The authors would like to thank Katelyn Lavrich for assistance with experimental design and manuscript edits.

1. Preparation of Cells
NOTE: This procedure describes the lentiviral transduction of an immortalized cell line (BEAS-2B; ATCC, Manassas, VA) to express the desired reporter (roGFP2 or HyPer). Other cell lines/types and/or methods of gene transfer, including transfection, can be utilized as long as they result in a level of reporter expression adequate to visualize a sufficient number of sensor-expressing cells per field of view (typically 5 - 10 cells). If using transfection methods, the procedure should be performed in the dish that will ultimately be used for the method of analysis (e.g., transfect the cells in the same dish that will be presented to the microscope). The steps described below provide details for preparing cell transductions to be performed in a 6-well cell culture plate. If this format is not an appropriate size for the desired application, other vessels can be substituted.
<ol>
	<li>Grow cells to approximately 40 - 60% confluence in normal growth media. 
	NOTE: This study utilized the human bronchial epithelial cell line, BEAS-2B, grown in keratinocyte growth medium (KGM).</li>
	<li>Just prior to transduction, replace the growth media on the cells with 500 &#181;L of serum-free media containing the appropriate amount of virus, as calculated by the formula: 
	<img alt="Equation 1" src="/files/ftp_upload/56945/56945eq1.jpg" />
	<ol>
		<li>For transduction of BEAS-2B cells, use serum-free keratinocyte basal medium (KBM) to replace the KGM growth media during viral incubations. 
		NOTE: The above calculation is for a single transduction of one well of cells in a 6-well format. If needed, perform transductions of several wells can be performed by adjusting the formula with a multiplication by the number of wells to be transduced. For lentiviral transductions, use a multiplicity of infection (MOI) between 5 and 20. For adenoviral transductions, use an MOI between 100 and 500.</li>
	</ol>
	</li>
	<li>Incubate cells with the viral mixture at 37 &#176;C for 4 h, redistributing the viral particles in the dish every 30 to 60 min with a brief rocking or swirling motion.</li>
	<li>Add 1 mL of complete growth media to the dish and incubate at 37 &#176;C for an additional 4 - 16 h.</li>
	<li>Remove all media and replace with fresh complete growth media.</li>
	<li>Continue to grow and passage cells, expand as needed. 
	NOTE: Cells should begin appreciably expressing the sensor within 12 - 48 h. If a lentiviral vector was used for the transduction, stable expression of the desired sensor should continue across passages.</li>
	<li>For microscopy-based assessments, seed cells into glass bottomed microscope dishes and grow to desired confluence (&#8805;70 - 80%) prior to imaging. 
	NOTE: Seeding density will depend on the size of the dish, growth rate of the cell-line being used, and the time of the assessment following seeding. For example, seed 300,000 BEAS-2B cells into a 35-mm dish to yield approximately 70 - 80% confluence after 1 day of growth.</li>
</ol>
2. Microscopy Set-up
NOTE: The protocol described below is performed using a confocal microscope equipped with laser lines at 404 and 488 nm. Other means of making fluorometric assessments of the sensors described within this protocol at their prescribed excitations/emission should also yield viable data. Importantly, equipment settings can vary greatly depending on the type, age, and condition of the instrument being used; thus, any instrument values mentioned may not be specific to the equipment used in other laboratories.
<ol>
	<li>Perform all imaging analysis using environmental controls to maintain an appropriate temperature (e.g., 37 &#176;C), humidity (typically &#62;95% relative humidity), and/or gas concentration (e.g., 5% CO2) suitable for the cells throughout the duration of the experiment.</li>
	<li>If using high values of relative humidity, keep any surfaces that may come in contact with the humidified atmosphere (e.g., the microscope objective) at or slightly above the temperature at which the humidity is generated in order to prevent condensation. This can be accomplished with the use of an objective heater and/or heating tape and an appropriate heater control.</li>
	<li>Turn on all microscope components and set up all equipment required for sequential excitation at 488 and 404 nm with emission at 510 nm. Ensure all components of the optical configuration are set appropriately for real-time acquisition.</li>
	<li>Set up a stage-top environmental chamber to maintain constant temperature at 37 &#176;C, 5% CO2 atmosphere, and &#62;95% humidity. Prior to starting image acquisition, equilibrate the environmental chamber for at least 10 min after the initial set-up of all environmental controls. 
	NOTE: Environmental conditions may be eliminated or adjusted depending on the experiment length, cell type, and exposure being used.</li>
	<li>Place the dish of cells (Step 1.7) on the stage-top within the environmental chamber.</li>
	<li>With the desired objective lens, find the focal plane of cells using the eyepiece and white light, and ensure normal morphology. 
	&#8203;NOTE: A 1.4 NA 60X violet-corrected, oil-immersed objective lens is commonly used, which permits identification of intracellular compartments while maximizing the optical resolution of the confocal system.</li>
	<li>Check the fluorescence expression of the cells in the field of view. Do it while visualizing the cells under wide-field fluorescence illumination with an appropriate filter set, such as fluorescein isothiocyanate (FITC). At this point, using wide-field illumination while looking through the eyepiece is more convenient because it is easier to move the dish to select a field of cells to study. In general, choose a field of view that contains at least 5 to 10 cells that are expressing the sensor, as indicated by green fluorescence.
	<ol>
		<li>Alternatively, perform this assessment confocally using laser excitation at a wavelength that is most compatible with the sensor being expressed (488 nm typically works best). 
		NOTE: Due to their intrinsic fluorescent properties, it will be more difficult to visualize cells expressing HyPer than those expressing roGFP2 using either FITC or at 488 nm. However, it should still be possible to see faint cells.</li>
	</ol>
	</li>
	<li>Once a desired field of view is found, close the environmental chamber. 
	NOTE: Use the focus-maintaining feature, often available on advanced microscope stands, to facilitate maintenance of a stable focal plane throughout the study.</li>
	<li>Set up acquisition parameters to ensure optimal assessment of the sensor of interest across the desired exposure period. Below are recommendations and approaches for confocal imaging of sensor responses:
	<ol>
		<li>Adjust the laser power for excitation at 488 nm and emission at 510 nm. Choose a laser power level that allows visualization of the cells, and keep this constant between samples or replicate dishes. 
		NOTE: For this study, 12% and 1.5% laser power were used for the 488 and 404 nm laser lines, respectively.</li>
		<li>Use the confocal controls of the acquisition software to ensure that the selected focal plane has been optimized for maximal fluorescence emission intensity in the center of the cells (z-axis) by scanning at 488 nm while adjusting the z-plane. This is made easier by using a high gain setting while searching for the z-plane that results in the most over-exposed cells. Once the appropriate focal plane has been found, return the gain to a setting that is most optimal for the fluorescence of the reporter being used without oversaturating the pixels being observed. 
		NOTE: The laser and gain settings that are appropriate for finding and observing cells during experimentation is completely dependent on the confocal system being utilized. In general, once the cells have been found in the field of view, it is recommended that a minimal amount of laser power be used (typically &#8804;20%), as excessive scanning of cells using high-powered laser light may induce oxidative changes detectable by the sensor.</li>
		<li>Use the gain to fine-tune the baseline fluorescence. With roGFP2, establish the baseline near the upper limit (&#8776; 90% relative intensity) of the instrument without over-saturation, as these cells will lose 510 nm fluorescence induced by 488 nm excitation when EGSH increases. In contrast, the fluorescence intensity with 488 nm excitation should be low (&#8776; 10% relative intensity) at baseline for HyPer expressing cells, as these cells will gain 510 nm fluorescence intensity when H2O2 is detected.</li>
		<li>Repeat steps 2.9.2 and 2.9.3 with excitation at 404 nm and emission at 510 nm. Gain settings for the 404 nm excitation wavelength are opposite to those used with 488 nm excitation for each sensor (i.e., low baseline fluorescence (&#8776; 10% relative intensity) at 404 nm for roGFP2, high baseline fluorescence (&#8776; 90% relative intensity) for the H2O2 sensor). 
		NOTE: In general, the fluorescence (510 emission) at 404 nm excitation will be considerably lower than that obtainable with 488 nm excitation in cells expressing both roGFP2 and HyPer, as the 404 nm peak is a relatively minor excitation maximum for both of these sensors.</li>
	</ol>
	</li>
</ol>
3. Data Acquisition
<ol>
	<li>Set up the acquisition software to sequentially excite the two excitation wavelengths (first 488 nm and then 404 nm) and collect emissions for both at 510 nm at a predetermined time interval throughout the desired length of the experiment (e.g. capture images every 60 s for 60 min).
	<ol>
		<li>Alternatively, acquire images manually before and after exposures if the approximate timing of changes in EGSH or H2O2 are known for the xenobiotic being tested. However, this may lead to loss of temporal resolution.</li>
	</ol>
	</li>
	<li>For real-time assessment of experimental parameters, choose at least 5 - 10 sensor-expressing cells in the field and establish them as regions of interest (&#34;ROIs&#34;) to monitor their fluorescence changes during the experiment. 
	&#8203;NOTE: This step is optional, and can be performed after the experiment if direct observation in real-time is undesirable or software limitations prevent continuous monitoring. Depending on the cell population, the selection of sensor expressing cells as ROIs might represent a range of expression levels across cells.
	<ol>
		<li>For these studies, add the environmental toxicant 9,10-phenanthrenequinone (9,10-PQ) or hydrogen peroxide (H2O2) after a 5-min baseline period.</li>
		<li>Prepare all reagents for later use in this protocol.&#160; Dissolve 9,10-PQ in dimethyl sulfoxide (DMSO) to a concentration of 15 mM, and dilute in basal cell media to yield a 250 &#181;M working solution.&#160; Additionally, prepare a working solution of hydrogen peroxide in water that will yield a final concentration of 1mM upon injection.</li>
	</ol>
	</li>
	<li>Once the experimental parameters have been defined, begin the time course acquisition. Establish a baseline period of at least 5 min (or 5 data points) prior to starting xenobiotic exposures.</li>
	<li>Expose the cells to the toxicant being investigated using conventional approaches for in vitro dosing. Prepare soluble compounds in either water or other appropriate solvent and inject directly into the media.
	<ol>
		<li>If organic solvents (e.g. dimethyl sulfoxide or ethanol) are required, keep the final solvent concentration at or below 0.1%. Verify that the solvent does not produce an effect on EGSH or H2O2 production with a vehicle control in a separate dish of cells.</li>
		<li>For compounds with lower solubility, add a mixing step using a micropipette to the protocol after the injection is made. Pump gaseous exposures directly into the environmental chamber using the appropriate carrier gas mixture.</li>
	</ol>
	</li>
	<li>Monitor changes in fluorescence during the exposure period. Perform subsequent injections as needed. Take great care not to shift the dish during injections, so that the same cells are followed throughout the entire time course while imaged in the same focal plane.</li>
	<li>At the end of the experiment, expose the cells to appropriate controls. For both genetically-encoded fluorogenic sensors, add specific concentrations of compounds known to oxidize and reduce these sensors in a pointed demonstration of their ratiometric responsiveness. H2O2 and dithiothreitol (DTT) serve as appropriate controls to oxidize and reduce both sensors. Typically, a final concentration of 100 - 1,000 &#181;M H2O2, followed by 1-5 mM dithiothreitol (DTT), allows the user to fully oxidize and reduce these sensors.
	<ol>
		<li>For these studies, use 1 mM H2O2 followed by 5 mM DTT to determine the maximum sensor response.</li>
		<li>Wait at least 5 min to allow the sensor to respond, and then inject a reducing agent, such as DTT, to reduce the senor and return it to a level of fluorescence at or near its established baseline. 
		NOTE: The use of controls in this step is crucial for normalization, and should be performed at the end of every experiment to determine the dynamic range of the sensor in each cell. For roGFP2, aldrithiol can also be added at the end of the experiment. Aldrithiol bypasses the roGFP2 redox relay to oxidize the sensor directly. This is useful to assess sensor function following xenobiotic exposure.</li>
	</ol>
	</li>
</ol>
4. Data Analysis
<ol>
	<li>If not already established, draw ROIs around the cells to be analyzed. Use the appropriate software to measure the fluorescence intensity of each ROI at each wavelength throughout the time course. Ensure that the cells did not move or the plate did not shift during the run. 
	NOTE: The establishment of ROIs is most easily accomplished by drawing an enclosed region that follows the fluorescent expression pattern of each cell. To further assist with this process, a transmitted light (or brightfield) image of the cells within the established field of view can be overlaid onto the fluorescent image in efforts to define cell boundaries. However, the use of a transmitted light image requires the user to capture an additional channel (set of images) throughout the course of the experiment. Alternatively, certain manufacturers incorporate algorithms into their imaging software that use specific parameters such as intensity thresholds to establish ROIs in a more quantitative, less subjective, method.</li>
	<li>Export the data to desired analysis software (e.g., electronic spreadsheet).</li>
	<li>Calculate the ratiometric sensor response for each ROI at each time point using the formulas: 
	<img alt="Equation 2" src="/files/ftp_upload/56945/56945eq2.jpg" /> 
	<img alt="Equation 3" src="/files/ftp_upload/56945/56945eq3.jpg" /></li>
	<li>For each time point, average the calculated ratiometric values of all ROIs.</li>
	<li>Calculate the baseline value, which will be used to normalize the data, by averaging the previously calculated ratiometric values (step 4.4) collected before addition of the xenobiotic (e.g., the first five-time points).</li>
	<li>Normalize the ratiometric values from step 4.4 by dividing each value by the baseline value calculated in 4.5. Normalized ratios will center around a value of 1 at baseline, while increases in EGSH or H2O2 following injections will cause the ratio to increase. Assessments of xenobiotic that induce oxidative changes tend to yield values in a range of 3 - 6 times the baseline value.</li>
	<li>To aid in comparing responses within and across experiments, express the normalized data as a percentage of maximal sensor response by defining the response to the control oxidant (e.g., 1 mM H2O2) as 100%. 
	NOTE: If expressing data as a percentage of maximal sensor response, it is critical to ensure that the same concentration of the control oxidant is used consistently across experiments. All data derived from live-cell imaging analysis is amenable to conventional pair-wise and group-wise statistical analysis to be chosen at the discretion of the investigator.</li>
</ol>

The research described in this article has been reviewed by the National Health and Environmental Effects Research Laboratory, U.S. Environmental Protection Agency, and approved for publication. The contents of this article should not be construed to represent agency policy, nor does mention of trade names or commercial products constitute endorsement or recommendation for use.

The use of roGFP2 and HyPer in detecting changes in intracellular EGSH and H2O2, respectively, has been well-described in previous physiological and toxicological studies 25,35,36,39,40,41,42,43. The protocol outlined here r...

The term &#34;oxidative stress&#34; is frequently cited as a mechanism in toxicology, yet rarely is this term described specifically. Oxidative stress can refer to several intracellular processes, including generation of reactive oxygen species, damage caused by free radicals, the oxidation of antioxidant molecules, and even the activation of specific signaling cascades. A broad range of environmental contaminants1,2 and pharmaceutical agents3,4 have been documented to induce oxidative stress by either direct action of the xenobiotic compound itself5,6 or secondarily by production of oxidant species as part of a cellular response7,8,9,10. It is therefore of great interest in toxicology to accurately observe and characterize the oxidative processes leading to adverse outcomes. Conventional methods of measuring oxidative stress involve identification of oxidized biomolecules11,12,13,14,15 or antioxidants16,17,18,19,20, or direct measurement of reactive species themselves21,22,23,24. However, these methods typically require cellular disruption, which often consumes the sample, eliminates spatial resolution, and potentially introduces artifacts25. The development of more sensitive and specific methods for the detection of oxidant species and markers of oxidative stress is broadly applicable to the investigation of the adverse effects of xenobiotic exposure.
Live-cell microscopy using a new generation of genetically-encoded fluorogenic sensors has emerged as a powerful tool to monitor the intracellular redox status of living cells. These sensors are typically expressed using a vector under the control of a viral promoter that is introduced via transfection or transduction methodologies. High expression efficiencies are not necessary, since cells expressing the fluorescent sensor can be easily identified visually. For toxicological assessments, cells expressing these sensors can be observed using fluorescence microscopy as they are exposed to xenobiotic compounds in real-time. This experimental design permits repeated measurements in the same cell, allowing each cell&#39;s established baseline to act as its own control. The high temporal resolution afforded by live-cell imaging is well-suited for the detection of oxidative events, particularly those that are modest in magnitude or transient in nature. In addition to being both sensitive and specific to their target molecules, the fluorescence of some of these sensors can be excited using two wavelengths of light. This phenomenon allows the fluorescent emission to be expressed as a ratio, which permits the discernment of signal changes associated with authentic sensor responses from those caused by artefacts such as variations in sensor expression, cell thickness, lamp intensity, photobleaching, and sensitivity of the fluorescence detector26. Another advantage of the use of fluorogenic sensors is that they can be targeted to specific cellular compartments, creating a level of spatial resolution that is unmatched by conventional methods25,26,27.
A large family of genetically-encoded sensors based on green fluorescent protein (GFP) have been developed and characterized to report on a broad variety of physiological markers, including pH, temperature, calcium concentrations, and the ATP/ADP ratio25,28,29,30,31. Included among these are sensors of the glutathione redox potential (EGSH) and hydrogen peroxide (H2O2). While these sensors were developed for applications in redox biology and physiology, they have also been adapted to study xenobiotic-induced oxidative stress. Specifically, the protocol outlined here describes the use of the EGSH sensor roGFP2 and the H2O2 sensor HyPer.
roGFP2 reports on the redox potential of intracellular reduced and oxidized glutathione (GSH/GSSG) through a redox relay involving glutathione peroxidase (GPx), glutaredoxin (Grx), and glutathione reductase (GR) (Figure 1)25,32,33. Glutathione is the predominant cellular antioxidant molecule and is present primarily in its reduced form (GSH) in millimolar concentrations in the cytosol25,34. While EGSH has not been linked to any functional outcome, it is recognized as an important indicator of intracellular oxidative status34. A relatively small increase in the concentration of GSSG results in an increase in EGSH that is detectable by roGFP2. Equally important, monitoring of EGSH using roGFP2 during xenobiotic exposures can potentially reveal much about the mechanism of action at several points in the redox relay and associated pathways, such as the pentose phosphate shunt (Figure 1)35. The second sensor discussed here, HyPer, is an intracellular H2O2 probe derived from the insertion of yellow fluorescent protein (YFP) into the regulatory domain of bacterial H2O2-sensitive transcription factor OxyR136. Although it has previously been considered a damaging reactive oxygen intermediate, H2O2 is increasingly being recognized as an important intracellular signaling molecule under physiological conditions37,38, suggesting that unrecognized roles for H2O2 exist in toxicology as well. For instance, excess H2O2 induced by a xenobiotic exposure could be a precursor to dysregulation in cellular signaling or a shift in bioenergetics.
Both genetically-encoded fluorogenic sensors have been expressed in several established cell lines, including the human epidermoid carcinoma cell line A431 and the human bronchial epithelial cell line BEAS-2B, to observe changes in EGSH and H2O2 in response to a variety of toxicological exposures. These include gaseous pollutants (ozone35), soluble components of particulate matter (1,2 naphthoquinone39,40 and zinc41), and nickel nanoparticles (unpublished data). These studies represent only a small subset of the possible applications of these two sensors. Theoretically, any cell type that is capable of receiving and expressing the DNA of these sensors through conventional molecular biology techniques can be utilized to assess the effects of xenobiotics suspected to alter the cellular oxidative state. To date, one or more of these sensors has been expressed in various prokaryotes and eukaryotes, including several mammalian, plant, bacterial, and yeast cell types25,26,36,42. The readout for both EGSH and H2O2 sensors is a change in the intensity of fluorescence emitted at 510 nm upon excitation with 488 and 404 nm light. This method is widely adaptable across fluorometric platforms, including various types of microscopy (confocal and wide-field) and plate readers. The method presented here allows for sensitive and specific observation of intracellular EGSH and H2O2 in in vitro toxicological systems.

<table><tbody><tr><td>roGFP2 Plasmid</td><td>University of Oregon</td><td>N/A</td><td>Generous gift of S.J. Remington</td></tr><tr><td>HyPer Plasmid</td><td>Evrogen</td><td>FP941</td><td></td></tr><tr><td>Hydrogen Peroxide</td><td>Sigma Aldrich</td><td>H1009</td><td></td></tr><tr><td>Dithiothreitol</td><td>Sigma Aldrich</td><td>10708984001</td><td></td></tr><tr><td>9,10-Phenanthrenequinone</td><td>Sigma Aldrich</td><td>156507</td><td></td></tr><tr><td>Black Wall Glass Bottomed Dishes</td><td>Ted Pella</td><td>14029-20</td><td></td></tr><tr><td>BEAS-2B cell line</td><td>American Type Culture Collection</td><td>CRL-9609</td><td></td></tr><tr><td>Keratinocyte Basal Medium</td><td>Lonza</td><td>192151</td><td></td></tr><tr><td>Keratinocyte Growth Medium BulletKit</td><td>Lonza</td><td>192060</td><td></td></tr><tr><td>Excel&amp;nbsp;</td><td>Microsoft Office Suite</td><td>N/A</td><td></td></tr><tr><td>NIS-Elements AR Imaging Software</td><td>Nikon</td><td>N/A</td><td></td></tr><tr><td>Nikon C1si Confocal Imaging System</td><td>Nikon</td><td>N/A</td><td></td></tr></tbody></table>

imaging approaches to assessments of toxicological oxidative stress using genetically-encoded fluorogenic sensors

While oxidative stress is a commonly cited toxicological mechanism, conventional methods to study it suffer from a number of shortcomings, including destruction of the sample, introduction of potential artifacts, and a lack of specificity for the reactive species involved. Thus, there is a current need in the field of toxicology for non-destructive, sensitive, and specific methods that can be used to observe and quantify intracellular redox perturbations, more commonly referred to as oxidative stress. Here, we present a method for the use of two genetically-encoded fluorogenic sensors, roGFP2 and HyPer, to be used in live-cell imaging studies to observe xenobiotic-induced oxidative responses. roGFP2 equilibrates with the glutathione redox potential (EGSH), while HyPer directly detects hydrogen peroxide (H2O2). Both sensors can be expressed into various cell types via transfection or transduction, and can be targeted to specific cellular compartments. Most importantly, live-cell microscopy using these sensors offers high spatial and temporal resolution that is not possible using conventional methods. Changes in the fluorescence intensity monitored at 510 nm serves as the readout for both genetically-encoded fluorogenic sensors when sequentially excited by 404 nm and 488 nm light. This property makes both sensors ratiometric, eliminating common microscopy artifacts and correcting for differences in sensor expression between cells. This methodology can be applied across a variety of fluorometric platforms capable of exciting and collecting emissions at the prescribed wavelengths, making it suitable for use with confocal imaging systems, conventional wide-field microscopy, and plate readers. Both genetically-encoded fluorogenic sensors have been used in a variety of cell types and toxicological studies to monitor cellular EGSH and H2O2 generation in real-time. Outlined here is a standardized method that is widely adaptable across cell types and fluorometric platforms for the application of roGFP2 and HyPer in live-cell toxicological assessments of oxidative stress.

The use of roGFP2 and HyPer in detecting changes in EGSH and intracellular H2O2 has been well described previously25,36,42,43 and is demonstrated here. Confocal images of cells expressing roGFP2 at baseline and following addition of H2O2 and DTT are shown in Figure 2. Data from ...

Watch this Scientific Journal Video about Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors at JoVE.com

Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors

This manuscript describes the use of genetically-encoded fluorogenic reporters in an application of live-cell imaging for the examination of xenobiotic-induced oxidative stress. This experimental approach offers unparalleled spatiotemporal resolution, sensitivity, and specificity while avoiding many of the shortcomings of conventional methods used for the detection of toxicological oxidative stress.

While oxidative stress is a commonly cited toxicological mechanism, conventional methods to study it suffer from a number of shortcomings, including ...

imaging-approaches-to-assessments-toxicological-oxidative-stress

Research

JoVE Journal

Cancer Research

7.4K Views.  University of North Carolina at Chapel Hill. Dit manuscript beschrijft het gebruik van genetisch-gecodeerde fluorogenic verslaggevers in een toepassing voor live-cel imaging voor de behandeling van xenobioticum-geïnduceerde oxidatieve stress. Deze experimentele aanpak biedt ongeëvenaarde Spatio resolutie, gevoeligheid en specificiteit terwijl het vermijden van veel van de tekortkomingen van conventionele methoden voor de detectie van toxicologische oxidatieve stress.

Video: Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors

Screening Assay for Oxidative Stress in a Feline Astrocyte Cell Line, G355-5

An often-suggested mechanism of virus induced neuronal damage is oxidative stress. Astrocytes have an important role in controlling oxidative stress of the Central Nervous System (CNS). Astrocytes help maintain a homeostatic environment for neurons as well as protecting neurons from Reactive Oxygen Species (ROS). CM-H2DCFDA is a cell-permeable indicator for the presence of ROS. CM-H2DCFDA enters the cell as a non-fluorescent compound, and becomes fluorescent after cellular esterases remove the acetate groups, and the compound is oxidized. The number of cells, measured by flow cytometry, that are found to be green fluorescing is an indication of the number of cells that are in an oxidative state. CM-H2DCFDA is susceptible to oxidation by a large number of different ROS. This lack of specificity, regarding which ROS can oxidize CM-H2DCFDA, makes this compound a valuable regent for use in the early stages of a pathogenesis investigation, as this assay can be used to screen for an oxidative cellular environment regardless of which oxygen radical or combination of ROS are responsible for the cellular conditions. Once it has been established that ROS are present by oxidation of CM-H2DCFDA, then additional experiments can be performed to determine which ROS or combination of ROSs are involved in the particular pathogenesis process. The results of this study demonstrate that with the addition of hydrogen peroxide an increase in CM-H2DCFDA fluoresce was detected relative to the saline controls, indicating that this assay is a valuable test for detecting an oxidative environment within G355-5 cells, a feline astrocyte cell line.

A screening method to detect oxidative cellular environments is to measure the oxidation of CM-H2DCFDA. Once oxidized within a cell, CM-H2DCFDA changes from non-fluorescent into a fluorescent compound. This change in fluorescence is measured by flow cytometry and indicates the number of cells in an oxidative environment.

An often-suggested mechanism of virus induced neuronal damage is oxidative stress. Astrocytes have an important role in controlling oxidative stress of ...

Neuroscience

Assessment of Cellular Oxidation using a Subcellular Compartment-Specific Redox-Sensitive Green Fluorescent Protein

Measuring the intracellular oxidation/reduction balance provides an overview of the physiological and/or pathophysiological redox status of an organism. Thiols are especially important for illuminating the redox status of cells via their reduced dithiol and oxidized disulfide ratios. Engineered cysteine-containing fluorescent proteins open a new era for redox-sensitive biosensors. One of them, redox-sensitive green fluorescent protein (roGFP), can easily be introduced into cells with adenoviral transduction, allowing the redox status of subcellular compartments to be evaluated without disrupting cellular processes. Reduced cysteines and oxidized cystines of roGFP have excitation maxima at 488 nm and 405 nm, respectively, with emission at 525 nm. Assessing the ratios of these reduced and oxidized forms allows the convenient calculation of redox balance within the cell. In this method article, immortalized human triple-negative breast cancer cells (MDA-MB-231) were used to assess redox status within the living cell. The protocol steps include MDA-MB-231 cell line transduction with adenovirus to express cytosolic roGFP, treatment with H2O2, and assessment of cysteine and cystine ratio with both flow cytometry and fluorescence microscopy.

This protocol describes the assessment of subcellular compartment-specific redox status within the cell. A redox-sensitive fluorescent probe allows convenient ratiometric analysis in intact cells.

Measuring the intracellular oxidation/reduction balance provides an overview of the physiological and/or pathophysiological redox status of an organism. ...

Biochemistry

Novel ROS Quantification Using Dihydroethidium in Liver Cancer Cells

High Throughput Screening Assessment of Reactive Oxygen Species (ROS) Generation using Dihydroethidium (DHE) Fluorescence Dye

Reactive oxygen species (ROS) play a key role in the regulation of cellular metabolism in physiological and pathological processes. Physiological ROS production plays a central role in the spatial and temporal modulation of normal cellular functions such as proliferation, signaling, apoptosis, and senescence. In contrast, chronic ROS overproduction is responsible for a wide spectrum of diseases, such as cancer, cardiovascular disease, and diabetes, among others. Quantifying ROS levels in an accurate and reproducible manner is thus essential to understanding normal cellular functionality. Fluorescence imaging-based methods to characterize intra-cellular ROS species is a common approach. Many of the imaging ROS protocols in the literature use 2&#39;-7&#39;-dichlorodihydrofluorescein diacetate (DCFH-DA) dye. However, this dye suffers from significant limitations in its usage and interpretability. The current protocol demonstrates the use of a dihydroethidium (DHE) fluorescent probe as an alternative method to quantify total ROS production in a high-throughput setting. The high throughput imaging platform, CX7 Cellomics, was used to measure and quantify the ROS production. This study was conducted in three hepatocellular cancer cell lines - HepG2, JHH4, and HUH-7. This protocol provides an in-depth description of the various procedures involved in the assessment of ROS within the cells, including - preparation of DHE solution, incubation of cells with DHE solution, and measurement of DHE intensity necessary to characterize the ROS production. This protocol demonstrates that DHE fluorescent dye is a robust and reproducible choice to characterize intracellular ROS production in a high-throughput manner. High throughput approaches to measure ROS production are likely to be helpful in a variety of studies, such as toxicology, drug screening, and cancer biology.

Author Spotlight: High-Throughput Measurement of Intracellular ROS Levels in Hepatocellular Lines

This protocol describes a novel method to quantify intracellular reactive oxygen species (ROS) using dihydroethidium (DHE) as a fluorescence dye probe using a high-throughput screening approach. The protocol describes the methods for quantitative assessment of intracellular reactive oxygen species (ROS) in the three different hepatocellular carcinoma cell lines.

Reactive oxygen species (ROS) play a key role in the regulation of cellular metabolism in physiological and pathological processes. Physiological ROS ...

Bioengineering

Analysis of Oxidative Stress in Zebrafish Embryos

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes oxidation of molecules (lipid, DNA, protein) and leads to cell death. Oxidative stress also impacts the progression of several pathological conditions such as diabetes, retinopathies, neurodegeneration, and cancer. Thus, it is important to define tools to investigate oxidative stress conditions not only at the level of single cells but also in the context of whole organisms. Here, we consider the zebrafish embryo as a useful in vivo system to perform such studies and present a protocol to measure in vivo oxidative stress. Taking advantage of fluorescent ROS probes and zebrafish transgenic fluorescent lines, we develop two different methods to measure oxidative stress in vivo: i) a &#8220;whole embryo ROS-detection method&#8221; for qualitative measurement of oxidative stress and ii) a &#8220;single-cell&#160;ROS detection method&#8221; for quantitative measurements of oxidative stress. Herein, we demonstrate the efficacy of these procedures by increasing oxidative stress in tissues by oxidant agents and physiological or genetic methods. This protocol is amenable for forward genetic screens and it will help address cause-effect relationships of ROS in animal models of oxidative stress-related pathologies such as neurological disorders and cancer.

Here we report a protocol to measure oxidative stress in living zebrafish embryos. This procedure allows reactive oxygen species (ROS) detection in both whole embryo tissues and single-cell populations. This protocol will accomplish both qualitative and quantitative analyses.

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes ...

Biology

Live Imaging of the Mitochondrial Glutathione Redox State in Primary Neurons using a Ratiometric Indicator

Mitochondrial redox homeostasis is important for neuronal viability and function. Although mitochondria contain several redox systems, the highly abundant thiol-disulfide redox buffer glutathione is considered a central player in antioxidant defenses. Therefore, measuring the mitochondrial glutathione redox potential provides useful information about mitochondrial redox status and oxidative stress. Glutaredoxin1-roGFP2 (Grx1-roGFP2) is a genetically encoded, green fluorescent protein (GFP)-based ratiometric indicator of the glutathione redox potential that has two redox-state-sensitive excitation peaks at 400 nm and 490 nm with a single emission peak at 510 nm. This article describes how to perform confocal live microscopy of mitochondria-targeted Grx1-roGFP2 in primary hippocampal and cortical neurons. It describes how to assess steady-state mitochondrial glutathione redox potential (e.g., to compare disease states or long-term treatments) and how to measure redox changes upon acute treatments (using the excitotoxic drug N-methyl-D-aspartate (NMDA) as an example). In addition, the article presents co-imaging of Grx1-roGFP2 and the mitochondrial membrane potential indicator, tetramethylrhodamine, ethyl ester (TMRE), to demonstrate how Grx1-roGPF2 can be multiplexed with additional indicators for multiparametric analyses. This protocol provides a detailed description of how to (i) optimize confocal laser scanning microscope settings, (ii) apply drugs for stimulation followed by sensor calibration with diamide and dithiothreitol, and (iii) analyze data with ImageJ/FIJI.

This article describes a protocol to determine differences in basal redox state and redox responses to acute perturbations in primary hippocampal and cortical neurons using confocal live microscopy. The protocol can be applied to other cell types and microscopes with minimal modifications.

Mitochondrial redox homeostasis is important for neuronal viability and function. Although mitochondria contain several redox systems, the highly abundant ...

Quantification of Reactive Oxygen Species Using 2&#8242;,7&#8242;-Dichlorofluorescein Diacetate Probe and Flow-Cytometry in M&#252;ller Glial Cells

The redox balance has an important role in maintaining cellular homeostasis. The increased generation of reactive oxygen species (ROS) promotes the modification of proteins, lipids, and DNA, which finally may lead to alteration in cellular function and cell death. Therefore, it is beneficial for cells to increase their antioxidant defense in response to detrimental insults, either by activating an antioxidant pathway like Keap1/Nrf2 or by improving redox scavengers (vitamins A, C, and E, &#946;-carotene, and polyphenols, among others). Inflammation and oxidative stress are involved in the pathogenesis and progression of retinopathies, such as diabetic retinopathy (DR) and retinopathy of prematurity (ROP). Since M&#252;ller glial cells (MGCs) play a key role in the homeostasis of neural retinal tissue, they are considered an ideal model to study these cellular protective mechanisms. In this sense, quantifying ROS levels with a reproducible and simple method is essential to assess the contribution of pathways or molecules that participate in the antioxidant cell defense mechanism. In this article, we provide a complete description of the procedures required for the measurement of ROS with DCFH-DA probe and flow cytometry in MGCs. Key steps for flow cytometry data processing with the software are provided here, so the readers will be able to measure ROS levels (geometric means of FITC) and analyze fluorescence histograms. These tools are highly helpful to evaluate not only the increase in ROS after a cellular insult but also to study the antioxidant effect of certain molecules that can provide a protective effect on the cells.

Here, we propose a systematized, accessible, and reproducible protocol to detect cellular reactive oxygen species (ROS) using 2&#8242;,7&#8242;-dichlorofluorescein diacetate probe (DCFH-DA) in M&#252;ller glial cells (MGCs). This method quantifies total cellular ROS levels with a flow cytometer. This protocol is very easy to use, suitable, and reproducible.

The redox balance has an important role in maintaining cellular homeostasis. The increased generation of reactive oxygen species (ROS) promotes the ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...

Measurements of Physiological Stress Responses in C. Elegans

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and physiology. Thus, organisms have evolved targeted and specific stress responses dedicated to repair damage and maintain homeostasis. These mechanisms include the unfolded protein response of the endoplasmic reticulum (UPRER), the unfolded protein response of the mitochondria (UPRMT), the heat shock response (HSR), and the oxidative stress response (OxSR). The protocols presented here describe methods to detect and characterize the activation of these pathways and their physiological consequences in the nematode, C. elegans. First, the use of pathway-specific fluorescent transcriptional reporters is described for rapid cellular characterization, drug screening, or large-scale genetic screening (e.g., RNAi or mutant libraries). In addition, complementary, robust physiological assays are described, which can be used to directly assess sensitivity of animals to specific stressors, serving as functional validation of the transcriptional reporters. Together, these methods allow for rapid characterization of the cellular and physiological effects of internal and external proteotoxic perturbations.

Measurements of Physiological Stress Responses in C. Elegans

Here, we characterize cellular proteotoxic stress responses in the nematode C. elegans by measuring the activation of fluorescent transcriptional reporters and assaying sensitivity to physiological stress.

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and ...

Analyzing Oxidative Stress in Murine Intestinal Organoids using Reactive Oxygen Species-Sensitive Fluorogenic Probe

Reactive oxygen species (ROS) play essential roles in intestinal homeostasis. ROS are natural by-products of cell metabolism. They are produced in response to infection or injury at the mucosal level as they are involved in antimicrobial responses and wound healing. They are also critical secondary messengers, regulating several pathways, including cell growth and differentiation. On the other hand, excessive ROS levels lead to oxidative stress, which can be deleterious for cells and favor intestinal diseases like chronic inflammation or cancer. This work provides a straightforward method to detect ROS in the intestinal murine organoids by live imaging and flow cytometry, using a commercially available fluorogenic probe. Here the protocol describes assaying the effect of compounds that modulate the redox balance in intestinal organoids and detect ROS levels in specific intestinal cell types, exemplified here by the analysis of the intestinal stem cells genetically labeled with GFP. This protocol may be used with other fluorescent probes.

The present protocol describes a method to detect reactive oxygen species (ROS) in the intestinal murine organoids using qualitative imaging and quantitative cytometry assays. This work can be potentially extended to other fluorescent probes to test the effect of selected compounds on ROS.

Reactive oxygen species (ROS) play essential roles in intestinal homeostasis. ROS are natural by-products of cell metabolism. They are produced in ...

Fluorescent Probe Imaging Assay: A Technique to Visualize Oxidative Stress in the Reactive Oxygen Species Inducer-Treated Cultured Intestinal Organoid Cells

Source: Stedman, A. et al., <a href="https://app.jove.com/v/62880">Analyzing Oxidative Stress in Murine Intestinal Organoids using Reactive Oxygen Species-Sensitive Fluorogenic Probe.</a> J. Vis. Exp. (2021).
This video describes the technique of visualizing oxidative stress in cultured organoids. This work helps in understanding the level of oxidative stress due to ROS accumulation in the various cell types present in an organoid after treatment with different ROS modulators.

Source: Stedman, A. et al., Analyzing Oxidative Stress in Murine Intestinal Organoids using Reactive Oxygen Species-Sensitive Fluorogenic Probe. J. Vis. ...

Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors

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