This work was supported by a grant from the Italian Ministry of Health (Ricerca Corrente 2023 to M.C.).

The protocol was approved by the Ethical Committee of the Institution (no.70/24, CE 18.06.24), and written informed consent was obtained from all blood donors. The protocol will guide the user through seven main steps (Figure 1). The first step addresses the procedure for performing blood sampling. Steps 2 and 3 outline the antibody titration protocol and sample preparation for the analysis of intracellular TF expression using a phycoerythrin (PE)-labeled &#945;CD142/HTF1 monoclonal antibody (mAb). Step 4 describes the protocol for evaluating the protein exposed on the platelet surface using an indirect staining method with &#945;CD142/HTF1 mAb. Due to the steric hindrance of the fluorochrome, the PE-labeled mAb used for intracellular staining does not bind to the catalytic domain of the surface-exposed protein. Therefore, a protocol for indirect staining with an Alexa Fluor 633-labeled secondary mAb is proposed. Steps 5 and 6 detail the phases of flow cytometry analysis. The final step outlines the procedure for preparing platelet-rich plasma (PRP) to avoid the loss of the large size&#160;platelet subpopulation. Details of the reagents and equipment used are listed in the Table of Materials.
1. Blood withdrawal
NOTE: Human blood collection should be performed only after obtaining Ethical Committee approval.
<ol>
	<li>Collect blood samples by peripheral venous or needle-cannula sampling from an arm not affected by peripheral venous infusions, preferably from the antecubital vein.
	<ol>
		<li>For peripheral venous sampling, apply the tourniquet and, after inserting a G19 butterfly needle into the vein, remove it to prevent platelet activation (Figure 2). 
		NOTE: Do not use a butterfly needle smaller than G21 to avoid platelet activation.</li>
		<li>For venous sampling with a needle-cannula, do not apply the tourniquet at any time during sampling. If necessary, wash the sample with 0.9% saline solution before taking it. Always discard the first 5 mL of blood before starting to fill tubes.</li>
	</ol>
	</li>
	<li>Draw 2 mL of blood into an EDTA vacutainer (to be used for blood cell count). Gently mix the tube three times to ensure complete mixing of blood and anticoagulant.</li>
	<li>Draw blood into a 0.129 M sodium citrate vacutainer for the analysis of platelet-associated TF expression. Carefully fill the vacutainer to the indicated level to maintain the correct blood/anticoagulant ratio. Gently invert the tubes five times to ensure thorough mixing of the blood with the anticoagulant, preventing clot formation and hemolysis of the samples. 
	NOTE: The use of 0.129 M citrate anticoagulant is recommended to optimize the detection of platelet-associated TF with the HTF1 clone of the &#945;CD142 mAb. If it is necessary to isolate platelets from whole blood, draw a second 0.129 M sodium citrate vacutainer for platelet-rich plasma (PRP) preparation.</li>
	<li>Keep vacutainers upright in a tube holder. Do not shake them on rotating or tilting plates, and avoid pneumatic conveying systems. Do not refrigerate the blood samples to prevent platelet activation. 
	NOTE: Blood samples should preferably be processed within 15 min after withdrawal if another granule-associated platelet-activation marker is to be measured along with TF. Otherwise, samples should be processed no later than 3 h from collection (Figure 3).</li>
</ol>
2. Titration of the anti-TF antibody (for intracellular TF evaluation)
NOTE: Determination of intracellularly stored TF is performed using the PE-&#945;CD142/HTF1 mAb.
<ol>
	<li>Antibody titration using beads 
	NOTE: The use of beads allows for defining the correct amount of antibody to be used (i.e., the antibody concentration producing the greatest separation of fluorescence peaks between positive and negative samples). This analysis is performed using two samples, one truly positive and one truly negative. The antibody concentration is not reported by the manufacturer (see Table of Materials), so antibody titration is performed using five dilution points.
	<ol>
		<li>Centrifuge the antibody at 17,000 x g for 5 min at 4 &#176;C. To prepare the five antibody dilution points, pipette 40 &#181;L of antibody into tube 1 and 20 &#181;L of PBS into tubes 2 through 5. Make four consecutive serial 1:2 dilutions by transferring 20 &#181;L from tube 1 to tube 2 and then onwards.</li>
		<li>Prepare one test tube for each antibody volume to test. Add one drop of negative beads and one drop of positive beads to each test tube. Add 20 &#181;L of each antibody dilution to the test tube and vortex immediately. Incubate in the dark at room temperature for 20 min.</li>
		<li>Add 1 mL of 1x PBS without Ca++ and Mg++, pH 7.4, to each tube and vortex.</li>
		<li>Acquire the samples on the flow cytometer according to the manufacturer&#39;s instructions and record a data file of 10,000 events in the &#34;beads&#34; gate. The optimal antibody concentration is the one that provides the greatest separation between positive and negative peaks.</li>
	</ol>
	</li>
	<li>Antibody titration on platelets 
	NOTE: As a rule, the optimal antibody concentration identified using the beads should be verified on the cells to be stained-in this case, platelets in whole blood.
	<ol>
		<li>Fixation
		<ol>
			<li>Gently invert the sodium citrate vacutainer five times to mix the blood and anticoagulant thoroughly. Transfer 50 &#181;L of blood into a 1.5 mL tube, add 950 &#181;L of 1% paraformaldehyde (PFA), gently mix, and incubate the sample for 90 min at room temperature.</li>
			<li>Centrifuge the sample at 1,500 x g for 5 min, with brake, at room temperature.</li>
			<li>Remove the supernatant and resuspend the pellet by gently pipetting in 1 mL of 1x PBS without Ca++ and Mg++, pH 7.4.</li>
		</ol>
		</li>
		<li>Permeabilization
		<ol>
			<li>Prepare one tube for each antibody volume to be tested (test the volume identified with the beads titration along with a higher and a lower dilution, e.g., 10 &#181;L, 5 &#181;L, 2.5 &#181;L if the identified volume is 5 &#181;L) and one tube for the Fluorescence Minus One (FMO) control.</li>
			<li>Dispense 100 &#181;L of fixed blood into each tube and centrifuge them at 1,500 x g for 5 min, with brake, at room temperature.</li>
			<li>Remove the supernatant, resuspend the pellet in 100 &#181;L of 0.1% Triton-PBS to permeabilize the samples, and incubate for 10 min at room temperature.</li>
		</ol>
		</li>
		<li>Staining
		<ol>
			<li>Centrifuge the antibody at 17,000 x g for 5 min at 4 &#176;C. Add the required antibody volumes to each test tube as determined in 2.2.2.1 and mix.</li>
			<li>Incubate the samples for 15 min in the dark at room temperature.</li>
			<li>Add 300 &#181;L of 1x PBS without Ca++ and Mg++, pH 7.4, to each tube and mix. Acquire the samples on the flow cytometer (see step 5).</li>
			<li>Calculate the stain index according to the following formula15: 
			&#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160;<img alt="Equation 1" src="/files/ftp_upload/67356/67356eq01.jpg" style="margin: auto;" /> 
			where, MFIpos is the Mean Fluorescence Intensity of the positive events, MFIneg is that of the negative events, and SD is the Standard Deviation of the negative population.</li>
			<li>Choose the lowest antibody concentration that provides the highest stain index. 
			NOTE: In the current experimental conditions, the highest stain index for the PE-&#945;CD142/HTF1 mAb is achieved using 5 &#181;L of antibody.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Sample preparation for flow cytometry analysis of intracellular TF expression
<ol>
	<li>Whole blood fixation and permeabilization
	<ol>
		<li>For intracellular TF analysis, fix and permeabilize whole blood according to the procedure outlined in step 2.</li>
	</ol>
	</li>
	<li>Staining procedure 
	NOTE: In the following protocol, the platelet population is identified using &#945;CD61-PerCP staining. Other population markers may be used at the operator&#39;s discretion.
	<ol>
		<li>Prepare one tube for the Fluorescence Minus One (FMO) control and one tube for TF staining. Add 100 &#181;L of fixed and permeabilized whole blood to each tube along with the antibodies listed in Table 1.</li>
		<li>Incubate the samples for 15 min in the dark at room temperature.</li>
		<li>Add 300 &#181;L of 1x PBS without Ca++ and Mg++, pH 7.4, to each tube and mix. Store the stained samples in the dark until flow cytometry analysis (see step 5). 
		NOTE: To calculate the compensation matrix, prepare the samples as described in Table 2.</li>
	</ol>
	</li>
</ol>
4. Sample preparation for flow cytometry analysis of cell surface-associated TF expression in resting and activated platelets
NOTE: For the detection of cell surface-associated TF, use unlabelled &#945;CD142/HTF1 mAb (1 mg/mL) and Alexa Fluor 633 goat anti-mouse IgG as the secondary antibody for signal detection. Adenosine 5&#39;-diphosphate (ADP) is used as a platelet agonist to induce the exposure of TF on the cell surface. A PE-labelled antibody is used as a platelet population marker to minimize spillover, which will be checked by signal compensation.
<ol>
	<li>TF staining
	<ol>
		<li>Dilute &#945;CD142/HTF1 mAb 1:100 in 1x PBS without Ca++ and Mg++, pH 7.4, to obtain a working concentration of 10 &#181;g/mL. Dilute Alexa Fluor 633 goat anti-mouse IgG 1:10 in 1x PBS without Ca++ and Mg++, pH 7.4.</li>
		<li>Prepare the three sample tubes and dispense 1x PBS without Ca++ and Mg++, pH 7.4, as indicated in Table 3.</li>
		<li>Dispense 7.5 &#181;L of diluted &#945;CD142/HTF1 mAb and add 5 &#181;L of ADP (200 &#181;M). Gently invert the citrate vacutainer 5 times to mix the blood and anticoagulant thoroughly, then dispense 5 &#181;L of whole blood into each tube. Gently mix the samples.</li>
		<li>Incubate the samples for 20 min at room temperature.</li>
	</ol>
	</li>
	<li>Sample fixation
	<ol>
		<li>Fix each sample by adding 300 &#181;L of 1% paraformaldehyde (PFA) and incubate the samples for 1 h at room temperature. 
		NOTE: Labelled samples fixed with 1% PFA and stored at 4 &#176;C remain stable for up to 4 days.</li>
		<li>Add 300 &#181;L of 1x PBS without Ca++ and Mg++, pH 7.4, to each tube and centrifuge at 1500 x g for 5 min with brake at room temperature.</li>
		<li>Remove the supernatant and resuspend the pellet in 90 &#181;L of 1x PBS without Ca++ and Mg++, pH 7.4, by gently pipetting.</li>
	</ol>
	</li>
	<li>Staining with the secondary antibody
	<ol>
		<li>Dispense 5 &#181;L of diluted Alexa Fluor 633 IgG and 5 &#181;L of &#945;CD41-PE into each tube (see Table 4). Incubate the samples for 15 min in the dark at room temperature.</li>
		<li>Add 300 &#181;L of 1x PBS without Ca++ and Mg++, pH 7.4, to each tube and mix.</li>
		<li>Store the stained samples in the dark until flow cytometry analysis (see step 6). 
		NOTE: To calculate the compensation matrix, prepare the samples according to Steps 4.1-4.3 and as outlined in Table 5 and Table 6.</li>
	</ol>
	</li>
</ol>
5. Flow cytometry analysis of intracellular TF expression
NOTE: The following procedure refers to acquisitions on a reference flow cytometer; thus, the reported settings and plots should be considered representative. The protocol can be applied to any high-sensitivity, new-generation flow cytometer. Switch on the flow cytometer well before sample acquisition to ensure laser stability. Thoroughly wash the fluidics and flow cell, as platelets are small, and any cell debris may interfere with the visualization of the platelet population in the forward scatter area/side scatter area (FSC-A/SSC-A) dot plot. Perform quality control with QC beads according to the manufacturer&#39;s instructions and wash the fluidics again to remove any residual beads.
<ol>
	<li>Acquisition setup
	<ol>
		<li>Create an FSC-A/SSC-A dot plot on a log scale displaying All Events (Figure 4A). Create a CD61-A/SSC-A dot plot on a log scale displaying All Events (Figure 4B).</li>
		<li>Adjust the PE (Y-585) and PerCP (B690) gain according to daily quality control. Set the threshold approximately to 2000 on SSC-H and to 1000 on PerCP (B690-H) to visualize the platelet population. Set the flow rate to LOW.</li>
		<li>Draw a region between 103&#160;and 105 called &#34;CD61+&#34; to identify the platelet population.</li>
		<li>Acquire single-stain controls to calculate the compensation matrix and then acquire the samples. Record a data file of 10,000 events within the &#34;CD61+&#34; gate (Figure 4D,E). Start recording 5-10 s after the beginning of the sample acquisition.</li>
	</ol>
	</li>
</ol>
6. Flow cytometry analysis of surface TF expression
<ol>
	<li>Make a&#160;FSC-A/SSC-A dot plot on a log scale displaying All Events (Figure 5A), and a CD41-A/SSC-A dot plot on a log scale displaying All Events (Figure 5B).</li>
	<li>Adjust the Alexa Fluor 633 gain and the PE (Y-585) gain according to daily quality control, set the threshold approximately to 1000 on SSC-H and to 2500 on PE (Y585-H) to visualize the platelet population as shown in Figure 5B. Set the flow rate to LOW.</li>
	<li>Draw a region between 104 and 105 called &#34;CD41+&#34; to identify the platelet population.</li>
	<li>Acquire single-stain controls to calculate the compensation matrix and then acquire the samples. Record a data file of 10,000 events within the &#34;CD41+&#34; gate (Figure 5D,E). Start recording 5-10 s after the beginning of the sample acquisition.</li>
</ol>
7. Analysis of TF-positive platelets in platelet-rich plasma
<ol>
	<li>Platelet-rich plasma (PRP) preparation 
	NOTE: Platelet isolation must be carried out at room temperature to prevent cell activation. Ensure that the temperature of the reagents and instruments is between 18 &#176;C and 22 &#176;C. Avoid fast pipetting and vigorous shaking throughout the procedure.
	<ol>
		<li>Gently mix the 0.129 M sodium citrate vacutainer 5 times and measure the platelet count twice on a blood cell counter to calculate the percentage of recovered platelets. 
		NOTE: Use the mean value of the two determinations as the platelet concentration.</li>
		<li>Centrifuge the 0.129 M sodium citrate vacutainer at 100 x g for 10 min, without brake, at room temperature (Figure 6). Collect the obtained PRP, transfer it to a 10 mL polypropylene tube, and record the volume collected. 
		NOTE: Do not use polystyrene tubes to avoid platelet activation.</li>
		<li>Count platelets twice in PRP using a blood cell counter and calculate the percentage of platelet recovery as follows: 
		<img alt="Equation 2" src="/files/ftp_upload/67356/67356eq02.jpg" style="margin: auto;" /> 
		&#8203;NOTE: If the recovery is lower than 65%, the platelet population is not fully representative of that in whole blood. Consider discarding the preparation.</li>
	</ol>
	</li>
	<li>Sample preparation for flow cytometry analysis of intracellular and surface-associated TF expression in PRP
	<ol>
		<li>To analyze intracellular and surface-associated TF expression in PRP, use the same protocol described for whole blood analysis (see steps 3-6).</li>
	</ol>
	</li>
</ol>

Assessing Intracellular and Surface TF-Positive Platelets via Flow Cytometry

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The authors declare no conflict of interest.

This manuscript reports detailed protocols for the evaluation of TF-positive platelets by whole blood flow cytometry, taking into account the analysis of the protein expressed both within and on the platelet surface.
During the past 20 years, the presence of TF in human platelets has been well established1,2,3,4,5,17,18. A subset of platelets, accounting for 20%-30% of the total circulating platelet population2, expresses the protein, reinforcing the concept of the platelet heterogeneity that is becoming increasingly relevant in the field19,20. It is worth mentioning that the TF-positive platelets have a high mean platelet volume, as previously shown by highly sensitive imaging flow cytometry2. It is of crucial importance to consider this feature, especially when TF expression has to be analyzed in platelets isolated from the other blood cell components. It is possible, indeed, that the centrifugation step used for platelet isolation may result in the loss of large-size platelets2. This pre-analytical issue is probably one of the main drivers of the controversy - which lasted until a few years ago - on the presence of TF in platelets7,12,13. Whole blood flow cytometry is an extremely useful approach in this regard, allowing the analysis of all platelet subsets without the need to isolate them.
A second reason that fostered the discussion on the presence of TF in platelets was the choice of the anti-human TF antibody used for the analysis. There are many commercially available antibodies that have been used for platelet-associated TF evaluation and that recognize different epitopes of the protein. Some antibodies, such as the TF9-10H10 3,21,22 and the VIC71,3 clones, bind to a non-functional region of TF without affecting FVII/FVIIa binding. Other antibodies, including the VD81,2,3 and the HTF118 clones -the latter used in the present protocols- recognize the catalytic site of the protein, competing with FVII/FVIIa for binding to TF, thus blocking its procoagulant function. It is important to note that, depending on the glycosylation state of the extracellular domain and on the conformation of the catalytic site of TF23, which could be modified in particular conditions (such as cell activation or disease), some of these antibodies could be differentially able to recognize, by flow cytometry, the surface expression of platelet-associated TF. Moreover, also the staining conditions, i.e., whether the sample to be analyzed is fresh, fixed and/or permeabilized, could affect the epitope conformation and, thus, the antigen-antibody binding14.
A further consideration that deserves attention is the choice of fluorochrome bound to the antibody. Since TF is expressed by only 20%-30% of the platelet population, low-emission fluorochromes, together with the use of low-sensitivity flow cytometers, may not be suitable to discriminate low positive signals. For this reason, antibodies labeled with bright fluorochromes, such as PE or APC, are to be preferred. It is important to note, however, that the large size of a fluorochrome such as PE can hinder the binding of the labeled antibody to the protein24. In order to overcome this issue, in the present protocol, the surface TF staining was performed as an indirect labeling with an unconjugated &#945;CD142/HTF1 mAb, then revealed through the use of a secondary antibody. The PE-steric hindrance does not affect &#945;CD142/HTF1 mAb binding to TF stored intracellularly where the catalytic site is more accessible to antibody binding. It is clear that assessment of the surface expression of platelet-associated TF by direct labeling, compared to the indirect one, would be preferable -if possible- to shorten the sample processing time. This approach, however, must take into account the critical issues mentioned above and related to the binding capacity of the several commercially available antibodies to platelet TF, due to the conformation of the protein itself, or to the staining conditions, as well as the steric hindrance of the different fluorochromes used. Therefore, in order to avoid false negative results, direct labeling of surface-expressed platelet-associated TF requires a mandatory careful optimization process. These aspects are the subject of an ongoing study aimed at evaluating the efficiency of direct staining with the different commercially available antibody clones labeled with all-in-use fluorochromes.
Several studies showed that the expression of TF is elevated in various pathological conditions characterized by a high thrombotic risk8,9,10,11,17. Recently, evidence has been provided that levels of TF-positive platelets, accurately quantified by flow cytometry analysis, can identify cardiovascular patients who, despite an optimal response to drug treatment with aspirin and P2Y12 antagonists, maintain a high thrombotic risk25. Indeed, platelet-associated TF is functionally active and thus potentially able to support thrombin generation1,3,4. In this context, the ability to perform flow cytometry analysis on a few microliters of blood is a significant advantage that allows platelet-associated TF analysis to be extended to the clinical setting. To this end, it is worth mentioning that the validation of platelet-associated TF measurement by flow cytometry has been the aim of an international project we recently coordinated, with the support of the International Society of Haemostasis and Thrombosis, to provide methodological guidance for its assessment.
Unlike other classical markers of platelet activation, such as P-selectin and platelet-leukocyte aggregates, which need to be analyzed immediately after blood sampling to avoid artifacts, TF expression is much more stable. In fact, the blood sample can be processed within a few hours after blood withdrawal without any significant change in platelet-TF expression; furthermore, the possibility of fixing the sample and labeling/processing it afterward makes this analysis suitable for many hospitals/laboratories. It is noteworthy that, although the protocol described in this manuscript is relatively simple to perform, the flow cytometry technique still requires specialized technical training. For this reason, technology is moving towards the development of increasingly simple cytometers intended for point-of-care assays.
In conclusion, the proposed protocol provides methods for the correct analysis of platelet-associated TF, emphasizing the advantages of a flow cytometry analysis-based technique for its determination. In addition, it highlights the pre-analytical variables that can influence the analysis of this protein, emphasizing the measures to be taken to avoid them.

Tissue Factor (TF), a transmembrane glycoprotein and key activator of the blood coagulation cascade, is expressed by a subset, ~20%-30%, of circulating human platelets1, particularly those with larger size2. Upon activation by classical platelet agonists such as adenosine diphosphate (ADP) and thrombin, platelets expose functionally active TF on their surface in a concentration-dependent manner, allowing it to bind factor VII/VIIa, ultimately leading to thrombin generation1,3,4,5,6. Notably, in pathological conditions such as coronary artery disease (CAD)7, severe acute respiratory syndrome-coronavirus-2 (SARS-CoV-2) infection8, migraine with aura associated with patent foramen ovale (MHA-PFO)9, cancer10, and sepsis11, the number of TF-positive platelets significantly increases, contributing to the greater thrombotic risk observed in these clinical settings.
Despite this compelling evidence, the detection of TF-positive platelets remains a topic of debate within the scientific community6,12,13. Methodological challenges, including pre-analytical variables, sample preparation, and antibody specificity, have been suggested as possible reasons for the contradictory results reported in the literature2,4,12,14. Therefore, it is crucial to establish a standardized assay based on a well-defined protocol for measuring TF in human platelets.
In this context, whole blood flow cytometry is the only technology that allows for the analysis of TF-positive platelets with minimal sample handling, which is crucial to avoid the loss of this platelet subset.
In this article, we propose a protocol for measuring platelet-associated TF using whole blood flow cytometry, assessing the protein (1) intracellularly in resting conditions and (2) on the cell surface in both resting and activated conditions. Instructions are also provided if the evaluation of TF-positive platelets in platelet-rich plasma (PRP) is required.

<table><tbody><tr><td>Adenosine 5&amp;rsquo;-diphosphate (ADP)</td><td>Sigma-Aldrich</td><td>A-2754</td><td>&amp;nbsp;Platelets agonist</td></tr><tr><td>Alexa Fluor 633 goat &amp;alpha;-mouse IgG</td><td>Invitrogen</td><td>A21052</td><td>Secondary antibody</td></tr><tr><td>Biotube 10 mL</td><td>Biosigma</td><td>BSP048</td><td>For PRP collection</td></tr><tr><td>Butterfly Needle G19</td><td>PIC solution</td><td>02044019030010</td><td>For blood withdrawal</td></tr><tr><td>CytExpert Software</td><td>Beckman Coulter</td><td></td><td>Flow cytometry data analysis</td></tr><tr><td>Cytoflex S</td><td>Beckman Coulter</td><td></td><td>Flow cytometer</td></tr><tr><td>Eppendorf tubes 1.5 mL</td><td>Eppendorf</td><td>1,20,086</td><td>&amp;nbsp;Blood fixation</td></tr><tr><td>ImageStreamX Mk II</td><td>Cytek Biosciences</td><td></td><td>Imaging flow cytometer</td></tr><tr><td>Kaluza Software</td><td>Beckman Coulter</td><td></td><td>Flow cytometry data analysis&amp;nbsp;</td></tr><tr><td>Paraformaldehyde&amp;nbsp;</td><td>Antibioticos &amp;ndash; Divisione Carlo Erba Reagenti</td><td>387507</td><td>To fix blood and platelets</td></tr><tr><td>PerCP-&amp;alpha;CD45</td><td>BD</td><td>345809</td><td>Leukocyte population marker</td></tr><tr><td>PerCP-&amp;alpha;CD61</td><td>BD</td><td>347408</td><td>Platelet poulation marker</td></tr><tr><td>PE-&amp;alpha;CD142</td><td>BD</td><td>550312</td><td>Anti-human Tissue Factor antibody</td></tr><tr><td>PE-&amp;alpha;CD41</td><td>Beckman Coulter</td><td>A07781</td><td>Platelet poulation marker</td></tr><tr><td>Phosphate Buffer Saline (PBS) 1x w/o Calcium and Magnesium</td><td>Gibco</td><td>10010-015</td><td>For antibodies and sample dilution</td></tr><tr><td>Pipet tips 10 &amp;micro;L&amp;nbsp;&amp;nbsp;</td><td>Gilson&amp;nbsp;</td><td>F161630</td><td>To prepare samples</td></tr><tr><td>Pipet tips 1000 &amp;micro;L</td><td>Gilson</td><td>F161670</td><td>To prepare samples</td></tr><tr><td>Pipet tips 200 &amp;micro;L</td><td>Gilson</td><td>F161930</td><td>To prepare samples</td></tr><tr><td>Round-Bottom Tube 5 mL</td><td>Corning</td><td>352052</td><td>For flow cytometer acquisition</td></tr><tr><td>Sodium Chloride 0.9 %</td><td>BD</td><td>306575</td><td>To wash the needle-cannula after blood withdrawal</td></tr><tr><td>Sysmex XN-450</td><td>Dasit</td><td></td><td>Blood cell analyser</td></tr><tr><td>Triton X-100</td><td>Carlo Erba Reagenti</td><td>9002-93-1</td><td>To permeabilise blood cells membrane</td></tr><tr><td>Vacutainer citrate 0.129M tubes</td><td>BD</td><td>363079</td><td>For blood collection</td></tr><tr><td>Vacutainer K2 EDTA tubes</td><td>BD</td><td>368857</td><td>For blood collection</td></tr><tr><td>Vacutainer multiple sample luer adapter</td><td>BD</td><td>367300</td><td>&amp;nbsp;For blood withdrawal</td></tr><tr><td>Vacutainer one use holder</td><td>BD</td><td>364815</td><td>&amp;nbsp;For blood withdrawal</td></tr><tr><td>Versacomp Antibody Capture beads kit</td><td>Beckman Coulter</td><td>B22804</td><td>Antibody titration</td></tr><tr><td>&amp;alpha;CD142 (clone HTF-1)</td><td>Invitrogen</td><td>16-1429-82</td><td>Anti-human Tissue Factor antibody</td></tr></tbody></table>

flow cytometry analysis of tissue factor expression in human platelets

A subset of circulating human platelets stores Tissue Factor (TF) intracellularly, the key activator of the blood coagulation cascade and thrombus formation. Upon platelet activation, TF is exposed on the cell membrane, where it binds to FVII, ultimately leading to thrombin generation. Considering that (1) levels of TF-positive platelets increase in various clinical settings, contributing to the patient&#39;s prothrombotic phenotype, and (2) different drugs can modulate platelet-associated TF expression, a standardized method for assessing TF-positive platelets is valuable, as its evaluation has been controversial in the past. Here, we outline a protocol for measuring the percentage of TF-positive platelets using flow cytometry in whole blood and platelet-rich plasma (PRP)/washed platelets. This protocol aims to provide detailed instructions for quantifying the percentage of TF-positive platelets by assessing the protein (1) intracellularly in resting conditions, and (2) on the cell surface, in both resting and activated conditions. The first section provides essential information for correctly performing blood withdrawal to ensure that pre-analytical procedures do not affect the results. Next, the protocol focuses on sample preparation and labeling procedures for flow cytometry analysis. Detailed steps for cell stimulation, labeling, fixation, and permeabilization -- where necessary -- are outlined. Finally, instructions for flow cytometry settings to correctly identify the platelet population and analyze TF-positive events are described. Lastly, the method includes the procedure for preparing PRP if TF-positive platelets are to be measured in isolated platelets. Since only a subset of platelets contains TF, it is important to ensure that these platelets are not lost during the centrifugation steps required to obtain PRP.

A summary of platelet-associated TF expression analysis is described in the flow diagram shown in Figure 1. The steps include: (1) Blood Collection: Collect blood from the human peripheral vein, preferably from the antecubital vein, using a G19 butterfly needle or a needle-cannula without a tourniquet and with anticoagulant-containing vacutainers. (2) Sample Processing: Process the samples according to the type of TF evaluation to be performed. (3) Sample Labeling: Label the samples with specific antibodies. (4) Flow Cytometry Analysis: Analyze the samples to detect surface and intracellular TF.
During blood withdrawal, particular attention must be paid to avoid platelet activation. To this end, two precautions are observed: removing the tourniquet shortly after inserting the needle into the vein, and discarding the blood in the first vacutainer drawn. Without the use of the tourniquet, the platelet population, displayed in an FSC-A/SSC-A dot plot, has a typical shape, as shown in Figure 2A. Conversely, with the use of the tourniquet, the physical properties of the platelet population result in an elongated shape resembling the morphology observed following stimulation with a classical platelet agonist, such as ADP (Figure 2B,C).
Experiments performed to assess the stability of platelet-associated TF expression over time showed that it was stable for 3 h from blood collection (Figure 3). When combining the measurement of TF expression with that of a marker of platelet activation, such as P-selectin, it is advised to process the blood sample as soon as possible to avoid platelet degranulation.
For the analysis of intracellularly stored TF, whole blood is fixed in 1% PFA, permeabilized with PBS-Triton, and then stained with a PE-labeled antibody. The platelet population is identified on a logarithmic FSC-A/SSC-A plot (Figure 4 A,D) and by &#945;CD61 mAb staining (Figure 4 B,E). Results from the analysis of intracellular TF in healthy subjects (n = 377) indicated that TF is present within 26.8% &#177; 4.8% of circulating platelets (Figure 4F).
Flow cytometry analysis of surface-associated TF expression has been performed in resting conditions and after cell stimulation with ADP 10 &#181;M. The whole blood platelet population, labeled with &#945;CD41 mAb, has been identified on a logarithmic FSC-A/SSC-A plot (Figure 5A,B). Results showed that in healthy subjects (n = 377), TF is expressed by 2.8% &#177; 1.4% of resting platelets (Figure 5D-F) and increases up to 24.5% &#177; 5.8% upon activation (Figure 5G-I). TF is expressed by the platelet subset with the largest size (Figure 5D,G, green dots). Furthermore, analysis by imaging flow cytometry highlights that TF signal is associated only with &#945;CD61 positive and not with &#945;CD45 positive events (Figure 7).
As previously mentioned, TF is stored in a subset of platelets characterized by the largest size within the whole population. Therefore, it is important not to lose this platelet fraction during PRP preparation. Following the indications provided in the method section (step 7), it is possible to isolate a platelet population within PRP that is representative of that present in whole blood. As shown in Figure 6, the flow cytometry dot plot of the platelet population obtained by 100 x g whole blood centrifugation is very similar to that of whole blood in terms of FSC distribution (Figure 6 A,B). In contrast, using a higher force and longer centrifugation time, as often reported in the literature12,14,16, the recovered platelet population has a lower FSC compared to that of whole blood (Figure 6C), suggesting the loss of the larger size platelets, which are those highly TF-positive.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67356/67356fig01.jpg" /> 
Figure 1: Schematic diagram for platelet-associated TF expression analysis and platelet isolation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67356/67356fig02.jpg" /> 
Figure 2: Representative density plots of the platelet population analyzed by whole blood flow cytometry. (A) Platelet population analyzed in a blood sample drawn without a tourniquet or (B) with the tourniquet. (C) Platelet population stimulated with ADP is reported for comparison. WBC: White Blood Cells; RBC: Red Blood Cells. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67356/67356fig03.jpg" /> 
Figure 3: Stability of cell surface-associated TF expression over time. Platelet-associated TF expression of resting platelets was analyzed at 4 different time points after blood withdrawal reported in the x-axis as follows: T0 = immediately after blood collection; T1 = 1 h after blood collection; T2 = 2 h after blood collection; T3 = 3 h after blood collection. Data are reported as mean &#177; SD, n = 3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67356/67356fig04.jpg" /> 
Figure 4: Representative flow cytometry analysis of intracellularly stored TF. Platelet population (red) in whole blood is identified in the FSC/SSC-A dot plot (A,D) and by &#945;CD61 mAb staining (B,E). TF expression is analyzed using a PE- &#945;CD142/HTF1 mAb (green dots). Histogram plots are used to identify negative (C) and positive (F) events. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/67356/67356fig5v2.jpg" /> 
Figure 5: Representative flow cytometry analysis of surface-associated TF-positive platelets. Fresh whole blood is labeled with &#945;CD142/HTF1 mAb, then fixed with 1% PFA and labeled with the secondary mAb and the &#945;CD41 mAb. Platelet population (red) in whole blood is identified in the FSC/SSC-A dot plot (A,D,G) and by &#945;CD41 mAb staining (B,E,H). TF expression in resting condition (D-F) and after ADP stimulation (G-I) is quantified by labeling with &#945;CD142/HTF1 mAb (green dots). Representative histogram plots of negative (C) and positive (F,I) samples are reported. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/67356/67356fig06v2.jpg" /> 
Figure 6: Effect of different centrifugation forces on PRP preparation. (A-C) Flow cytometry analysis of the platelet population in whole blood, PRP prepared at 100 x g and at 350 x g. The reference range of platelet population on FSC-H/SSC-H is indicated by dotted lines.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig06largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/67356/67356fig07.jpg" /> 
Figure 7: Evaluation of TF expression in platelets and leukocytes by imaging flow cytometry. Representative channel images of brightfield (BF, grey), Tissue Factor (TF, green), &#945;CD61 (red), &#945;CD45 (grey), nuclei staining with Hoechst (magenta), and the composite image (merge), acquired at 60x magnification, are shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67356/67356fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample tube ID</td>
			<td>Fix/perm whole blood (&#181;L)</td>
			<td>&#945;CD142-PE (&#181;L)</td>
			<td>&#945;CD61-PerCP (&#181;L)</td>
		</tr>
		<tr>
			<td>FMO</td>
			<td>100</td>
			<td>-</td>
			<td>15</td>
		</tr>
		<tr>
			<td>TF</td>
			<td>100</td>
			<td>5</td>
			<td>15</td>
		</tr>
	</tbody>
</table>
Table 1:&#160;Staining protocol for flow cytometry analysis of intracellular TF expression.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample tube ID</td>
			<td>Fix/perm whole blood (&#181;L)</td>
			<td>&#945;CD142-PE (&#181;L)</td>
			<td>&#945;CD61-PerCP (&#181;L)</td>
		</tr>
		<tr>
			<td>Unstained</td>
			<td>100</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>&#945;CD142-PE</td>
			<td>95</td>
			<td>5</td>
			<td>-</td>
		</tr>
		<tr>
			<td>&#945;CD61-PerCP</td>
			<td>85</td>
			<td>-</td>
			<td>15</td>
		</tr>
	</tbody>
</table>
Table 2:&#160;Single stain control for compensation matrix of intracellular TF expression.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample tube ID</td>
			<td>PBS 1x (&#181;L)</td>
			<td>&#945;CD142 (&#181;L)</td>
			<td>ADP (&#181;L)</td>
			<td>Fresh whole blood (&#181;L)</td>
			<td>PFA 1% (&#181;L)</td>
		</tr>
		<tr>
			<td>FMO</td>
			<td>95</td>
			<td>-</td>
			<td>-</td>
			<td>5</td>
			<td>300</td>
		</tr>
		<tr>
			<td>Resting</td>
			<td>87.5</td>
			<td>7.5</td>
			<td>-</td>
			<td>5</td>
			<td>300</td>
		</tr>
		<tr>
			<td>ADP Activated</td>
			<td>82.5</td>
			<td>7.5</td>
			<td>5</td>
			<td>5</td>
			<td>300</td>
		</tr>
	</tbody>
</table>
Table 3:&#160;Staining protocol for flow cytometry analysis of surface TF expression.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample tube ID</td>
			<td>PBS 1x (&#181;L)</td>
			<td>Alexa Fluor 633 IgG (&#181;L)&#160;</td>
			<td>&#945;CD41-PE (&#181;L)</td>
		</tr>
		<tr>
			<td>FMO</td>
			<td>90</td>
			<td>5</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Resting</td>
			<td>90</td>
			<td>5</td>
			<td>5</td>
		</tr>
		<tr>
			<td>ADP Activated</td>
			<td>90</td>
			<td>5</td>
			<td>5</td>
		</tr>
	</tbody>
</table>
Table 4:&#160;Secondary antibody staining protocol for surface-associated TF analysis.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample tube ID</td>
			<td>PBS 1x (&#181;L)</td>
			<td>&#945;CD142 (&#181;L)&#160;</td>
			<td>ADP (&#181;L)</td>
			<td>Fresh whole blood (&#181;L)</td>
			<td>PFA 1% (&#181;L)</td>
		</tr>
		<tr>
			<td>Unstained</td>
			<td>90</td>
			<td>-</td>
			<td>5</td>
			<td>5</td>
			<td>300</td>
		</tr>
		<tr>
			<td>TF + Alexa Fluor 633 IgG</td>
			<td>82.5</td>
			<td>7.5</td>
			<td>5</td>
			<td>5</td>
			<td>300</td>
		</tr>
		<tr>
			<td>&#945;CD41-PE</td>
			<td>90</td>
			<td>-</td>
			<td>5</td>
			<td>5</td>
			<td>300</td>
		</tr>
	</tbody>
</table>
Table 5: Compensation matrix preparation for intracellular TF analysis.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Sample tube ID</td>
			<td>PBS 1x (&#181;L)</td>
			<td>Alexa Fluor 633 IgG (&#181;L)</td>
			<td>&#945;CD41-PE (&#181;L)</td>
		</tr>
		<tr>
			<td>Unstained</td>
			<td>100</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>TF + Alexa Fluor 633 IgG</td>
			<td>95</td>
			<td>5</td>
			<td>-</td>
		</tr>
		<tr>
			<td>&#945;CD41-PE</td>
			<td>95</td>
			<td>-</td>
			<td>5</td>
		</tr>
	</tbody>
</table>
Table 6: Compensation matrix preparation for surface-associated TF analysis.

Flow Cytometry Analysis of Tissue Factor Expression in Human Platelets

This article outlines the protocol for quantifying the percentage of Tissue Factor (TF)-positive platelets using whole blood flow cytometry, assessing the protein: (1) intracellularly in resting conditions, and (2) on the cell surface, in both resting and activated conditions. Guidance is also provided for evaluating TF-positive platelets in platelet-rich plasma.

A subset of circulating human platelets stores Tissue Factor (TF) intracellularly, the key activator of the blood coagulation cascade and thrombus ...

flow-cytometry-analysis-of-tissue-factor-expression-in-human-platelets

Research

JoVE Journal

Biology

395 次观看.  Centro Cardiologico Monzino IRCCS. This article outlines the protocol for quantifying the percentage of Tissue Factor (TF)-positive platelets using whole blood flow cytometry, assessing the protein: (1) intracellularly in resting conditions, and (2) on the cell surface, in both resting and activated conditions. Guidance is also provided for evaluating TF-positive platelets in platelet-rich plasma.

视频: Flow Cytometry Analysis of Tissue Factor Expression in Human Platelets

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

流动小鼠减数分裂细胞的流式细胞仪净化

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

科研

生物学

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

流式细胞仪分离纯化<em&gt; S.酵母</em&gt;合子

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro and in vivo.1-5 Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
<ol>
	<li>stem and progenitor cell quiescence, proliferation and/or differentiation6-8</li>
	<li>antigen-driven membrane transfer9 and/or precursor cell proliferation3,4,10-18 and</li>
	<li>immune regulatory and effector cell function1,18-21.</li>
</ol>
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24.
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
<ol>
	<li>Failure to achieve bright, uniform, reproducible labeling. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.</li>
	<li>Suboptimal fluorochrome combinations and/or failure to include critical compensation controls. Tracking dye fluorescence is typically 102 - 103 times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.</li>
	<li>Failure to obtain a good fit with peak modeling software. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.</li>
</ol>
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.

Successful use of cell tracking dyes to monitor immune cell function and proliferation involves several critical steps. We describe methods for: 1) obtaining bright, uniform, reproducible label-ing with membrane dyes; 2) selecting fluorochromes and data acquisition conditions; and 3) choosing a model to quantify cell proliferation based on dye dilution.

优化细胞分裂监测细胞示踪染料的染色和扩散建模方法

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of ...

Analysis of Gene Function and Visualization of Cilia-Generated Fluid Flow in Kupffer's Vesicle

Internal organs such as the heart, brain, and gut develop left-right (LR) asymmetries that are critical for their normal functions1. Motile cilia are involved in establishing LR asymmetry in vertebrate embryos, including mouse, frog, and zebrafish2-6. These 'LR cilia' generate asymmetric fluid flow that is necessary to trigger a conserved asymmetric Nodal (TGF-&beta; superfamily) signaling cascade in the left lateral plate mesoderm, which is thought to provide LR patterning information for developing organs7. Thus, to understand mechanisms underlying LR patterning, it is essential to identify genes that regulate the organization of LR ciliated cells, the motility and length of LR cilia and their ability to generate robust asymmetric flow.
In the zebrafish embryo, LR cilia are located in Kupffer's vesicle (KV)2,4,5. KV is comprised of a single layer of monociliated epithelial cells that enclose a fluid-filled lumen. Fate mapping has shown that KV is derived from a group of ~20-30 cells known as dorsal forerunner cells (DFCs) that migrate at the dorsal blastoderm margin during epiboly stages8,9. During early somite stages, DFCs cluster and differentiate into ciliated epithelial cells to form KV in the tailbud of the embryo10,11. The ability to identify and track DFCs&mdash;in combination with optical transparency and rapid development of the zebrafish embryo&mdash;make zebrafish KV an excellent model system to study LR ciliated cells.
Interestingly, progenitors of the DFC/KV cell lineage retain cytoplasmic bridges between the yolk cell up to 4 hr post-fertilization (hpf), whereas cytoplasmic bridges between the yolk cell and other embryonic cells close after 2 hpf8. Taking advantage of these cytoplasmic bridges, we developed a stage-specific injection strategy to deliver morpholino oligonucleotides (MO) exclusively to DFCs and knockdown the function of a targeted gene in these cells12. This technique creates chimeric embryos in which gene function is knocked down in the DFC/KV lineage developing in the context of a wild-type embryo. To analyze asymmetric fluid flow in KV, we inject fluorescent microbeads into the KV lumen and record bead movement using videomicroscopy2. Fluid flow is easily visualized and can be quantified by tracking bead displacement over time.
Here, using the stage-specific DFC-targeted gene knockdown technique and injection of fluorescent microbeads into KV to visualize flow, we present a protocol that provides an effective approach to characterize the role of a particular gene during KV development and function.

Cilia-generated fluid flow in Kupffer&#x2019;s Vesicle (KV) controls left-right patterning of the zebrafish embryo. Here, we describe a technique to modulate gene function specifically in KV cells. In addition, we show how to deliver fluorescent beads into KV to visualize fluid flow.

基因功能分析和可视化的纤毛产生的流体流动的在枯否囊泡中

Internal organs such as the  heart, brain, and gut develop left-right (LR) asymmetries that are critical for  their normal functions1. Motile cilia are ...

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Spermatozoa are male reproductive cells especially designed to reach, recognize and fuse with the egg. To perform these tasks, sperm cells must be prepared to face a constantly changing environment and to overcome several physical barriers. Being in essence transcriptionally and translationally silent, these motile cells rely profoundly on diverse signaling mechanisms to orient themselves and swim in a directed fashion, and to contend with challenging environmental conditions during their journey to find the egg. In particular, Ca2+-mediated signaling is pivotal for several sperm functions: activation of motility, capacitation (a complex process that prepares sperm for the acrosome reaction) and the acrosome reaction (an exocytotic event that allows sperm-egg fusion). The use of fluorescent dyes to track intracellular fluctuations of this ion is of remarkable importance due to their ease of application, sensitivity, and versatility of detection. Using one single dye-loading protocol we utilize four different fluorometric techniques to monitor sperm Ca2+ dynamics. Each technique provides distinct information that enables spatial and/or temporal resolution, generating data both at single cell and cell population levels.

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Intracellular Ca2+ dynamics are very important in sperm physiology and Ca2+-sensitive fluorescent dyes constitute a versatile tool to study them. Population experiments (fluorometry and stopped flow fluorometry) and single cell experiments (flow cytometry and single cell imaging) are used to track spatio-temporal [Ca2+] changes in human sperm cells.

测量细胞内Ca<sup&gt; +</sup在人类精子的变化采用4项技术:传统的荧光，停流荧光，流式细胞术及单细胞成像

Spermatozoa  are male reproductive cells especially designed to reach, recognize and fuse  with the egg. To perform these tasks, sperm cells must be ...

High Efficiency Differentiation of Human Pluripotent Stem Cells to Cardiomyocytes and Characterization by Flow Cytometry

There is an urgent need to develop approaches for repairing the damaged heart, discovering new therapeutic drugs that do not have toxic effects on the heart, and improving strategies to accurately model heart disease. The potential of exploiting human induced pluripotent stem cell (hiPSC) technology to generate cardiac muscle &#8220;in a dish&#8221; for these applications continues to generate high enthusiasm. In recent years, the ability to efficiently generate cardiomyogenic cells from human pluripotent stem cells (hPSCs) has greatly improved, offering us new opportunities to model very early stages of human cardiac development not otherwise accessible. In contrast to many previous methods, the cardiomyocyte differentiation protocol described here does not require cell aggregation or the addition of Activin A or BMP4 and robustly generates cultures of cells that are highly positive for cardiac troponin I and T (TNNI3, TNNT2), iroquois-class homeodomain protein IRX-4 (IRX4), myosin regulatory light chain 2, ventricular/cardiac muscle isoform (MLC2v) and myosin regulatory light chain 2, atrial isoform (MLC2a) by day 10 across all human embryonic stem cell (hESC) and hiPSC lines tested to date. Cells can be passaged and maintained for more than 90 days in culture. The strategy is technically simple to implement and cost-effective. Characterization of cardiomyocytes derived from pluripotent cells often includes the analysis of reference markers, both at the mRNA and protein level. For protein analysis, flow cytometry is a powerful analytical tool for assessing quality of cells in culture and determining subpopulation homogeneity. However, technical variation in sample preparation can significantly affect quality of flow cytometry data. Thus, standardization of staining protocols should facilitate comparisons among various differentiation strategies. Accordingly, optimized staining protocols for the analysis of IRX4, MLC2v, MLC2a, TNNI3, and TNNT2 by flow cytometry are described.

The article describes the detailed methodology to efficiently differentiate human pluripotent stem cells into cardiomyocytes by selectively modulating the Wnt pathway, followed by flow cytometry analysis of reference markers to assess homogeneity and identity of the population.

人类多能干细胞的高效分化为心肌细胞，并鉴定流式细胞仪

There is an urgent need to develop approaches for repairing the damaged heart, discovering new therapeutic drugs that do not have toxic effects on the ...

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes.

Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach.

对于外小泡的使用流式细胞仪分析技术

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help ...

LED Thermo Flow &#8212; Combining Optogenetics with Flow Cytometry

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the controlled delivery of light to the cell sample and hence the most popular tools for optogenetic studies are microscopy-based cell analyses and in vitro experiments. The flow cytometer has major advantages over a microscope, including the ability to rapidly measure thousands of cells at single cell resolution. However, it is not yet widely used in optogenetics. Here, we present a device that combines the power of optogenetics and flow cytometry: the LED Thermo Flow. This device illuminates cells at specific wavelengths, light intensities and temperatures during flow cytometric measurements. It can be built at low cost and be used with most common flow cytometers. To demonstrate its utility, we characterized the photoswitching kinetics of Dronpa proteins in vivo and in real time. This protocol can be adapted to almost all optically controlled substances and substantially expands the set of possible experiments. More importantly, it will greatly simplify the discovery and development of new optogenetic tools.

Optically controlled substances are powerful tools to study signaling pathways. To expand the spectrum of possible experiments, we developed a device for studying optically controlled substances in real time using flow cytometry: the LED Thermo Flow.

LED热流量 - 结合光遗传学与流式细胞仪

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the ...

Determination of the Relative Cell Surface and Total Expression of Recombinant Ion Channels Using Flow Cytometry

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these multi-spanning transmembrane proteins, with or without change in channel gating, is often postulated to contribute significantly in this process. It has thus become critical to develop a method to quantify the change of the relative cell surface expression of cardiac ion channels on a large scale. Herein, a detailed protocol is provided to determine the relative total and cell surface expression of cardiac L-type calcium channels CaV1.2 and membrane-associated subunits in tsA-201 cells using two-color fluorescent cytometry assays. Compared with other microscopy-based or immunoblotting-based qualitative methods, flow cytometry experiments are fast, reproducible, and large-volume assays that deliver quantifiable end-points on large samples of live cells (ranging from 104 to 106 cells) with similar cellular characteristics in a single flow. Constructs were designed to constitutively express mCherry at the intracellular C-terminus (thus allowing a rapid assessment of the total protein expression) and express an extracellular-facing hemagglutinin (HA) epitope to estimate the cell surface expression of membrane proteins using an anti-HA fluorescence conjugated antibody. To avoid false negative, experiments were also conducted in permeabilized cells to confirm the accessibility and proper expression of the HA epitope. The detailed procedure provides: (1) design of tagged DNA (deoxyribonucleic acid) constructs, (2) lipid-mediated transfection of constructs in tsA-201 cells, (3) culture, harvest, and staining of non-permeabilized and permeabilized cells, and (4) acquisition and analysis of fluorescent signals. Additionally, the basic principles of flow cytometry are explained and the experimental design, including the choice of fluorophores, titration of the HA antibody and control experiments, is thoroughly discussed. This specific approach offers objective relative quantification of the total and cell surface expression of ion channels that can be extended to study ion pumps and plasma membrane transporters.

Inherited cardiac arrhythmias are often caused by mutations that alter the surface delivery of one or more ion channels. Here, we adapt flow cytometry assays to provide a quantification of the relative total and cell surface protein expression of recombinant ion channels expressed in tsA-201 cells.

重组离子通道的相对细胞表面表达的总用流式细胞仪测定

Inherited or de novo mutations in cation-selective channels may lead to sudden cardiac death. Alteration in the plasma membrane trafficking of these ...

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

通过光谱流式细胞仪由固态组织中分离出悬浮细胞系的分析

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...