This work was made possible by funding from the German Federal Ministry of Education and Research (BMBF, 1315883) as well as by the S&#228;chsische Aufbaubank (SAB) project 100116526 from a funding of the European Regional Development Fund (ERDF).

All human blood samples were obtained from healthy volunteers, and the applied leukocyte enrichment protocol follows the guidelines of ethics commission of the Medical Faculty of the University of Leipzig. The experiments with rat blood were approved by the responsible local ethical committee (Landesdirektion Sachsen, Referat 24), according to the German guidelines on animal care and use.
1. Experimental Setup
NOTE: As the hypotonic lysis procedure for the depletion of erythrocytes from the blood samples is a time-critical procedure, prepare all necessary equipment (e.g., buffers) for this part of the protocol in advance.
<ol>
	<li>Label one 15 ml centrifuge tube for hypotonic lysis and one 1.5 ml sample tube for subsequent aminophenyl fluorescein (APF) staining per blood sample.
	<ol>
		<li>If the leukocyte enrichment will be performed under sterile conditions, perform this labeling under a flow box.</li>
	</ol>
	</li>
	<li>Prepare about 12 ml phosphate-buffered saline (PBS, 10 mM) per sample. Depending on whether the leukocytes shall be isolated under sterile conditions or not, prepare PBS either by using sterile ready solution from a supplier or by dissolving PBS tablets in distilled water. Check the pH value and, if necessary, adjust to pH 7.4 by using small amounts of 0.1 M HCl and NaOH solutions.
	<ol>
		<li>Dissolve PBS tablets (see Table of Materials) in 200 ml distilled water to obtain a non-sterile buffer solution sufficient for about 16 samples. If desired, sterilize this solution by sterile filtration.</li>
	</ol>
	</li>
	<li>Prepare about 1 ml Hanks balanced salt solution (HBSS) supplemented with Ca2+ per sample.
	<ol>
		<li>Dissolve 970 mg of HBSS salt in 100 ml (final volume) double-distilled water. Before filling up to the final volume, check the pH value and adjust it to pH 7.4. Due to its low buffer capacity, prepare this buffer freshly each day and perform the pH adjustment with special care by using small amounts of 0.1 M HCl and NaOH solutions. 
		NOTE: Sterile solution may be used, depending on the experiment. The solution may be sterilized by sterile filtration.</li>
	</ol>
	</li>
	<li>Besides the two buffers, label and prepare a tube (e.g., 50 ml centrifugation tube) or flask (e.g., 250 ml measuring cup) with double-distilled water for the hypotonic lysis procedure in advance. Prepare about 10 ml water per sample.</li>
	<li>Prepare a container with crushed ice for the APF staining (section 3), which is performed on ice.</li>
	<li>Prepare aliquots of APF in advance to allow the quick preparation of working solutions used during the staining (step 3.2) and to avoid repeatedly freezing and thawing of the APF solution. 
	NOTE: APF is commonly obtained as a 5 mg/ml (11.81 mM) stock solution in methyl acetate.
	<ol>
		<li>Freeze aliquots of 10-100 &#181;l in 0.5 ml tubes. For the APF staining a final dye concentration of 10 &#181;M is used (step 3.8).</li>
	</ol>
	</li>
</ol>
2. Hypotonic Lysis of the Erythrocytes
NOTE: Perform the hypotonic lysis steps (except the centrifugation steps) under a laminar flow bench to avoid contamination. Perform the whole procedure at room temperature. As the hypotonic lysis is a time critical process, adjust the pipettes for water (2 ml) and PBS (5 ml) addition in advance and open the water and PBS flasks before starting the procedure. Store the PBS solution and the distilled water at room temperature before usage.
<ol>
	<li>From each blood sample transfer 100 &#181;l to the appropriate 15 ml centrifugation tube. 
	NOTE: In our experiments blood samples usually contain 10 U/ml of heparin in order to avoid coagulation. Yet due to different sources of the sample material the nature and concentration of the anti-coagulant may differ. Its influence on the APF staining should be tested by comparing the results to results obtained from blood samples supplemented with other types or concentrations of anti-coagulant.</li>
	<li>Add 2 ml distilled water and mix the samples by using a vortex mixer set to a medium level. Incubate the samples for 60 sec, then add 5 ml PBS per sample and mix using the vortex mixer. 
	NOTE: Up to about 5 samples can be prepared in parallel. Keep the sample order the same for the addition of water and PBS to achieve comparable incubation times.</li>
	<li>After regeneration of isotonic conditions by PBS addition (step 2.2) pellet the remaining intact cells in all samples by centrifugation for 6 min at room temperature and 450 x g.</li>
	<li>Remove the supernatant by pouring it into a table waste bin. Remove as much liquid as possible by inverting the centrifugation tube and tapping the opening onto a paper towel. Ensure that the cell pellet is as dry as possible.</li>
	<li>Repeat the hypotonic lysis procedure as described before (step 2.2). 
	NOTE: In principle this hypotonic lysis procedure (steps 2.3 to 2.5) could be repeated more than once, depending on the purity of the mixed leukocyte fraction to be obtained. After two lysis steps the share of remaining erythrocytes in the obtained mixed cell fraction is about 25%.</li>
	<li>Pellet the remaining intact cells by centrifugation for 6 min at room temperature and 450 x g. Remove the supernatant (see step 2.4). Add 500 &#181;l HBSS to the pellet and gently dissolve it until a homogenous solution is obtained. Transfer each sample to the accordingly labeled 1.5 ml sample tube.</li>
	<li>Depending on the experiment directly analyze obtained samples (e.g., via flow cytometry)25, stain with fluorescence-labeled antibodies (for identification of single cell types)32, incubate with cell stimuli (for cell activation experiments)33 or stored on ice and use later. Depending on the aim of the study, immediately perform subsequent experiments on the mixed leukocyte fraction since granulocytes have a fairly short half-life of about 20 hr. 
	NOTE: This also holds for the peroxidase activity staining described below.</li>
</ol>
3. Halogenating Peroxidase Activity Staining
NOTE: HOCl- and HOBr-production by the blood-derived heme peroxidases MPO and EPO is quantified by using APF, which is oxidized to fluorescein by the named hypohalous acids. Therefore if cell labeling with fluorescence-labeled antibodies is performed in combination with APF staining, avoid fluorophores that interfere with the emission signal of fluorescein.
<ol>
	<li>Perform APF staining at 4 &#176;C on ice.</li>
	<li>For preparing a working solution of APF thaw an appropriate aliquot of the dye (e.g., 10 &#181;l, 11.81 mM) and dilute it with HBSS to exactly 1 mM (e.g., add 108.1 &#181;l buffer to the 10 &#181;l).
	<ol>
		<li>For each sample (500 &#181;l cell solution) prepare 5 &#181;l of the described APF working solution. Accordingly, use one 10 &#181;l aliquot of APF (11.81 mM), which yields 118.1 &#181;l APF working solution (1 mM) to prepare about 20 samples. Due to loss during pipetting prepare about 10% more APF working solution than calculated.</li>
	</ol>
	</li>
	<li>In order to check the peroxidase specificity of the APF staining, prepare control samples with the heme peroxidase inhibitor 4-aminobenzoic acid hydrazide (4-ABAH)35.
	<ol>
		<li>Prepare a 1 M stock solution of 4-ABAH (151.17 g/mol) in dimethyl sulfoxide (DMSO) by diluting 75.6 mg 4-ABAH in 500 &#181;l solvent. Further dilute 4-ABAH 1/10 in HBSS.</li>
	</ol>
	</li>
	<li>For the preparation of a H2O2 working solution label two 1.5 ml centrifugation tubes with &#34;70 mM H2O2&#34; and &#34;7 mM H2O2&#34;, respectively.
	<ol>
		<li>In the tube labelled &#34;70 mM H2O2&#34;, freshly dilute 10 &#181;l of a 30% stock solution (8.8 mM) of H2O2 using 990 &#181;l distilled water to obtain a concentration of about 88 mM. Store on ice and use within 4 hr. 
		NOTE: H2O2 is typically delivered as a 30% stock solution by the suppliers. When stored at 4 &#176;C, it is stable for about 3 years. Perform dilutions of H2O2 in distilled water to avoid decomposition. H2O2 dilutions could also be prepared in 0.1 M NaOH solution.</li>
		<li>For the determination of the exact H2O2 concentration prepare a further 1 to 10 dilution from the first H2O2 stock solution (about 88 mM) by adding 100 &#181;l from this solution to 900 &#181;l distilled water in the 1.5 ml tube labeled &#34;7 mM H2O2&#34;.</li>
		<li>Record a spectrum in the near UV range (e.g., 200-300 nm) by using an UV-Vis photospectrometer and use the absorbance at 240 nm (&#949;240 = 34 M-1 cm-1) to determine the actual H2O2 concentration in the second H2O2 stock solution34. Ensure that the reference cuvette contains only distilled water. For the measurement use quartz cuvettes as a spectrum in the UV range is recorded.</li>
		<li>From this result, calculate the exact concentrations of both H2O2 stock solutions. Adjust the concentration of the first stock solution in the &#34;70 mM H2O2&#34; sample tube to exactly 70 mM by adding an appropriate amount of distilled water. Store the obtained working solution on ice and use within one hour.</li>
	</ol>
	</li>
	<li>Add 5 &#181;l of the 4-ABAH working solution (100 mM) to the appropriate samples for a final concentration of 1 mM. Incubate the samples for 15 min at 37 &#176;C in the incubator before APF addition.</li>
	<li>Add 5 &#181;l APF working solution (1 mM) to each 500 &#181;l sample to obtain a final dye concentration of 10 &#181;M. Mix the samples gently (e.g., by using the vortex mixer on a medium setting) and incubate for 30 min at 37 &#176;C.</li>
	<li>Add 5 &#181;l H2O2 working solution (70 mM) to each 500 &#181;l sample for a final concentration of 700 &#181;M. Gently mix the samples and incubate them for 60 min at 37 &#176;C. Perform appropriate control measurements in parallel. 
	NOTE: Control measurements showed that 700 &#181;M H2O2 are not toxic for the cells. Nor were any significant cellular responses observed. 
	NOTE: The APF staining may also be performed by omitting the addition of H2O2, depending on the scientific question. In the absence of hydrogen peroxide only the basal chlorinating MPO and EPO activity is determined while the addition of H2O2 allows the detection of the maximal HOCl and HOBr production by the cells. The latter means a significantly higher fluorescent signal.</li>
	<li>For pelleting the cells, centrifuge the samples for 10 min at room temperature and 400 x g. Thoroughly remove the supernatant without destroying the pellet and add 250 &#181;l HBSS to resuspend the cells. 
	NOTE: The samples are now ready for analysis via flow cytometry25.</li>
	<li>Store the samples in the dark until analysis to avoid bleaching. After excitation at 480-490 nm detect the emission of fluorescein at about 525 nm37.</li>
</ol>

The authors have nothing to disclose.

As neutrophils are the most abundant leukocytes in human blood the isolation of peroxidase-positive cells often only focuses on these cells and includes a separation of neutrophils from other leukocytes by density gradient centrifugation38. Yet as neutrophils are much less abundant in murine blood samples39 for the latter more complicated methods have to be used40. Moreover both methods also lead to the removal of peroxidase-positive monocytes from the samples and, due to the need of larg...

Polymorphonuclear leukocytes (PMNs, also called granulocytes) and monocytes represent important cellular components of the innate immune system in the blood1,2. They contribute to the primary defense against pathogens as well as to the activation of the acquired immune system and the initiation of a systemic inflammatory response2-4. Yet especially neutrophils, the most abundant type of granulocytes, and monocytes also significantly contribute to the regulation and termination of acute inflammatory events5. Therefore these cells may also play an important role in chronic inflammatory diseases like rheumatoid arthritis6,7. In fact, asthma, a chronic inflammatory airway disease, is characterized by an impaired apoptosis of eosinophils, the second most granulocyte type in the blood8. Yet the apoptosis of granulocytes and their quick removal by macrophages are two essential steps during the cellular termination of inflammation9-11.
In the named immune cells two closely related enzymes, namely myeloperoxidase (MPO, neutrophils and monocytes) and eosinophil peroxidase (EPO, eosinophils) can be found12,13. These heme peroxidases are classically related to the humoral immune response as they two-electronically oxidize (pseudo-)halides to the corresponding hypo(pseudo)halous acids which are known for their bactericidal properties14-16. Under physiological conditions MPO mainly forms hypochlorous acid (HOCl) and hypothiocyanite (-OSCN) while the latter and hypobromous acid (HOBr) are formed by EPO17-19. New results suggest that this (pseudo-)halogenating enzyme activity may also contribute to the regulation of inflammatory responses and to the termination of immune reactions20,21. In fact, the HOCl production by MPO and derived products were shown to suppress T cell-based adaptive immune responses22-24.
In order to gain more insights into the immunological role of leukocytes from the innate immune system at chronic inflammatory diseases and to determine the contribution of MPO and EPO to this physiological function we developed a method to quickly enrich leukocytes from small blood samples for a subsequent specific determination of the halogenating peroxidase activity in these cells. For erythrocyte depletion we have chosen a standardized method including two-subsequent hypotonic lysis steps with distilled water, which leads to a quick leukocyte enrichment at low material costs. For the subsequent determination of the halogenating MPO and EPO activity the HOCl- and HOBr-specific dye aminophenyl fluorescein (APF) was used25-27. In contrast to the application of unspecific peroxidase staining methods28,29, this approach allows the selective detection of the halogenating peroxidase activity, which is often impaired at severe inflammation30,31.

<table><tbody><tr><td>&lt;em&gt;Materials/Equipment&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>15 ml centrifugation tubes</td><td>VWR/Corning</td><td>734-0451</td><td>-</td></tr><tr><td>1.5 ml sample tubes</td><td>VWR/Eppendorf</td><td>211-2130DE</td><td>-</td></tr><tr><td>Pipettes for volumes up to 5 ml</td><td>Eppendorf</td><td>&lt;em&gt;e.g.&lt;/em&gt;, 3120000070</td><td>We are using Eppendorf Resarch plus pipettes with adjustable volumes in the range 1-10 &amp;micro;l, 10-100 &amp;micro;l, 100-1,000 &amp;micro;l an 500-5,000 &amp;micro;l</td></tr><tr><td>laminar flow bench</td><td>Thermo Electron Corperation</td><td>HeraSafe</td><td>-</td></tr><tr><td>Vortex mixer</td><td>Bender &amp;amp; Hobein AG</td><td>Vortex Genie 2</td><td>-</td></tr><tr><td>Tabletop centrifuge</td><td>Kendro Laboratory Products</td><td>Laborfuge 400R</td><td>The centrifuge should be able to be used at 450 x g</td></tr><tr><td>Small centrifuge</td><td>Eppendorf</td><td>5415D</td><td>The centrifuge should be able to be used at 400 x g</td></tr><tr><td>Incubator</td><td>Heraeus</td><td>cytoperm 2</td><td>Settings: 37 &amp;deg;C, 95% humidity, 5% CO&lt;sub&gt;2&lt;/sub&gt; content</td></tr><tr><td>UV-Vis spectrophotometer</td><td>Varian</td><td>Cary 50 bio</td><td>A spectrum between 200 and 300 nm has to be recorded. Thus quartz cuvettes have to be applied</td></tr><tr><td>Flow cytometer</td><td>Becton, Dickinson</td><td>BD Facs Calibur</td><td>Any flow cytometer can be used which is equiped with a laser suitable for the excitation of fluorescein (&lt;em&gt;e.g.&lt;/em&gt;, 488 nm argon laser)</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;em&gt;Chemicals&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>amresco</td><td>K812</td><td>sterile solution, ready to use</td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Sigma-Aldrich</td><td>P4417</td><td>tablets for solving in 200 ml millipore water</td></tr><tr><td>Hanks balanced salt solution (HBSS) with Ca&lt;sup&gt;2+&lt;/sup&gt;</td><td>Sigma-Aldrich</td><td>H1387</td><td>970 mg/100 ml, carefully check and adjust the pH value to 7.4</td></tr><tr><td>Hydrochloric acid</td><td>Merck Millipore</td><td>1.09057.1000</td><td>1 M solution</td></tr><tr><td>Sodium hydroxide</td><td>Riedel-deHa&amp;euml;n</td><td>30620</td><td>Solid pellets. For a 1 M solution solve 4 g/100 ml Millipore water</td></tr><tr><td>Aminophenyl fluorescein</td><td>Cayman</td><td>10157</td><td>5 mg/ml solution (11.81 mM) in methyl acetate, aliquots (&lt;em&gt;e.g.&lt;/em&gt; 100 &amp;micro;l) should be prepared and stored at -20 &amp;deg;C</td></tr><tr><td>Hydrogen peroxide</td><td>Sigma-Aldrich</td><td>H1009</td><td>This 30% stock solution corresponds to a concentration of about 8.8 M. Further dilutions have to be freshly prepared in distilled water immediately prior to use and quantified by absorbance measurements</td></tr><tr><td>4-aminobenzoic acid hydrazide (4-ABAH)</td><td>Sigma-Aldrich</td><td>A41909</td><td>A first stock solution of 1 M should be prepared in DMSO a second one of 100 mM by 1:10 dilution in HBSS</td></tr><tr><td>DMSO</td><td>VWR chemicals</td><td>23500.26</td><td>-</td></tr></tbody></table>

fast and specific assessment of the halogenating peroxidase activity in leukocyte-enriched blood samples

In this paper a protocol for the quick and standardized enrichment of leukocytes from small whole blood samples is described. This procedure is based on the hypotonic lysis of erythrocytes and can be applied to human samples as well as to blood of non-human origin. The small initial sample volume of about 50 to 100 &#181;l makes this method applicable to recurrent blood sampling from small laboratory animals. Moreover, leukocyte enrichment is achieved within minutes and with low material efforts regarding chemicals and instrumentation, making this method applicable in multiple laboratory environments.
Standardized purification of leukocytes is combined with a highly selective staining method to evaluate halogenating peroxidase activity of the heme peroxidases, myeloperoxidase (MPO) and eosinophil peroxidase (EPO), i.e., the formation of hypochlorous and hypobromous acid (HOCl and HOBr). While MPO is strongly expressed in neutrophils, the most abundant immune cell type in human blood as well as in monocytes, the related enzyme EPO is exclusively expressed in eosinophils. The halogenating activity of these enzymes is addressed by using the almost HOCl- and HOBr-specific dye aminophenyl fluorescein (APF) and the primary peroxidase substrate hydrogen peroxide. Upon subsequent flow cytometry analysis all peroxidase-positive cells (neutrophils, monocytes, eosinophils) are distinguishable and their halogenating peroxidase activity can be quantified. Since APF staining may be combined with the application of cell surface markers, this protocol can be extended to specifically address leukocyte sub-fractions. The method is applicable to detect HOCl and HOBr production both in human and in rodent leukocytes.
Given the widely and diversely discussed immunological role of these enzymatic products in chronic inflammatory diseases, this protocol may contribute to a better understanding of the immunological relevance of leukocyte-derived heme peroxidases.

As reported previously the method described above turned out to be applicable both to human and to non-human material32. Moreover as shown for mice with asthmatic symptoms the APF staining may be a suitable tool to detect differences in the systemic pro-inflammatory status. Therefore in a subsequent study we used this protocol to repeatedly evaluate the halogenating activity of MPO (and EPO) in female Dark Agouti rats with pristane-induced arthritis (PIA). A representa...

Watch this Scientific Journal Video about Fast and Specific Assessment of the Halogenating Peroxidase Activity in Leukocyte-enriched Blood Samples at JoVE.com

Fast and Specific Assessment of the Halogenating Peroxidase Activity in Leukocyte-enriched Blood Samples

This protocol describes the quick enrichment of leukocytes from small blood samples for a subsequent specific determination of the halogenating peroxidase activity within the cells. The method can be applied to human and non-human material and may contribute to the evaluation of new inflammatory markers.

In this paper a protocol for the quick and standardized enrichment of leukocytes from small whole blood samples is described. This procedure is based on ...

fast-specific-assessment-halogenating-peroxidase-activity-leukocyte

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JoVE Journal

Immunology and Infection

10.3K Views.  University of Leipzig. This protocol describes the quick enrichment of leukocytes from small blood samples for a subsequent specific determination of the halogenating peroxidase activity within the cells. The method can be applied to human and non-human material and may contribute to the evaluation of new inflammatory markers.

Video: Fast and Specific Assessment of the Halogenating Peroxidase Activity in Leukocyte-enriched Blood Samples

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.

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 ...

Cancer Research

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Mitochondrial dysfunction is known to play a significant role in a number of pathological conditions such as atherosclerosis, diabetes, septic shock, and neurodegenerative diseases but assessing changes in bioenergetic function in patients is challenging.&#160;Although diseases such as diabetes or atherosclerosis present clinically with specific organ impairment, the systemic components of the pathology, such as hyperglycemia or inflammation, can alter bioenergetic function in circulating leukocytes or platelets. This concept has been recognized for some time but its widespread application has been constrained by the large number of primary cells needed for bioenergetic analysis.&#160;This technical limitation has been overcome by combining the specificity of the magnetic bead isolation techniques, cell adhesion techniques, which allow cells to be attached without activation to microplates, and the sensitivity of new technologies designed for high throughput microplate respirometry.&#160;An example of this equipment is the extracellular flux analyzer. Such instrumentation typically uses oxygen and pH sensitive probes to measure rates of change in these parameters in adherent cells, which can then be related to metabolism. Here we detail the methods for the isolation and plating of monocytes, lymphocytes, neutrophils and platelets, without activation, from human blood and the analysis of mitochondrial bioenergetic function in these cells. In addition, we demonstrate how the oxidative burst in monocytes and neutrophils can also be measured in the same samples. Since these methods use only 8-20 ml human blood they have potential for monitoring reactive oxygen species generation and bioenergetics in a clinical setting.

Blood leukocytes and platelets can be used as a marker of overall bioenergetic health of an individual and so have the potential to monitor pathological processes and the impact of treatments. Here&#160;we describe a method to isolate and measure mitochondrial function and the oxidative burst in these cells.

Mitochondrial dysfunction is known to play a significant role in a number of pathological conditions such as atherosclerosis, diabetes, septic shock, and ...

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

Cell-free Biochemical Fluorometric Enzymatic Assay for High-throughput Measurement of Lipid Peroxidation in High Density Lipoprotein

Low high-density lipoprotein cholesterol (HDL-C) levels are one of the most powerful independent negative predictors of atherosclerotic cardiovascular disease (CVD). The structure and function of HDL rather than HDL-C may more accurately predict atherosclerosis. Several HDL protein and lipid compositional changes that impair HDL function occur in inflammatory states such as atherosclerosis. HDL function is usually determined by cell based assays such as cholesterol efflux assay but these assays have numerous drawbacks lack of standardization. Cell-free assays may give more robust measures of HDL function compared to cell-based assays. HDL oxidation impairs HDL function. HDL has a major role in lipid peroxide transport and high amount of lipid peroxides is related to abnormal HDL function. Lipid-probe interactions should be considered when interpreting the results of non-enzymatic fluorescence assays for measuring the lipid oxidative state. This motivated us to develop a cell-free biochemical enzymatic method to assess HDL lipid peroxide content (HDLox) that contributes to HDL dysfunction. This method is based on the enzyme horseradish peroxidase (HRP) and the fluorochrome Amplex Red that can quantify (without cholesterol oxidase) the lipid peroxide content per mg of HDL-C. Here a protocol is describedfor determination of HDL-lipid peroxidation using the fluorochrome reagent. Assay variability can be reduced by strict standardization of experimental conditions. Higher HDLox values are associated with reduced HDL antioxidant function. The readout of this assay is associated with readouts of validated cell-based assays, surrogate measures of cardiovascular disease, systemic inflammation, immune dysfunction, and associated cardiovascular and metabolic risk phenotypes. This technical approach is a robust method to assess HDL function in human disease where systemic inflammation, oxidative stress and oxidized lipids have a key role (such as atherosclerosis).

We describe here a fluorometric cell-free biochemical assay for determination of HDL-lipid peroxidation. This rapid and reproducible assay can be used to determine HDL function in large scale studies and can contribute to our understanding of HDL function in human disease.

Low high-density lipoprotein cholesterol (HDL-C) levels are one of the most powerful independent negative predictors of atherosclerotic cardiovascular ...

GSH/GSSG Measurement &#8211; A Refined OPA Assay for Antioxidant Research

Rapid Quantification of Oxidized and Reduced Forms of Glutathione Using Ortho -phthalaldehyde in Cultured Mammalian Cells In Vitro

Glutathione has long been considered a key biomarker for determining the antioxidant response of the cell. Hence, it is a primary marker for reactive oxygen species studies. The method utilizes Ortho-phthalaldehyde (OPA) to quantify the cellular concentration of glutathione(s). OPA conjugates with reduced glutathione (GSH) via sulfhydryl binding to subsequently form an isoindole, resulting in a highly fluorescent conjugate. To attain an accurate result of both oxidized glutathione (GSSG) and GSH, a combination of masking agents and reducing agents, which have been implemented in this protocol, are required. Treatments may also impact cellular viability. Hence, normalization via protein assay is presented in this multiparametric assay. The assay demonstrates a pseudo-linear detection range of 0.234 - 30&#181;M (R2=0.9932&#177;0.007 (N=12)) specific to GSH. The proposed assay also allows for the determination of oxidized glutathione with the addition of the masking agent N-ethylmaleimide to bind reduced glutathione, and the reducing agent tris(2-carboxyethyl) phosphine is introduced to cleave the disulfide bond in GSSG to produce two molecules of GSH. The assay is used in combination with a validated bicinchoninic acid assay for protein quantification and an adenylate kinase assay for cytotoxicity assessment.

Author Spotlight: Innovating Thiol Quantification and Biomarker Detection for Oxidative Stress Research 

Quantification of both oxidized and reduced forms of glutathione (GSSG and GSH, respectively) has been achieved through the use of Ortho-phthalaldehyde (OPA). OPA becomes highly fluorescent once conjugated to GSH but is unable to conjugate GSSG until reduced. Here, we describe a multiparametric assay to quantify both using protein quantification for normalization.

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Detecting, Visualizing and Quantitating the Generation of Reactive Oxygen Species in an Amoeba Model System

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or eliminated either catalytically or spontaneously. Due to the short life spans of most ROS and the diversity of their sources and subcellular localizations, a complete picture can be obtained only by careful measurements using a combination of protocols. Here, we present a set of three different protocols using OxyBurst Green (OBG)-coated beads, or dihydroethidium (DHE) and Amplex UltraRed (AUR), to monitor qualitatively and quantitatively various ROS in professional phagocytes such as Dictyostelium. We optimised the beads coating procedures and used OBG-coated beads and live microscopy to dynamically visualize intraphagosomal ROS generation at the single cell level. We identified lipopolysaccharide (LPS) from E. coli as a potent stimulator for ROS generation in Dictyostelium. In addition, we developed real time, medium-throughput assays using DHE and AUR to quantitatively measure intracellular superoxide and extracellular H2O2 production, respectively.

We adapted a set of protocols for the measurement of reactive oxygen species (ROS) that can be applied in various amoeba and mammalian cellular models for qualitative and quantitative studies.

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or ...

Biology

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

Detection of Superoxide Anion in Platelets Using Fluorescence Imaging and EPR

Alternative Methods for the Detection of Superoxide Anion Generation in Platelets

Reactive oxygen species (ROS) are highly unstable oxygen-containing molecules. Their chemical instability makes them extremely reactive and gives them the ability to react with important biological molecules such as proteins, nucleic acids, and lipids. Superoxide anions are important ROS generated by the reduction of molecular oxygen reduction (i.e., acquisition of one electron). Despite their initial implication exclusively in aging, degenerative, and pathogenic processes, their participation in important physiological responses has recently become apparent. In the vascular system, superoxide anions have been shown to modulate the differentiation and function of vascular smooth muscle cells, the proliferation and migration of vascular endothelial cells in angiogenesis, the immune response, and the activation of platelets in hemostasis. The role of superoxide anions is particularly important in the dysregulation of platelets and the cardiovascular complications associated with a plethora of conditions, including cancer, infection, inflammation, diabetes, and obesity. It has, therefore, become extremely relevant in cardiovascular research to be able to effectively measure the generation of superoxide anions by&#160;human platelets,&#160;understand the redox-dependent mechanisms regulating the balance between hemostasis and thrombosis and, eventually,&#160;identify novel pharmacological tools for the modulation of platelet responses leading to thrombosis and cardiovascular complications. This study presents three experimental protocols successfully adopted for the detection of superoxide anions in platelets and the study of the redox-dependent mechanisms regulating hemostasis and thrombosis: 1) dihydroethidium (DHE)-based superoxide anion detection by flow cytometry; 2) DHE-based superoxide anion visualization and analysis by single platelet imaging; and 3) spin probe-based quantification of superoxide anion output in platelets by electron paramagnetic resonance (EPR).

Author Spotlight: Innovative Techniques for ROS Detection and Implications for Platelet Research

The generation of superoxide anion is essential for the stimulation of platelets and, if dysregulated, critical for thrombotic diseases. Here, we present three protocols for the selective detection of superoxide anions and the study of redox-dependent platelet regulation.

Reactive oxygen species (ROS) are highly unstable oxygen-containing molecules. Their chemical instability makes them extremely reactive and gives them the ...

Evaluation of a Point-of-Care Testing Analyzer for Measuring Peripheral Blood Leukocytes

White blood cell (WBC) is an important indicator of inflammation in the body, and it can help distinguish between bacterial and viral infections. At present, most primary medical institutions in China have a poor percentage of adoption of blood-testing technology, and a hematology detection system with a high price to performance ratio and easy operation is urgently needed in primary healthcare centers. This paper introduces the principle and operation procedures of a point-of-care testing (POCT) card-based leukocyte analyzer (evaluated system), which was used to detect WBC indexes such as neutrophils, lymphocytes, and intermediate group cells (including eosinophils, basophils, and monocytes) in whole blood. The results from the evaluated system were compared to those from two commercial automatic hematology analyzers (reference system). The correlation and consistency between the evaluated system and the commercial reference systems were analyzed. The results showed that WBC count and number of granulocytes detected by the evaluated and reference systems showed a strong positive correlation (rs = 0.972 and 0.973, respectively), while the number of lymphocytes showed a relatively low correlation (rs = 0.851). A Bland-Altman plot showed that the major difference between the values detected by the evaluated system and the reference systems is within 95% limits of agreement (LoA), indicating that the two systems are in good agreement. In conclusion, the evaluated system has an excellent correlation, robust consistency, and a reliable comparison with the results of the widely used automatic hematology analyzers. It is ideal for WBC detection in primary medical institutions where a full-automatic five-category hematology analyzer is unavailable, especially during the COVID-19 normalized prevention and control period.

A protocol to measure peripheral blood leukocytes using a POCT card-based leukocyte analyzer is presented here. Same blood samples were tested by two automated hematology analyzers to evaluate the consistency and accuracy of the results. The results showed that the evaluated analyzer had a good correlation with the reference system.

White blood cell (WBC) is an important indicator of inflammation in the body, and it can help distinguish between bacterial and viral infections. At ...

Medicine

Analysis of Semen for Peroxidase-Positive Leukocyte Cells

The Examination of Peroxidase-Positive Leukocytes in Semen

Leukocytospermia can lead to decreased spermatozoa motility, increased spermatozoa morphological abnormalities, elevated spermatozoa DNA fragmentation index, impairment of the spermatozoa acrosome function, and even affected embryonic development. It is a common andrological disease in clinical practice and one of the important causes of male infertility. When determining whether male reproductive tract inflammation exists, andrologists often choose to examine round cells or seminal plasma elastase in the semen as a clinical diagnostic basis. However, the examination of round cells is easily influenced by sloughed spermatogenic cells and reproductive tract epithelial cells, which do not contribute to reducing the indiscriminate and unnecessary use of antibiotics. At the same time, the detection process of elastase is relatively complicated, time-consuming, and slow in reporting results, which is not beneficial for early diagnosis and treatment of diseases such as male genital tract infections (MGTIs). We have innovatively applied the examination of peroxidase-positive leukocytes in semen assisted by a computer-assisted semen analysis (CASA) system as a diagnostic criterion for leukocytospermia, successfully solving these problems. This examination only requires the addition of the operating fluid consisting of four reagents into the specimen, and the total reaction time at room temperature can be controlled within 20-30 min. With the subsequent smear and microscopic examination, the concentration of peroxidase-positive leukocytes in semen can be obtained within a total of 60 min, which can be used to diagnose whether the inflammation of the male reproductive tract existed.

Author Spotlight: A Simple and Cost-Effective Method for Leukocyte Analysis in Semen 

This paper presents an economical and efficient protocol for examining peroxidase-positive leukocytes in semen. With the assistance of a computer-assisted semen analysis (CASA) system, the concentration of peroxidase-positive leukocytes in semen can be obtained within a total of 60 min, effectively improving the efficiency of andrology laboratory and andrologists.

Leukocytospermia can lead to decreased spermatozoa motility, increased spermatozoa morphological abnormalities, elevated spermatozoa DNA fragmentation ...