Scientific bright-Line and Hy-Lite Counting Chambers</td>
			<td >Hausser Scientific</td>
			<td >267110</td>
			<td ></td>
		</tr>
		<tr >
			<td >HBSS (no calcium, no magnesium)</td>
			<td >Gibco</td>
			<td >14170120</td>
			<td >Used for primary cell materials described in Section 2.2.1 and 2.2.2</td>
		</tr>
		<tr >
			<td >HEPES</td>
			<td >Gibco</td>
			<td>15630080</td>
			<td ></td>
		</tr>
		<tr >
			<td >Huygens Professional deconvolution software (V. 20.10)</td>
			<td >Scientific Volume Imaging (SVI)</td>
			<td >--</td>
			<td >The deconvolution software used in this protocol and described in Section 5</td>
		</tr>
		<tr >
			<td >IX83 inverted microscope with Continuous Autofocus</td>
			<td >Olympus</td>
			<td >--</td>
			<td >This paper uses a STED Infinity Line system built around an Olympus IX83 inverted microscope, described in Section 4</td>
		</tr>
		<tr >
			<td >Lightbox software (V. 16.3.16118)</td>
			<td >Abberior</td>
			<td >--</td>
			<td >Vendor software used for STED image acquisition, described in Section 4</td>
		</tr>
		<tr >
			<td >Live Orange Mito dye</td>
			<td >Abberior</td>
			<td >LVORANGE-0146-30NMOL</td>
			<td >Live cell imaging IMM-targeting dye described in Discussion</td>
		</tr>
		<tr >
			<td >Neurobasal media</td>
			<td >Gibco</td>
			<td >21103049</td>
			<td >Used for primary cell materials referred to in Section 2.1.2</td>
		</tr>
		<tr >
			<td >Nunc Lab-Tek II 2-well chambered coverglass</td>
			<td >Nunc</td>
			<td >155379</td>
			<td >Can purchase a variety of chambers but make sure the coverglass is #1.5</td>
		</tr>
		<tr >
			<td >Pasteur Pipets (Fisherbrand)</td>
			<td >Thermo Fisher Scientific</td>
			<td >22183632</td>
			<td ></td>
		</tr>
		<tr >
			<td >Penicillin-Streptomycin (10,000 U/mL)</td>
			<td >Gibco</td>
			<td >15140122</td>
			<td ></td>
		</tr>
		<tr >
			<td >PKmito Orange dye</td>
			<td >Spirochrome</td>
			<td >SC053</td>
			<td ></td>
		</tr>
		<tr >
			<td >Poly-D-lysine</td>
			<td >Gibco</td>
			<td >A3890401</td>
			<td ></td>
		</tr>
		<tr >
			<td >SH-SY5Y Cell line</td>
			<td >ATCC</td>
			<td >CRL2266</td>
			<td ></td>
		</tr>
		<tr >
			<td >Sodium pyruvate (100 mM)</td>
			<td >Gibco</td>
			<td >11360070</td>
			<td >Used for primary cell materials described in Section 2</td>
		</tr>
		<tr >
			<td >Spectrometer (GENESYS 180 UV-Vis)</td>
			<td >Thermo Fisher Scientific</td>
			<td >840309000</td>
			<td ></td>
		</tr>
		<tr >
			<td >STED Expert Line microscope</td>
			<td >Abberior</td>
			<td >--</td>
			<td >STED setup can be customized, but at time of purchase instrument was considered Abberior&#8217;s Expert Line; brief description of setup is available in Section 4 of protocol</td>
		</tr>
		<tr >
			<td >T25 flask (TC-treated, filter cap)</td>
			<td >Thermo Fisher Scientific</td>
			<td >156367</td>
			<td >Other culture vessels and sizes available</td>
		</tr>
	</tbody>
</table>

using live cell sted imaging to visualize mitochondrial inner membrane ultrastructure in neuronal cell models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of reactive oxygen species (ROS). These mitochondria-mediated processes take on specialized roles in neurons, coordinating aerobic metabolism to meet the high energy demands of these cells, modulating Ca2+ signaling, providing lipids for axon growth and regeneration, and tuning ROS production for neuronal development and function. Mitochondrial dysfunction is therefore a central driver in neurodegenerative diseases. Mitochondrial structure and function are inextricably linked. The morphologically complex inner membrane with structural infolds called cristae harbors many molecular systems that perform the signature processes of the mitochondrion. The architectural features of the inner membrane are ultrastructural and therefore, too small to be visualized by traditional diffraction-limited resolved microscopy. Thus, most insights on mitochondrial ultrastructure have come from electron microscopy on fixed samples. However, emerging technologies in super-resolution fluorescence microscopy now provide resolution down to tens of nanometers, allowing visualization of ultrastructural features in live cells. Super-resolution imaging therefore offers an unprecedented ability to directly image fine details of mitochondrial structure, nanoscale protein distributions, and cristae dynamics, providing fundamental new insights that link mitochondria to human health and disease. This protocol presents the use of stimulated emission depletion (STED) super-resolution microscopy to visualize the mitochondrial ultrastructure of live human neuroblastoma cells and primary rat neurons. This procedure is organized into five sections: (1) growth and differentiation of the SH-SY5Y cell line, (2) isolation, plating, and growth of primary rat hippocampal neurons, (3) procedures for staining cells for live STED imaging, (4) procedures for live cell STED experiments using a STED microscope&#160;for reference, and (5) guidance for segmentation and image processing using examples to measure and quantify morphological features of the inner membrane.

This protocol describes cell growth conditions for cultured and primary cells with a focus on live cell STED imaging and subsequent analyses of mitochondrial cristae. Projections made with ImageJ of mitochondria from undifferentiated SH-SY5Y (Figure 3A) and RA-differentiated SH-SY5Y (Figure 3B) cells can be collected as z-stacks with traditional confocal and STED, and the raw STED images can then be deconvolved. Timelapse imaging can also be performed and subsequently deconvolved (Figure 3C,D). Using slightly different imaging parameters for primary rat hippocampal neurons (Table 1), confocal and raw STED images can be acquired as z-stacks, and the raw STED images can be deconvolved (Figure 4A). Timelapse imaging of mitochondria from primary neurons is also possible (Figure 4B). In general, the time-lapse images should be able to show mitochondrial dynamic events.
When raw STED and deconvolved STED z-stack projections from the samples used for segmentation appear consistent, quantitative measurements are performed. The TWS plugin uses the deconvolved STED image to segment to make a probability mask, which is then used to create a binary mask of the cristae to obtain size and shape parameters (Figure 5A). The regions from this mask are saved in the ROI manager and can be applied to the raw STED image if desired to measure differences in relative intensity. The deconvolved STED projections can also be used to determine the cristae periodicity and density in a given area (Figure 5B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65561/65561fig01.jpg" /> 
Figure 1: Mitochondrial morphology. Mitochondria have a two-membrane system that defines different subcompartments (A). Cristae are infolds of the inner membrane with defined features (B). Abbreviations: OMM, outer mitochondrial membrane; ICS, intracristal space; IMS, intermembrane space; CM, cristae membrane; IBM, inner boundary membrane; IMM, inner mitochondrial membrane; CT, cristae tip; CJ, cristae junction. <a href="https://www.jove.com/files/ftp_upload/65561/65561fig01large.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/65561/65561fig02.jpg" /> 
Figure 2: Schematic of workflow. SH-SY5Y cells or primary rat hippocampal neurons are grown on a PDL-coated coverglass. SH-SY5Y cells are grown in parallel to remain undifferentiated or subjected to RA differentiation over the course of six days. Primary rat hippocampal neurons were grown on a PDL-coated coverglass after being isolated from hippocampal sections for seven days. Once ready to be imaged, cells were stained with PKMO and imaged with STED. Raw STED images are then deconvolved, and the deconvolved images are processed in FIJI to obtain size and shape measurements, such as cristae density, area, perimeter, circularity, and aspect ratio. <a href="https://www.jove.com/files/ftp_upload/65561/65561fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65561/65561fig03.jpg" /> 
Figure 3: Imaging of mitochondria in SH-SY5Y cells. Representative confocal (left), raw STED (center), and Huygens deconvolved STED image z-stack projections (right) of mitochondria from non-differentiated (A) and&#160;RA-differentiated (B) SH-SY5Y cells with PKMO staining are shown. A timelapse with 30 s intervals and 5 iterations of RA-differentiated SH-SY5Y cells is shown (C) with selected regions (white boxes) expanded upon (D) using scaled images of those regions without interpolation. Scale bars: A,B, 250 nm; C,D, 1 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65561/65561fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65561/65561fig04.jpg" /> 
Figure 4: Imaging of mitochondria in primary rat hippocampal neurons. Representative confocal (left), raw STED (center), and Huygens deconvolved STED (right) image z-stack projections of mitochondria from primary rat hippocampal neurons are shown (A). A timelapse with 25 s intervals and 5 iterations of mitochondria in these neurons is shown (B). Scale bars: A, 250 nm; B, 1 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65561/65561fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65561/65561fig05.jpg" /> 
Figure 5: Processing of deconvolved STED images in ImageJ. Representative use of the Trainable Weka Segmentation plugin to measure cristae size and shape is shown (A). From left to right, the following images are shown: the deconvolved STED image, the probability map based on segmentation from the TWS plugin, the mask from thresholding in FIJI using the probability map as input, the mask with the ROIs outlined, and the ROIs overlayed onto the original deconvolved STED image. The resulting area, perimeter, circularity and aspect ratio measurements corresponding to these objects can be found in Supplementary Table 1.&#160;A line plot using the deconvolved STED image to measure peak-to-peak distances as a readout for cristae density is shown (B). Scale bars: 0.5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65561/65561fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Pixel size (nm)</td>
			<td>Dwell time (&#181;s)</td>
			<td>Line acc.</td>
			<td>561 nm excitation during STED acquisition (%)</td>
			<td>775 nm STED depletion power (%)</td>
			<td>Step size (nm)</td>
			<td>Pinhole (AU)</td>
			<td>Timelapse interval (s)</td>
			<td>Timelapse iterations</td>
		</tr>
		<tr>
			<td>Undiffer-entiated SH-SY5Y</td>
			<td>20</td>
			<td>4</td>
			<td>10</td>
			<td>15</td>
			<td>20</td>
			<td>200</td>
			<td>1.0</td>
			<td>30</td>
			<td>5</td>
		</tr>
		<tr>
			<td>RA-Differe-ntiated SH-SY5Y</td>
			<td>20-25</td>
			<td>4</td>
			<td>10-12</td>
			<td>15</td>
			<td>20-22</td>
			<td>150-200</td>
			<td>1.0</td>
			<td>30</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Primary Neurons</td>
			<td>20-25</td>
			<td>4</td>
			<td>10</td>
			<td>10</td>
			<td>25</td>
			<td>200</td>
			<td>0.7</td>
			<td>30</td>
			<td>5</td>
		</tr>
		<tr>
			<td colspan="10">NOTE: Pixel size can vary based on imaging requirements and intent to deconvolve images. Proper sampling is required for deconvolution. Pixel sizes for raw STED images without deconvolution can go up to 30 nm.</td>
		</tr>
	</tbody>
</table>
Table 1: Summary of STED acquisition parameters. The settings used for 2D STED imaging for each cell type, undifferentiated SH-SY5Y, RA-differentiated SH-SY5Y, and primary rat hippocampal neurons, are displayed. For all time-lapses, 5 iterations were taken with varying intervals based on ROI size.
Supplementary Figure 1: Imaging of SH-SY5Y cells with amyloid-&#946; (A&#946;) addition. Representative confocal (left), raw STED (center), and deconvolved STED (right) images of RA-differentiated SH-SY5Y cells with PKMO stain (top) and A&#946;-HiLyte647 (bottom) are shown (A). Merged z-stack projections of raw PKMO STED (green) with raw&#160;A&#946; STED (magenta) (B) or deconvolved PKMO STED (green) with deconvolved A&#946; STED (magenta) (C) are shown. Scale bars: 0.5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65561/Suppl Fig.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Table 1: Size and shape measurements of segmented cristae. The size and shape measurements of area (&#181;m2), perimeter (&#181;m), circularity, and aspect ratio, corresponding to the objects outlined in Figure 5A from segmented mitochondria, are shown. <a href="https://www.jove.com/files/ftp_upload/65561/65561_Supplementary Table 1.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary Table 2: Summary of acquisition parameters with amyloid-&#946; samples. The settings used for 2D STED imaging of PKMO and A&#946;-HiLyte647 in undifferentiated and RA-differentiated SH-SY5Y cells are displayed. Confocal of A&#946;-HiLyte647 may be used alone as there is no specific structure to resolve; STED images of A&#946;-HiLyte647 are shown here for smaller particle sizes. <a href="https://www.jove.com/files/ftp_upload/65561/65561_Supplementary Table 2.xlsx" target="_blank">Please click here to download this File.</a>
Supplementary File 1: Amyloid-&#946; treatment protocol. <a href="https://www.jove.com/files/ftp_upload/65561/65561_Supplementary File 1.docx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Decoding Mitochondrial Aging at JoVE.com

Author Spotlight: Decoding Mitochondrial Aging 

This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of ...

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Research

JoVE Journal

Neuroscience

3.4K Views.  University of Connecticut. This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Video: Author Spotlight: Decoding Mitochondrial Aging 

Automated Sholl Analysis of Digitized Neuronal Morphology at Multiple Scales

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3. Specifically, it affects the ability of neurons to receive inputs from other cells2 and contributes to the propagation of action potentials4,5. The morphology of the neurites also affects how information is processed. The diversity of dendrite morphologies facilitate local and long range signaling and allow individual neurons or groups of neurons to carry out specialized functions within the neuronal network6,7. Alterations in dendrite morphology, including fragmentation of dendrites and changes in branching patterns, have been observed in a number of disease states, including Alzheimer's disease8, schizophrenia9,10, and mental retardation11. The ability to both understand the factors that shape dendrite morphologies and to identify changes in dendrite morphologies is essential in the understanding of nervous system function and dysfunction.
Neurite morphology is often analyzed by Sholl analysis and by counting the number of neurites and the number of branch tips. This analysis is generally applied to dendrites, but it can also be applied to axons. Performing this analysis by hand is both time consuming and inevitably introduces variability due to experimenter bias and inconsistency. The Bonfire program is a semi-automated approach to the analysis of dendrite and axon morphology that builds upon available open-source morphological analysis tools. Our program enables the detection of local changes in dendrite and axon branching behaviors by performing Sholl analysis on subregions of the neuritic arbor. For example, Sholl analysis is performed on both the neuron as a whole as well as on each subset of processes (primary, secondary, terminal, root, etc.) Dendrite and axon patterning is influenced by a number of intracellular and extracellular factors, many acting locally. Thus, the resulting arbor morphology is a result of specific processes acting on specific neurites, making it necessary to perform morphological analysis on a smaller scale in order to observe these local variations12.
The Bonfire program requires the use of two open-source analysis tools, the NeuronJ plugin to ImageJ and NeuronStudio. Neurons are traced in ImageJ, and NeuronStudio is used to define the connectivity between neurites. Bonfire contains a number of custom scripts written in MATLAB (MathWorks) that are used to convert the data into the appropriate format for further analysis, check for user errors, and ultimately perform Sholl analysis. Finally, data are exported into Excel for statistical analysis. A flow chart of the Bonfire program is shown in Figure 1.

We have developed a computer program to analyze neuronal morphology. In combination with two existing open source analysis tools, our program performs Sholl analysis and determines the number of neurites, branch points, and neurite tips. The analyses are performed so that local changes in neurite morphology can be observed.

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3.  Specifically, it affects the ability of neurons to ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of existing mouse lines for genetic cell lineage tracing: a tamoxifen-inducible Cre line and a Cre reporter line expressing dsRed upon Cre-mediated recombination. By using a relatively low level of tamoxifen, we were able to induce recombination in a small number of cells, permitting us to follow individual cell divisions. Additionally, we observed the transcriptional response to Sonic Hedgehog (Shh) signaling using an Olig2-eGFP transgenic line 1-3 and we monitored formation of cilia by infecting the cultured slice with virus expressing the cilia marker, Sstr3-GFP 4. In order to image the neuroepithelium, we harvested embryos at E8.5, isolated the neural tube, mounted the neural slice in proper culturing conditions into the imaging chamber and performed time-lapse confocal imaging. Our ex vivo live imaging method enables us to trace single cell divisions to assess the relative timing of primary cilia formation and Shh response in a physiologically relevant manner. This method can be easily adapted using distinct fluorescent markers and provides the field the tools with which to monitor cell behavior in situ and in real time.

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5 neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate decisions, will be crucial to design therapeutic strategies for many diseases affecting the nervous system. Current methods to track cell populations rely on their final outcomes in still images and they generally fail to provide sufficient temporal resolution to identify behavioral features in single cells. Moreover, variations in cell death, behavioral heterogeneity within a cell population, dilution, spreading, or the low efficiency of the markers used to analyze cells are all important handicaps that will lead to incomplete or incorrect read-outs of the results. Conversely, performing live imaging and single cell tracking under appropriate conditions represents a powerful tool to monitor each of these events. Here, a time-lapse video-microscopy protocol, followed by post-processing, is described to track neural populations with single cell resolution, employing specific software. The methods described enable researchers to address essential questions regarding the cell biology and lineage progression of distinct neural populations.

A robust protocol to monitor neural populations by time-lapse video-microscopy followed by software-based post-processing is described. This method represents a powerful tool to identify biological events in a selected population during live imaging experiments.

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate ...

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 ...