The research was supported with Professional Development and Research Startup Funds from The University of Ontario Institute of Technology (UOIT) to Dr. John A. Samis and a Canadian Institutes of Health Research Health Professional Student Research Award to Irina Levit. The authors acknowledge Shannon Everett (Teaching and Learning Centre, UOIT) for her assistance preparing the video. The authors acknowledge Dr. Michael E. Nesheim (Department of Biochemistry, Queen's University, Kingston, ON) for his mentorship, insight, and helpful discussions. The authors also acknowledge Dr. Cheng Hock Toh (Roald Dahl Haemostasis and Thrombosis Centre, Royal Liverpool University Hospital, Liverpool, UK) for providing the DIC patient plasmas used in the study.

1. Preparation of FV-deficient Human Plasma
<ol>
 <li>This method is a modification of the procedure originally described by Bloom et al 4. </li>
 <li>Obtain whole human blood from a consenting healthy adult volunteer (Male or Female, age 18 - 50 yrs) that is not taking any medication. Wear latex gloves and a laboratory coat throughout the procedure.</li>
 <li>Prepare 100 ml of 3.2% (w/v) trisodium citrate in distilled water. Load 6.0 ml of this solution into separate 60 ml syringes. Remove air by depressing plunger carefully to the 6.0 ml mark on the syringe. </li>
 <li>Use an alcohol swab to clean area of cubital vein between bicep and forearm. Connect 20 gauge butterfly apparatus to a 3 ml syringe. Insert butterfly needle into cubital vein and draw gently on plunger until blood fills to the 3 ml mark. Discard syringe with blood. This step is performed to remove any tissue factor released from the damaged endothelium at the needle injury site that may activate the coagulation process and interfere with preparation of FV-deficient plasma. </li>
 <li>Connect butterfly needle to 60 ml syringe 'loaded' with 6.0 ml trisodium citrate and draw gently on the plunger until the blood reaches the 60 ml mark on the syringe. (Final citrate concentration 10.9mM). </li>
 <li>Remove syringe from butterfly needle and gently mix syringe while keeping a gloved finger or thumb on the syringe outlet. </li>
 <li>Continue drawing blood into 60 ml syringes each filled with 6.0 ml of 3.2% trisodium citrate as described above. One should anticipate recovery of approx 50% of the citrated blood volume as plasma after centrifugation and that 50 &mu;l of FV-deficient plasma is required per sample in the microplate assay. Our laboratory routinely processes 300-420 ml of citrated blood at a time using 5-7 x 60 ml syringes each with 6.0 ml of 3.2% trisodium citrate/syringe. This is sufficient FV-deficient plasma for approximately 3,000 microplate assays (See below). </li>
 <li>Remove butterfly needle from volunteer's arm and press a sterile gauze pad over site of vein puncture for 1.0 min. Cover site of vein puncture with a sterile bandage. </li>
 <li>Centrifuge citrated blood at 3,300 x g for 20 min at room temperature. Recover upper yellow plasma layer to a plastic beaker with a stir bar using a plastic transfer pipette taking care not to disturb the lower red and white blood cell layer. </li>
 <li>Measure volume of plasma in ml and retain 100 &mu;l of plasma before EDTA addition on ice for future prothrombin clot time (PT) testing (See below). </li>
 <li>Slowly add solid EDTA to citrated plasma with gentle stirring at room temperature to achieve 5mM (Final concentration). Once the EDTA has dissolved, adjust pH to 7.0 by dropwise addition of 1.0M NaOH with gentle stirring. </li>
 <li>Cover beaker with Saran Wrap and incubate for 8-10 hr without stirring at 37 &deg;C. </li>
 <li>Every 2 hr, recover 100 &mu;l of the EDTA-treated plasma on ice and determine the PT at the time of incubation with EDTA and compare to the PT of plasma prior to EDTA addition (See above). </li>
 <li>For the PT determinations, place removable 8 well immunomodule strips into plastic template holder and insert into microplate reader. </li>
 <li>Orient immunomodule strips in microplate reader as: empty, empty, sample, empty, empty, sample, empty, and empty from left to right in the template holder to minimize light scatter interference from neighbouring sample-containing strips. Also, use only wells 2-7 from top to bottom in each 8 well immunomodule strip to minimize light scatter interference from edges of plastic template holder. </li>
 <li>The FV microplate assay is capable of measuring fibrin clot formation in at least 12 different samples simultaneously (6 samples per immunomodule strip over 2 immunomodule strips). </li>
 <li>Using a multichannel pipette, add 50 &mu;l of HBS (20mM HEPES, 150mM NaCl, pH 7.4) to each microplate well for each of sample to be assayed. Using a single pipette, add 50 &mu;l of normal human pooled reference plasma (NHP) or plasma before addition of EDTA or at various incubation times after adding EDTA to separate microplate wells. </li>
 <li>Using a multichannel pipette, add 50 &mu;l thromboplastin (See below for preparation method) to all microplate wells with plasma to be assayed. Incubate in microplate reader at 25 &deg;C for 1 min and program the microplate reader to shake the plate during the 15-25 sec of the 1 min incubation. </li>
 <li>Using the computer interfaced with the microplate reader, access SoftMax Pro software microplate application. Set the reader to Absorbance, Wavelength to 405nm, mode to Kinetic, temperature to 25 &deg;C, and program the microplate reader to shake for 5 sec, and read the absorbance at 405nm every 5 sec for 6 min. </li>
 <li>Using a multichannel pipette, add calcium chloride (50 &mu;l of 25mM in distilled water; 2.1mM final concentration) to all of the microplate wells, wait 15 sec, and press the 'Start' icon on the microplate SoftMax Pro menu to begin the assay. </li>
 <li>Determine the PTs from the Absorbance vs. Time profiles in SoftMax Pro as the time to reach the half maximal increase in absorbance at 405nm (The mid-point of the sigmoidal Absorbance vs. Time curve produced after thromboplastin and calcium chloride addition. See below, Figure 1).</li>
 <li>Calculate the FV 1-stage activity from the Log Clot Time (sec) vs. Log FV Activity (Units/ml) as illustrated below in Figure 2A. For quantitation purposes, 1Unit of FV activity is defined as the activity present in 1.0 ml NHP prior to intentional activation with added thrombin (See below, Assaying human plasma samples with the FV 1-stage and 2-stage microplate coagulation assay). </li>
 <li>Prepare 4.0l of 20mM HEPES containing 0.15M NaCl in distilled water with gentle stirring. Adjust pH to 7.4 with slow dropwise addition of 5.0M NaOH (HBS, pH 7.4). </li>
 <li>Obtain a sufficient length of dialysis tubing and rinse thoroughly with distilled water. Tie a knot in one end of the tubing. Transfer EDTA treated plasma to dialysis tubing with a plastic transfer pipette, remove air, and tie a knot in the other end of the tubing. </li>
 <li>Carefully hang the EDTA-treated plasma in the tubing into the HBS with gentle stirring. Cover with Saran Wrap and dialyze for 14-16h with gentle stirring at 4 &deg;C. </li>
 <li>Carefully remove the EDTA-treated/dialyzed plasma in 3.0 ml aliquots to 15.0 ml screw capped tubes and retain 100 &mu;l for PT assay of 'final' FV-deficient plasma on ice. Store the FV-deficient plasma in at -80 &deg;C until use. Under the conditions described above, the PTs increase from 10-15 sec before EDTA addition to 90-100 sec 8-10 hr after EDTA addition. The FV-deficient plasma prepared by this method routinely contains less than 0.1% of the active FV present in the starting fresh human plasma. </li>
</ol>
2. Preparation of Thromboplastin
<ol>
 <li>Dialysis is required to remove any calcium from this reagent in order to permit precisely timing fibrin clot formation from the time of calcium chloride addition to initiate coagulation process. </li>
 <li>Prepare 100 ml and 4.0l of 20mM HEPES and 0.15M NaCl in distilled water in a plastic beaker with gentle stirring. Adjust pH to 7.4 with slow dropwise addition of 5.0M NaOH (HBS, pH 7.4). </li>
 <li>Add 6.0 ml of HBS, pH 7.4 to each vial of thromboplastin reagent, replace lid, and shake gently to dissolve. Incubate reagent in capped vials for 15 min at room temperature to dissolve dried material. Shake gently to completely dissolve. </li>
 <li>Cut a sufficient length of dialysis tubing, rinse well with distilled water, and tie off one end of the tubing with a knot. Transfer thromboplastin to dialysis tubing, remove air, and tie off other end of tubing with a knot. </li>
 <li>Carefully hang dialysis tubing in 4.0l of HBS, pH 7.4 and gently stir covered with Saran Wrap at 4 &deg;C for 14-16 hr. </li>
 <li>Transfer 3.0 ml aliquots of the dialyzed thromboplastin to 15 ml screw capped tubes and store at -80 &deg;C until used. </li>
</ol>
3. Preparation of FV 1-stage Microplate Coagulation Assay Activity Standard Curves
<ol>
 <li>Thaw FV-deficient plasma, NHP, and thromboplastin at 37 &deg;C for 10 min. Gently mix FV-deficient plasma and thromboplastin independently, transfer to separate plastic washing trays, and incubate on ice. Store the NHP on ice after gently mixing. Store calcium chloride (25mM in distilled water) in a separate washing tray at room temperature. </li>
 <li>Prepare 200 &mu;l each of 2-fold serial dilutions of NHP from 0-, 2-, 4-, 8-, 16-, 32-, 64-, 128-, 256-, 512-fold in HBS, pH 7.4 using 1.5 ml microcentrifuge tubes. Vortex well and incubate on ice. </li>
 <li>Place removable 8 well immunomodule strips into plastic template holder and insert into microplate reader. </li>
 <li>Orient immunomodule strips in microplate reader as: empty, empty, sample, empty, empty, sample, empty, and empty from left to right in the template holder to minimize light scatter interference from neighbouring sample-containing strips. Also, use only wells 2-7 from top to bottom in each 8 well immunomodule strip to minimize light scatter interference from edges of plastic template holder. </li>
 <li>The FV microplate assay is capable of measuring fibrin clot formation in at least 12 different samples simultaneously (6 samples per immunomodule strip over 2 immunomodule strips).</li>
 <li>Turn on microplate reader and access SoftMax Pro microplate application software. </li>
 <li>Using a multichannel pipette, make simultaneous additions of FV-deficient plasma (50 &mu;l) to all sample microplate wells. Use a single pipette, add 50 &mu;l of the 10 serial dilutions of NHP prepared above to separate microplate wells. </li>
 <li>Using a multichannel pipette, add thromboplastin (50 &mu;l) to all sample wells. Incubate in the microplate reader at 25 &deg;C for 1 min and program the microplate reader to shake the plate during the 15-25 sec of the 1 min incubation. </li>
 <li>Using the computer interfaced with the microplate reader, access SoftMax Pro software microplate application. Set the reader to Absorbance, Wavelength to 405nm, mode to Kinetic, temperature to 25 &deg;C, and program the microplate reader to shake for the first 5 sec after calcium chloride addition, and measure absorbance at 405nm every 5 sec for 6 min thereafter. The 5 sec shaking interval is not included in the calculated clot times (See below).</li>
 <li>Using a multichannel pipette, add calcium chloride (50 &mu;l of 25mM in distilled water; final concentration 3.5mM) to all sample microplate wells, wait 15 sec, press the 'Start' icon in the microplate application in SoftMax Pro from computer to begin the assay. </li>
 <li>The clot times are calculated as the time in seconds to reach the midpoint between the minimum and maximum absorbance at 405nm after thromboplastin and calcium chloride addition. </li>
 <li>The initial rates of clot formation are calculated as the rate of increase in absorbance at 405nm (mUnits/min) that encompass the first 5 time points of clot formation in the linear portion of the Absorbance vs. Time curve. </li>
 <li>The extent of clot formation was calculated as the difference between the maximal and minimal absorbance at 405nm (Units) achieved during the clot formation event. </li>
</ol>
4. Assaying Human Plasma Samples with the FV 1-stage and 2-stage Microplate Coagulation Assay
<ol>
 <li>Thaw FV-deficient plasma, NHP, sample plasmas, and thromboplastin at 37 &deg;C for 10 min. Gently mix FV-deficient plasma and thromboplastin independently, transfer to separate plastic ELISA washing trays, and incubate on ice. Store the NHP and sample plasmas on ice after gently mixing. Store calcium chloride (25mM in distilled water) in a separate ELISA washing tray at room temperature. </li>
 <li>Prepare 200 &mu;l each of 40-fold dilutions of NHP and sample plasmas in HBS, pH 7.4 using 1.5 ml microcentrifuge tubes. Vortex well and incubate on ice. </li>
 <li>Insert removable 8 well immunomodule strips into plastic template holder and insert into microplate reader. </li>
 <li>Alternate empty, empty, sample, empty, empty, sample, empty, and empty immunomodule strips from left to right in the template holder to minimize light scatter interference from neighbouring sample-containing strips. Use only wells 2-7 from top to bottom in each immunomodule strip to minimize light scatter interference from edges of plastic template holder. </li>
 <li>Turn on microplate reader and access SoftMax Pro microplate application software. </li>
 <li>Using a multichannel pipette, add FV-deficient plasma (50 &mu;l) to all sample microplate wells. Using a single pipette, add 50 &mu;l of 40-fold diluted NHP or sample plasmas prepared above to separate microplate wells. </li>
 <li>Using a multichannel pipette, add thromboplastin (50 &mu;l) to all sample wells. Incubate in the microplate reader at 25 &deg;C for 1 min and program the microplate reader to shake the plate during the 15-25 sec of the 1 min incubation. </li>
 <li>Using the computer interfaced with the microplate reader, access SoftMax Pro software microplate application. Set the reader to Absorbance, Wavelength to 405nm, mode to Kinetic, temperature to 25 &deg;C, and program the microplate reader to shake for the first 5 sec after calcium chloride addition, and measure absorbance at 405nm every 5 sec for 6 min thereafter. The 5 sec shaking interval is not included in the calculated clot times (See below). </li>
 <li>Using a multichannel pipette, add calcium chloride (50 &mu;l of 25mM in distilled water; 6.25mM final concentration) to all sample microplate wells and immediately press the 'Start' icon in the microplate application in SoftMax Pro from computer to begin the assay.</li>
 <li>At the end of the run, determine the time, initial rate, and extent of clot formation for 40-fold diluted NHP and sample plasmas from the absorbance vs. time profiles in SoftMax Pro.</li>
 <li>Taking into account the 40-fold dilution, use the clot time for each sample to calculate the FV activity in Units/ml from the FV 1-stage activity standard curve of Log Clot Time vs. Log FV Activity (Figure 2A). This represents the FV-1 stage activity or that without 'intentional' activation by thrombin. </li>
 <li>Since FV is activated with thrombin to the active cofactor, FVa 1, a second assay after addition and incubation with thrombin is critical to accurately measure the FV 2-stage activity of a plasma sample (With 'intentional' activation by added thrombin). </li>
 <li>For the FV 2-stage assay, prepare 200-300 &mu;l of purified thrombin 18 at 100nM in HBS, pH 7.4 and incubate on ice. Plan to use 10 &mu;l of diluted thrombin for each plasma sample to be assayed with the FV 2-stage assay. </li>
 <li>Dilute 120 &mu;l of the above NHP and sample plasmas from 40-fold to 100-fold with 170 &mu;l of HBS, pH 7.4 containing 2.8mM calcium chloride. Add 10 &mu;l thrombin (10nM final concentration) to each sample. Vortex well to mix and incubate at 37 &deg;C for 1 min. </li>
 <li>Dilute NHP and sample plasmas to 500-fold by adding 400 &mu;l of HBS, pH 7.4, vortex well, and assay 50 &mu;l of each plasma sample in the FV 1-stage microplate assay as described above. </li>
 <li>Determine the clot time for NHP and each plasma sample assayed from the Absorbance vs. Time profiles in SoftMax Pro. Taking into account the 500-fold dilution, use the clot time for each sample to calculate the FV 2-stage activity from the FV 1-stage standard curve of Log Clot Time vs. Log FV Activity (Figure 2A). </li>
 <li>The total FV activity may then be calculated as FV 2-stage activity - FV 1-stage activity. </li>
</ol>

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No conflicts of interest declared.

The kinetic microplate assay method outlines the development of a novel, rapid, inexpensive, and convenient technique for measurement of FV coagulation activity in samples of human plasma which have not (FV 1-stage assay) or have been intentionally activated with added thrombin (FV 2-stage assay). The assay utilizes a kinetic microplate reader and all the materials and reagents required are commercially available or may be made in-house. The assay continuously monitors the change in the light scatter of plasma during fibrin clot formation at 405nm. The microplate assay has the advantage of using small plasma sample volumes (&mu;l) and is amenable for analysis of multiple samples simultaneously (up to 12). These assay attributes are advantageous when expensive equipment and reagents are used (Automated analyzers and Factor-deficient plasmas); only small sample volumes are available (Patient plasmas) or analyzing a large number of samples (During FV purification from plasma).
The FV microplate assay has the added advantages that it is convenient, fast, and does not require the isolation and purification of the required constituent components. Compared with manual tilt-tube 3, 4 and automated 5-7 FV assays that have been reported, the FV microplate assay has comparable useful range of clot times (25-75 sec) and corresponding FV levels in the sample (0.5-0.005 Units/ml), and sensitivity/detection limits (20-80pM). Compared with manual tilt-tube FV assays which suffer from a subjective visual assessment of clot formation 3, 4, the FV microplate assay permits objective and quantitative measurement of the clot time, initial rate, and extent of fibrin clot formation. Compared with automated FV assays which suffer from the requirement of expensive analyzers and reagents 5-7, the FV microplate assay reagents and materials are inexpensive and all the required materials may be purchased commercially or made in-house. Manual tilt-tube 3, 4 and automated 5-7 FV assays generally only provide the time for clot formation; not the time, initial rate, and extent of fibrin clot formation which are all accurately measured spectrophotometrically with the FV microplate assay. Finally, manual tilt-tube 3,4 and automated 5-7 FV assays suffer from only being able to measure the FV activity in plasma samples one at a time; whereas the FV microplate assay is amenable to high-throughput and 12 samples may be assayed simultaneously.
The microplate assay was used to demonstrate that the FV in 9 DIC patient plasmas was less active than in NHP because of a combination of delayed clot times and lower initial rates of clot formation in the FV 1-stage assay. The initial rates of clot formation in the FV 2-stage assay in the DIC patient plasmas were not changed from NHP. The extents of clot formation in the DIC patient plasmas were also not changed from NHP in the FV 1-stage and 2-stage assays. Decreased FV 1-stage, 2-stage, and total activities may have been due to increased FV consumption 19 and/or inactivation 6 in accordance with other studies during the pathogenesis of this acquired blood disorder. Given the extent of clot formation in the DIC plasmas was the same as observed with NHP, this variable was most likely a result of the fibrinogen concentration in the FV-deficient plasma and not related to the characteristics of the patient plasmas.
The results from the microplate assay indicated that measurement of FV activity in samples of human plasma may be obtained from the time and initial rate of clot formation from the FV 1-stage assay and the time of clot formation and total activity from the FV 2-stage assay. It was not possible to obtain quantitative measurement of FV activity in a plasma sample based on the extent of clot formation in the FV 1- and 2-stage assays or the initial rate of clot formation in the FV 2-stage assay. Given all reactions reached approximately the same extent of clot formation of 0.35 - 0.45 Units between the initial absorbance before and the maximal absorbance after thromboplastin and calcium chloride addition, measurement of the extent of clot formation does not provide a quantitative assessment of FV activity in a given plasma sample.
The novel microplate assay will find use in research and clinical settings for measurement of FV activity in samples of human plasma and fractions during its purification from human and animal plasma. The microplate assay may be used to measure the time, initial rate, and extent of clot formation; parameters not generally monitored simultaneously in manual tilt-tube assays and automated coagulation analyzers. This information provides more of a complete quantitative assessment of the involvement of FV during the clot formation event in human plasma than has been previously reported 3-7. This information is especially important in both research and clinical laboratories when measuring significant changes in FV activity in samples of plasma or with purified FV or FVa. The FV microplate assay may also be used to characterize and measure compounds that may activate or inactivate FV and FVa, measure the FV activity in patients at risk for venous thrombosis as a result of the FV Leiden mutation 20, 21 and to monitor the activity of FV during its purification from plasma.
The FV microplate assay is easily adaptable to measure the activity of any coagulation factor in the extrinsic, intrinsic, or common pathway using the appropriate factor-deficient plasma and initiating reagents for fibrin formation. The assay will increase our understanding of FV biochemistry through a more accurate and complete measurement of its activity in research and clinical laboratories. Ultimately, this information will positively impact healthcare environments through earlier diagnosis and development of more effective treatments for individuals afflicted with coagulation disorders, such as DIC.

<table border="1">
 <tbody>
 <tr>
 <td>Name of reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (Optional)</td>
 </tr>
 <tr>
 <td>Kinetic microplate reader</td>
 <td>Molecular Devices</td>
 <td>Spectra Max 190</td>
 <td>SOFTmax PRO 4.3 LS software</td>
 </tr>
 <tr>
 <td>Normal pooled human reference plasma</td>
 <td>Precision Biologicals</td>
 <td>CCN-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Trisodium citrate</td>
 <td>Becton Dickenson</td>
 <td>B10242</td>
 <td></td>
 </tr>
 <tr>
 <td>EDTA</td>
 <td>Becton Dickenson</td>
 <td>B10093</td>
 <td></td>
 </tr>
 <tr>
 <td>FV-deficient human plasma4</td>
 <td></td>
 <td></td>
 <td>Made from fresh human plasma and stored in aliquots at -70 &deg;C.</td>
 </tr>
 <tr>
 <td>3 and 60 ml syringes</td>
 <td>Becton Dickenson</td>
 <td>309585 and 136898</td>
 <td></td>
 </tr>
 <tr>
 <td>Butterfly apparatus</td>
 <td>Vacutainer</td>
 <td>367251</td>
 <td>20 gauge</td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Sigma/Aldrich</td>
 <td>H-3375</td>
 <td></td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>Fisher</td>
 <td>BP358-212</td>
 <td></td>
 </tr>
 <tr>
 <td>Thromboplastin</td>
 <td>Trinity Biotech</td>
 <td>T1106</td>
 <td>Dialyzed vs. HBS, pH 7.4 and stored in aliquots at -70 &deg;C.</td>
 </tr>
 <tr>
 <td>Calcium chloride</td>
 <td>Becton Dickenson</td>
 <td>B10070</td>
 <td></td>
 </tr>
 <tr>
 <td>Human thrombin15</td>
 <td></td>
 <td></td>
 <td>From human plasma; stored at -20 &deg;C in 50% (v/v) glycerol.</td>
 </tr>
 <tr>
 <td>Dialysis membrane</td>
 <td>Fisher</td>
 <td>21-152-6</td>
 <td>6-8kDa molecular weight cutoff; 50mm x 3m.</td>
 </tr>
 <tr>
 <td>Template holder and removable 8 well strips.</td>
 <td>Nunc</td>
 <td>14-245-63 and 469957</td>
 <td></td>
 </tr>
 <tr>
 <td>ELISA washing trays</td>
 <td>BioRad</td>
 <td>224-4872</td>
 <td></td>
 </tr>
 <tr>
 <td>1.5 ml microfuge and 15 ml screw capped tubes.</td>
 <td>Sarstedt</td>
 <td>72690 and 62554205</td>
 <td></td>
 </tr>
 <tr>
 <td>Multichannel pipette</td>
 <td>Fisher</td>
 <td>2703630</td>
 <td>8 channel model.</td>
 </tr>
 </tbody>
 </table>

measurement of factor v activity in human plasma using a microplate coagulation assay

In response to injury, blood coagulation is activated and results in generation of the clotting protease, thrombin. Thrombin cleaves fibrinogen to fibrin which forms an insoluble clot that stops hemorrhage. Factor V (FV) in its activated form, FVa, is a critical cofactor for the protease FXa and accelerator of thrombin generation during fibrin clot formation as part of prothrombinase 1, 2. Manual FV assays have been described 3, 4, but they are time consuming and subjective. Automated FV assays have been reported 5-7, but the analyzer and reagents are expensive and generally provide only the clot time, not the rate and extent of fibrin formation. The microplate platform is preferred for measuring enzyme-catalyzed events because of convenience, time, cost, small volume, continuous monitoring, and high-throughput 8, 9. Microplate assays have been reported for clot lysis 10, platelet aggregation 11, and coagulation Factors 12, but not for FV activity in human plasma. The goal of the method was to develop a microplate assay that measures FV activity during fibrin formation in human plasma.
This novel microplate method outlines a simple, inexpensive, and rapid assay of FV activity in human plasma. The assay utilizes a kinetic microplate reader to monitor the absorbance change at 405nm during fibrin formation in human plasma (Figure 1) 13. The assay accurately measures the time, initial rate, and extent of fibrin clot formation. It requires only &mu;l quantities of plasma, is complete in 6 min, has high-throughput, is sensitive to 24-80pM FV, and measures the amount of unintentionally activated (1-stage activity) and thrombin-activated FV (2-stage activity) to obtain a complete assessment of its total functional activity (2-stage activity - 1-stage activity).
Disseminated intravascular coagulation (DIC) is an acquired coagulopathy that most often develops from pre-existing infections 14. DIC is associated with a poor prognosis and increases mortality above the pre-existing pathology 15. The assay was used to show that in 9 patients with DIC, the FV 1-stage, 2-stage, and total activities were decreased, on average, by 54%, 44%, and 42%, respectively, compared with normal pooled human reference plasma (NHP).
The FV microplate assay is easily adaptable to measure the activity of any coagulation factor. This assay will increase our understanding of FV biochemistry through a more accurate and complete measurement of its activity in research and clinical settings. This information will positively impact healthcare environments through earlier diagnosis and development of more effective treatments for coagulation disorders, such as DIC.

Representative Results - Preparation of FV 1-stage microplate coagulation assay activity standard curves
A representative example of fibrin clot formation in the FV assay over time generated by the microplate reader is illustrated in Figure 1. The FV assay accurately measures the time, initial rate, and extent of fibrin clot formation. Inspection of the wells upon assay completion confirmed that clot formation occurred. All reactions reached approximately the same extent of clot formation with a general change in absorbance at 405nm of 0.35 - 0.45 Units between the starting absorbance before and the maximal absorbance after thromboplastin and calcium chloride addition. Representative examples of FV 1-stage activity standard curves of Log Clot Time (in seconds) vs. Log FV Activity (Units/ml) and Log Initial Rate of Clot Formation (in mUnits/min) vs. Log FV Activity (Units/ml) for serial dilutions of NHP is shown in Figure 2A and Figure 2B, respectively. Fitting the Log-Log plot of Clot Time vs. FV 1-Activity indicated a strong relationship between these variables after linear regression analysis (Figure 2A; R2 = 0.980). Fitting the Log-Log plot of the Initial Rate of Clot Formation vs. FV Activity also indicated a strong relationship between these variables after linear regression analysis (Figure 2B; R2 = 0.983). The relationship of both Log Clot Time and Log Initial Rate of Clot Formation vs. Log FV Activity remained linear for NHP diluted up to 512-fold. Since FV circulates in NHP at approx 12-40nM 16, the microplate assay is sensitive to approx 24-80pM FV in NHP. Given that the dissociation constant of the interaction of FVa-FXa-lipid in prothrombinase is approximately 1nM 17, the FV microplate assay is entirely suitable for measurement of FV levels in the physiologically relevant nM range for FVa function in prothrombinase.
Using the FV microplate assay, it was also determined that the normal range of FV activity in the FV 1-stage activity assay of 15 healthy control plasmas (Male and Female, Age 18-20yrs) was (Mean &plusmn; Standard deviation; Range): 0.96 &plusmn; 0.14U/ml; 0.68-1.11U/ml. This agrees well with the normal healthy FV activity and range (0.66-1.14U/ml) reported by Cutler et al. for the FV 1-stage activity determined with an automated analyzer 7. The intra-assay variability of the time, extent, and initial rate of clot formation in the FV 1-stage assay in 6 wells on 8 different days was 3.4%, 4.4%, and 3.1%, respectively. The inter-assay variability of the time, extent, and initial rate of clot formation in the FV 1-stage assay in 6 wells of 8 different experiments on 8 different days was 7.1%, 7.8%, and 9.2%, respectively. Thus, the intra- and inter-assay variability of these three measured variables was at a low and acceptable level for robust assay performance within and between microplate assay of multiple samples simultaneously (Up to 12).
Representative Results - Assaying human plasma samples with the FV 1-stage and 2-stage microplate coagulation assay
The standard curve of Log Clot Time vs. Log FV Activity (Figure 2A) was used to measure the FV activity in NHP and 9 DIC patient plasmas which were not intentionally activated with added thrombin (FV 1-stage activity) or were intentionally activated with added thrombin (FV 2-stage activity) and the results are shown in Table 1. All 9 DIC patient plasmas exhibited FV 1-stage activities and initial rates of clot formation that were decreased on average, by 54% and 18%, respectively, from NHP. The extents of clot formation in the FV 1-stage assay in the DIC patients were not largely different from NHP, and increased on average, by 13% from NHP.
 Activation of NHP with thrombin generated an approximate 8-fold increase in FV 2-stage activity above the FV 1-stage activity (Table 1). This indicates that the FV in NHP was mainly present in its unactivated form and agrees with the previously reported results using manual tilt-tube FV assays 3, 4 and automated FV assays 5-7. The FV 2-stage and total activity were also decreased in the DIC patients on average, by 44% and 42%, respectively, from NHP. The initial rates and extents of clot formation in the FV 2-stage assay in the DIC patients were not significantly different from NHP, and varied on average, by approx 9% and 4%, respectively, from that observed with NHP. These results indicated that compared with the FV in NHP, the FV in the DIC patient plasmas resulted in a prolonged time and decreased rate of fibrin clot formation and that the patient FV was on average, only 56% as activatable with thrombin as well. 
<img src="/files/ftp_upload/3822/3822fig1.jpg" alt="Figure 1" /> 
Figure 1. Clot formation in normal pooled human reference plasma measured with the kinetic microplate FV 1-stage coagulation assay. Fibrin clot formation in NHP was continuously monitored at 405nm using a kinetic microplate reader. The plot is the microplate reader output of a 6 min reaction of 32-fold diluted NHP, FV-deficient plasma, thromboplastin, and calcium chloride in a microplate well. The vertical axis represents the change in absorbance at 405nm that occurred as a result of fibrin formation in plasma. The time of fibrin formation was defined as the time to reach the half maximal increase in absorbance or the midpoint of curve (36.40 sec). The initial rate of clot formation was defined as the rate of change of absorbance at 405nm over the first 5 time points of the linear increase of the absorbance portion of the curve (611.88 mUnits/min). The extent of clot formation was defined as the difference between the maximum and minimum absorbance at 405 nm (0.35 Units). 
<img src="/files/ftp_upload/3822/3822fig2.jpg" alt="Figure 2" /> 
Figure 2. Standard curves of time and initial rate of clot formation vs. Factor V activity in normal pooled human reference plasma using the FV 1-stage microplate assay. NHP was serially diluted (0- to 512-fold in HBS) and assayed with the FV 1-stage microplate assay as described in the protocol. Log-Log plots of the time and initial rate of clot formation vs. FV activity in the FV 1-stage microplate assay are shown after linear regression modeling of the data in panels A and B, respectively.
<table border="1">
 <tbody>
 <tr>
 <td>Sample</td>
 <td>1-stage assay Activity (Units/ml)</td>
 <td>1-stage assay Extent (Units)</td>
 <td>1-stage assay Initial Rate (mUnits/min)</td>
 <td>2-stage assay Activity (Units/ml)</td>
 <td>2-stage assay Extent (Units)</td>
 <td>2-stage assay Initial Rate (mUnits/min)</td>
 <td>Total activity (Units/ml)</td>
 </tr>
 <tr>
 <td>NHP</td>
 <td>1.02</td>
 <td>0.363</td>
 <td>744.96</td>
 <td>7.93</td>
 <td>0.433</td>
 <td>375.84</td>
 <td>6.91</td>
 </tr>
 <tr>
 <td>Patient 1</td>
 <td>0.36</td>
 <td>0.404</td>
 <td>583.80</td>
 <td>3.84</td>
 <td>0.432</td>
 <td>249.96</td>
 <td>3.48</td>
 </tr>
 <tr>
 <td>Patient 2</td>
 <td>0.67</td>
 <td>0.416</td>
 <td>704.04</td>
 <td>5.28</td>
 <td>0.469</td>
 <td>323.16</td>
 <td>4.61</td>
 </tr>
 <tr>
 <td>Patient 3</td>
 <td>0.31</td>
 <td>0.435</td>
 <td>562.08</td>
 <td>3.92</td>
 <td>0.453</td>
 <td>294.96</td>
 <td>3.61</td>
 </tr>
 <tr>
 <td>Patient 4</td>
 <td>0.39</td>
 <td>0.435</td>
 <td>617.04</td>
 <td>4.14</td>
 <td>0.462</td>
 <td>372.60</td>
 <td>3.75</td>
 </tr>
 <tr>
 <td>Patient 5</td>
 <td>0.73</td>
 <td>0.401</td>
 <td>641.40</td>
 <td>5.91</td>
 <td>0.433</td>
 <td>354.24</td>
 <td>5.18</td>
 </tr>
 <tr>
 <td>Patient 6</td>
 <td>0.45</td>
 <td>0.403</td>
 <td>600.72</td>
 <td>4.16</td>
 <td>0.445</td>
 <td>393.96</td>
 <td>3.71</td>
 </tr>
 <tr>
 <td>Patient 7</td>
 <td>0.49</td>
 <td>0.395</td>
 <td>575.40</td>
 <td>4.89</td>
 <td>0.449</td>
 <td>357.48</td>
 <td>4.40</td>
 </tr>
 <tr>
 <td>Patient 8</td>
 <td>0.19</td>
 <td>0.448</td>
 <td>489.00</td>
 <td>2.89</td>
 <td>0.450</td>
 <td>330.48</td>
 <td>2.70</td>
 </tr>
 <tr>
 <td>Patient 9</td>
 <td>0.64</td>
 <td>0.423</td>
 <td>699.48</td>
 <td>5.11</td>
 <td>0.455</td>
 <td>417.24</td>
 <td>4.47</td>
 </tr>
 </tbody>
</table>
Table 1 FV Activity in NHP and in 9 Patients That Developed Disseminated Intravascular Coagulation (DIC). The FV 1-stage, 2-stage, and total activity in NHP and 9 DIC patient plasmas were determined from the FV 1-stage microplate assay standard curve of time of clot formation vs. FV activity (Figure 2A) and are given in Units/ml. The initial rates (Initial rate of increase in A405nm over first five time points in mUnits/min) and extents (Maximum A405nm - Minimum A405nm) of clot formation in the FV 1- and 2-stage microplate assay are also shown.

Watch this Scientific Journal Video about Measurement of Factor V Activity in Human Plasma Using a Microplate Coagulation Assay at JoVE.com

Measurement of Factor V Activity in Human Plasma Using a Microplate Coagulation Assay

This study describes a novel microplate assay that measures FV coagulation activity during fibrin clot formation in human plasma which has not been reported previously. The method uses a kinetic microplate reader to continuously measure the change in absorbance at 405nm during fibrin clot formation in human plasma.

 In response to injury, blood coagulation is activated and results in generation of the clotting protease, thrombin. Thrombin cleaves fibrinogen to fibrin ...

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Immunology and Infection

18.9K Views.  University of Ontario Institute of Technology. This study describes a novel microplate assay that measures FV coagulation activity during fibrin clot formation in human plasma which has not been reported previously. The method uses a kinetic microplate reader to continuously measure the change in absorbance at 405nm during fibrin clot formation in human plasma.

Video: Measurement of Factor V Activity in Human Plasma Using a Microplate Coagulation Assay

Co-culture Models of Pseudomonas aeruginosa Biofilms Grown on Live Human Airway Cells

Bacterial biofilms have been associated with a number of different human diseases, but biofilm development has generally been studied on non-living surfaces. In this paper, we describe protocols for forming Pseudomonas aeruginosa biofilms on human airway epithelial cells (CFBE cells) grown in culture. In the first method (termed the Static Co-culture Biofilm Model), P. aeruginosa is incubated with CFBE cells grown as confluent monolayers on standard tissue culture plates. Although the bacterium is quite toxic to epithelial cells, the addition of arginine delays the destruction of the monolayer long enough for biofilms to form on the CFBE cells. The second method (termed the Flow Cell Co-culture Biofilm Model), involves adaptation of a biofilm flow cell apparatus, which is often used in biofilm research, to accommodate a glass coverslip supporting a confluent monolayer of CFBE cells. This monolayer is inoculated with P. aeruginosa and a peristaltic pump then flows fresh medium across the cells. In both systems, bacterial biofilms form within 6-8 hours after inoculation. Visualization of the biofilm is enhanced by the use of P. aeruginosa strains constitutively expressing green fluorescent protein (GFP). The Static and Flow Cell Co-culture Biofilm assays are model systems for early P. aeruginosa infection of the Cystic Fibrosis (CF) lung, and these techniques allow different aspects of P. aeruginosa biofilm formation and virulence to be studied, including biofilm cytotoxicity, measurement of biofilm CFU, and staining and visualizing the biofilm.

Co-culture Models of Pseudomonas aeruginosa Biofilms Grown on Live Human Airway Cells

This paper describes different methods of growing Pseudomonas aeruginosa biofilms on cultured human airway epithelial cells. These protocols can be adapted to study different aspects of biofilm formation, including visualization of the biofilm, staining of the biofilm, measuring the colony forming units (CFU) of the biofilm, and studying biofilm cytotoxicity.

Bacterial biofilms have been associated with a number of different human diseases, but biofilm development has generally been studied on non-living ...

Expansion, Purification, and Functional Assessment of Human Peripheral Blood NK Cells

Natural killer (NK) cells play an important role in immune surveillance against a variety of infectious microorganisms and tumors. Limited availability of NK cells and ability to expand in vitro has restricted development of NK cell immunotherapy. Here we describe a method to efficiently expand vast quantities of functional NK cells ex vivo using K562 cells expressing membrane-bound IL21, as an artificial antigen-presenting cell (aAPC).
NK cell adoptive therapies to date have utilized a cell product obtained by steady-state leukapheresis of the donor followed by depletion of T cells or positive selection of NK cells. The product is usually activated in IL-2 overnight and then administered the following day 1. Because of the low frequency of NK cells in peripheral blood, relatively small numbers of NK cells have been delivered in clinical trials.
The inability to propagate NK cells in vitro has been the limiting factor for generating sufficient cell numbers for optimal clinical outcome. Some expansion of NK cells (5-10 fold over 1-2 weeks) has be achieved through high-dose IL-2 alone 2. Activation of autologous T cells can mediate NK cell expansion, presumably also through release of local cytokine 3. Support with mesenchymal stroma or artificial antigen presenting cells (aAPCs) can support the expansion of NK cells from both peripheral blood and cord blood 4. Combined NKp46 and CD2 activation by antibody-coated beads is currently marketed for NK cell expansion (Miltenyi Biotec, Auburn CA), resulting in approximately 100-fold expansion in 21 days.
Clinical trials using aAPC-expanded or -activated NK cells are underway, one using leukemic cell line CTV-1 to prime and activate NK cells5 without significant expansion. A second trial utilizes EBV-LCL for NK cell expansion, achieving a mean 490-fold expansion in 21 days6. The third utilizes a K562-based aAPC transduced with 4-1BBL (CD137L) and membrane-bound IL-15 (mIL-15)7, which achieved a mean NK expansion 277-fold in 21 days. Although, the NK cells expanded using K562-41BBL-mIL15 aAPC are highly cytotoxic in vitro and in vivo compared to unexpanded NK cells, and participate in ADCC, their proliferation is limited by senescence attributed to telomere shortening8. More recently a 350-fold expansion of NK cells was reported using K562 expressing MICA, 4-1BBL and IL159.
Our method of NK cell expansion described herein produces rapid proliferation of NK cells without senescence achieving a median 21,000-fold expansion in 21 days.

Here we describe a method to efficiently expand and purify large numbers of human NK cells and assess their function.

Natural killer (NK) cells play an important role in immune surveillance against a variety of infectious microorganisms and tumors. Limited availability of ...

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes. The majority of them belong to the superfamily of G protein-coupled receptors (GPCRs). We have focused on characterizing arthropod kinin receptors from the tick and mosquito. Arthropod kinins are multifunctional neuropeptides with myotropic, diuretic, and neurotransmitter function. Here, a method for systematic analyses of structure-activity relationships of insect kinins on two heterologous kinin receptor-expressing systems is described. We provide important information relevant to the development of biostable kinin analogs with the potential to disrupt the diuretic, myotropic, and/or digestive processes in ticks and mosquitoes.
The kinin receptors from the southern cattle tick, Boophilus microplus (Canestrini), and the mosquito Aedes aegypti (Linnaeus), were stably expressed in the mammalian cell line CHO-K1. Functional analyses of these receptors were completed using a calcium bioluminescence plate assay that measures intracellular bioluminescence to determine cytoplasmic calcium levels upon peptide application to these recombinant cells. This method takes advantage of the aequorin protein, a photoprotein isolated from luminescent jellyfish. We transiently transfected the aequorin plasmid (mtAEQ/pcDNA1) in cell lines that stably expressed the kinin receptors. These cells were then treated with the cofactor coelenterazine, which complexes with intracellular aequorin. This bond breaks in the presence of calcium, emitting luminescence levels indicative of the calcium concentration. As the kinin receptor signals through the release of intracellular calcium, the intensity of the signal is related to the potency of the peptide.
This protocol is a synthesis of several previously described protocols with modifications; it presents step-by-step instructions for the stable expression of GPCRs in a mammalian cell line through functional plate assays (Staubly et al., 2002 and Stables et al., 1997). Using this methodology, we were able to establish stable cell lines expressing the mosquito and the tick kinin receptors, compare the potency of three mosquito kinins, identify critical amino acid positions for the ligand-receptor interaction, and perform semi-throughput screening of a peptide library. Because insect kinins are susceptible to fast enzymatic degradation by endogenous peptidases, they are severely limited in use as tools for pest control or endocrinological studies. Therefore, we also tested kinin analogs containing amino isobutyric acid (Aib) to enhance their potency and biostability. This peptidase-resistant analog represents an important lead in the development of biostable insect kinin analogs and may aid in the development of neuropeptide-based arthropod control strategies.

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito Aedes aegypti and Tick Rhipicephalus microplus G Protein-coupled Receptors

This protocol provides instructions for clonal-cell line selection and a calcium bioluminescence assay to analyze the structure-activity relationships of synthesized arthropod neuropeptides on their cognate GPCRs. This assay can be used for receptor deorphanization and structure-activity relationship studies for synthetic analog design and peptide/drug-lead discovery.

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes.  The ...

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

A Simple and Efficient Method to Detect Nuclear Factor Activation in Human Neutrophils by Flow Cytometry

Neutrophils are the most abundant leukocytes in peripheral blood. These cells are the first to appear at sites of inflammation and infection, thus becoming the first line of defense against invading microorganisms. Neutrophils possess important antimicrobial functions such as phagocytosis, release of lytic enzymes, and production of reactive oxygen species. In addition to these important defense functions, neutrophils perform other tasks in response to infection such as production of proinflammatory cytokines and inhibition of apoptosis. Cytokines recruit other leukocytes that help clear the infection, and inhibition of apoptosis allows the neutrophil to live longer at the site of infection. These functions are regulated at the level of transcription. However, because neutrophils are short-lived cells, the study of transcriptionally regulated responses in these cells cannot be performed with conventional reporter gene methods since there are no efficient techniques for neutrophil transfection. Here, we present a simple and efficient method that allows detection and quantification of nuclear factors in isolated and immunolabeled nuclei by flow cytometry. We describe techniques to isolate pure neutrophils from human peripheral blood, stimulate these cells with anti-receptor antibodies, isolate and immunolabel nuclei, and analyze nuclei by flow cytometry. The method has been successfully used to detect NF-&kappa;B and Elk-1 nuclear factors in nuclei from neutrophils and other cell types. Thus, this method represents an option for analyzing activation of transcription factors in isolated nuclei from a variety of cell types.

Neutrophils are the most abundant leukocytes in blood. Neutrophils possess transcriptionally regulated functions such as production of proinflammatory cytokines and inhibition of apoptosis. These functions can be studied with the method presented here, which allows detection and quantification of nuclear factors by flow cytometry in isolated nuclei

Neutrophils  are the most abundant leukocytes in peripheral blood. These cells are the first  to appear at sites of inflammation and infection, thus ...

A Microplate Assay to Assess Chemical Effects on RBL-2H3 Mast Cell Degranulation: Effects of Triclosan without Use of an Organic Solvent

Mast cells play important roles in allergic disease and immune defense against parasites. Once activated (e.g. by an allergen), they degranulate, a process that results in the exocytosis of allergic mediators. Modulation of mast cell degranulation by drugs and toxicants may have positive or adverse effects on human health. Mast cell function has been dissected in detail with the use of rat basophilic leukemia mast cells (RBL-2H3), a widely accepted model of human mucosal mast cells3-5. Mast cell granule component and the allergic mediator &#946;-hexosaminidase, which is released linearly in tandem with histamine from mast cells6, can easily and reliably be measured through reaction with a fluorogenic substrate, yielding measurable fluorescence intensity in a microplate assay that is amenable to high-throughput studies1. Originally published by Naal et al.1, we have adapted this degranulation assay for the screening of drugs and toxicants and demonstrate its use here.
Triclosan is a broad-spectrum antibacterial agent that is present in many consumer products and has been found to be a therapeutic aid in human allergic skin disease7-11, although the mechanism for this effect is unknown. Here we demonstrate an assay for the effect of triclosan on mast cell degranulation. We recently showed that triclosan strongly affects mast cell function2. In an effort to avoid use of an organic solvent, triclosan is dissolved directly into aqueous buffer with heat and stirring, and resultant concentration is confirmed using UV-Vis spectrophotometry (using &#949;280 = 4,200 L/M/cm)12. This protocol has the potential to be used with a variety of chemicals to determine their effects on mast cell degranulation, and more broadly, their allergic potential.

Mast cell degranulation, the release of allergic mediators, is important in allergy, asthma, and parasite defense. Here we demonstrate techniques1 for assessing effects of drugs and toxicants on degranulation, methodology recently utilized to exhibit the powerful inhibitory effect of antibacterial agent triclosan2.

Mast cells play important roles in allergic disease and immune defense against parasites. Once activated (e.g. by an allergen), they degranulate, a ...

Activation and Measurement of NLRP3 Inflammasome Activity Using IL-1&#946; in Human Monocyte-derived Dendritic Cells

Inflammatory processes resulting from the secretion of Interleukin (IL)-1 family cytokines by immune cells lead to local or systemic inflammation, tissue remodeling and repair, and virologic control1,2 . Interleukin-1&#946; is an essential element of the innate immune response and contributes to eliminate invading pathogens while preventing the establishment of persistent infection1-5.
Inflammasomes are the key signaling platform for the activation of interleukin 1 converting enzyme (ICE or Caspase-1). The NLRP3 inflammasome requires at least two signals in DCs to cause IL-1&#946; secretion6. Pro-IL-1&#946; protein expression is limited in resting cells; therefore a priming signal is required for IL-1&#946; transcription and protein expression. A second signal sensed by NLRP3 results in the formation of the multi-protein NLRP3 inflammasome. The ability of dendritic cells to respond to the signals required for IL-1&#946; secretion can be tested using a synthetic purine, R848, which is sensed by TLR8 in human monocyte derived dendritic cells&#160;(moDCs) to prime cells, followed by activation of the NLRP3 inflammasome with the bacterial toxin and potassium ionophore, nigericin.
Monocyte derived DCs&#160;are easily produced in culture and provide significantly more cells than purified human myeloid DCs. The method presented here differs from other inflammasome assays in that it uses in vitro human, instead of mouse derived, DCs thus allowing for the study of the inflammasome in human disease and infection.

Activation and Measurement of NLRP3 Inflammasome Activity Using IL-1&amp;#946; in Human Monocyte-derived Dendritic Cells

Dendritic cells (DCs)&#160;secrete IL-1&#946; in response to TLR8 recognition of synthetic purine, R848, followed by NLRP3 inflammasome activation with nigericin, therefore, IL-1&#946; can be used to measure NLRP3 inflammasome activity. Intracellular cytokine staining, immunoblotting, and ELISA are used to accurately measure NLRP3 inflammasome priming and activation via IL-1&#946; expression.

Inflammatory processes resulting from the secretion of Interleukin (IL)-1 family cytokines by immune cells lead to local or systemic inflammation, tissue ...

Culture of Macrophage Colony-stimulating Factor Differentiated Human Monocyte-derived Macrophages

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. For initiation of experiments, fresh or frozen monocytes are cultured in flasks for 1 week with M-CSF to induce their differentiation into macrophages. Then, the macrophages can be harvested and seeded into culture wells at required cell densities for carrying out experiments. The use of defined numbers of macrophages rather than defined numbers of monocytes to initiate macrophage cultures for experiments yields macrophage cultures in which the desired cell density can be more consistently attained. Use of cryopreserved monocytes reduces dependency on donor availability and produces more homogeneous macrophage cultures.

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. The protocol utilizes cryopreservation of monocytes coupled with their bulk differentiation into macrophages. Then harvested macrophages can then be seeded into culture wells at required cell densities for carrying out experiments.

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. For initiation ...

Measuring Granulocyte and Monocyte Phagocytosis and Oxidative Burst Activity in Human Blood

The granulocyte and monocyte phagocytosis and oxidative burst (OB) activity assay can be used to study the innate immune system. This manuscript provides the necessary methodology to add this assay to an exercise immunology arsenal. The first step in this assay is to prepare two aliquots ("H" and "F") of whole blood (heparin). Then, dihydroethidium is added to the H aliquot, and both aliquots are incubated in a warm water bath followed by a cold water bath. Next, Staphylococcus aureus (S. aureus) is added to the H aliquot and fluorescein isothiocyanate-labeled S. aureus is added to the F aliquot (bacteria:phagocyte = 8:1), and both aliquots are incubated in a warm water bath followed by a cold water bath. Then, trypan blue is added to each aliquot to quench extracellular fluorescence, and the cells are washed with phosphate-buffered saline. Next, the red blood cells are lysed, and the white blood cells are fixed. Finally, a flow cytometer and appropriate analysis software are used to measure granulocyte and monocyte phagocytosis and OB activity. This assay has been used for over 20 years. After heavy and prolonged exertion, athletes experience a significant but transient increase in phagocytosis and an extended decrease in OB activity. The post-exercise increase in phagocytosis is correlated with inflammation. In contrast to normal weight individuals, granulocyte and monocyte phagocytosis is chronically elevated in overweight and obese participants, and is modestly correlated with C-reactive protein. In summary, this flow cytometry-based assay measures the phagocytosis and OB activity of phagocytes and can be used as an additional measure of exercise- and obesity-induced inflammation.

The purpose of this manuscript is to present a method for measuring monocyte and granulocyte phagocytosis and oxidative burst activity in human blood samples.

The granulocyte and monocyte phagocytosis and oxidative burst (OB) activity assay can be used to study the innate immune system. This manuscript provides ...

Measurement of In Vitro Integration Activity of HIV-1 Preintegration Complexes

HIV-1 envelope proteins engage cognate receptors on the target cell surface, which leads to viral-cell membrane fusion followed by the release of the viral capsid (CA) core into the cytoplasm. Subsequently, the viral Reverse Transcriptase (RT), as part of a namesake nucleoprotein complex termed the Reverse Transcription Complex (RTC), converts the viral single-stranded RNA genome into a double-stranded DNA copy (vDNA). This leads to the biogenesis of another nucleoprotein complex, termed the pre-integration complex (PIC), composed of the vDNA and associated virus proteins and host factors. The PIC-associated viral integrase (IN) orchestrates the integration of the vDNA into the host chromosomal DNA in a temporally and spatially regulated two-step process. First, the IN processes the 3&#39; ends of the vDNA in the cytoplasm and, second, after the PIC traffics to the nucleus, it mediates integration of the processed vDNA into the chromosomal DNA. The PICs isolated from target cells acutely infected with HIV-1 are functional in vitro, as they are competent to integrate the associated vDNA into an exogenously added heterologous target DNA. Such PIC-based in vitro integration assays have significantly contributed to delineating the mechanistic details of retroviral integration and to discovering IN inhibitors. In this report, we elaborate upon an updated HIV-1 PIC assay that employs a nested real-time quantitative Polymerase Chain Reaction (qPCR)-based strategy for measuring the in vitro integration activity of isolated native PICs.

Measurement of In Vitro Integration Activity of HIV-1 Preintegration Complexes

This report describes an in vitro assay for measuring the human immunodeficiency virus type 1 (HIV-1) preintegration complex (PIC)-associated integration activity in the cytoplasmic extracts of acutely infected cells. The integration of PIC-associated HIV-1 DNA into heterologous target DNA is quantified by using a nested real-time polymerase chain reaction strategy.

HIV-1 envelope proteins engage cognate receptors on the target cell surface, which leads to viral-cell membrane fusion followed by the release of the ...

Methods Article

Measurement of Factor V Activity in Human Plasma Using a Microplate Coagulation Assay