Journal of Immunological Methods 381 (2012) 9–13

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Journal of Immunological Methods

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Research paper

A flow cytometric protocol for enumeration of endothelial progenitor cellsand monocyte subsets in human blood

Mihail Hristov a,⁎, Susanne Schmitz a, Frans Nauwelaers c, Christian Weber a,b

a Institute for Cardiovascular Prevention, Ludwig-Maximilians-University (LMU), Munich, Germanyb Munich Heart Alliance, Munich, Germanyc BD Biosciences Europe, Erembodegem, Belgium

a r t i c l e i n f o

⁎ Corresponding author at: Institute for CardiovascMunich, Pettenkoferstr. 9, 80336 Munich, Germany. Tefax: +49 89 5160 4352.

E-mail address: mihail.hristov@med.uni-muenche

0022-1759/$ – see front matter © 2012 Elsevier B.V. Adoi:10.1016/j.jim.2012.04.003

a b s t r a c t

Article history:Received 26 January 2012Received in revised form 3 April 2012Accepted 5 April 2012Available online 13 April 2012

Background: Accumulating evidence intensively advises circulating endothelial progenitor cells(EPCs) and monocyte subsets as surrogate cellular biomarkers in cardiovascular and cancerdisease. However, a general standard on their quantification is still elusive, thus precluding aroutine monitoring and comparative interpretation of clinical studies.Objective: We intend to develop an advanced and express flow cytometric protocol for properex vivo quantification of monocyte subsets and EPCs in human blood.Methods: We employ now lyse/no-wash procedure and bead-based determination of absolutecell counts. We use three-color antibody panels at appropriate compensation. Analysis of rareevents and low antigen expression in the EPC experiment is strengthening by sequential gatingwith exclusion of dead cells, as well as by matching high-intensity fluorochromes to low-densitymarkers and by implementing the fluorescence-minus-one control.Results: Analysis of peripheral blood of ten healthy donors revealed median (IQR) value of1.88 (1.35–2.85) viable CD45dimCD34+VEGFR2+ EPCs per microliter. Analysis of monocytesrevealed 329.5 (264.5–374.8), 16.0 (8.0–22.2) and 26.5 (19.8–36.3) cells per microliter forclassical CD14++(high)CD16−, intermediate CD14++CD16+(mid) and non-classical CD14+(low)

CD16++ monocytes.Conclusion: Our current protocol provides quantitative information under a simple gating logicwhile using commonly accepted fluorochromes. This assay is therefore highly adapted for routineuse.

© 2012 Elsevier B.V. All rights reserved.

Keywords:Hematopoietic stem cellsLeukocytesEndotheliumCellular biomarker

1. Introduction

Wehave previously designed a flow cytometric protocol forex vivo assessment of angiogenic Tie2+ monocytes and endo-thelial progenitor cells (EPCs) in human peripheral blood(Hristov et al., 2009). At that time, we intended to update andimprove existing assays that substantially differ in methodol-ogy. Based on this protocol, we have reported about pilotreference values of cells-of-interest in patients with stable

ular Prevention, LMUl.: +49 89 5160 4672;

n.de (M. Hristov).

ll rights reserved.

coronary artery disease (CAD) and evaluated the relation ofmonocyte subsets to established cardiovascular risk factors(Hristov et al., 2009, 2010). Indeed, the differential monitoringof CD14/CD16 monocyte subsets may identify patients at highcardiovascular risk but also subjects with subclinical athero-sclerosis (Hristov and Weber, 2011). A recent study furtherintroduced standardized single-platform assay for the enumer-ation of humanmonocyte subpopulations using four-color panelof CD45, CD14, CD16 and HLA-DR determination (Heimbeck etal., 2010).

Accumulating evidence further advises circulating EPCs andendothelial microparticles as surrogate biomarkers for endo-thelial integrity in different pathological disorders such as car-diovascular and cancer disease (Werner and Nickenig, 2006;

10 M. Hristov et al. / Journal of Immunological Methods 381 (2012) 9–13

Bertolini et al., 2006). Nonetheless, a general agreement ondefinition and measurement of the “true” EPC appear stillpuzzling, as diverse marker combinations and protocols wereused to assess the amount of (endothelial) progenitor cells inperipheral blood (Duda et al., 2007; Van Craenenbroeck et al.,2008; Estes et al., 2010). Very recent studies have furtherdeveloped an optimized quantitation of EPCs by flow cytometryin patients with CAD or cancer (Schmidt-Lucke et al., 2010;Farace et al., 2011). However, in theseworks dead cell exclusionand absolute counts were not considered (Schmidt-Lucke et al.,2010), or EPCs were assessed in mononuclear cell preparationsafter density gradient centrifugation and not direct in wholeblood (Farace et al., 2011). As a substantial advantage however,both studies used CD45dimCD34+VEGFR2+ as consistent defi-nition of circulating EPCs. Moreover, exact this cell subset wasreduced in CAD patients as compared to healthy controls andcorrelated inversely with the number of cardiovascular riskfactors (Schmidt-Lucke et al., 2010). Despite controversy but inline with several earlier findings including also sorting andfunctional experiments, this surface profile appears as precisemarker combination of the circulating putative EPC (Ingram etal., 2005; Bertolini et al., 2006; Case et al., 2007; Timmermans etal., 2007).

Since methodology is constantly evolving and novel con-cepts or applications are being implemented, we have upgradedour previously published protocol (Hristov et al., 2009). Asmajor modification we employ now lyse/no-wash procedureand determination of absolute cell counts by simple gating logicand antibody panels with commonly accepted dyes. The currentassay is therefore highly adapted for routine use.

2. Materials and methods

2.1. Blood samples

After informed consent was obtained, peripheral blood wascollected into EDTA tubes (Sarstedt, no. 05.1167) by asepticvenipuncture from ten (four females, six males) healthyvolunteers, age range of 24–37 years. In order to test assayreliability, two separate blood samples at two time pointswere collected in a new group of five healthy donors. The bloodsamples were stored at 4 °C and the immunostaining wasperformed no later than 2 h after blood drawing.

2.2. Immunostaining

Allmonoclonal antibodies (mAbs)were fromBDBiosciences.For analysis of EPCs, 20 μl CD45-FITC (clone 2D1, no. 345808),5 μl CD34-APC (clone 8 G12, no. 345804), 20 μl CD309/VEGFR2-PE (no. 560494) mAb and 5 μl 7-AAD (BD Biosciences, no.559925)were pipetted into a Trucount tube (BD Biosciences, no.340334). For the fluorescence-minus-one (FMO) control, 20 μlCD45-FITC and 5 μl CD34-APC mAb were pipetted into a poly-styrene round-bottom snap cap tube. For assessment of mono-cytes, 20 μl CD45-PerCP (clone 2D1, no. 345809), 20 μl CD14-FITC (no. 345784) and 20 μl CD16-PE (clone 3 G8, no. 555407)mAb were pipetted into a Trucount tube. Fifty microliters ofwell-mixed bloodwere added to each tube by reverse pipetting.Sample tubeswere incubated for 20 min at room temperature inthe dark. Red blood cells were lysed by adding 450 μl of 1×lysing solution (BD Biosciences, no. 555899) to each tube and

incubated for 10 min at room temperature in the dark. After thistime, tubes were placed immediately on wet ice protected fromlight until ready to measure.

2.3. Flow cytometric analysis

Sample acquisitionwas donewithin 1 h of red cell lysis on aFACSCanto II flow cytometer (2-laser, 6-color configuration)with FACSDiva 6.1.2 software (BD Biosciences). Instrumentperformance check was completed daily with CST beads (BDBiosciences, no. 641319). In addition, the flow cytometer wasused at least 30 min after fluidics start-up to allow reliablestabilization and warming-up of the system. Fluorescencecompensation was assessed with anti-mouse Igκ CompBeads(BD Biosciences, no. 552843) and the respective antibodies.Compensation values were calculated automatically. Onlyvoltages for forward scatter (FSC) and side scatter (SSC) wereadditionally fine-tuned in the current experiment. The FSCthreshold was not adjusted. Amplifier settings for FSC and SSCwere used in linear mode and those for fluorescence channelswere used in logarithmicmode. Sample tubes were acquired ata flow rate b10,000 events/s by setting acquisition gates on theCD45+ cells or on the monocyte scatter population. At least200,000 CD45+ events and 10,000 monocytes were recorded.

2.4. Data analysis

FCS files were saved after successfully recording a sampletube. All experiments were exported and stored in the BD FACSdatabase. Data analysis was performed with the FACSDivasoftware.

2.4.1. Analysis of EPCsFor analysis of EPCs (Fig. 1), all leukocytes are first gated on

a FSC/SSC dot plot (P1). Then, the P1 content is displayed on aSSC/7-AAD dot plot and all 7-AAD- events are gated as P2.This P2 population is shown on a SSC/CD45 dot plot and theCD45+ events are gated as P3. Then, the P3 cell set is shownon a SSC/CD34 dot plot following gating of the CD34+ events asP4. The events of P4 are subsequently presented on a FSC/SSCdot plot and gated as P5 in order to confirm the lymph-blastscatter region and to remove residual debris. The events of P5are then shown on a SSC/CD45 dot plot and only the CD45dim

cells are gated as P6. Gating of the FSC/SSC lymph-blast region(Fig. 1e) and back-scattering of CD34+ events to a SSC/CD45dot plot (Fig. 1f) are essential steps to confirm cell size andclustering of bona fide CD45dimCD34+ cells. Optionally and asstated in the original ISHAGE protocol (Sutherland et al., 1996),a gate preferentially including the lymphocytes with lowestSSC of P3 (Fig. 1c) can be used to confirm the boundaries of thelymph-blast region (Fig. 1e). Finally, the CD45dimCD34+ eventsare shown on a CD34/VEGFR2 dot plot and the cut-off forVEGFR2+ events (P7) is assessed by the FMO control. For beadcounting, the bead region (P8) is gated on a CD45/CD34 dotplot showing all events (Fig. 1i). Calculation is carried out by thefollowing equation: [(# of cells in P7):(# of events in P8)]×[(#of beads per tube):(sample volume in microliter)]=absolutecell count per microliter.

a b c d

e f g

j

h

i

Fig. 1.Multicolor flow cytometric analysis of human endothelial progenitor cells in whole blood. All leukocytes are first gated on a FSC/SSC dot plot as P1 (a). ThisP1 is displayed on a SSC/7-AAD dot plot and all 7-AAD- events are gated as P2 (b). The CD45+ events of P2 are gated on a SSC/CD45 dot plot as P3 (c). Then, the P3content is shown on a SSC/CD34 dot plot following gating of the CD34+ events as P4 (d). The events of P4 are subsequently presented on a FSC/SSC dot plot andgated as P5 in order to confirm the lymph-blast scatter region and to remove residual debris in front of the population (e). The events of P5 are then plotted againon a SSC/CD45 dot plot and only the CD45dim cells are gated as P6 (f). These CD45dimCD34+ events are shown on a CD34/VEGFR2 dot plot (g) and the cut-off forVEGFR2+ events (P7) is assessed by the FMO control (h). The bead count is obtained from an ungated CD45/CD34 dot plot (P8 in i). (j) Population hierarchy.

11M. Hristov et al. / Journal of Immunological Methods 381 (2012) 9–13

2.4.2. Analysis of monocyte subsetsFor analysis of monocyte subsets (Fig. 2), cell debris are first

removed by gating all leukocytes on a FSC/SSC dot plot (P1).Then, the P1 is displayed on a SSC/CD45 dot plot and all CD45+

events are gated as P2. These CD45+ events are then shown ona FSC/SSC density plot followed by tightly gating of just themonocyte scatter population (P3). This is a crucial step to avoidpotential “contamination” with non-monocytic CD16+ events.Nevertheless, when reviewing as a SSC/CD45 dot plot, someevents from P3 tend to appear very close to the lymphocytescatter signals. This is because non-classical monocytes areusually less dense and smaller in size as compared to classicalmonocytes (Heimbeck et al., 2010; Hristov and Weber, 2011).The P3 population is finally shown on a CD14/CD16 dot plot andthe three major monocyte subsets are consequently defined byusing a quadrant marker: (Q1) for “classical” CD14++(high)

CD16−, (Q2) for “intermediate” CD14++CD16+(mid) and (Q4)for “non-classical” CD14+(low)CD16++ monocytes. Absolute

counts are calculated from the above formula after obtainingthe bead number from an ungated CD45/CD16 dot plot of allevents (gate P4 in Fig. 2e).

2.5. Statistics

Statistical data analysiswas performedusingGraphPad Prism5.04 software (GraphPad Software, La Jolla, CA). Cell number andpercentage were presented as median and interquartile range(IQR).

3. Results and discussion

Median (IQR) value of EPCs was 1.88 (1.35–2.85) cells permicroliter (Table 1). The average proportion of EPCs as related toall CD45+ leukocytes was 0.03% (0.02–0.04%). The test for EPCassay reliability revealed a coefficient of variation at 6.8% and8.2% between aliquots and time points, respectively (Table 2).

a b c

d e f

Fig. 2. Multicolor flow cytometric analysis of human monocyte subsets. Following scatter exclusion of debris (a), the sequential analysis continues with gating ofCD45+ leukocytes (P2 in green in b) and the monocyte scatter population (P3 in grey in c). Then, the three major monocyte subsets are separated on a CD14/CD16 dot plot by using quadrant marker (d). Counting beads are gated on a separate SSC/CD45 dot plot showing all events (P4 in blue in e). (f) Populationhierarchy.

12 M. Hristov et al. / Journal of Immunological Methods 381 (2012) 9–13

Analysis of monocyte subsets resulted in 329.5 (264.5–374.8),16.0 (8.0–22.2) and 26.5 (19.8–36.3) cells per microliter forclassical, intermediate and non-classical monocytes (Table 1).The respective proportion for each subset persisted at 85.4%(83.2–88.1%), 4.1% (2.5–4.9%) and 7.5% (6.4–8.3%) of totalmonocytes. The coefficients of variation between blood sampleswere 4.6%, 13.5% and 8.5% for classical, intermediate and non-classical monocytes (Table 2). Corresponding coefficients ofvariation between the two time points of blood collectionremained by 9.2%, 19.4% and 18.8%.

As major advancementwe use now lyse/no-wash approachand determination of absolute cell numbers by counting beads.Analysis of EPCs is partially adapted to the sequential gating ofthe ISHAGE guidelines for flow cytometric enumeration ofCD34+ hematopoietic stem cells (Sutherland et al., 1996). Thelyse/no-wash procedure further prevents potential cell lossor activation as this may occur during washing/centrifugation.In addition, analysis of monocytes is now complemented byinitially gating-out cell debris and setting a gate on CD45+ leu-kocytes. We also do not apply tandem dyes anymore. Instead,

Table 1Absolute number and proportion of human EPCs and monocyte subsets.

Cell subsets Number per μ

CD45dimCD34+VEGFR2+ EPCs 1.88 (1.35–2Classical CD14++CD16- monocytes 329.5 (264.5–Intermediate CD14++CD16+ monocytes 16.0 (8.0–22Non-classical CD14+CD16++ monocytes 26.5 (19.8–3

Values are median (IQR).

standard fluorochromes are used as three-color panels atappropriate compensation. This may extend our protocol alsoto flow cytometers with a basic 2-laser-4-color configurationstimulating users to adapt our assay to their own equipment.Flow cytometric assessment of circulating EPCs belongs tothe “rare events analysis”, as these cells display weak antigenexpression and their frequency at steady-state conditions israther low. Such an analysis requires certain efforts in order toobtain reliable data (Dahmen et al., 2002). For instance, wematch high-intensity fluorochromes (PE, APC) to low-densitymarkers (VEGFR2, CD34) and use the FMO control to properlyassess the cut-off for true positive cells (Maecker et al., 2004;Maecker and Trotter, 2006; Hristov et al., 2009). Our rare eventanalysis is further supported by regular calibration and longerwarming-up time of the cytometer to compensate for varia-tions in the environment of the instrument. Moreover, oursequential gating precisely confirms cell size, clustering andviability. This gating also eliminates background noise andpotential unspecific antibody binding by platelets, lysed redblood cells or other debris. Finally, we use mAb clones that are

l Fraction

.85) 0.03% (0.02–0.04%) of CD45+ leukocytes374.8) 85.4% (83.2–88.1%) of total monocytes.2) 4.1% (2.5–4.9%) of total monocytes6.3) 7.5% (6.4–8.3%) of total monocytes

Table 2Absolute cell number per microliter as measured in two separate blood samples obtained at two time points.

Monocytes EPCs

Time point I Time point II Timepoint I

Timepoint II

Cl Int Ncl Cl Int Ncl

Donor 1 Sample 1 423 12 14 362 6 13 0.95 1.02Sample 2 420 11 15 426 8 16 0.93 1.14

Donor 2 Sample 1 257 16 35 348 12 18 3.10 3.25Sample 2 283 14 39 337 13 22 3.31 3.69

Donor 3 Sample 1 432 18 18 381 14 29 3.18 2.77Sample 2 445 11 15 410 13 33 2.84 2.98

Donor 4 Sample 1 379 8 17 456 12 18 1.09 0.91Sample 2 434 12 18 460 14 20 0.99 1.20

Donor 5 Sample 1 301 8 16 246 8 16 1.97 1.73Sample 2 282 10 16 256 8 14 1.93 1.83

Cl = classical, Int = intermediate, Ncl = non-classical.

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well established for reliable detection in hematological assays.Together, these strategies increase the sensitivity and specific-ity of our rare event EPC experiment.

As an advantage to the protocol proposed by Schmidt-Luckeet al. (2010), we use FMO control and a fluorochrome with ahigher stain-index for the anti-CD34 mAb, together with deadcell exclusion and counting beads. Comparing our assay to themethod by Farace et al. (2011), we use the lyse/no-washwholeblood procedure with sequential gating and FMO—instead ofisotype control. Enhancing the protocol by Heimbeck et al.(2010), we initially exclude cell debris and do not analyzeCD16 monocytes as a pooled population but separate “non-classical” CD14+(low)CD16++ monocytes from the “interme-diate” CD14++CD16+(mid) subset (Hristov and Weber, 2011;Ziegler-Heitbrock et al., 2010). We have also omitted the anti-HLA-DR antibody and instead tightened the monocyte gate inthe scatter density plot to exclude potential “contamination”by granulocytes or NK cells.

In conclusion, our modified protocol becomes now quantita-tive,more sensitive, rapid and adapted for commonpractical use.Since accumulating evidence advises the monitoring of circulat-ing EPCs and monocyte subsets as novel surrogate prognostic/diagnostic cellular biomarker, we believe that this advancedversion of our assay should be of broad clinical relevance.

Acknowledgments

This work was supported by the Deutsche Forschungsge-meinschaft (FOR809, HR 18/1-1) and the August-Lenz-Stiftung.

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