SUMMARY
Background:
Platelet activation and subsequent aggregation is a vital component of atherothrombosis resulting in acute myocardial infarction. Therefore, quantifying platelet aggregation is a valuable measure for elucidating the pathogenesis of acute coronary syndromes (ACS). Circulating platelet-monocyte conjugates (PMC) as determined by flow cytometry (FCM) are an important measure of in vivo platelet aggregation. However, the influence of sample handling on FCM measurement of PMC is not well-studied.
Objectives:
The changes in FCM measurement of PMC with variation in sample handling techniques were evaluated. The stability of PMC concentrations over time with changes in fixation and immunolabeling intervals was assessed.
Patients/Methods:
The effect of Time-to-Fix and Time-to-Stain on FCM PMC measurements was investigated in five healthy volunteers. Time-to-Fix (i.e., interval between phlebotomy and sample fixation) was performed at 3, 30, and 60 minutes. Time-to-Stain (i.e., time of fixed sample storage to staining) was performed at 1, 24, and 48 hours.
Results:
Increasing Time-to-Stain from 1 to 24 or 48 hours resulted in lower PMC measures (p<0.0001). A statistically significant difference in PMC measurement with increasing Time-to-Fix was not observed (p<0.41).
Conclusion:
Postponement of sample staining has deleterious effects on the measurement of PMC via FCM. Delays in immunolabeling of fixed samples compromised measurement of PMC by 30% over the first 24 hours. Staining/FCM should be completed within an hour of collection.
Keywords: Flow Cytometry, Methods, Platelets, Monocytes, Specimen Handling
INTRODUCTION
Acute coronary syndrome (ACS) most frequently results from the destabilization, disruption, or rupture of atherosclerotic plaques [1]. With rupture of an atherosclerotic plaque, the subsequent vascular damage may lead to thrombotic occlusion of a coronary artery, causing acute myocardial infarction (AMI), or thrombotic occlusion of a cerebral artery, resulting in acute ischemic stroke. Cardiac troponins and other common markers of myocardial necrosis fail to provide information about plaque disruption [2, 3]. Platelet surface P-selectin has previously been regarded as an indicator of platelet activation in clinical settings; however, Michelson et al [4] demonstrated this approach may not fully account for activated platelets that have shed their surface P-selectin yet continue to circulate as functional platelets in vivo. Circulating platelet-monocyte conjugates (PMC) and platelet-leukocyte conjugates (PLeC) serve as a more sensitive reflection of atherothrombosis [2]. Platelet aggregation in response to plaque rupture is a vital component of atherothrombosis; therefore, in vivo quantification of platelet aggregation is an essential tool for the study of atherothrombotic AMI [5].
Whole blood flow cytometry (FCM) is an established method for quantifying circulating PMCs [6–8]. Cells are fluorescently tagged with a conjugated monoclonal antibody and are passed through the focused beam of a laser, emitting the light scattering properties of each cell [7]. Flow cytometry has the advantage of directly analyzing individual cells in their physiological state with minimal artifactual activation [7]. Other advantages include a high degree of sensitivity for the detection of platelet subpopulations, a miniscule sample volume (~2 μl) required for testing, and the capacity to evaluate the activation state as well as the reactivity of circulating platelets [7, 9]. The major obstacle when using this technique is the sensitivity to variations in sample collection and processing in the laboratory, especially during the critical time period between phlebotomy and specimen testing [10]. Sample fixation is utilized to preserve the aggregatory status of a platelet at the time of phlebotomy and minimize in vitro platelet activation [8]. This method is especially advantageous in clinical settings where immediate access to a flow cytometer is limited. Shattil et al [9] demonstrated that immediate fixation of the blood after venipuncture with 1% paraformaldehyde prevented spontaneous platelet activation and stabilized the sample. However, this variable must be controlled for when fixing prior to immunolabeling because the binding of monoclonal antibodies to fixed platelets is activation-dependent and decreases over time in comparison to unfixed platelets [8]. Previous research [11] exhibited a 1.7% increase of PMC measurement in unfixed samples anticoagulated with sodium citrate for every ten minute delay to immunolabeling and fixation. This highlights the necessity for expeditious sample processing as well as the standardization of time intervals. The first variable of interest in this study evaluates the effect of time of phlebotomy to sample fixation (Time-to-Fix) on the measurement of PMC.
In addition to the use of fixatives to preserve the in vivo state of activation, sample stability between fixation and antibody labeling, or staining, has not been rigorously evaluated. In some studies [8, 9, 12], no significant difference in the intensity of fluorescence was observed during flow cytometric analysis upon immediate fixation or within 24 hours of fixation. Other publications, on the other hand, contradict this conclusion and propose a minimal time of delay between fixation, labeling, and analyzing. This calls into question the stability of fixed samples over time and its effect on PMC quantification. Delays after fixation and pre-staining could be due to logistical circumstances (transport of samples in clinical settings), availability of skilled operators, and access to shared flow cytometers. As such, the second variable of interest evaluated the effect of time of fixed samples to staining with monoclonal antibodies (Time-to-Stain) on the measured levels and stability of PMC measurement. Although flow cytometric protocols for the measurement of PMC have previously been described [8, 9, 11, 12], the consequences of variations in processing methodology, sample manipulation, and PMC stability over time have not been adequately characterized. Blood from five healthy volunteers was obtained, and four heterotypic conjugate phenotypes were studied: platelet-monocyte conjugates (PMC), platelet-lymphocyte conjugates (PLC), platelet-granulocyte conjugates (PGC), and total platelet-leukocytes conjugates (PLeC). [4, 13–15]. Our objective is to evaluate the effect of Time-to-Fix and Time-to-Stain on PMC measurement via FCM.
METHODS
Changes in PMCs were quantified using whole blood flow cytometry. Two experimental manipulations of an established lab protocol (DeFilippis, University of Louisville) were generated to evaluate sample stability (Table 2). These variables, Time-to-Fix and Time-to-Stain were chosen based on the inconsistency and ambiguity pertaining to the methodological considerations of these two factors and their potential to produce inconsistent and inaccurate results. Time-to-Fix is defined as the time between sample collection (phlebotomy, venipuncture) and fixation with 1.2% paraformaldehyde. For this study, Time-to-Fix value was standardized at 3, 30, and 60 minutes after the final tube of blood was collected. After fixation, each sample was incubated on ice for 30 minutes, washed, and stored at 4°C until its respective staining time. Time-to-Stain was assessed at 1, 24, and 48 hours, and indicates the amount of time between post fixation sample storage and staining time. FCM was performed on each sample within 15 minutes of staining. Samples were evaluated in triplicate. All materials, operators, and processing techniques were standardized to minimize confounding.
Table 2:
Analysis of Deviance of Four Main Cellular Platelet-Conjugate Phenotypes
| Process | P-Value | |||
|---|---|---|---|---|
| Leukocytes | Granulocytes | Lymphocytes | Monocytes | |
| Fixation | 0.004 | 0.01 | 0.001 | 0.41 |
| Staining | 0.0001 | <0.0001 | 0.002 | <0.0001 |
| Fixation × Staining | 0.52 | 0.50 | 0.74 | 0.31 |
Study Population
Five healthy volunteers, age 18–50, were enrolled under a University of Louisville Institutional Review Board (IRB) approved Biorepository Study after informed consent was obtained. Demographics and medical history were acquired via direct interview. Each volunteer met a set of preliminary enrollment criteria, including no significant cardiac history, no use of medication known to affect platelet function within 7 days of participation, no tobacco use, and no diabetes. Participant age, sex, and race were recorded in Table 1.
Table 1:
Characteristics of Study Participants
| Subject ID | Age | Gender | Race |
|---|---|---|---|
| Subject 1002 | 20 | Female | Asian |
| Subject 1003 | 22 | Male | Caucasian |
| Subject 1004 | 34 | Male | Asian |
| Subject 1005 | 31 | Female | Caucasian |
| Subject 1006 | 44 | Male | Caucasian |
Sample Collection
Thirty-two mL of venous blood was collected at a single time point by venipuncture (21G, BD Push Bottom) with the use of a light tourniquet into seven 4.5 mL tubes containing 3.2% sodium citrate (BD Vacutainer). The first vial of blood was collected and discarded as a waste tube to minimize red cell hemolysis, which has been shown to precipitate artifactual platelet activation [16]. T0 is defined as the time the final sample vial was collected. Each vial was gently inverted eight times and kept stationary, upright, and at room temperature until the assigned Time-to-Fix interval was reached (3 min, 30 min, 60 min) from T0.
Sample Preparation
The sample preparation scheme is depicted in Figure 1. Upon the Time-to-Fix interval, four mL of citrate-anticoagulated whole blood was added to a 50 mL conical vial (VWR) containing 17.2 mL of a freshly prepared fixative composed of paraformaldehyde and buffered saline. 5.2 mL of a 4% paraformaldehyde aqueous solution (Electron Microscopy Sciences) was diluted with 12 mL of Phosphate-Buffered Saline (Hardy Diagnostics) to achieve a final paraformaldehyde concentration of 1.2%. Paraformaldehyde is widely accepted as a reliable fixative that terminates platelet reactivity and is recommended when sample analysis is delayed for more than 2 hours [17]. The fixed sample stock solution ensured the standardization of Time-to-Fix across all Time-to-Stain samples. 5.3 mL of the stock solution was aliquoted into three separate conical vials (VWR) assigned with a Time-to-Stain interval.
Figure 1:
Sample Preparation Scheme
Irrespective of the Time-to-Fix interval, all fixed samples were immediately placed on ice and incubated for 30 minutes to simulate transport conditions in a clinical study. At the end of the incubation period (30 minutes post Time-to-Fix), 24 mL of deionized (DI) water was added to lyse red blood cells in the fixed sample. The tube was inverted eight times, and underwent centrifugation (400 g, 10 mins, 20°C). The supernatant was discarded, and the pellet was resuspended in 1 mL of Tyrode’s/bovine serum albumin (BSA) buffer (1 mg BSA per 1 mL Tyrode’s bicarbonate buffer, Boston Bioproducts). This solution was stored at 4°C until stained with monoclonal antibodies at their assigned Time-to-Stain intervals (1, 24, and 48 hours).
Sample staining began with centrifugation (1000 g, 5 min, 20°C), followed by a 1 mL Tyrode’s/BSA wash and centrifugation (1000 g, 5 min, 20°C), then incubation with Fc Receptor Binding Inhibitor (eBiosciences) for 10 min on ice. APC Mouse Anti-Human CD45 (BD Bioscience) and FITC Mouse Anti-Human CD41a (BD Bioscience) were added to the sample and incubated for 30 minutes on ice. After washing with 300uL Tyrode’s/BSA and centrifuging (1000 g, 5 mins, 20°C), the cell pellet was resuspended in 300ul Tyrode’s/BSA solution and analyzed immediately in the flow cytometer. All stained samples were measured in triplicate. Stored stained samples were not evaluated in this study.
FCM Analysis
Platelet activation is determined by a combination of specific monoclonal antibodies [7, 8]. This experiment used APC Mouse Anti-Human CD45 (BD Biosciences) and FITC Mouse Anti-Human CD41a (BD Biosciences) to label the populations of interest. CD45, also known as the leukocyte common antigen, is present on all human leukocytes [18]. APC anti-CD45 was used to label all granulocytes, monocytes, and lymphocytes. CD41a, also known as integrin alpha IIb (ITGA2B), is a platelet membrane glycoprotein [19]. FITC anti-CD41a was used to label platelets bound to leukocytes, and is widely recognized as a useful antibody in the study of platelet aggregation [19].
Samples were analyzed using a BD LSR II, Special Order System (Serial # H47206039), with BD FACS Diva v6.1 software. The flow cytometer was calibrated daily using BD Cytometer Setup and Tracking Beads to ensure sample reproducibility. No electronic color compensation was set because the antibodies are bound to fluorochromes that do not overlap wavelengths on the emission spectra. Immunolabeled samples were analyzed at a low setting and flow rate. An unstained sample was used to establish gating parameters. Platelets are detected in whole blood by light scatter only, and the two color/two antibody technique is recommended [7–9].
FlowJo software [BD Biosciences] was used to gate all events. Side scatter versus CD45 was analyzed first to gate granulocytes, monocytes, and lymphocytes. Next, CD41a versus forward scatter was analyzed for each of the different leukocyte populations to gate platelet conjugates. Events positive for both CD45 and CD41 were quantified as conjugates. PMC events were measured as the percentage of proportionate aggregates of monocytes.
Outcome Measures
The four heterotypic conjugate phenotypes under study were platelet-monocyte conjugates (PMC), platelet-lymphocyte conjugates (PLC), platelet-granulocyte conjugates (PGC), and total platelet-leukocytes conjugates (PLeC). Time-to-Fix (3 minutes, 30 minutes, and 60 minutes) and Time-to-Stain (1 hour, 24 hours, 48 hours) were experimentally manipulated. This constituted a two-factor design.
For each combination, five biological samples (each from different study participants) and three technical replicates of each sample were evaluated. The outcome measure was the proportion of counted cells that were conjugated to platelets. As the outcome measure could only take continuous values in [0, 1], the logit transformation was applied. A linear mixed effects model was then fit. Fixed effects included Time-to-Fix, Time-to-Stain, and fixation×staining interaction with a random intercept included for each study subject. In each model the interaction term was non-significant and was removed for estimating odds ratios. Odds ratios and 95% confidence intervals are presented relative to fixation at 3 minutes and staining at 1 hour.
RESULTS
PMC, PLC, PGC, and PLeC were assessed in five healthy subjects with three technical replicates of each sample for a Time-to-Fix of 3, 30, and 60 minutes; and Time-to-Stain of 1, 24, and 48 hours. For each cell phenotype, Time-to-Stain had a significant effect on the odds an identified cell was in a heterotypic cell conjugate (Table 2). For each cell phenotype excluding monocytes, Time-to-Fix had a significant effect on the odds a cell was conjugated with a platelet (Table 2). No interaction between Time-to-Fix and Time-to-Stain was observed (Table 2). Figure 2 presents the model estimated odds ratio for a counted cell being in conjugation with platelets relative to fixation at 3 minutes and staining at 1 hour.
Figure 2:
Model Estimated Odds Ratio for Four Main Cellular Platelet-Conjugate Phenotypes
Fixation at 3 minutes and staining at 1 hour serve as the reference points
The odds a cell was in aggregate with platelets were lower among all four tested platelet-cell conjugate types with increasing Time-to-Fix and Time-to-Stain with no evidence of an interaction between Time-to-Fix and Time-to-Stain (Figure 2, Table 2). Odds ratios and 95% confidence intervals (CI) are presented relative to fixation at 3 minutes and staining at 1 hour. We define the number of platelet-cell type interactions as compared to the total number of cell types across different experimental variables. The odds are defined as the ratio of a cell type conjugated with a platelet (platelet-leukocyte, platelet-granulocyte, platelet-lymphocyte, platelet-monocyte) over the number of that cell type of interest not conjugated with platelets (leukocyte, granulocyte, lymphocyte, monocyte). The reference odds value is represented in the first block (1.00) of each cell type quadrant and is attributed with experimental conditions of Time-to-Fix at 3 mins and Time-to-Stain at 1 hour. The odds of experimental condition outcomes, or the change of Time-to-Fix and Time-to-Stain, were compared to this baseline value normalized to 1.00 (Figure 2). When Time-to-Fix changes from 3 mins to 30 mins, the odds of measuring a monocyte in aggregate with a platelet decreases from 1.00 to 0.98 (95% CI: 0.84, 1.10). When the Time-to-Fix variable changes from 3 mins to 60 mins, the odds of a measured PMC decreases from 1.00 to 0.92 (95% CI: 0.79, 1.10). When samples were fixed at 3 mins but stained 1 hr or 24 hrs later (Time-to-Stain), the odds of PMCs measured decreased from 1.00 to 0.77 (95% CI: 0.66, 0.90). Further increase in Time-to-Stain to 48 hrs resulted in a 34% average decrease in detected heterotypic cell conjugates in comparison to staining at 1 hour when the fixation time is held at 3 mins. With manipulation of both Time-to-Fix from 3 mins to 60 mins and Time-to-Stain from 1 hr to 48 hrs, the odds of measuring a platelet-monocyte conjugate decreased from 1.00 to 0.60 (95% CI: 0.49, 0.75). This pattern was consistent among all the heterotypic cell conjugates and is illustrated in Figure 2.
Assessment of individual subject data is depicted in Figure 3. Shorter time intervals to fix and stain demonstrated higher PMC aggregates. Observable PMC proportions were higher when Time-to-Fix × Time-to-Stain was 3 mins × 1 hr in comparison to each individual subject’s Time-to-Fix × Time-to-Stain value at 60 mins × 48 hrs. The average coefficient of variance (%) over the technical replicates ranged from 9.14%−22.71% for PMC, 2.96%−4.93% for PGC, 2.33%−4.18% for PLeC, and 3.84%−7.20% for PLC (Table 3).
Figure 3:
Measurement Time-Course for Each Study Subject
Table 3:
Average Coefficient of Variance (%) Over the Technical Replicates
| Cell Phenotype | Time-to-Staining (Hours) | Fixation at 3 Minutes | Fixation at 30 Minutes | Fixation at 60 Minutes |
|---|---|---|---|---|
| Granulocytes | 1 | 3.11 | 1.85 | 3.46 |
| 24 | 4.28 | 4.27 | 4.93 | |
| 48 | 4.28 | 2.96 | 5.19 | |
| Leukocytes | 1 | 2.57 | 2.33 | 3.80 |
| 24 | 4.18 | 3.37 | 3.51 | |
| 48 | 3.98 | 2.57 | 2.97 | |
| Lymphocytes | 1 | 7.20 | 5.69 | 6.50 |
| 24 | 3.96 | 3.84 | 6.04 | |
| 48 | 7.13 | 4.93 | 6.90 | |
| Monocytes | 1 | 9.14 | 10.02 | 13.25 |
| 24 | 12.47 | 13.82 | 13.58 | |
| 48 | 22.71 | 14.87 | 16.81 |
The analysis of deviance in Table 2 showed a significant effect (p < 0.05) of Time-to-Stain for all heterotypic aggregates. The effect of Time-to-Fix was significant in PLeC (p<0.004), PGC (p<0.01), and PLC (p<0.001); however, evidence of a significant effect of Time-to-Fix was not observed for PMC (p<0.41). Evidence of a significant compounding or moderating effect of Time-to-Fix × Time-to-Stain was not observed in any of the heterotypic conjugates.
DISCUSSION
Current clinically available diagnostic tools for patients with AMI are limited to markers of myocardial necrosis if ST-segment abnormalities are absent on an electrocardiogram [3]. Indicators such as cardiac troponins fail to measure the cause and therapeutic target of AMI—atherothrombosis. [20, 21]. Platelets are a vital component of atherothrombosis [22], and a greater understanding of platelets in AMI has shown promise for differentiating etiologically distinct types of AMI that necessitate different treatment strategies to improve care of patients suspected of AMI [23–25].
While platelet surface P-selectin has previously been considered a marker of platelet activation [11], Michelson et al [4] have shown that platelet surface P-selectin sheds rapidly in vivo while functioning platelets continue to circulate. Platelet-monocyte aggregates are a more sensitive marker of platelet activation as well as a reliable indicator of AMI and other thrombotic responses [4, 11, 13, 15]. Currently, limited knowledge regarding PMC processing and methodological techniques is available for laboratory protocol development. In this study, the in vitro stability of PMC over time was evaluated in blood samples from healthy donors.
Two variables, Time-to-Fix (3, 30, and 60 minutes) and Time-to-Stain (1, 24, and 48 hours), were assessed due to their vulnerability to variation during sample acquisition and processing. Previous research has demonstrated that sample fixation preserves the status of PMCs in vitro by preventing subsequent artifactual platelet activation, thus representing the in vivo conditions more accurately [7, 8]. Studies by Harding et al. demonstrated that the percentage of PMCs in unfixed samples increased with every ten-minute delay to sample fixation [11]. This prompted the use of shorter time intervals to evaluate the effect of Time-to-Fix while maintaining logistical feasibility.
When samples are fixed before labeling, Michelson et al. demonstrated that there are no significant differences in samples analyzed immediately and samples analyzed within 24 hours of staining [8]. Harding et al. showed that fixed samples stored at 4C did not alter PMCs over a 24 hr period [11]. As such, evaluating the effect of Time-to-Stain at intervals within a 24-hour period was considered inconsequential. Time-to-Stain at 1 hour (immediately), 24 hours, and 48 hours was measured with the intention of evaluating a difference in detected values after the 24-hour period. Data on longer Time-to-Stain intervals is important to many clinical and research sites that do not have access to a flow cytometer laboratories or expertise, requiring that their samples be shipped to another laboratory in order to complete the measurements (e.g., “send out lab”) [7, 8].
There were two major cytometric findings that will have great impact on clinical research. First, for each cell phenotype excluding monocytes (PLC, PGC, PLeC), Time-to-Fix had a significant effect on the odds a cell in a heterotypic conjugate was measured. The effect of increasing the Time-to-Fix from 3 to 30 minutes did not significantly impact measurement of the number of PMCs measured when holding Time-to-Stain constant. This is notable because whole blood in sodium citrate remains relatively stable for 60 mins prior to fixation, while the Time-to-Stain has a more substantial effect on the measurement of PMCs over a 48-hour period. However, a significant variance in these estimates was observed. This may be attributed to the lower number events observed when quantifying monocytes as opposed to total leukocytes. The higher CV value in the monocyte phenotype in comparison to leukocytes (Time-to-Fix × Time-to-Stain [3 mins × 1 hr]: 9.14% [PMC], 2.57% [PLC]) suggests that the increasing specificity of the cell may be more difficult to evaluate with FCM. Additionally, the inconsistent variability could undermine results in future experiments; therefore, protocols should be developed by research laboratories independently, prior to any experimental analysis.
Second, for each cell phenotype, Time-to-Stain had a significant effect on the odds of measuring a cell in a platelet-leukocyte conjugate. A statistically significant change in all measured conjugate interactions was observed when manipulating the Time-to-Stain interval. Samples in fixative have previously been described to remain stable over a 24-hour period [7, 8]. However, the results of this study suggest that Time-to-Stain is more consequential in the methodological approach. PMC measurement stability decreased over time, but the Time-to-Fix was not significant. It may be discerned that the relative stability of PMC measurement remains stable over a 48 hr period prior to sample fixation. This is in contrast to conclusions found by Furman et al [3, 26] in which sample stability post fixation did not result in artifactual platelet activation. The odds ratio for the Time-to-Stain variable decreased over the experimental time period and challenges the notion that samples in fixative remain stable for an extended period of time prior to immunolabeling and analysis. Circulating PMCs decreased an average of 2% within the first 30 minutes and 8% within 60 minutes, suggesting an immediate or bedside fixation may not be as crucial. This is distinct from the results regarding Time-to-Stain. Achieving an optimal fixation time of 3 minutes, yet a sample staining time at 24 hours decreased the odds of detected platelet-monocyte conjugate by an average of 24%. Thus, we recommend that samples are fixed within 30–60 minutes and stained at a standardized time point as soon as possible.
The relative increase in detected platelet-cell conjugates over the initial 24-hr period presented in Figure 3 may be attributed to multiple washing and centrifugation steps, which can induce artifactual PLC/PMC activation [6–8]. This artifactual elevation could be more apparent when analyzing samples of patients with ACS who already have higher levels of circulating PMC in comparison to the healthy subject participants of the present study [13–15, 27, 28].
The limitations of the present study include the small sample size (n=5), multiple washing and centrifugation steps, and the use of healthy blood donors. However, three technical replicates were performed for the conditions evaluated and a mixed-effects model was utilized to account for the within-subject dependence to maximize statistical power from the five human participants. Confounding from differences in participant characteristics was eliminated by evaluating the impact of variable fixation and staining times in aliquots from the same participant. All comparisons for differences in platelet-cell conjugate measures are made from the same sample via preparation of multiple aliquots at the time of phlebotomy; thus, each participant served as their own control. It is possible that the impact of variations in Time-to-Fix and Time-to-Stain on the detection of platelet-cell conjugates is influenced by differences in the characteristics of the participant for which the measurements are being made (e.g., age, sex). Additional research will be required to answer this question. While PMCs can be quantified with flow cytometry, it should be noted that FCM measures an individual particle’s amount of fluorescence, regardless of its identity as a single platelet or part of an aggregate with an unknown amount of platelets [5, 7, 8, 10, 29]. Non-adherent monocytes and platelets fortuitously near each other during the laser’s evaluation may be indistinguishable from true PMCs [5, 8, 29].
Sample processing and handling techniques can influence platelet-monocyte conjugation and in vitro platelet activation [8, 11]. Limited research has been done to establish optimal protocols that minimize artifactual platelet-monocyte conjugation. These measurements are susceptible to fluctuation as a result of daily lab practices, handling techniques, and logistical impediments such as sample transport or restricted access to flow cytometer. Not only minimizing, but standardizing variables such as Time-to-Fix and Time-to-Stain will enable the most representative and accurate results. When using a fixation first approach, it is recommended that samples for experimental evaluation be fixed at an immediate, standardized time interval not exceeding 60 minutes. More importantly, samples should be stained within 1 hour, assuming they are analyzed by FCM immediately (within 15 minutes of sample immunolabeling) in order to accurately quantify heterotypic cell conjugates.
The findings of this study are challenging to contextualize in current literature. Due to varying protocols, techniques, and sample measurement (FCM and corresponding software), such results are difficult to compare directly with other laboratory experiments. We propose that PMC stability decreases over time in vitro, irrespective of the lab protocol and methodology, and a deliberate effort should be made to minimize artifactual platelet activation. Michelson et al [7, 8] previously recommended that standardizing fixation time, minimizing washing and centrifugation, using a light tourniquet, discarding the first 2 mL of blood, and immediately labeling the blood with monoclonal antibodies for subsequent analysis will more reliably quantify levels of circulating PMCs. This study further suggests that all samples for platelet-cell conjugate evaluation should be fixed at an immediate, standardized time interval not exceeding 60 minutes. More importantly, samples should be stained within 1 hour, and analyzed by FCM immediately (within 15 minutes of sample immunolabeling) in order to accurately quantify heterotypic cell conjugates.
Assessing the stability of PMCs over time is relevant in laboratories that strive to quantify the in vivo status of circulating platelet activation in vitro. The time dependent decrease in detected PMCs can be a result of delays in sample processing and handling techniques. This numerical difference can be minimized with standardized time intervals in order to optimize the number of detected aggregates. This research suggests that the importance of an immediate fixation is questionable, allowing more flexibility in fixation time previously cautioned against in other research studies [7, 11]. Previous research has shown that fixed samples remain stable over a 24-hour period, but our research indicates a significant difference in detection of PMAs with staining at 1 hour versus 24 hours (when fixation is standardized at 3 minutes) [7, 8]. This may implicate data dependent on numerical aggregate proportions and its clinical relevance exists but has not been fully characterized. However, to move from the bench to the bedside for use as a clinical tool, standardization of qualitative and quantitative PMC measurement will be required.
ESSENTIALS.
Platelet-monocyte conjugates (PMC) are a sensitive marker of platelet activation.
PMC measurements are influenced by sample handling.
In healthy blood, measured PMC significantly decreased when staining was delayed over 24 hrs.
Immediate sample fixation and staining within 1 hr reduces PMC measurement variability.
Acknowledgements:
This study was financially supported by the American Heart Association (11CRP7300003) and National Institute of Health (1P20 GM103492). We thank Jacob Schultz, Mallory Hatfield, and Joey Shaw for their assistance with sample acquisition and processing; Aaron Puckett (UofL Diabetes and Obesity Center Core Lab) for helping with flow cytometry; Allison Smith for editing the manuscript; Yong Siow and Tim O’Toole for intellectual guidance.
Grant Support: American Heart Association (11CRP7300003) and National Institutes of Health (1P20 GM103492)
Footnotes
Publisher's Disclaimer: This Author Accepted Manuscript is a PDF file of an unedited peer-reviewed manuscript that has been accepted for publication but has not been copyedited or corrected. The official version of record that is published in the journal is kept up to date and so may therefore differ from this version.
Disclosures: All authors have no conflict of interest to disclose.
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