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 larger blood volumes (e.g., 400 &#181;l41), are not applicable to micro-samples obtained during recurrent blood sampling from small laboratory animals38,40. The purification of murine neutrophils from peritoneal exudates also need the sacrifice of the animals and, moreover, does not yield blood-derived granulocytes38,40. Recently developed methods to obtain highly purified granulocyte fractions from murine blood (e.g., the application of antibodies) are often expensive, complicated and time consuming40,42,43 and again only focus on the purification of one peroxidase-positive leukocyte type.
Thus the method presented here has a couple of advantages over other methods applied for the purification of peroxidase-positive leukocytes. As it is based on small blood samples the obtained cells represent the situation in the circulation. Due to the small initial sample volume needed for the protocol the method is applicable to both human and non-human blood, despite the species-specific differences in the abundance of neutrophils. In fact, we were even able to apply the described method to initial blood volumes as small as 50 &#181;l (data not shown). The small blood sample needed for the method also makes it suitable for repeated blood withdrawal from small laboratory animals or for special medical applications (e.g., newborn diagnosis). As the leukocyte enrichment is based on the depletion of erythrocytes all peroxidase-positive cells (neutrophils, eosinophils, monocytes) are included in the obtained mixed leukocyte fraction and can be separately analyzed by using flow cytometry. The method is fast, reliable and has low requirements regarding chemicals and instrumentation, making the protocol suitable for multiple scientific environments.
As neutrophils are easily activated one critical step during the described method is the precise adjustment of the pH value of the buffer solutions used (PBS and HBSS, see steps 1.2 and 1.3 of the protocol section). Furthermore the incubation time of the blood cells with distilled water should be no longer than 60 sec before restoring normosmotic conditions (see step 2.2 of the protocol section. The (almost) completely removal of solvent from the cell pellet (step 2.4) before the addition of water for the second hypotonic lysis is another critical step as buffer residues will diminish the efficiency of the hypotonic lysis procedure. Another critical step refers to the application of H2O2 during the APF staining. Although the applied amount should not be cytotoxic, cell vitality should be checked. During our studies we typically apply the dye 5,5&#39;,6,6&#39;-tetrachloro-1,1&#39;,3,3&#39;-tetraethylbenzimi-dazolylcarbocyanine iodide (JC-1) for the detection of early apoptotic events.
Moreover, there are a couple of limitations in the described simplified leukocyte enrichment method. While the technique can be applied to blood samples from several species (man, rat and mouse have been tried), the leukocyte amount obtained after the lysis of erythrocytes depends on their initial abundance in the blood. The latter certainly varies between species39 and is also dependent on the inflammatory state of the probed individual. Moreover, while the lysis procedure can be done with up to one hundred samples in parallel the intermediate centrifugation steps pose a limitation as, depending on the centrifuge used, only up 30 samples can be processed in parallel. Furthermore, while APF readily detects HOCl and HOBr in the presence of SCN-, the halogenating activity of MPO and EPO also affects to the latter pseudo-halide. Thereby hypothiocyanite (-OSCN) is formed which is unable to oxidize APF. Another limitation comes from the properties of the APF dye used for the determination of HOCl- and HOBr-production by MPO and EPO. As the dye is oxidized to fluorescein, the staining cannot be combined with fluorescent antibodies with an emission spectrum in the same range. Of course any interferences between the APF-derived fluorescence and the signal of additional fluorescent signals have to be checked and compensated for in the flow cytometer.
Otherwise, if the detection of the halogenating peroxidase activity is not the scope of the study, after application of the erythrocyte depletion method other analytical methods may be used instead of APF. As the described hypotonic lysis procedure leads only to a partial depletion of erythrocytes and as all leukocyte are still present in the obtained mixed cell fraction, only methods which allow a single cell analysis should be applied. Methods which include the lysis of cells (e.g., Western plot analysis) will not yield reliable results. The small initial sample volume may be another obstacle for such analytical methods.
Regarding the investigation of MPO (and EPO) activity in leukocytes often only the general oxidant production by the cells is addressed instead of specifically evaluating the heme peroxidase activity44,45. Moreover often no attempts are made to distinguish between the halogenating and the peroxidase activity of MPO and EPO46,47. Methods, which specifically address the chlorinating MPO activity by quantifying HOCl-derived products like chlorotyrosine do not detect over-oxidized and/or metabolized products and, thus, often lead to an underestimation of the real HOCl-production48. Moreover this method as well as the detection of HOCl-derived 2-chloroethidium are also time-consuming, need costly analytical equipment and include lysis of cells48-50.There are many reports about new dyes for the specific determination of the chlorinating MPO activity within vital cells44,51,52. Yet to date only APF and its related compound hydroxyphenyl fluorescein (HPF) are commercially available and therefore routinely used in research25,33.
In fact, by using APF we could show that the chlorinating MPO activity cannot only be quantified by using the isolated enzyme53 but also by applying the dye to living cells26,33. Thus by combining the quick leukocyte enrichment from blood micro-samples stated above with an APF staining we developed a protocol, which is suitable to specifically and simultaneously address the halogenating activity of MPO and EPO in all peroxidase positive leukocytes, i.e., neutrophils, monocytes and eosinophils32. Moreover the method does not require cell lysis, which allows a broad variety of analytical methods, including flow cytometry and confocal fluorescent microscopy. Moreover this peroxidase activity staining can be combined with the application of cell surface markers, which allows the specific analysis of leukocyte sub-fractions, depending on the scientific task. Yet it has to be considered that the applied fluorescence-labeled antibodies do not interfere with the fluorescein-based fluorescence signal used to quantify the HOCl- and HOBr-derived APF oxidation.
In summary this method may provide a new tool to get more insights into the physiological role of the halogenating peroxidase activity of the immunological relevant blood-derived peroxidases MPO and EPO, which is still not well understood8,20,23. Moreover in an animal study on rheumatoid arthritis we could recently observe that this protocol may lead to the evaluation of the MPO-derived HOCl production as a new clinical marker at chronic inflammatory diseases (unpublished data).

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 border="0" cellpadding="0" cellspacing="0" width="738">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">materials/equipment</td>
			<td style="width:203px;"></td>
			<td style="width:120px;"></td>
			<td style="width:203px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">15 ml centrifugation tubes</td>
			<td style="width:203px;">VWR/Corning</td>
			<td style="width:120px;">734-0451</td>
			<td style="width:203px;">-</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">1.5 ml sample tubes</td>
			<td style="width:203px;">VWR/Eppendorf</td>
			<td style="width:120px;">211-2130DE</td>
			<td style="width:203px;">-</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:213px;">Pipettes for volumes up to 5 ml</td>
			<td style="width:203px;">Eppendorf</td>
			<td style="width:120px;">e.g. 3120000070</td>
			<td style="width:203px;">We are using Eppendorf Resarch plus pipettes with adjustable volumes in the range 1-10 &#181;l, 10-100&#181;l, 100-1000 &#181;l an 500-5000&#181;l</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">laminar flow bench</td>
			<td style="width:203px;">Thermo Electron Corperation</td>
			<td style="width:120px;">HeraSafe</td>
			<td style="width:203px;">-</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">Vortex mixer</td>
			<td style="width:203px;">Bender &#38; Hobein AG</td>
			<td style="width:120px;">Vortex Genie 2</td>
			<td style="width:203px;">-</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Tabletop centrifuge</td>
			<td style="width:203px;">Kendro Laboratory Products</td>
			<td style="width:120px;">Laborfuge 400R</td>
			<td style="width:203px;">The centrifuge should be able to be used at 450x g</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Small centrifuge</td>
			<td style="width:203px;">eppendorf</td>
			<td style="width:120px;">5415D</td>
			<td style="width:203px;">The centrifuge should be able to be used at 400x g</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Incubator</td>
			<td style="width:203px;">Heraeus</td>
			<td style="width:120px;">cytoperm 2</td>
			<td style="width:203px;">Settings: 37 &#176;C, 95% humidity, 5 % CO2 content</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;">UV-Vis spectrophotometer</td>
			<td style="width:203px;">Varian</td>
			<td style="width:120px;">Cary 50 bio</td>
			<td style="width:203px;">A spectrum between 200 and 300 nm has to be recorded. Thus quartz cuvettes have to be applied</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;">Flow cytometer</td>
			<td style="width:203px;">Becton, Dickinson</td>
			<td style="width:120px;">BD Facs Calibur</td>
			<td style="width:203px;">Any flow cytometer can be used which is equiped with a laser suitable for the excitation of fluorescein (e.g 488 nm argon laser)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Name</td>
			<td style="width:203px;">Company</td>
			<td style="width:120px;">Catalog Number</td>
			<td style="width:203px;">Comments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">Chemicals</td>
			<td style="width:203px;"></td>
			<td style="width:120px;"></td>
			<td style="width:203px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:213px;">Phosphate buffered saline (PBS)</td>
			<td style="width:203px;">amresco</td>
			<td style="width:120px;">K812</td>
			<td style="width:203px;">sterile solution, ready to use</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:213px;"></td>
			<td style="width:203px;">Sigma-Aldrich</td>
			<td style="width:120px;">P4417</td>
			<td style="width:203px;">tablets for solving in 200 ml millipore water</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:213px;">Hanks balanced salt solution (HBSS) with Ca(2+)</td>
			<td style="width:203px;">Sigma-Aldrich</td>
			<td style="width:120px;">H1387</td>
			<td style="width:203px;">970 mg/100 ml, carefully check and adjust the pH value to 7.4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:213px;">Hydrochloric acid</td>
			<td style="width:203px;">Merck Millipore</td>
			<td style="width:120px;">1.09057.1000</td>
			<td style="width:203px;">1M solution</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:213px;">Sodium hydroxide</td>
			<td style="width:203px;">Riedel-deHa&#235;n</td>
			<td style="width:120px;">30620</td>
			<td style="width:203px;">Solid pellets. For a 1M solution solve 4 g/100 ml Millipore water</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:213px;">Aminophenyl fluorescein</td>
			<td style="width:203px;">Cayman</td>
			<td style="width:120px;">10157</td>
			<td style="width:203px;">5 mg/ml solution (11.81 mM) in methyl acetate, aliquotes of e.g. 100 &#181;l should be prepared and stored at -20 &#176;C</td>
		</tr>
		<tr height="168">
			<td height="168" style="height:168px;width:213px;">Hydrogen peroxide</td>
			<td style="width:203px;">Sigma-Aldrich</td>
			<td style="width:120px;">H1009</td>
			<td style="width:203px;">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 height="84">
			<td height="84" style="height:84px;width:213px;">4-aminobenzoic acid hydrazide (4-ABAH)</td>
			<td style="width:203px;">Sigma-Aldrich</td>
			<td style="width:120px;">A41909</td>
			<td style="width:203px;">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 height="21">
			<td height="21" style="height:21px;width:213px;">DMSO</td>
			<td style="width:203px;">VWR chemicals</td>
			<td style="width:120px;">23500.26</td>
			<td style="width:203px;">-</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 representative example of leukocyte enrichment and staining performed during this experiment is shown in Figure 1. About 200 &#181;l whole blood was obtained from the retrobulbar venous plexus of the animal under anesthesia and heparinized. The erythrocyte depletion and a subsequent APF staining were performed by using 100 &#181;l sample volume. Afterwards the sample was analyzed by flow cytometry.
As shown by plotting cell size versus cell granularity (Figure 1A), a strong depletion of erythrocytes from the sample was achieved. Furthermore different leukocyte types were clearly discernible, which were identified by applying fluorescence-labeled cell surface markers (not shown). Briefly the erythrocyte (CD235a-positive)/cell debris region only accounted for 22% of the events while about 48% of the events were identified as CD16-positive neutrophils. Furthermore 3% of the events were each identified as CCR3-positive eosinophils and CD14-positive monocytes and about 19% of the events accounted for CD5/CD19-positive B and T lymphocytes, respectively. The intensity distribution of the APF-derived fluorescence (Figure 1B) clearly allowed the discrimination of peroxidase-negative erythrocytes and lymphocytes while monocytes and eosinophils led to a moderate APF oxidation and neutrophils were strongly peroxidase-positive under the chosen experimental conditions. These results are in line with the high abundance of MPO in the latter cells as compared to the enzyme concentration in monocytes20. The relatively weak response of the eosinophils can be explained by the fact, that no bromide was added, which may have prompted the EPO-derived formation of HOBr. Preliminary studies on isolated eosinophils surprisingly did not show a big influence of externally added bromide on the APF oxidation by these cells. Still, oxidation of APF by the eosinophils was observed which may be explained by the introduction of small amounts of Br- via the buffer solutions used (PBS and HBSS). Alternatively EPO may also produce small amounts of HOCl, at least under acidic conditions (e.g., in phagolysosomes).
<img alt="Figure 1" src="/files/ftp_upload/54484/54484fig1.jpg" /> 
Figure 1: Leukocyte enrichment and APF-derived fluorescence in a blood micro-sample from a rat. An amount of about 200 &#181;l whole blood was obtained from the retrobulbar venous plexus of a female Dark Agouti rat. After application of the described hemolysis procedure to 100 &#181;l blood and a subsequent APF staining the mixed cell fraction was analyzed by flow cytometry. As shown by the FSC-/SSC- Plot (A) the erythrocytes were strongly depleted from the sample and the different leukocyte types could easily be distinguished. The distribution of the APF-derived fluorescence intensity (B) clearly showed heme peroxidase-negative cells (erythrocytes and lymphocytes) as well as leukocytes with moderate (eosinophils and monocytes) or strong (neutrophils) halogenating peroxidase activity. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54484/54484fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

10.2K 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

Depletion of Specific Cell Populations by Complement Depletion

The purification of immune cell populations is often required in order to study their unique functions. In particular, molecular approaches such as real-time PCR and microarray analysis require the isolation of cell populations with high purity. Commonly used purification strategies include fluorescent activated cell sorting (FACS), magnetic bead separation and complement depletion. Of the three strategies, complement depletion offers the advantages of being fast, inexpensive, gentle on the cells and a high cell yield. The complement system is composed of a large number of plasma proteins that when activated initiate a proteolytic cascade culminating in the formation of a membrane-attack complex that forms a pore on a cell surface resulting in cell death1. The classical pathway is activated by IgM and IgG antibodies and was first described as a mechanism for killing bacteria. With the generation of monoclonal antibodies (mAb), the complement cascade can be used to lyse any cell population in an antigen-specific manner. Depletion of cells by the complement cascade is achieved by the addition of complement fixing antigen-specific antibodies and rabbit complement to the starting cell population. The cells are incubated for one hour at 37&deg;C and the lysed cells are subsequently removed by two rounds of washing. MAb with a high efficiency for complement fixation typically deplete 95-100% of the targeted cell population. Depending on the purification strategy for the targeted cell population, complement depletion can be used for cell purification or for the enrichment of cell populations that then can be further purified by a subsequent method.

To effectively study the function of immune cell populations their purification is often required. Complement depletion is a fast and inexpensive technique for the isolation of immune cell populations with high purity.

The purification of immune cell populations is often required in order to study their unique functions.  In particular, molecular approaches such as ...

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting (FACS)

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known phenotype.  Other bulk methods of purification include panning, complement depletion and magnetic bead separation.  However, FACS has several advantages over other available methods.  FACS is the preferred method when very high purity of the desired population is required, when the target cell population expresses a very low level of the identifying marker or when cell populations require separation based on differential marker density.  In addition, FACS is the only available purification technique to isolate cells based on internal staining or intracellular protein expression, such as a genetically modified fluorescent protein marker.  FACS allows the purification of individual cells based on size, granularity and fluorescence.  In order to purify cells of interest, they are first stained with fluorescently-tagged monoclonal antibodies (mAb), which recognize specific surface markers on the desired cell population (1).  Negative selection of unstained cells is also possible.  FACS purification requires a flow cytometer with sorting capacity and the appropriate software.  For FACS, cells in suspension are passed as a stream in droplets with each containing a single cell in front of a laser.  The fluorescence detection system detects cells of interest based on predetermined fluorescent parameters of the cells.  The instrument applies a charge to the droplet containing a cell of interest and an electrostatic deflection system facilitates collection of the charged droplets into appropriate collection tubes (2).  The success of staining and thereby sorting depends largely on the selection of the identifying markers and the choice of mAb.  Sorting parameters can be adjusted depending on the requirement of purity and yield.  Although FACS requires specialized equipment and personnel training, it is the method of choice for isolation of highly purified cell populations.

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS

For many scientific studies requiring a biological and chemical analysis of cell populations the cells must be in a high state of purity. Fluorescence activated cell sorting (FACS) is a superior method in which to obtain pure cell populations.

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known ...

Introducing Shear Stress in the Study of Bacterial Adhesion

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the initial and determining step of the pathogenesis. Although experimentally adhesion is mostly studied in static conditions adhesion actually takes place in the presence of flowing liquid. First encounters between bacteria and their host often occur at the mucosal level, mouth, lung, gut, eye, etc. where mucus flows along the surface of epithelial cells. Later in infection, pathogens occasionally access the blood circulation causing life-threatening illnesses such as septicemia, sepsis and meningitis. A defining feature of these infections is the ability of these pathogens to interact with endothelial cells in presence of circulating blood. The presence of flowing liquid, mucus or blood for instance, determines adhesion because it generates a mechanical force on the pathogen. To characterize the effect of flowing liquid one usually refers to the notion of shear stress, which is the tangential force exerted per unit area by a fluid moving near a stationary wall, expressed in dynes/cm2. Intensities of shear stress vary widely according to the different vessels type, size, organ, location etc. (0-100 dynes/cm2). Circulation in capillaries can reach very low shear stress values and even temporarily stop during periods ranging between a few seconds to several minutes 1. On the other end of the spectrum shear stress in arterioles can reach 100 dynes/cm2 2. The impact of shear stress on different biological processes has been clearly demonstrated as for instance during the interaction of leukocytes with the endothelium 3. To take into account this mechanical parameter in the process of bacterial adhesion we took advantage of an experimental procedure based on the use of a disposable flow chamber 4. Host cells are grown in the flow chamber and fluorescent bacteria are introduced in the flow controlled by a syringe pump. We initially focused our investigations on the bacterial pathogen Neisseria meningitidis, a Gram-negative bacterium responsible for septicemia and meningitis. The procedure described here allowed us to study the impact of shear stress on the ability of the bacteria to: adhere to cells 1, to proliferate on the cell surface 5and to detach to colonize new sites 6 (Figure 1). Complementary technical information can be found in reference 7. Shear stress values presented here were chosen based on our previous experience1 and to represent values found in the literature. The protocol should be applicable to a wide range of pathogens with specific adjustments depending on the objectives of the study.

During the infection process, a key step is the adhesion of pathogens with host cells. In most instances this adhesion step occurs in the presence of mechanical stress generated by flowing liquid. We describe a technique that introduces shear stress as an important parameter in the study of bacterial adhesion.

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the ...

Determining the Phagocytic Activity of Clinical Antibody Samples

Antibody-driven phagocytosis is induced via the engagement of Fc receptors on professional phagocytes, and can contribute to both clearance as well as pathology of disease. While the properties of the variable domains of antibodies have long been considered critical to in vivo function, the ability of antibodies to recruit innate immune cells via their Fc domains has become increasingly appreciated as a major factor in their efficacy, both in the setting of recombinant monoclonal antibody therapy, as well as in the course of natural infection or vaccination1-3.
Importantly, despite its nomenclature as a constant domain, the antibody Fc domain does not have constant function, and is strongly modulated by IgG subclass (IgG1-4) and glycosylation at Asparagine 2974-6. Thus, this method to study functional differences of antigen-specific antibodies in clinical samples will facilitate correlation of the phagocytic potential of antibodies to disease state, susceptibility to infection, progression, or clinical outcome.
Furthermore, this effector function is particularly important in light of the documented ability of antibodies to enhance infection by providing pathogens access into host cells via Fc receptor-driven phagocytosis7. Additionally, there is some evidence that phagocytic uptake of immune complexes can impact the Th1/Th2 polarization of the immune response8.
Here, we describe an assay designed to detect differences in antibody-induced phagocytosis, which may be caused by differential IgG subclass, glycan structure at Asn297, as well as the ability to form immune complexes of antigen-specific antibodies in a high-throughput fashion. To this end, 1 &mu;m fluorescent beads are coated with antigen, then incubated with clinical antibody samples, generating fluorescent antigen specific immune complexes. These antibody-opsonized beads are then incubated with a monocytic cell line expressing multiple Fc&gamma;Rs, including both inhibitory and activating. Assay output can include phagocytic activity, cytokine secretion, and patterns of Fc&gamma;Rs usage, and are determined in a standardized manner, making this a highly useful system for parsing differences in this antibody-dependent effector function in both infection and vaccine-mediated protection9.

We present a high-throughput flow cytometric assay to determine the phagocytic activity of antigen-specific antibodies from clinical samples, utilizing fluorescent antigen-coated beads and a monocytic cell line expressing multiple Fc receptors&#x2014;providing receptor usage and phagocytic activity determinations in a standardized and reproducible fashion for any antigen of interest.

Antibody-driven phagocytosis is induced via the engagement of Fc receptors on professional phagocytes, and can contribute to both clearance as well as ...

Quantitative Imaging of Lineage-specific Toll-like Receptor-mediated Signaling in Monocytes and Dendritic Cells from Small Samples of Human Blood

Individual variations in immune status determine responses to infection and contribute to disease severity and outcome. Aging is associated with an increased susceptibility to viral and bacterial infections and decreased responsiveness to vaccines with a well-documented decline in humoral as well as cell-mediated immune responses1,2. We have recently assessed the effects of aging on Toll-like receptors (TLRs), key components of the innate immune system that detect microbial infection and trigger antimicrobial host defense responses3. In a large cohort of healthy human donors, we showed that peripheral blood monocytes from the elderly have decreased expression and function of certain TLRs4 and similar reduced TLR levels and signaling responses in dendritic cells (DCs), antigen-presenting cells that are pivotal in the linkage between innate and adaptive immunity5. We have shown dysregulation of TLR3 in macrophages and lower production of IFN by DCs from elderly donors in response to infection with West Nile virus6,7. 
Paramount to our understanding of immunosenescence and to therapeutic intervention is a detailed understanding of specific cell types responding and the mechanism(s) of signal transduction. Traditional studies of immune responses through imaging of primary cells and surveying cell markers by FACS or immunoblot have advanced our understanding significantly, however, these studies are generally limited technically by the small sample volume available from patients and the inability to conduct complex laboratory techniques on multiple human samples. ImageStream combines quantitative flow cytometry with simultaneous high-resolution digital imaging and thus facilitates investigation in multiple cell populations contemporaneously for an efficient capture of patient susceptibility. Here we demonstrate the use of ImageStream in DCs to assess TLR7/8 activation-mediated increases in phosphorylation and nuclear translocation of a key transcription factor, NF-&kappa;B, which initiates transcription of numerous genes that are critical for immune responses8. Using this technology, we have also recently demonstrated a previously unrecognized alteration of TLR5 signaling and the NF-&kappa;B pathway in monocytes from older donors that may contribute to altered immune responsiveness in aging9.

We describe use of ImageStream technology (<a href="http://www.amnis.com/" target="_blank">www.amnis.com</a>), which combines quantitative flow cytometry with simultaneous high-resolution digital imaging, to quantify cellular mechanisms of primary immune cells from well-defined patient cohorts. Our studies provide a blueprint for translational investigations to quantify lineage specific cellular responses in small samples from subject cohorts.

Individual variations in immune status determine responses to infection and contribute to disease severity and outcome.  Aging is associated with an ...

Generation of Human Alloantigen-specific T Cells from Peripheral Blood

The study of human T lymphocyte biology often involves examination of responses to activating ligands. T cells recognize and respond to processed peptide antigens presented by MHC (human ortholog HLA) molecules through the T cell receptor (TCR) in a highly sensitive and specific manner. While the primary function of T cells is to mediate protective immune responses to foreign antigens presented by self-MHC, T cells respond robustly to antigenic differences in allogeneic tissues. T cell responses to alloantigens can be described as either direct or indirect alloreactivity. In alloreactivity, the T cell responds through highly specific recognition of both the presented peptide and the MHC molecule. The robust oligoclonal response of T cells to allogeneic stimulation reflects the large number of potentially stimulatory alloantigens present in allogeneic tissues. While the breadth of alloreactive T cell responses is an important factor in initiating and mediating the pathology associated with biologically-relevant alloreactive responses such as graft versus host disease and allograft rejection, it can preclude analysis of T cell responses to allogeneic ligands. To this end, this protocol describes a method for generating alloreactive T cells from naive human peripheral blood leukocytes (PBL) that respond to known peptide-MHC (pMHC) alloantigens. The protocol applies pMHC multimer labeling, magnetic bead enrichment and flow cytometry to single cell in vitro culture methods for the generation of alloantigen-specific T cell clones. This enables studies of the biochemistry and function of T cells responding to allogeneic stimulation.

This article describes a method for the generation and propagation of human T cell clones that specifically respond to a defined alloantigen. This protocol can be adapted for cloning human T cells specific for a variety of peptide-MHC ligands.

The study of human T lymphocyte biology often involves examination of responses to activating ligands. T cells recognize and respond to processed peptide ...

Flow Cytometric Analysis of Particle-bound Bet v 1 Allergen in PM10

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in diameter. In addition to a characterization of forward and sideward scatter properties, it enables the use of fluorescent labeled markers like antibodies to detect respective structures. Using indirect antibody staining, flow cytometry is employed here to quantify birch pollen allergen (precisely Bet v 1)-loaded particles of 0.5 to 10 &#181;m in diameter in inhalable particulate matter (PM10, particle size &#8804;10 &#181;m in diameter). PM10 particles may act as carriers of adsorbed allergens possibly transporting them to the lower respiratory tract, where they could trigger allergic reactions.
So far the allergen content of PM10 has been studied by means of enzyme linked immunosorbent assays (ELISAs) and scanning electron microscopy. ELISA measures the dissolved and not the particle-bound allergen. Compared to scanning electron microscopy, which can visualize allergen-loaded particles, flow cytometry may additionally quantify them. As allergen content of ambient air can deviate from birch pollen count, allergic symptoms might perhaps correlate better with allergen exposure than with pollen count. In conjunction with clinical data, the presented method offers the opportunity to test in future experiments whether allergic reactions to birch pollen antigens are associated with the Bet v 1 allergen content of PM10 particles &#62;0.5 &#181;m.

Here, we present a protocol to quantify allergen-loaded particles by flow cytometry. Ambient particulate matter particles may act as carriers of adsorbed allergens. We show here that flow cytometry, a method widely used to characterize suspended solids &#62;0.5 &#181;m in diameter, can be used to measure these allergen-loaded particles.

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in ...

Flow Cytometric Analysis of Natural Killer Cell Lytic Activity in Human Whole Blood

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and reproducibility of this assay can be considered unstable, either because of user&#39;s errors or because of the sensitivity of NK cells to experimental manipulation. To eliminate these issues, a workflow that reduces them to a minimum was established and is presented here. To illustrate, we obtained blood samples, at various time points, from runners (n = 4) that were submitted to an intense bout of exercise. First, NK cells are simultaneously identified and isolated through CD56 tagging and magnetic-based sorting, directly from whole blood and from as little as one milliliter. The sorted NK cells are removed of any reagent or capping antibodies. They can be counted in order to establish an accurate NK cell count per milliliter of blood. Secondly, the sorted NK cells (effectors cells or E) can be mixed with 3,3&#39;-Diotadecyloxacarbocyanine Perchlorate (DiO) tagged K562 cells (target cells or T) at an assay-optimal 1:5 T:E ratio, and analyzed using an imaging flow-cytometer that allows for the visualization of each event and the elimination of any false positive or false negatives (such as doublets or effector cells). This workflow can be completed in about 4 h, and allows for very stable results even when working with human samples. When available, research teams can test several experimental interventions in human subjects, and compare measurements across several time points without compromising the data&#39;s integrity.

This work describes an advanced workflow for the accurate and fast determination of NK (Natural Killer) cell count and NK cell cytotoxicity in human blood samples.

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and ...

Dextran Enhances the Lentiviral Transduction Efficiency of Murine and Human Primary NK Cells

The efficient transduction of specific genes into natural killer (NK) cells has been a major challenge. Successful transductions are critical to defining the role of the gene of interest in the development, differentiation, and function of NK cells. Recent advances related to chimeric antigen receptors (CARs) in cancer immunotherapy accentuate the need for an efficient method to deliver exogenous genes to effector lymphocytes. The efficiencies of lentiviral-mediated gene transductions into primary human or mouse NK cells remain significantly low, which is a major limiting factor. Recent advances using cationic polymers, such as polybrene, show an improved gene transduction efficiency in T cells. However, these products failed to improve the transduction efficiencies of NK cells. This work shows that dextran, a branched glucan polysaccharide, significantly improves the transduction efficiency of human and mouse primary NK cells. This highly reproducible transduction methodology provides a competent tool for transducing human primary NK cells, which can vastly improve clinical gene delivery applications and thus NK cell-based cancer immunotherapy.

The goal of this study was to formulate technologies that allow for successful gene transduction in primary natural killer (NK) cells. The dextran-mediated lentiviral transduction of human or mouse primary NK cells results in higher gene expression efficiencies. This method of gene transduction will vastly improve NK cell genetic manipulation.

The efficient transduction of specific genes into natural killer (NK) cells has been a major challenge. Successful transductions are critical to defining ...

Magnetic Stirrer Method for the Detection of Trichinella Larvae in Muscle Samples

Trichinellosis is a debilitating disease in humans and is caused by the consumption of raw or undercooked meat of animals infected with the nematode larvae of the genus Trichinella. The most important sources of human infections worldwide are game meat and pork or pork products. In many countries, the prevention of human trichinellosis is based on the identification of infected animals by means of the artificial digestion of muscle samples from susceptible animal carcasses.
There are several methods based on the digestion of meat but the magnetic stirrer method is considered the gold standard. This method allows the detection of Trichinella larvae by microscopy after the enzymatic digestion of muscle samples and subsequent filtration and sedimentation steps. Although this method does not require special and expensive equipment, internal controls cannot be used. Therefore, stringent quality management should be applied throughout the test. The aim of the present work is to provide detailed handling instructions and critical control points of the method to analysts, based on the experience of the European Union Reference Laboratory for Parasites and the National Reference Laboratory of Germany for Trichinella.

Magnetic Stirrer Method for the Detection of Trichinella Larvae in Muscle Samples

The prevention of human trichinellosis in many countries worldwide is based on the laboratory examination of muscle samples from susceptible animals by methods of digestion, of which the magnetic stirrer method is considered the gold standard.

Trichinellosis is a debilitating disease in humans and is caused by the consumption of raw or undercooked meat of animals infected with the nematode ...