> A unique feature of the P2X7 receptor observed under physiological conditions is the slow further increase in permeability that develops after the initial opening of the P2X7 channel, which is readily studied by flow cytometry (Wiley *et al.*, 1998). Ethidium bromide is a phenanthridinium intercalator which binds both DNA and RNA and is generally excluded from viable cells. It has been used previously to assess cell permeabilization by ATP (Gomperts, 1983). Once bound to nucleic acids, the fluorescence is enhanced 20~30 fold. The excitation maximum is shifted to 512 nm and the emission maximum is shifted to 605 nm. This long emission wavelength allows simultaneous detection of ethidium+ influx on the FL2 photomultiplier (570 to 610 nm) in the presence of FITC-labeled antibodies which are detected on the FL1 photomultiplier (525 nm). Time-resolved flow cytometry generates the mean fluorescence intensity of analysed cells over a certain time period. This technique

Broad Applications of Multi-Colour Time-Resolved Flow Cytometry 193

*(to calculate the average linear mean channel of basal line in the same period before ATP is added into*

Function(C5)=AVERAGE(C\$7:INDIRECT(ADDRESS(D4+5,COLUMN(),4,1),1))

Function(D5) =AVERAGE(D\$7:INDIRECT(ADDRESS(D4+5,COLUMN(),4,1),1))

Fig. 6. Typical dotplots and gating strategy used to analyse ethidium+ uptake by CD14

**Ethidium+ Uptake**

Fig. 7. An example of spreadsheet (left) and a typical curve of ethidum+ uptake (right)

(Linear MFI/cell)

0

50

**ATP**

100

150

200

Time (min) 0123456

Basal ATP

*(to calculate the average linear mean channel of basal line before ATP is added)*

*the test tube)*

positive monocytes

Function(D4) =MATCH(0,D\$7:D\$30,0) *(to locate in which interval that ATP is added)*

allows a sensitive measurement of the initial rate of ethidium uptake, which is essentially unidirectional because of binding of this permeant cation to nucleic acids. By using timeresolved flow cytometry, our group has shown that there is large variation of P2X7 function among individuals (Gu *et al.*, 2000) mainly due to genetic polymorphisms which alter the function of this receptor (Gu *et al.*, 2004; Gu *et al.*, 2001; Shemon *et al.*, 2006; Skarratt *et al.*, 2005; Wiley *et al.*, 2003)

Previous methods used to measure surface P2X7 function have been semi-quantitative and unable to distinguish sub-populations within the overall cell suspension as well as being unable to distinguish live and dead cells. The two-colour time resolved flow cytometry methods described here allow quantitative assessment of the abundance of functional P2X7 receptors on the surface of different subtypes of leukocytes as well as excluding dead cells from analysis (Gu *et al.*, 2000; Jursik *et al.*, 2007).

#### **3.2 Method**

Mononuclear cells (2x106) pre-labeled with FITC-conjugated cell markers are washed once and resuspended in 100 μL Na medium with 0.1 mM Ca2+ at room temperature. Following the addition of 900 μL K medium, a small magnetic stir bar is added to the tube before it is inserted into the Time-Zero System which controls temperature and allows magnetic stirring. The FL2 voltage is set at around 595V with a gain of 5.0, at which the linear mean channel fluorescence intensity for Quantum PE standard beads with MESF 300747 is 48±1 (256 linear scale) and the peak channel for right reference standard PE high level beads (MESF ~560,000) is 100±1 (256 linear scale). The compensation of FL1-FL2 and FL2-FL1 is 7% and 8% respectively. Ethidium bromide (25 μM) is added, followed 40 s later by addition of 1.0 mM ATP. Mononuclear cells are acquired at approximately 1000 events per second by a Becton Dickinson FACSCalibur flow cytometer and the data comprising forward scatter (log mode), side scatter (log mode), FL1 (log mode), FL2 (linear mode, 256 channels) and time (5 sec intervals) for each event are collected. The data of each run (approximately 380 sec) is saved into a listmode file.

#### **3.3 Gating and calculation**

The listmode file is analysed by WinMDI. Cells are gated by forward and side scatter and by cell type specific antibodies (Fig. 6). Cells with maximum ethidium+ uptake (254 channels or over) are considered as fully permeable necrotic cells and therefore excluded from the assay. The kinetic linear mean channel of fluorescence intensity for each gated subpopulation over successive 5 sec intervals is plotted against time and saved into a text file.

To quantify ethidium+ uptake, arbitrary unit of area under the uptake curve in the first 5 min after addition of ATP is calculated. The basal line (without ATP) is firstly adjusted to the same level as the test line (with ATP added after 40 sec). The area under the ethidium uptake curve between the basal and test lines is considered as the sum of 60 trapezoids (Fig. 1a).

The text file is then imported into *Microsoft* Excel, and the area under the ethidium+ uptake curve is calculated using the following functions:

Function(C5)=AVERAGE(C\$7:INDIRECT(ADDRESS(D4+5,COLUMN(),4,1),1)) *(to calculate the average linear mean channel of basal line in the same period before ATP is added into the test tube)*

Function(D4) =MATCH(0,D\$7:D\$30,0) *(to locate in which interval that ATP is added)*

192 Flow Cytometry – Recent Perspectives

allows a sensitive measurement of the initial rate of ethidium uptake, which is essentially unidirectional because of binding of this permeant cation to nucleic acids. By using timeresolved flow cytometry, our group has shown that there is large variation of P2X7 function among individuals (Gu *et al.*, 2000) mainly due to genetic polymorphisms which alter the function of this receptor (Gu *et al.*, 2004; Gu *et al.*, 2001; Shemon *et al.*, 2006; Skarratt *et al.*,

Previous methods used to measure surface P2X7 function have been semi-quantitative and unable to distinguish sub-populations within the overall cell suspension as well as being unable to distinguish live and dead cells. The two-colour time resolved flow cytometry methods described here allow quantitative assessment of the abundance of functional P2X7 receptors on the surface of different subtypes of leukocytes as well as excluding dead cells

Mononuclear cells (2x106) pre-labeled with FITC-conjugated cell markers are washed once and resuspended in 100 μL Na medium with 0.1 mM Ca2+ at room temperature. Following the addition of 900 μL K medium, a small magnetic stir bar is added to the tube before it is inserted into the Time-Zero System which controls temperature and allows magnetic stirring. The FL2 voltage is set at around 595V with a gain of 5.0, at which the linear mean channel fluorescence intensity for Quantum PE standard beads with MESF 300747 is 48±1 (256 linear scale) and the peak channel for right reference standard PE high level beads (MESF ~560,000) is 100±1 (256 linear scale). The compensation of FL1-FL2 and FL2-FL1 is 7% and 8% respectively. Ethidium bromide (25 μM) is added, followed 40 s later by addition of 1.0 mM ATP. Mononuclear cells are acquired at approximately 1000 events per second by a Becton Dickinson FACSCalibur flow cytometer and the data comprising forward scatter (log mode), side scatter (log mode), FL1 (log mode), FL2 (linear mode, 256 channels) and time (5 sec intervals) for each event are collected. The data of each run (approximately 380 sec) is

The listmode file is analysed by WinMDI. Cells are gated by forward and side scatter and by cell type specific antibodies (Fig. 6). Cells with maximum ethidium+ uptake (254 channels or over) are considered as fully permeable necrotic cells and therefore excluded from the assay. The kinetic linear mean channel of fluorescence intensity for each gated subpopulation over

To quantify ethidium+ uptake, arbitrary unit of area under the uptake curve in the first 5 min after addition of ATP is calculated. The basal line (without ATP) is firstly adjusted to the same level as the test line (with ATP added after 40 sec). The area under the ethidium uptake curve between the basal and test lines is considered as the sum of 60 trapezoids (Fig.

The text file is then imported into *Microsoft* Excel, and the area under the ethidium+ uptake

successive 5 sec intervals is plotted against time and saved into a text file.

curve is calculated using the following functions:

2005; Wiley *et al.*, 2003)

saved into a listmode file.

**3.3 Gating and calculation**

1a).

**3.2 Method**

from analysis (Gu *et al.*, 2000; Jursik *et al.*, 2007).

Function(D5) =AVERAGE(D\$7:INDIRECT(ADDRESS(D4+5,COLUMN(),4,1),1)) *(to calculate the average linear mean channel of basal line before ATP is added)*

Fig. 6. Typical dotplots and gating strategy used to analyse ethidium+ uptake by CD14 positive monocytes

Fig. 7. An example of spreadsheet (left) and a typical curve of ethidum+ uptake (right)

Broad Applications of Multi-Colour Time-Resolved Flow Cytometry 195

Leukocyte subtypes are identified by monoclonal antibodies and we observed that unconjugated anti-CD14, CD3, CD19 or CD16 antibody did not affect ATP-induced ethidium+ uptake. Thus, with the correct FL2-FL1 compensation, the positively gated population should give a similar value of area under ethidium+ uptake curve as the negatively gated population. While the extremely bright fluorescence of ethidium+ makes for accuracy in the uptake measurement, it does make the correct compensation between FL1 and FL2 essential. While under-compensation of FL2-FL1 leads to high basal level of ethidium+ uptake, any over-compensation of FL2-FL1 dramatically reduces the area under ethidium+ uptake curves. Meanwhile, if the FL1-FL2 compensation is too high or too low, the R2 gated population (Fig. 6) will incline towards left or right, respectively. Ethidium+ uptake can also exclude the necrotic or apoptotic cells from the live cells in the suspension, simply by excluding fluorescent events at the maximum fluorescent channel intensity at

Phagocytosis is a fundamental aspect of the innate immune system which is preserved in specialized cells of all metazoans. Human monocytes suspended in saline media (without serum) rapidly phagocytose a range of foreign particles which are internalized into a phagosome via rearrangement of the actin-myosin cytoskeleton. This innate immune function requires recognition of foreign particles by one or more scavenger receptors on the monocyte surface. We have recently shown that an intact P2X7-nonmuscle myosin heavy chain IIA complex in monocyte/macrophages can regulate the phagocytosis of a range of non-opsonized particles including latex beads, and live and dead *Staphylococcus aureus* and

Technical advances in the assessment of phagocytosis have allowed rapid advances in our knowledge of molecular interactions associated with engulfment of particles by phagocytes. Confocal microscopy has shown that internal membranes within the cell fuse with plasma membrane during the course of particle ingestion, and that recycling endosomes are the primary source of membrane for enlargement of phagocytic cup (Touret *et al.*, 2005). Flow cytometric assessment of particle engulfment has to some extent replaced microscopic observation (Steinkamp *et al.*, 1982) particularly as the kinetics of uptake of fluorescent targets by phagocytes can be followed by instruments capable of time-resolved measurements in a stirred cuvette at 37ºC (Gu *et al.*, 2010). These assays of phagocytosis by flow cytometry usually include measurements of fluorescence particle uptake by cells preincubated with cytochalasin D (CytD), an inhibitor of F-actin polymerization and phagocytic cup formation and this control condition allows the assay to distinguish engulfment of particles from adhesion. Particle size is also important in flow studies which generally

Various methods have been employed in phagocytosis studies. However, no published method reflects the quantitative particle uptake in real time although this parameter is important to fully assess the engulfment ability of phagocytes. In this section, we describe a quantitative method to measure the phagocytic ability of human monocytes using time-

zero time which defines the fully permeabilized cells present.

employ fluorescent particles ranging from 1 to 3 μm in diameter.

**4. Phagocytosis of fluorescent latex beads**

**4.1 Introduction**

*Escherichia coli* (Gu *et al.*, 2010).

resolved two-colour flow cytometry.

Function(D6)

```
=2.5*(SUM(INDIRECT(ADDRESS(D4+8,COLUMN(),4,1),1):INDIRECT(ADDRESS(D4+5*12
+8,COLUMN(),4,1),1))+SUM(INDIRECT(ADDRESS(D4+9,COLUMN(),4,1),1):INDIRECT(A
DDRESS(D4+5*12+7,COLUMN(),4,1),1))-SUM(INDIRECT(ADDRESS(D4+8,COLUMN()-
1,4,1),1):INDIRECT(ADDRESS(D4+5*12+8,COLUMN()-1,4,1),1))-
SUM(INDIRECT(ADDRESS(E4+9,COLUMN()-
1,4,1),1):INDIRECT(ADDRESS(D4+5*12+7,COLUMN()-1,4,1),1))-(D5-C5)*120)
(to calculate uptake curve in the first 5 min after area under ethidium+ uptake curve after addition of
ATP)
```
The arbitrary unit obtained in Function(D6) is used to quantify the P2X7 pore function. Time frame chosen for calculation of ATP-induced ethidium+ uptake is 5 min. This is simply because the linear response after addition of ATP is about 5 min. However, in monocytes from some healthy subjects with high P2X7 function, ATP may induce a linear response only in the first 2 or 3 min before the fluorescence of cell-associated ethidium reaches the maximum channel number. The calculation model can be readily adapted to shorter time points on the ethidium+ uptake curve. To do this, the time point in above function (5\*) can be replaced by 2\*, 3\* or 4\* for 2, 3 or 4 min respectively.

#### **3.4 Notes for technique**

The peak channel of Standard PE Reference Beads measured on linear mode in FL2 can be as large as ±20% from one day to another. Therefore it is essential to calibrate the instrument daily before each set of experiments. Furthermore, ethidium+ concentration, temperature and the composition of the suspending buffer are critical parameters. The area under the ethidium+ uptake curve increases proportionally with the ethidium+ concentration up to about 100 µM. Excess ethidium+ however might bind to mitochondrial membranes and lead to inaccurate results. Moreover, P2X7 function is dependent on temperature. We have shown that P2X7 agonists fail to induce ethidium+ uptake at 12ºC, which is gradually restored by a temperature rise up to a physiological value of 37ºC. Results are further altered by the absence or presence of Ca2+, Cl- and Na+ ions in the acquisition medium. Small amounts of extracellular Ca2+ (~10 μM) in the medium are always included to maintain cell membrane integrity. However, high concentration of Ca2+ may reduce the ATP activity by binding to its active species, ATP4- or by direct competition for permeation (Wiley *et al.*, 1996). Isotonic K+ or sucrose media remove the inhibitory effect of Na+ and/or Cl- ions on P2X7 function, which facilitates the detection of differences in P2X7 function among individuals. Since multiple factors cause large alterations in the area under the ethidium uptake curve, it is essential to keep standardized assay conditions.

ATP-induced ethidium+ uptake is directly proportional to the concentration of ethidium cation over the range 1 to 100 μM, consistent with permeation of ethidium+ through a dilated pore. 25 μM ethidium bromide is routinely used for measurement of uptakes over a 3 to 5 minute time course. The pore formation by activated P2X7 is also sensitive to temperature. P2X7 function decreases at lower temperature while at 12ºC or lower, ATPinduced ethidium+ uptake is almost completely abolished. It has been reported that removal of extracellular Cl as well as extracellular Na+ enhances permeability responses and stimulates the function of the P2X7 receptor (Michel *et al.*, 1999). P2X7 function measured by ATP-induced ethidium+ uptake is greatest in Na+ free, Cl- free sucrose medium, less in KCl medium, and least in NaCl medium.

Leukocyte subtypes are identified by monoclonal antibodies and we observed that unconjugated anti-CD14, CD3, CD19 or CD16 antibody did not affect ATP-induced ethidium+ uptake. Thus, with the correct FL2-FL1 compensation, the positively gated population should give a similar value of area under ethidium+ uptake curve as the negatively gated population. While the extremely bright fluorescence of ethidium+ makes for accuracy in the uptake measurement, it does make the correct compensation between FL1 and FL2 essential. While under-compensation of FL2-FL1 leads to high basal level of ethidium+ uptake, any over-compensation of FL2-FL1 dramatically reduces the area under ethidium+ uptake curves. Meanwhile, if the FL1-FL2 compensation is too high or too low, the R2 gated population (Fig. 6) will incline towards left or right, respectively. Ethidium+ uptake can also exclude the necrotic or apoptotic cells from the live cells in the suspension, simply by excluding fluorescent events at the maximum fluorescent channel intensity at zero time which defines the fully permeabilized cells present.

#### **4. Phagocytosis of fluorescent latex beads**

#### **4.1 Introduction**

194 Flow Cytometry – Recent Perspectives

=2.5\*(SUM(INDIRECT(ADDRESS(D4+8,COLUMN(),4,1),1):INDIRECT(ADDRESS(D4+5\*12 +8,COLUMN(),4,1),1))+SUM(INDIRECT(ADDRESS(D4+9,COLUMN(),4,1),1):INDIRECT(A DDRESS(D4+5\*12+7,COLUMN(),4,1),1))-SUM(INDIRECT(ADDRESS(D4+8,COLUMN()-

*(to calculate uptake curve in the first 5 min after area under ethidium+ uptake curve after addition of*

The arbitrary unit obtained in Function(D6) is used to quantify the P2X7 pore function. Time frame chosen for calculation of ATP-induced ethidium+ uptake is 5 min. This is simply because the linear response after addition of ATP is about 5 min. However, in monocytes from some healthy subjects with high P2X7 function, ATP may induce a linear response only in the first 2 or 3 min before the fluorescence of cell-associated ethidium reaches the maximum channel number. The calculation model can be readily adapted to shorter time points on the ethidium+ uptake curve. To do this, the time point in above function (5\*) can

The peak channel of Standard PE Reference Beads measured on linear mode in FL2 can be as large as ±20% from one day to another. Therefore it is essential to calibrate the instrument daily before each set of experiments. Furthermore, ethidium+ concentration, temperature and the composition of the suspending buffer are critical parameters. The area under the ethidium+ uptake curve increases proportionally with the ethidium+ concentration up to about 100 µM. Excess ethidium+ however might bind to mitochondrial membranes and lead to inaccurate results. Moreover, P2X7 function is dependent on temperature. We have shown that P2X7 agonists fail to induce ethidium+ uptake at 12ºC, which is gradually restored by a temperature rise up to a physiological value of 37ºC. Results are further altered by the absence or presence of Ca2+, Cl- and Na+ ions in the acquisition medium. Small amounts of extracellular Ca2+ (~10 μM) in the medium are always included to maintain cell membrane integrity. However, high concentration of Ca2+ may reduce the ATP activity by binding to its active species, ATP4- or by direct competition for permeation (Wiley *et al.*, 1996). Isotonic K+ or sucrose media remove the inhibitory effect of Na+ and/or Cl- ions on P2X7 function, which facilitates the detection of differences in P2X7 function among individuals. Since multiple factors cause large alterations in the area under the

ethidium uptake curve, it is essential to keep standardized assay conditions.

ATP-induced ethidium+ uptake is directly proportional to the concentration of ethidium cation over the range 1 to 100 μM, consistent with permeation of ethidium+ through a dilated pore. 25 μM ethidium bromide is routinely used for measurement of uptakes over a 3 to 5 minute time course. The pore formation by activated P2X7 is also sensitive to temperature. P2X7 function decreases at lower temperature while at 12ºC or lower, ATPinduced ethidium+ uptake is almost completely abolished. It has been reported that removal

stimulates the function of the P2X7 receptor (Michel *et al.*, 1999). P2X7 function measured by ATP-induced ethidium+ uptake is greatest in Na+ free, Cl- free sucrose medium, less in KCl

as well as extracellular Na+ enhances permeability responses and

1,4,1),1):INDIRECT(ADDRESS(D4+5\*12+8,COLUMN()-1,4,1),1))-

1,4,1),1):INDIRECT(ADDRESS(D4+5\*12+7,COLUMN()-1,4,1),1))-(D5-C5)\*120)

SUM(INDIRECT(ADDRESS(E4+9,COLUMN()-

be replaced by 2\*, 3\* or 4\* for 2, 3 or 4 min respectively.

Function(D6)

*ATP)*

**3.4 Notes for technique**

of extracellular Cl-

medium, and least in NaCl medium.

Phagocytosis is a fundamental aspect of the innate immune system which is preserved in specialized cells of all metazoans. Human monocytes suspended in saline media (without serum) rapidly phagocytose a range of foreign particles which are internalized into a phagosome via rearrangement of the actin-myosin cytoskeleton. This innate immune function requires recognition of foreign particles by one or more scavenger receptors on the monocyte surface. We have recently shown that an intact P2X7-nonmuscle myosin heavy chain IIA complex in monocyte/macrophages can regulate the phagocytosis of a range of non-opsonized particles including latex beads, and live and dead *Staphylococcus aureus* and *Escherichia coli* (Gu *et al.*, 2010).

Technical advances in the assessment of phagocytosis have allowed rapid advances in our knowledge of molecular interactions associated with engulfment of particles by phagocytes. Confocal microscopy has shown that internal membranes within the cell fuse with plasma membrane during the course of particle ingestion, and that recycling endosomes are the primary source of membrane for enlargement of phagocytic cup (Touret *et al.*, 2005). Flow cytometric assessment of particle engulfment has to some extent replaced microscopic observation (Steinkamp *et al.*, 1982) particularly as the kinetics of uptake of fluorescent targets by phagocytes can be followed by instruments capable of time-resolved measurements in a stirred cuvette at 37ºC (Gu *et al.*, 2010). These assays of phagocytosis by flow cytometry usually include measurements of fluorescence particle uptake by cells preincubated with cytochalasin D (CytD), an inhibitor of F-actin polymerization and phagocytic cup formation and this control condition allows the assay to distinguish engulfment of particles from adhesion. Particle size is also important in flow studies which generally employ fluorescent particles ranging from 1 to 3 μm in diameter.

Various methods have been employed in phagocytosis studies. However, no published method reflects the quantitative particle uptake in real time although this parameter is important to fully assess the engulfment ability of phagocytes. In this section, we describe a quantitative method to measure the phagocytic ability of human monocytes using timeresolved two-colour flow cytometry.

Broad Applications of Multi-Colour Time-Resolved Flow Cytometry 197

The area under the YG bead uptake curve is calculated over the time frame of 6.5 min, because the uptake of beads is usually maximal at this time point. However, the calculation model can be readily adapted to different time points on the YG bead uptake curve. To do this, the time point in above function (ie 6.5) can be replaced by the desired time in minutes.

**YG Beads Uptake**

(Linear MFI / cell)

Fig. 9. An example of spreadsheet (left) and a typical curve of YG bead uptake (right)

neutrophils which contaminate the Ficoll separated mononuclear population.

is used and the flow cytometer is carefully calibrated from day-to-day.

**4.4 Notes for technique**

from above factors.

0

The time resolved flow cytometry based method described here allows real-time quantitative assessment of the phagocytic function of peripheral blood monocytes labeled with APC conjugated CD14 antibody. The excitation and emission wavelength of APC is well separated from the YG-beads containing fluorescein (APC: Ex. 633nm, Em. 660nm; YG: Ex. 441nm, Em. 486nm), thus the compensation setting of the flow cytometer can be ignored. Although lymphocytes do not phagocytose 1 µm YG bead in this setting (Gu *et al*., 2010), the CD14+ gating is necessary to exclude uptake of YG beads by the small number of

Since the bead size is much less than cell size, multiple beads transit through the laser beam together with the cell, leading to a considerable fluorescence background. In addition, some YG beads may also adhere to cell surface non-specifically. Thus cytochalasin D (a potent phagocytosis inhibitor) is used to assess fluorescence intensity due to background resulting

Many factors can affect this phagocytosis assay including the nature of bead surface. The total uptake of negatively charged carboxylate beads is higher than the uptake of uncharged beads suggesting the negatively charged carboxyl groups on the bead surface facilitates their uptake by monocytes. The bead size, cell:bead ratio, pH and temperature are critical parameters affecting uptake. Uptake of 1 µm beads is maximal at a pH 6.5-7.5 and 37°C in serum-free Na medium with 0.1 mM Ca2+. This method can be used to compare the phagocytic ability of monocytes from different individuals, especially if a standard protocol

100

200

300

400

500

Basal ATP Cytochalasin D

**YG beads**

600

Time (min)

01234

#### **4.2 Method**

Mononuclear cells (2×106 in 100 μL) are pre-labeled with APC conjugated-anti CD14 followed by addition of 900 μL Na medium with 0.1 mM Ca2+ with a small magnetic stir bar. The tube is inserted into the Time-Zero System (from Cytek Development, Fremont, CA, USA) which monitors temperature (37ºC) and allows magnetic stirring. 5 to 10 μL yellowgreen carboxylated fluorescent polystyrene latex microspheres (YG bead, 1 µm, from Polyscience, Warrington, PA) are added 20 sec later. Linear MFI of YG fluorescence is collected in FL1 (voltage: 380-420, gain: 2.0). Events are acquired at about 1500 events per second by a Becton Dickinson FACSCalibur flow cytometer. The data of each run are collected for about 7 min and saved into a listmode file.

#### **4.3 Gating and calculation**

The listmode file is analysed by WinMDI. Cells are gated by forward and side scatter and by cell type specific antibodies (Fig. 8). The linear mean channel of fluorescence intensity for each gated subpopulation over successive 10 sec intervals is plotted against time to yield kinetic data which is saved into a text file.

Fig. 8. Typical dotplots and gating strategy used to analyse YG bead uptake by CD14+ monocytes

To quantify bead uptake, arbitrary units of area under the uptake curve in first 6.5 min after addition of beads is calculated. The area is considered as the sum of 39 trapezoids, which is calculated by the following mathematical model. The text file is then imported into *Microsoft*  Excel, and mathematical calculation of the area under the bead uptake curve is programmed using the following functions:

C5, D5 and E5 are the lowest value among C6, D6 and E6, represent the threshold to exclude the background fluorescence intensity.

Function(C6)=INDIRECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+10,COLUMN(),4,1),1) *(To calculate initial levels of fluorescence intensity)*

Function(C7)=5\*(C\$6+2\*SUM(INDIRECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+11,COLUMN (),4,1),1):INDIRECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+6.5\*6+9,COLUMN(),4,1),1))+INDI RECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+6.5\*6+10,COLUMN(),4,1),1))-C\$6\*6.5\*6\*10 *(To calculate area under YG bead uptake curve in the first 6.5 min)*

The area under the YG bead uptake curve is calculated over the time frame of 6.5 min, because the uptake of beads is usually maximal at this time point. However, the calculation model can be readily adapted to different time points on the YG bead uptake curve. To do this, the time point in above function (ie 6.5) can be replaced by the desired time in minutes.

Fig. 9. An example of spreadsheet (left) and a typical curve of YG bead uptake (right)

#### **4.4 Notes for technique**

196 Flow Cytometry – Recent Perspectives

Mononuclear cells (2×106 in 100 μL) are pre-labeled with APC conjugated-anti CD14 followed by addition of 900 μL Na medium with 0.1 mM Ca2+ with a small magnetic stir bar. The tube is inserted into the Time-Zero System (from Cytek Development, Fremont, CA, USA) which monitors temperature (37ºC) and allows magnetic stirring. 5 to 10 μL yellowgreen carboxylated fluorescent polystyrene latex microspheres (YG bead, 1 µm, from Polyscience, Warrington, PA) are added 20 sec later. Linear MFI of YG fluorescence is collected in FL1 (voltage: 380-420, gain: 2.0). Events are acquired at about 1500 events per second by a Becton Dickinson FACSCalibur flow cytometer. The data of each run are

The listmode file is analysed by WinMDI. Cells are gated by forward and side scatter and by cell type specific antibodies (Fig. 8). The linear mean channel of fluorescence intensity for each gated subpopulation over successive 10 sec intervals is plotted against time to yield

Fig. 8. Typical dotplots and gating strategy used to analyse YG bead uptake by CD14+

To quantify bead uptake, arbitrary units of area under the uptake curve in first 6.5 min after addition of beads is calculated. The area is considered as the sum of 39 trapezoids, which is calculated by the following mathematical model. The text file is then imported into *Microsoft*  Excel, and mathematical calculation of the area under the bead uptake curve is programmed

C5, D5 and E5 are the lowest value among C6, D6 and E6, represent the threshold to exclude

Function(C7)=5\*(C\$6+2\*SUM(INDIRECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+11,COLUMN (),4,1),1):INDIRECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+6.5\*6+9,COLUMN(),4,1),1))+INDI RECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+6.5\*6+10,COLUMN(),4,1),1))-C\$6\*6.5\*6\*10

Function(C6)=INDIRECT(ADDRESS(MATCH(30,C\$9:C\$32,1)+10,COLUMN(),4,1),1)

collected for about 7 min and saved into a listmode file.

**4.2 Method**

monocytes

using the following functions:

the background fluorescence intensity.

*(To calculate initial levels of fluorescence intensity)*

*(To calculate area under YG bead uptake curve in the first 6.5 min)*

**4.3 Gating and calculation**

kinetic data which is saved into a text file.

The time resolved flow cytometry based method described here allows real-time quantitative assessment of the phagocytic function of peripheral blood monocytes labeled with APC conjugated CD14 antibody. The excitation and emission wavelength of APC is well separated from the YG-beads containing fluorescein (APC: Ex. 633nm, Em. 660nm; YG: Ex. 441nm, Em. 486nm), thus the compensation setting of the flow cytometer can be ignored. Although lymphocytes do not phagocytose 1 µm YG bead in this setting (Gu *et al*., 2010), the CD14+ gating is necessary to exclude uptake of YG beads by the small number of neutrophils which contaminate the Ficoll separated mononuclear population.

Since the bead size is much less than cell size, multiple beads transit through the laser beam together with the cell, leading to a considerable fluorescence background. In addition, some YG beads may also adhere to cell surface non-specifically. Thus cytochalasin D (a potent phagocytosis inhibitor) is used to assess fluorescence intensity due to background resulting from above factors.

Many factors can affect this phagocytosis assay including the nature of bead surface. The total uptake of negatively charged carboxylate beads is higher than the uptake of uncharged beads suggesting the negatively charged carboxyl groups on the bead surface facilitates their uptake by monocytes. The bead size, cell:bead ratio, pH and temperature are critical parameters affecting uptake. Uptake of 1 µm beads is maximal at a pH 6.5-7.5 and 37°C in serum-free Na medium with 0.1 mM Ca2+. This method can be used to compare the phagocytic ability of monocytes from different individuals, especially if a standard protocol is used and the flow cytometer is carefully calibrated from day-to-day.

Broad Applications of Multi-Colour Time-Resolved Flow Cytometry 199

=0.5\*5\*(INDIRECT(ADDRESS(C\$4+7,COLUMN(),4,1),1)+2\*SUM(INDIRECT(ADDRESS(C\$ 4+8,COLUMN(),4,1),1):INDIRECT(ADDRESS(C\$4+12\*12+6,COLUMN(),4,1),1))+INDIRECT

*To transform the log MFI to artificial linear MFI. The rest data in Column D are transformed using*

=0.5\*5\*(INDIRECT(ADDRESS(D\$4+7,COLUMN(),4,1),1)+2\*SUM(INDIRECT(ADDRESS(D\$ 4+8,COLUMN(),4,1),1):INDIRECT(ADDRESS(D\$4+12\*12+6,COLUMN(),4,1),1))+INDIRECT

Fig. 10. Typical dotplots and gating strategy used to analyse protein-protein interaction. **a.** The FSC and SSC of HEK-293 cells gated by gate R1. **b.** The AcGFP+DsRed+ HEK-293 cells gated by R2. **c.** Changes in fluorescence of AcGFP tagged NMMHC-IIA. **d.** Kinetic P2X7

Function (C5)=AVERAGE(C\$7:INDIRECT(ADDRESS(C\$4+5,COLUMN(),4,1),1))

(ADDRESS(C\$4+12\*12+7,COLUMN(),4,1),1))-C\$5\*12\*12\*5

(ADDRESS(D\$4+12\*12+7,COLUMN(),4,1),1))-D\$5\*12\*12\*5

(a) (b)

(c) (d)

*To calculate area above/under curve (Log mode)*

*the same function by "Ctrl-D" command.*

Funciton(D7)=IF(C7=0,0,POWER(10,(C7+1)/256))

*To calculate area above/under curve (Linear mode)*

with time of DsRed tagged fluorescence

*To calculate the base line.*

Function(C6)

Function(D6)

#### **5. Protein complex dissociation**

#### **5.1 Principle**

Fluorescence resonance energy transfer (FRET) is one of the major techniques used for protein interaction studies. Confocal or fluorescent microscopes with photobleach or fluorescence lifetime modules are the common instruments which perform FRET based measurements. However, microscopy can only detect fluorescence signals in a single cell, and cannot measure multiple fluorescence emission wavelengths simultaneously. This limitation on confocal microscope makes flow cytometer an attractive alternative for FRET based measurement. The time-resolved fluorescence decay measurement by flow cytometry has been used to study molecule interactions (Condrau *et al.*, 1994a; Condrau *et al.*, 1994b; Deka & Steinkamp, 1996). However, this technique requires major modification to a flow cytometer since a pulsed laser has to be used in order to measure fluorescence lifetime.

The P2X7 receptor has been shown to form a protein complex with nonmuscle myosin heavy chain IIA (NMMHC-IIA) in its unactivated state, and this complex dissociates following activation of P2X7 by extracellular ATP (Gu *et al.*, 2009). To study the interaction of these two proteins in a large cohort of live cells, we developed a time-resolved FRET (TR-FRET) flow cytometry method. The HEK-293 cells are transfected with AcGFP-tagged NMMHC-IIA and DsRed-tagged P2X7. The NMMHC-P2X7 interaction is assessed by monitoring the fluorescence intensity changes of AcGFP and DsRed in double positive cells over 10-15 minutes after addition of P2X7 agonist. This method does not require any modification of the flow cytometer, and can be readily adapted to study the interaction within the intact cell of virtually any two tagged proteins expressed in a large cohort of transfected live cells.

#### **5.2 Method**

HEK-293 cells co-transfected with *pAcGFP-N1-MYH9* (the gene encoding NMMHC-IIA) and *pDsRed-monomer-N1-P2RX7* (1:4) are resuspended in Na medium with 0.1 mM CaCl2 at a concentration of 2.0x106/mL. The cell suspension (2 mL) is stirred and temperature maintained at 37ºC using a Time Zero module. ATP (1.0 mM) is added 2 min after the tube is inserted. Cells are analysed at about 1500 events/s on a FACSCalibur flow cytometer. The voltage settings and compensation are established using non-transfected cell and cells transfected with either AcGFP or DsRed. The FSC, SSC, log FL1, log FL3 and the time are collected into a FCS 2.0 file.

#### **5.3 Gating and calculation**

Fluorescent events are gated by forward and side scatter and by AcGFP and DsRed fluorescence intensity. The log mean channel of fluorescence intensity for each gated subpopulation over successive 5-s intervals is analysed by WinMDI software and saved into a text file.

To quantify levels of protein dissociation, arbitrary unit of area under/above the kinetic fluorescence curve in first 12 min after addition of ATP is calculated. The text file is then imported into *Microsoft* Excel spreadsheet, and mathematical calculation of the area under/above curve is programmed using the following functions:

Function(C4)=MATCH(0,C\$7:C\$40,0) ; *To calculate when ATP is added.*

Function (C5)=AVERAGE(C\$7:INDIRECT(ADDRESS(C\$4+5,COLUMN(),4,1),1)) *To calculate the base line.*

Function(C6)

198 Flow Cytometry – Recent Perspectives

Fluorescence resonance energy transfer (FRET) is one of the major techniques used for protein interaction studies. Confocal or fluorescent microscopes with photobleach or fluorescence lifetime modules are the common instruments which perform FRET based measurements. However, microscopy can only detect fluorescence signals in a single cell, and cannot measure multiple fluorescence emission wavelengths simultaneously. This limitation on confocal microscope makes flow cytometer an attractive alternative for FRET based measurement. The time-resolved fluorescence decay measurement by flow cytometry has been used to study molecule interactions (Condrau *et al.*, 1994a; Condrau *et al.*, 1994b; Deka & Steinkamp, 1996). However, this technique requires major modification to a flow cytometer since a pulsed laser has to be used in order to measure fluorescence lifetime.

The P2X7 receptor has been shown to form a protein complex with nonmuscle myosin heavy chain IIA (NMMHC-IIA) in its unactivated state, and this complex dissociates following activation of P2X7 by extracellular ATP (Gu *et al.*, 2009). To study the interaction of these two proteins in a large cohort of live cells, we developed a time-resolved FRET (TR-FRET) flow cytometry method. The HEK-293 cells are transfected with AcGFP-tagged NMMHC-IIA and DsRed-tagged P2X7. The NMMHC-P2X7 interaction is assessed by monitoring the fluorescence intensity changes of AcGFP and DsRed in double positive cells over 10-15 minutes after addition of P2X7 agonist. This method does not require any modification of the flow cytometer, and can be readily adapted to study the interaction within the intact cell of virtually any two tagged proteins expressed in a large cohort of transfected live cells.

HEK-293 cells co-transfected with *pAcGFP-N1-MYH9* (the gene encoding NMMHC-IIA) and *pDsRed-monomer-N1-P2RX7* (1:4) are resuspended in Na medium with 0.1 mM CaCl2 at a concentration of 2.0x106/mL. The cell suspension (2 mL) is stirred and temperature maintained at 37ºC using a Time Zero module. ATP (1.0 mM) is added 2 min after the tube is inserted. Cells are analysed at about 1500 events/s on a FACSCalibur flow cytometer. The voltage settings and compensation are established using non-transfected cell and cells transfected with either AcGFP or DsRed. The FSC, SSC, log FL1, log FL3 and the time are

Fluorescent events are gated by forward and side scatter and by AcGFP and DsRed fluorescence intensity. The log mean channel of fluorescence intensity for each gated subpopulation over successive 5-s intervals is analysed by WinMDI software and saved into

To quantify levels of protein dissociation, arbitrary unit of area under/above the kinetic fluorescence curve in first 12 min after addition of ATP is calculated. The text file is then imported into *Microsoft* Excel spreadsheet, and mathematical calculation of the area

under/above curve is programmed using the following functions: Function(C4)=MATCH(0,C\$7:C\$40,0) ; *To calculate when ATP is added.*

**5. Protein complex dissociation**

**5.1 Principle**

**5.2 Method**

a text file.

collected into a FCS 2.0 file.

**5.3 Gating and calculation**

=0.5\*5\*(INDIRECT(ADDRESS(C\$4+7,COLUMN(),4,1),1)+2\*SUM(INDIRECT(ADDRESS(C\$ 4+8,COLUMN(),4,1),1):INDIRECT(ADDRESS(C\$4+12\*12+6,COLUMN(),4,1),1))+INDIRECT (ADDRESS(C\$4+12\*12+7,COLUMN(),4,1),1))-C\$5\*12\*12\*5 *To calculate area above/under curve (Log mode)*

#### Funciton(D7)=IF(C7=0,0,POWER(10,(C7+1)/256))

*To transform the log MFI to artificial linear MFI. The rest data in Column D are transformed using the same function by "Ctrl-D" command.*

Function(D6)

=0.5\*5\*(INDIRECT(ADDRESS(D\$4+7,COLUMN(),4,1),1)+2\*SUM(INDIRECT(ADDRESS(D\$ 4+8,COLUMN(),4,1),1):INDIRECT(ADDRESS(D\$4+12\*12+6,COLUMN(),4,1),1))+INDIRECT (ADDRESS(D\$4+12\*12+7,COLUMN(),4,1),1))-D\$5\*12\*12\*5 *To calculate area above/under curve (Linear mode)*

Fig. 10. Typical dotplots and gating strategy used to analyse protein-protein interaction. **a.** The FSC and SSC of HEK-293 cells gated by gate R1. **b.** The AcGFP+DsRed+ HEK-293 cells gated by R2. **c.** Changes in fluorescence of AcGFP tagged NMMHC-IIA. **d.** Kinetic P2X7 with time of DsRed tagged fluorescence

Broad Applications of Multi-Colour Time-Resolved Flow Cytometry 201

The new generation of flow cytometers, such as Accuri C6 from BD, MACSQuant Analyzer from Miltenyi Biotec, Cyan from Beckman Coulter, Attune Acoustic Focusing Cytometer from Invitrogen, uses "plugs" to deliver sample for analysis (Edwards *et al.*, 2001a). This analyses precise defined volume and the sample vessel need not be pressurized. However, none of these cytometers can perform TR-FCM applications described in this chapter, although all have the time parameter for acquisition. Manufacturers have not realized the importance of TR-FCM, and its marketing value. In fact, with small modification of the sample platform and the acquisition software, these new generation of flow cytometers can

Condrau, M.A., Schwendener, R.A., Niederer, P. & Anliker, M. (1994a). Time-resolved flow

Condrau, M.A., Schwendener, R.A., Zimmermann, M., Muser, M.H., Graf, U., Niederer, P. &

Deka, C. & Steinkamp, J.A. (1996). Time-resolved fluorescence-decay measurement and analysis on single cells by flow cytometry. *Appl Opt*,Vol. 35,No.22, pp.4481-9 Edwards, B.S., Kuckuck, F.W., Prossnitz, E.R., Okun, A., Ransom, J.T. & Sklar, L.A. (2001a).

Edwards, B.S., Kuckuck, F.W., Prossnitz, E.R., Ransom, J.T. & Sklar, L.A. (2001b). HTPS flow

Gomperts, B.D. (1983). Involvement of guanine nucleotide-binding protein in the gating of

Gu, B.J., Rathsam, C., Stokes, L., McGeachie, A.B. & Wiley, J.S. (2009). Extracellular ATP

Gu, B.J., Sluyter, R., Skarratt, K.K., Shemon, A.N., Dao-Ung, L.P., Fuller, S.J., Barden, J.A.,

Gu, B.J., Zhang, W., Worthington, R.A., Sluyter, R., Dao-Ung, P., Petrou, S., Barden, J.A. &

characterization. *Journal of biomolecular screening*,Vol. 6,No.2, pp.83-90 Förster, T. (1948). Intermolecular energy migration and fluorescence. *Annalen der Physik*,Vol.

pore formation. *Am J Physiol Cell Physiol*,Vol. 297,No.2, pp.C430-439 Gu, B.J., Saunders, B.M., Jursik, C. & Wiley, J.S. (2010). The P2X7-nonmuscle myosin

theoretical evaluation. *Cytometry*,Vol. 16,No.3, pp.187-94

pharmacology. *Cytometry*,Vol. 43,No.3, pp.211-6

Ca2+ by receptors. *Nature*,Vol. 306,No.5938, pp.64-6

*Journal of Biological Chemistry*,Vol. 279,No.30, pp.31287-95

*Immunology*,Vol. 187,No.5, pp.2365-2375

*Cytometry*,Vol. 16,No.3, pp.195-205

cytometry for the measurement of lanthanide chelate fluorescence: I. Concept and

Anliker, M. (1994b). Time-resolved flow cytometry for the measurement of lanthanide chelate fluorescence: II. Instrument design and experimental results.

Plug flow cytometry extends analytical capabilities in cell adhesion and receptor

cytometry: a novel platform for automated high throughput drug discovery and

dissociates nonmuscle myosin from P2X7 complex: this dissociation regulates P2X7

membrane complex regulates phagocytosis of non-opsonized particles and bacteria by a pathway attenuated by extracellular ATP. *Blood*,Vol. 115,No.8, pp.1621-1631 Gu, B.J., Saunders, B.M., Petrou, S. & Wiley, J.S. (2011). P2X7 is a scavenger receptor for

apoptotic cells in the absence of its ligand extracellular ATP. *Journal of*

Clarke, A.L., Petrou, S. & Wiley, J.S. (2004). An Arg307 to Gln polymorphism within the ATP-binding site causes loss of function of the human P2X7 receptor.

Wiley, J.S. (2001). A Glu-496 to Ala polymorphism leads to loss of function of the human P2X7 receptor. *Journal of Biological Chemistry*,Vol. 276,No.14, pp.11135-42 Gu, B.J., Zhang, W.Y., Bendall, L.J., Chessell, I.P., Buell, G.N. & Wiley, J.S. (2000). Expression

of P2X7 purinoceptors on human lymphocytes and monocytes: evidence for

easily be upgraded for TR-FCM.

437,No.2, pp.55-75

**7. References**

Fig. 11. An example of spreadsheet (left) and a typical curve of protein dissociation (right)

#### **5.4 Notes for technique**

The compensation setting in this application is critical. HEK-293 transfected with either AcGFP- or DsRed-tagged protein should be used to establish correct compensation. Although data on FL2 is not collected, the voltage should still be set at a similar level to FL1 to enable compensation between FL1 and FL3.

AcGFP and DsRed is a good pair of FRET partners for flow cytometry. Both AcGFP and DsRed are monomers, thus possible self-polymerization can be avoided. A regular 488 laser is optimized to excite AcGFP fluorescence signal but has gives little excitation of DsRed. The emission wavelength of AcGFP is in the optimum range for excitation of DsRed. Therefore, the DsRed emission signal is mainly due to the FRET effect. With careful compensation settings, the AcGFP and DsRed emissions can be readily studied in a real time flow cytometry system using only a 488nm laser. This method allows a quick and reliable assessment of protein dissociation. The dissociation rate can be compared between samples in the same experiment. However, this method cannot provide an estimated value for FRET efficiency derived according to Förster formulae (Förster, 1948).

#### **6. Discussion**

Although flow cytometry is a mature technique and time-resolved flow cytometry (TR-FCM) has been widely used in cellular kinetic study, the great potential of TR-FCM has not been fully realized by most researchers. In part this is due to the requirement of modification of existing instruments as well as data analysis. The four applications described in this chapter are simple and feasible, requiring only minimum modification of the flow cytometer. The Time-Zero module can be easily set up in less than 5 min. If this module is not available, a small beaker with 37ºC water can substitute for temperature control. The analysis software is free and the example of MS Excel templates are available. We hope above four examples may encourage the use of TR-FCM in more cellular kinetic studies.

The new generation of flow cytometers, such as Accuri C6 from BD, MACSQuant Analyzer from Miltenyi Biotec, Cyan from Beckman Coulter, Attune Acoustic Focusing Cytometer from Invitrogen, uses "plugs" to deliver sample for analysis (Edwards *et al.*, 2001a). This analyses precise defined volume and the sample vessel need not be pressurized. However, none of these cytometers can perform TR-FCM applications described in this chapter, although all have the time parameter for acquisition. Manufacturers have not realized the importance of TR-FCM, and its marketing value. In fact, with small modification of the sample platform and the acquisition software, these new generation of flow cytometers can easily be upgraded for TR-FCM.

#### **7. References**

200 Flow Cytometry – Recent Perspectives

**ATP**

Linear MFI

**5.4 Notes for technique**

**6. Discussion**

to enable compensation between FL1 and FL3.

efficiency derived according to Förster formulae (Förster, 1948).

encourage the use of TR-FCM in more cellular kinetic studies.

0

Fig. 11. An example of spreadsheet (left) and a typical curve of protein dissociation (right)

The compensation setting in this application is critical. HEK-293 transfected with either AcGFP- or DsRed-tagged protein should be used to establish correct compensation. Although data on FL2 is not collected, the voltage should still be set at a similar level to FL1

AcGFP and DsRed is a good pair of FRET partners for flow cytometry. Both AcGFP and DsRed are monomers, thus possible self-polymerization can be avoided. A regular 488 laser is optimized to excite AcGFP fluorescence signal but has gives little excitation of DsRed. The emission wavelength of AcGFP is in the optimum range for excitation of DsRed. Therefore, the DsRed emission signal is mainly due to the FRET effect. With careful compensation settings, the AcGFP and DsRed emissions can be readily studied in a real time flow cytometry system using only a 488nm laser. This method allows a quick and reliable assessment of protein dissociation. The dissociation rate can be compared between samples in the same experiment. However, this method cannot provide an estimated value for FRET

Although flow cytometry is a mature technique and time-resolved flow cytometry (TR-FCM) has been widely used in cellular kinetic study, the great potential of TR-FCM has not been fully realized by most researchers. In part this is due to the requirement of modification of existing instruments as well as data analysis. The four applications described in this chapter are simple and feasible, requiring only minimum modification of the flow cytometer. The Time-Zero module can be easily set up in less than 5 min. If this module is not available, a small beaker with 37ºC water can substitute for temperature control. The analysis software is free and the example of MS Excel templates are available. We hope above four examples may

50

100

150

200

250

300

Time (min) 0 2 4 6 8 10 12 14

NMMHC-IIA-AcGFP P2X7-DsRed

**11**

*Poland*

**Application of Flow Cytometry**

**in the Studies of Microparticles**

Monika Baj-Krzyworzeka, Jarek Baran, Rafał Szatanek and Maciej Siedlar *Department of Clinical Immunology, Jagiellonian University Medical College, Cracow,*

Many cell types including leukocytes, platelets and endothelial cells release small membrane fragments called microparticles (MP). MP are shed during cell growth, activation, proliferation, senescence and apoptosis. MP contain proteins (intracellular as well as surface markers), mRNA and miRNA of the cells they have originated from. Based on the release mechanism, size and phenotype, MP are frequently divided into two categories: exosomes and ectosomes called also microvesicles. There is no doubt that the biological significance of MP has been largely overlooked for many years, regarding them as merely cellular fragments or debris. Nowadays, MP are being recognized as an important regulator of cellular interactions under physiological and pathological conditions. MP are present in all body fluids and physiologically serve various functions like blood clotting, enhance cell adhesiveness, increase cell aggregation, etc. They mediate cell-to-cell communication by transferring cell surface receptors, mRNA, and miRNA from the cell of origin to target cells. The growing body of literature regarding the role of MP in many pathologies has recently progressed from describing the association of elevated MP number with disease stage (e.g. cancer, sepsis) through understanding how MP may contribute to thrombosis, preeclampsia and tumor progression, and finally, to using MP as a source of antigens in new

Flow cytometry is a preferred method in the studies of MP because of its ability to quantitate the absolute number of particles and multicolor analysis attributes, allowing detection of several markers simultaneously. However, despite its usefulness, flow cytometry has some limitations in this field. The definition of MP using flow cytometry is still an area of great debate. In this review we propose a comprehensive summary of the possibilities, advantages and disadvantages of flow cytometry as a "gold standard" in the studies of MP.

MP are defined as a mixture of heterogeneous vesicles size-wise, and there is a number of schemes trying to classify them by considering their different characteristics (i.e. origin, size, distinct cell surface and/or internal determinant patterns, etc.), which may become confusing

forms of vaccines against infectious or malignant diseases.

**2. Overview of different forms of microparticles**

**2.1 Definition of various MP – plenty is a plaque**

**1. Introduction**

at times.

nonfunctional P2X7 receptors. *American Journal of Physiology - Cell Physiology*,Vol. 279,No.4, pp.C1189-97

### **Application of Flow Cytometry in the Studies of Microparticles**

Monika Baj-Krzyworzeka, Jarek Baran, Rafał Szatanek and Maciej Siedlar *Department of Clinical Immunology, Jagiellonian University Medical College, Cracow, Poland*

#### **1. Introduction**

202 Flow Cytometry – Recent Perspectives

Humphreys, B.D. & Dubyak, G.R. (1996). Induction of the P2Z/P2X7 nucleotide receptor and

Michel, A.D., Chessell, I.P. & Humphrey, P.P. (1999). Ionic effects on human recombinant

Nolan, J.P. & Sklar, L.A. (1998). The emergence of flow cytometry for sensitive, real-time

Ramirez, S., Aiken, C.T., Andrzejewski, B., Sklar, L.A. & Edwards, B.S. (2003). High-

killing by macrophages. *Journal Of Biological Chemistry*,Vol. 281,No.4, pp.2079 Skarratt, K.K., Fuller, S.J., Sluyter, R., Dao-Ung, L.P., Gu, B.J. & Wiley, J.S. (2005). A 5 '

Sklar, L.A., Carter, M.B. & Edwards, B.S. (2007). Flow cytometry for drug discovery,

Steinkamp, J.A., Wilson, J.S., Saunders, G.C. & Stewart, C.C. (1982). Phagocytosis: flow

Touret, N., Paroutis, P., Terebiznik, M., Harrison, R.E., Trombetta, S., Pypaert, M., Chow, A.,

Wiley, J.S., Chen, J.R., Snook, M.S., Gargett, C.E. & Jamieson, G.P. (1996). Transduction

Wiley, J.S., Chen, R., Wiley, M.J. & Jamieson, G.P. (1992). The ATP4- receptor-operated ion

Wiley, J.S., Gargett, C.E., Zhang, W., Snook, M.B. & Jamieson, G.A. (1998). Partial agonists

the Caucasian population. *Febs Letters*,Vol. 579,No.12, pp.2675

279,No.4, pp.C1189-97

359,No.2, pp.102-9

7,No.5, pp.527-34

123,No.1, pp.157-70

discussion 160-5

pp.C1224-C1231

*Chemistry*,Vol. 278,No.19, pp.17108-17113

nonfunctional P2X7 receptors. *American Journal of Physiology - Cell Physiology*,Vol.

associated phospholipase D activity by lipopolysaccharide and IFN-gamma in the human THP-1 monocytic cell line. *Journal of Immunology*,Vol. 157,No.12, pp.5627-37 Jursik, C., Sluyter, R., Georgiou, J.G., Fuller, S.J., Wiley, J.S. & Gu, B.J. (2007). A quantitative

method for routine measurement of cell surface P2X(7) receptor function in leucocyte subsets by two-colour time-resolved flow cytometry. *J Immunol Methods*,Vol. 32567-77

P2X7 receptor function. *Naunyn-Schmiedebergs Archives of Pharmacology*,Vol.

measurements of molecular interactions. *Nature Biotechnology*,Vol. 16,No.7, pp.633-8

throughput flow cytometry: validation in microvolume bioassays. *Cytometry Part A: the journal of the International Society for Analytical Cytology*,Vol. 53,No.1, pp.55-65 Shemon, A.N., Sluyter, R., Fernando, S.L., Clarke, A.L., Dao-Ung, L.P., Skarratt, K.K., Saunders,

B.M., Tan, K.S., Gu, B.J., Fuller, S.J., Britton, W.J., Petrou, S. & Wiley, J.S. (2006). A Thr(357) to Ser polymorphism in homozygous and compound heterozygous subjects causes absent or reduced P2X(7) function and impairs ATP-induced mycobacterial

intronic splice site polymorphism leads to a null allele of the P2X(7) gene in 1-2% of

receptor pharmacology and high-throughput screening. *Curr Opin Pharmacol*,Vol.

cytometric quantitation with fluorescent microspheres. *Science*,Vol. 215,No.4528, pp.64-6

Jiang, A., Shaw, J., Yip, C., Moore, H.P., van der Wel, N., Houben, D., Peters, P.J., de Chastellier, C., Mellman, I. & Grinstein, S. (2005). Quantitative and dynamic assessment of the contribution of the ER to phagosome formation. *Cell*,Vol.

mechanisms of P2Z purinoceptors. *Ciba Foundation Symposium*,Vol. 198149-60;

channel of human lymphocytes: inhibition of ion fluxes by amiloride analogs and by extracellular sodium ions. *Archives of Biochemistry & Biophysics*,Vol. 292,No.2, pp.411-8 Wiley, J.S., Dao-Ung, L.-P., Li, C., Shemon, A.N., Gu, B.J., Smart, M.L., Fuller, S.J., Barden,

J.A., Petrou, S. & Sluyter, R. (2003). An Ile-568 to Asn Polymorphism Prevents Normal Trafficking and Function of the Human P2X7 Receptor. *Journal of Biological*

and antagonists reveal a second permeability state of human lymphocyte P2Z/P2X7 channel. *American Journal of Physiology - Cell Physiology*,Vol. 44,No.5, Many cell types including leukocytes, platelets and endothelial cells release small membrane fragments called microparticles (MP). MP are shed during cell growth, activation, proliferation, senescence and apoptosis. MP contain proteins (intracellular as well as surface markers), mRNA and miRNA of the cells they have originated from. Based on the release mechanism, size and phenotype, MP are frequently divided into two categories: exosomes and ectosomes called also microvesicles. There is no doubt that the biological significance of MP has been largely overlooked for many years, regarding them as merely cellular fragments or debris. Nowadays, MP are being recognized as an important regulator of cellular interactions under physiological and pathological conditions. MP are present in all body fluids and physiologically serve various functions like blood clotting, enhance cell adhesiveness, increase cell aggregation, etc. They mediate cell-to-cell communication by transferring cell surface receptors, mRNA, and miRNA from the cell of origin to target cells. The growing body of literature regarding the role of MP in many pathologies has recently progressed from describing the association of elevated MP number with disease stage (e.g. cancer, sepsis) through understanding how MP may contribute to thrombosis, preeclampsia and tumor progression, and finally, to using MP as a source of antigens in new forms of vaccines against infectious or malignant diseases.

Flow cytometry is a preferred method in the studies of MP because of its ability to quantitate the absolute number of particles and multicolor analysis attributes, allowing detection of several markers simultaneously. However, despite its usefulness, flow cytometry has some limitations in this field. The definition of MP using flow cytometry is still an area of great debate. In this review we propose a comprehensive summary of the possibilities, advantages and disadvantages of flow cytometry as a "gold standard" in the studies of MP.

#### **2. Overview of different forms of microparticles**

#### **2.1 Definition of various MP – plenty is a plaque**

MP are defined as a mixture of heterogeneous vesicles size-wise, and there is a number of schemes trying to classify them by considering their different characteristics (i.e. origin, size, distinct cell surface and/or internal determinant patterns, etc.), which may become confusing at times.

Application of Flow Cytometry in the Studies of Microparticles 205

sample preparation such as collecting, processing temperature, etc. as well as how the

The most commonly used methodology for MP isolation is differential centrifugation which employs a number of different centrifugation steps characterized by different centrifugal forces and centrifugation times. The concept behind this type of methodology is to purify the sample in such a manner as to obtain the correct MP population. Although, differences between the protocols exist, there seems to be a general agreement on the purification of the MP sample from cells and other larger cellular fragments that remain after a sample collection. This is regardless of the sample origin, whether it is culture supernatants, plasma or any other biological fluid. Thus, the initial purification step is set at a relatively low centrifugal force (around 200-500xg) for a short time period (app. 10 min.) to get rid of the larger fragments/cells from the sample [Orozco et al., 2010, Baran et al., 2010]. The next step, which centrifugal force ranges between 10,000-17,000xg and is set between 30 min. to 1 hour, is designed either to obtain the MP or to further purify the sample of unwanted cellular fragments or MP [Ayers et al., 2011, Baran et al., 2010, Gelderman & Simak, 2008]. One has to consider its interest because at this speed some of the MP can be lost which might be of significance. Thus, if platelet-derived MP (PMP), whose presence is predominant in plasma samples, is a subject of the study then the pellet obtained after this centrifugation step will mainly consist of them. At this step, one has to also consider the impact of size with regards to MP because if exosomes (part of MP population), which are considered to be smaller vesicles, are the subject of the study then they will remain in the supernatant, and the next centrifugation step/steps is/are considered to be crucial to obtain them (centrifugal force up

to 150,000xg) [Grant et al., 2011, Baran et al., 2010, Al-Nedawi et al., 2008].

with respect to the surface determinant profile.

The adopted form of verifying the individual steps of the different centrifugation protocols employs flow cytometry. Staining the pellet or supernatant samples obtained during the different phases of the protocol with appropriate antibodies and then analyzing them using flow cytometry seems to be the method of choice by many groups. The idea behind it is to trace the MP fractions, whether present in the pellets or supernatants that come off during each step of the centrifugation protocol. Thus, for example, if platelet-free sample is required, one would stain the pellet and supernatant obtained after the initial and the 10,000-17,000xg centrifugation steps to check for platelet markers (i.e. CD41, CD61) [Baran et al., 2010]. The depletion of CD41+ or CD61+ entities would then signal that sample purification was successful and that further sample processing can be initiated (Fig. 2 represents an example of a marker tracing (CD61) on MP in the supernatant fractions of a plasma sample obtained during different centrifugation steps). Analogically, flow cytometry is used when the final MP population is obtained and needs verification or characterization

Another form of centrifugation, which is also commonly used in MP isolation, involves the application of a sucrose gradient [Lamparski et al., 2002, Keller et al., 2011, Zhong et al., 2011]. In the case of the sucrose gradient filtration method, a sucrose gradient is created by gently overlaying a sucrose layer of lesser concentration on top of the layer with higher sucrose concentration in some defined concentration increments in an appropriate test tube. Next, the sample is placed on top of the sucrose density gradient and subjected to a high centrifugal force (app. 150,000xg) for an extended time period. During the centrifugation step, the sample particles move through the gradient until they reach the sucrose density

samples should be stored and prepared for future use.

One of the most routinely used schemes in defining MP is their cellular origin [Heijnen et al., 1999, Hess et al., 1999, Dumaswala et al., 1984, Ginestra et al., 1998, Zitvogel et al., 1998]. This method utilizes flow cytometry to compare the cell membrane determinant composition as well as the internal cargo of MP with that of the original cell. Scientists that chose this method do not restrict themselves to just one surface/cellular determinant, but use many, in order to define the MP most precisely. It is very desirable in this case to have a unique determinant (present on/in the original cell and its MP) that would definitely establish the MP origin. Based on this classification method, scientists then use terminology that stresses out the MP origin, i.e. dendritic cell-derived microvesicles, erythrocyte-derived microvesicles, platelet-derived microvesicles, etc. [Heijnen et al., 1999, Hess et al., 1999, Dumaswala et al., 1984, Ginestra et al., 1998, Zitvogel et al., 1998].

Another way of defining MP considers two populations of MP, ectosomes and exosomes, depending on the place of origin within the same cell [Pilzer et al., 2005, Rak et al., 2010]. Thus, ectosomes are considered to be vesicles which are formed upon plasma membrane vesiculation, where as exosomes are generated within endosomal structures inside a cell [Al-Nedawi et al., 2009]. Ectosomes are relatively larger than exosomes with their size ranging from 100-1000 nm in diameter, and their outer membrane is rich in phosphatidylserine (PS) residues [Al-Nedawi et al., 2009, Del Conde et al., 2005]. Ectosomes are also associated with the formation of lipid rafts, membrane regions containing high levels of cholesterol and signaling complexes. Exosomes, on the other hand, have a lower abundance of PS residues compared to ectosomes and are usually smaller (30-100 nm) in diameter [Simpson et al., 2009]. They also seem to transport a different type of cargo than other MV originating from the same cell, showing more transporting selectivity for intracellular proteins/molecules.

Another group of MP that is often regarded as a separate population is derived from tumor cells. The most important criterion in defining TMV seems to be the assessment of their tumor origin [Yu et al., 2005, Bergmann et al., 2009, Kim et al., 2003, Huber et al., 2005]. Here, again, flow cytometry is used as the means of establishing the determinant composition (surface and/or internal) of TMV which is then compared to that of the tumor cells.

Considering the different ways of MP classification it has to be kept in mind that there is no clear, well-defined approach that could be applied for MP differentiation. Many of the methods mentioned above tend to overlap, which when used separately, could result in defining the same MP population. There is a growing need of trying to establish clear-cut guidelines that would help properly define the different types of MP. Figure 1 is a representation of the heterogeneity of MP and their different nomenclature. It also depicts some possible determinants that can be transported by particular MP population.

#### **2.2 Isolation of MP from biological fluids or culture supernatants – The devil is in the details**

There is no real consensus or a uniform protocol on MP isolation from different types of biological fluids or culture supernatants. Most of the time, people who try to isolate MP develop their own protocols which tend to incorporate their particular interests as well as the availability of equipment in their laboratory setting. The other side to this problem is

One of the most routinely used schemes in defining MP is their cellular origin [Heijnen et al., 1999, Hess et al., 1999, Dumaswala et al., 1984, Ginestra et al., 1998, Zitvogel et al., 1998]. This method utilizes flow cytometry to compare the cell membrane determinant composition as well as the internal cargo of MP with that of the original cell. Scientists that chose this method do not restrict themselves to just one surface/cellular determinant, but use many, in order to define the MP most precisely. It is very desirable in this case to have a unique determinant (present on/in the original cell and its MP) that would definitely establish the MP origin. Based on this classification method, scientists then use terminology that stresses out the MP origin, i.e. dendritic cell-derived microvesicles, erythrocyte-derived microvesicles, platelet-derived microvesicles, etc. [Heijnen et al., 1999, Hess et al., 1999,

Another way of defining MP considers two populations of MP, ectosomes and exosomes, depending on the place of origin within the same cell [Pilzer et al., 2005, Rak et al., 2010]. Thus, ectosomes are considered to be vesicles which are formed upon plasma membrane vesiculation, where as exosomes are generated within endosomal structures inside a cell [Al-Nedawi et al., 2009]. Ectosomes are relatively larger than exosomes with their size ranging from 100-1000 nm in diameter, and their outer membrane is rich in phosphatidylserine (PS) residues [Al-Nedawi et al., 2009, Del Conde et al., 2005]. Ectosomes are also associated with the formation of lipid rafts, membrane regions containing high levels of cholesterol and signaling complexes. Exosomes, on the other hand, have a lower abundance of PS residues compared to ectosomes and are usually smaller (30-100 nm) in diameter [Simpson et al., 2009]. They also seem to transport a different type of cargo than other MV originating from the same cell, showing more transporting selectivity for

Another group of MP that is often regarded as a separate population is derived from tumor cells. The most important criterion in defining TMV seems to be the assessment of their tumor origin [Yu et al., 2005, Bergmann et al., 2009, Kim et al., 2003, Huber et al., 2005]. Here, again, flow cytometry is used as the means of establishing the determinant composition (surface and/or internal) of TMV which is then compared to that of the tumor

Considering the different ways of MP classification it has to be kept in mind that there is no clear, well-defined approach that could be applied for MP differentiation. Many of the methods mentioned above tend to overlap, which when used separately, could result in defining the same MP population. There is a growing need of trying to establish clear-cut guidelines that would help properly define the different types of MP. Figure 1 is a representation of the heterogeneity of MP and their different nomenclature. It also depicts

**2.2 Isolation of MP from biological fluids or culture supernatants – The devil is in the**

There is no real consensus or a uniform protocol on MP isolation from different types of biological fluids or culture supernatants. Most of the time, people who try to isolate MP develop their own protocols which tend to incorporate their particular interests as well as the availability of equipment in their laboratory setting. The other side to this problem is

some possible determinants that can be transported by particular MP population.

Dumaswala et al., 1984, Ginestra et al., 1998, Zitvogel et al., 1998].

intracellular proteins/molecules.

cells.

**details**

sample preparation such as collecting, processing temperature, etc. as well as how the samples should be stored and prepared for future use.

The most commonly used methodology for MP isolation is differential centrifugation which employs a number of different centrifugation steps characterized by different centrifugal forces and centrifugation times. The concept behind this type of methodology is to purify the sample in such a manner as to obtain the correct MP population. Although, differences between the protocols exist, there seems to be a general agreement on the purification of the MP sample from cells and other larger cellular fragments that remain after a sample collection. This is regardless of the sample origin, whether it is culture supernatants, plasma or any other biological fluid. Thus, the initial purification step is set at a relatively low centrifugal force (around 200-500xg) for a short time period (app. 10 min.) to get rid of the larger fragments/cells from the sample [Orozco et al., 2010, Baran et al., 2010]. The next step, which centrifugal force ranges between 10,000-17,000xg and is set between 30 min. to 1 hour, is designed either to obtain the MP or to further purify the sample of unwanted cellular fragments or MP [Ayers et al., 2011, Baran et al., 2010, Gelderman & Simak, 2008]. One has to consider its interest because at this speed some of the MP can be lost which might be of significance. Thus, if platelet-derived MP (PMP), whose presence is predominant in plasma samples, is a subject of the study then the pellet obtained after this centrifugation step will mainly consist of them. At this step, one has to also consider the impact of size with regards to MP because if exosomes (part of MP population), which are considered to be smaller vesicles, are the subject of the study then they will remain in the supernatant, and the next centrifugation step/steps is/are considered to be crucial to obtain them (centrifugal force up to 150,000xg) [Grant et al., 2011, Baran et al., 2010, Al-Nedawi et al., 2008].

The adopted form of verifying the individual steps of the different centrifugation protocols employs flow cytometry. Staining the pellet or supernatant samples obtained during the different phases of the protocol with appropriate antibodies and then analyzing them using flow cytometry seems to be the method of choice by many groups. The idea behind it is to trace the MP fractions, whether present in the pellets or supernatants that come off during each step of the centrifugation protocol. Thus, for example, if platelet-free sample is required, one would stain the pellet and supernatant obtained after the initial and the 10,000-17,000xg centrifugation steps to check for platelet markers (i.e. CD41, CD61) [Baran et al., 2010]. The depletion of CD41+ or CD61+ entities would then signal that sample purification was successful and that further sample processing can be initiated (Fig. 2 represents an example of a marker tracing (CD61) on MP in the supernatant fractions of a plasma sample obtained during different centrifugation steps). Analogically, flow cytometry is used when the final MP population is obtained and needs verification or characterization with respect to the surface determinant profile.

Another form of centrifugation, which is also commonly used in MP isolation, involves the application of a sucrose gradient [Lamparski et al., 2002, Keller et al., 2011, Zhong et al., 2011]. In the case of the sucrose gradient filtration method, a sucrose gradient is created by gently overlaying a sucrose layer of lesser concentration on top of the layer with higher sucrose concentration in some defined concentration increments in an appropriate test tube. Next, the sample is placed on top of the sucrose density gradient and subjected to a high centrifugal force (app. 150,000xg) for an extended time period. During the centrifugation step, the sample particles move through the gradient until they reach the sucrose density

Application of Flow Cytometry in the Studies of Microparticles 207

erythrocytes

PS erythrocyte-derived MP

platelets

platelet-derived MP

dendritic cells

tumor cells

dendritic cell-derived MP

**MICROPARTICLES (MICROVESICLES)**

tumor cell-derived MP (oncosomes)

CD37 CD81 CD82

TF Flotillin-1 PSGL1 CD61 CD41

glycoprotein A hemoglobin ESA

ECTOSOMES

**- site of generation: plasma membrane - generation type: constitutive - generation mechanism: budding**

> β1 integrin IL-1β IL-8 Caspases ? MMP2 MMP9 EMMPRIN MUC1 CB1

Fig. 1. Heterogeneity of microparticles

**- size: 100-1000nm - high PS at the surface**

 **and regulated**

EXOSOMES **- size: 30-100nm - low PS at the surface - site of generation: MVB generation type: regulated - generation mechanism:**

 **secretion?**

CD9 CD63 CD81 HSP70 HSP90 Caveolin-1 Rab-5b Rab27A Alix

that matches their own, where they stop. At the end of this step, the obtained fractions that contain MP according to their different densities, can be removed and utilized in further tests.

Another method for isolating MP which is gaining interest is referred to as microfluidic immunoaffinity method [Chen et al., 2010, Hsu et al., 2008]. The propagators of this procedure point out that this method is much faster in comparison to differential centrifugation or sucrose gradient filtration and that it yields higher MP recovery rates [Chen et al., 2010]. Another major advantage to this method seems to point out to the fact that smaller sample volumes are being used as well as the isolated MP remain relatively "untouched" (the impact of centrifugal force being excluded) thus resembling more accurately their native state i.e. shape, determinant surface profile, etc. The principle behind this method is to selectively bind MP populations to antibody-coated surfaces [Chen et al., 2010, Hsu et al., 2008, Choi et al., 2011, Cheng et al., 2007] . The method utilizes a flow channel of different dimensions, depending on the initial sample volume to be processed, which surface is chemically modified in order to later coat it with appropriate antibodies. The outcome of such modification results in the preparation of a column that is coated with an antibody that recognizes a surface determinant characteristic for particular MP population. Next, a sample is injected into the channel and the appropriate MP are immobilized on its surface. This is followed by a washing step which purpose is to elute the immobilized MP which then can be used for further tests (protein analysis, RNA isolation, etc.).

 Another aspect of MP isolation that has a major impact on MP quality is sample collection, preparation and storage [Dey-Hazra et al., 2010, Trummer et al., 2009, Shah et al., 2008]. Unfortunately, here again no uniform protocols exist that would state the proper way to address these issues. As with different isolation protocols, the sample collection, preparation and/or storage depend on individual needs and settings characteristic of the study. However, there seems to be a general understanding that freshly obtained samples are the best to work with and that multiple "freezing and thawing" has a substantial impact on both the MP number as well as the surface determinant expression, which suggest that, if possible, should be avoided (see section 2.1.1) [Dey-Hazra et al., 2010, Trummer et al., 2009, Shah et al., 2008].

#### **2.3 Sizing of MP and estimation of their absolute number by flow cytometry – Role of instrumentation**

Flow cytometric analysis of MP appears to be the most favored method used for their characterization [Jy et al. 2004]. Typically, MP are identified as particles with a forward scatter smaller than an internal standard consisting of 1-1.5 µm sized latex particles [Shet et al., 2003]. Light scattering is a basic phenomenon for detecting and characterizing particles in modern flow cytometry. A light beam directed at a particle can interact with it through reflective, refractive and diffractive effects. Then, information about a particle or aggregates of particles can be derived from the changes in direction and intensity of the scattered light [Kim & Ligler 2010]. Collecting scattered light at various angles from the incident beam has been reported to provide different types of information about the particle, including both size and density [Shvalov et al., 1999]. Typically, forward scattered light (0.5-5º) can provide approximate information about the size of particle [Shapiro 2004]. It should, however, be

that matches their own, where they stop. At the end of this step, the obtained fractions that contain MP according to their different densities, can be removed and utilized in further

Another method for isolating MP which is gaining interest is referred to as microfluidic immunoaffinity method [Chen et al., 2010, Hsu et al., 2008]. The propagators of this procedure point out that this method is much faster in comparison to differential centrifugation or sucrose gradient filtration and that it yields higher MP recovery rates [Chen et al., 2010]. Another major advantage to this method seems to point out to the fact that smaller sample volumes are being used as well as the isolated MP remain relatively "untouched" (the impact of centrifugal force being excluded) thus resembling more accurately their native state i.e. shape, determinant surface profile, etc. The principle behind this method is to selectively bind MP populations to antibody-coated surfaces [Chen et al., 2010, Hsu et al., 2008, Choi et al., 2011, Cheng et al., 2007] . The method utilizes a flow channel of different dimensions, depending on the initial sample volume to be processed, which surface is chemically modified in order to later coat it with appropriate antibodies. The outcome of such modification results in the preparation of a column that is coated with an antibody that recognizes a surface determinant characteristic for particular MP population. Next, a sample is injected into the channel and the appropriate MP are immobilized on its surface. This is followed by a washing step which purpose is to elute the immobilized MP which then can be used for further tests (protein analysis, RNA isolation,

 Another aspect of MP isolation that has a major impact on MP quality is sample collection, preparation and storage [Dey-Hazra et al., 2010, Trummer et al., 2009, Shah et al., 2008]. Unfortunately, here again no uniform protocols exist that would state the proper way to address these issues. As with different isolation protocols, the sample collection, preparation and/or storage depend on individual needs and settings characteristic of the study. However, there seems to be a general understanding that freshly obtained samples are the best to work with and that multiple "freezing and thawing" has a substantial impact on both the MP number as well as the surface determinant expression, which suggest that, if possible, should be avoided (see section 2.1.1) [Dey-Hazra et al., 2010, Trummer et al., 2009,

**2.3 Sizing of MP and estimation of their absolute number by flow cytometry – Role of**

Flow cytometric analysis of MP appears to be the most favored method used for their characterization [Jy et al. 2004]. Typically, MP are identified as particles with a forward scatter smaller than an internal standard consisting of 1-1.5 µm sized latex particles [Shet et al., 2003]. Light scattering is a basic phenomenon for detecting and characterizing particles in modern flow cytometry. A light beam directed at a particle can interact with it through reflective, refractive and diffractive effects. Then, information about a particle or aggregates of particles can be derived from the changes in direction and intensity of the scattered light [Kim & Ligler 2010]. Collecting scattered light at various angles from the incident beam has been reported to provide different types of information about the particle, including both size and density [Shvalov et al., 1999]. Typically, forward scattered light (0.5-5º) can provide approximate information about the size of particle [Shapiro 2004]. It should, however, be

tests.

etc.).

Shah et al., 2008].

**instrumentation**

Fig. 1. Heterogeneity of microparticles

Application of Flow Cytometry in the Studies of Microparticles 209

(A) (B) (C)

Fig. 2. Tracking of CD61 expression to follow platelets and PMP elimination in a gastric patient plasma sample subjected to differential centrifugation. A- initial plasma; B- plasma subjected to centrifugation at 15,000xg; C- plasma subjected to centrifugation at 50,000xg

samples as well as high fluorescence sensitivity making their use even more practical. One of such product is manufactured by Apogee Flow Systems and allows detection of particles less than 150 nm using 3 light scatter detectors. However, for appropriate MP detection an even lower detection limit is required as compared to microspheres [Chandler, et al. 2011]. For smaller particles, like exosomes (< 100 nm in diameter) some other approaches can be introduced to allow their analysis by flow cytometry. Caby et al. in a very elegant study have presented detection and characterization of peripheral blood exosomes by flow cytometry analysis using CD63-coated latex beads, which can be easily detected by standard instrumentation [Caby et al., 2005]. Other approaches introduce electrical detection systems to detect beads with attached MP/exosomes. Those are based on the work of Wallace Coulter, who demonstrated that electrical charge can be used to detect, size and count particles in solution. Advances in nanotechnologies have drawn many research groups to develop Coulter counters on chip-based platforms [Kim & Ligler 2010]. Holmes et al. has demonstrated a microfabricated flow cytometer for rapid analysis of microspheres using impedance for particle detection [Holmes et al., 2007]. Such cytometers could measure impedance at high (10 MHz) and low (0.5 MHz) frequencies to distinguish mixed bead populations [Kim & Ligler, 2010]. These beads may be coated with antibodies capturing

Multicolor flow cytometry analysis of MP, using monoclonal antibodies, opened a new way of extensive investigation and characterization of MP. Multicolor analysis is used to detect the cellular origin of MP based on their phenotype. Gelderman and Simak have developed a three-color flow cytometric assay for immunophenotyping MP that are present in plasma [Gelderman & Simak, 2008]. The assay has been used to study MP present in plasma of healthy donors and in patients with paroxysmal nocturnal hemoglobinuria, sickle cell anemia, and in patients with acute ischemic stroke [Simak et al., 2002, Simak et al., 2004, Simak et al., 2002a]. With the use of monoclonal antibodies conjugated to different fluorochromes, a combination of three or even more antigens can be analyzed on a single particle. In addition, annexin V conjugated to a fluorochrome can be used to detect PS on

autofluorescence

MP/exosomes of different origin.

**2.4 Multicolor flow cytometry analysis of MP**

noted that the intensity of FSC is not related directly to particle size [Becker et al., 2002]. On the other hand, side-scattered light (15-150º) is often collected at 90º angle and provides information about smaller particles or granularity of internal structures. Measuring sidescattered and forward scattered light has become a standard in biomedical research, enabling cells to be distinguished by size and granularity.

A problem with MP in a standard flow cytometry is that they cannot reliably be detected when setting a forward scatter as a trigger, as they are smaller than the wavelength of a laser light used, and cannot always be discriminated from the background noise. Moreover, forward scatter is the most variable signal between instruments of different manufacturers and its proper alignment is crucial. It is affected by refractive index mismatches between sheath fluid and sample, beam geometry, polarization, beam stop position and collection angle [Nebe-von-Caron, 2009]. Unfortunately, there are a lot of data published in peer reviewed journals based on the assumption that a forward scatter signal of certain size beads represents similar size in MP. The problems with this approach have been already highlighted, and nowadays it is recommended to use log side scatter for comparative analysis of MP and beads, as all the instruments show good correlation with regards to side scatter response, being capable of reproducing the same level of sensitivity against the particular 190 nm latex particles [Nebe-von-Caron, 2009]. While the identification of MP on the basis of the light scattered parameters tests the limit of sensitivity of flow cytometry, some investigators have overcome this problem by setting the parameters of the instrument to detect fluorescence as a trigger (Horstman et al, 1994). Thus while analyzing MP one should look at log side scatter versus log fluorescence of the selected staining triggered on fluorescence and side scatter at the instrument noise level (if triggered on two channels) or either of the two, depending on the analysis needed [Nebe-von-Caron 2011].

#### **Enumeration of MP by flow cytometry**

Flow cytometry can also be used to enumerate MP (usually in the plasma) by adding, as an internal standard, a known number of fluorescent latex beads (Flow-Count Fluorospheres, Beckman-Coulter) or using tubes containing already predefined number of them (TruCount tubes, BD Biosciences). The number of MP present in the sample is derived from the following formula (1) and adjusted for the final dilution of the original sample [Baran et al, 2010]:

$$\text{MP count} = \frac{\text{No. of events in region containing MP}}{\text{No. of events in absolute count head region}} \times \frac{\text{No. of heads per test}}{\text{Test volume [\mu l]}} \tag{1}$$

Alternatively, if the flow cytometry instrument delivers the sample to the optics by screwdriven syringe at a known rate, then the sample MP count can be calculated [Orozco & Lewis, 2010].

#### **Role of instrumentation**

Recently, a class of flow cytometers has been developed which allows the detection of particles of down to 100 nm in size. The developers of these cytometers state that the unique optical design of the apparatus eliminates the unwanted light thus giving the best signal to noise ratio. Due to these adjustments, the cytometers are supposedly representing the highest light scatter sensitivity and resolution available making it an attractive tool in MP research. Additionally, they also offer the volumetric-based absolute count ability of

noted that the intensity of FSC is not related directly to particle size [Becker et al., 2002]. On the other hand, side-scattered light (15-150º) is often collected at 90º angle and provides information about smaller particles or granularity of internal structures. Measuring sidescattered and forward scattered light has become a standard in biomedical research,

A problem with MP in a standard flow cytometry is that they cannot reliably be detected when setting a forward scatter as a trigger, as they are smaller than the wavelength of a laser light used, and cannot always be discriminated from the background noise. Moreover, forward scatter is the most variable signal between instruments of different manufacturers and its proper alignment is crucial. It is affected by refractive index mismatches between sheath fluid and sample, beam geometry, polarization, beam stop position and collection angle [Nebe-von-Caron, 2009]. Unfortunately, there are a lot of data published in peer reviewed journals based on the assumption that a forward scatter signal of certain size beads represents similar size in MP. The problems with this approach have been already highlighted, and nowadays it is recommended to use log side scatter for comparative analysis of MP and beads, as all the instruments show good correlation with regards to side scatter response, being capable of reproducing the same level of sensitivity against the particular 190 nm latex particles [Nebe-von-Caron, 2009]. While the identification of MP on the basis of the light scattered parameters tests the limit of sensitivity of flow cytometry, some investigators have overcome this problem by setting the parameters of the instrument to detect fluorescence as a trigger (Horstman et al, 1994). Thus while analyzing MP one should look at log side scatter versus log fluorescence of the selected staining triggered on fluorescence and side scatter at the instrument noise level (if triggered on two channels) or

either of the two, depending on the analysis needed [Nebe-von-Caron 2011].

Flow cytometry can also be used to enumerate MP (usually in the plasma) by adding, as an internal standard, a known number of fluorescent latex beads (Flow-Count Fluorospheres, Beckman-Coulter) or using tubes containing already predefined number of them (TruCount tubes, BD Biosciences). The number of MP present in the sample is derived from the following formula (1) and adjusted for the final dilution of the original sample [Baran et al,

No. of events in region containing MP No. of beads per test MP count No. of events in absolute count bead region Test volume [ l] <sup>=</sup> <sup>×</sup> <sup>μ</sup>

Alternatively, if the flow cytometry instrument delivers the sample to the optics by screwdriven syringe at a known rate, then the sample MP count can be calculated [Orozco &

Recently, a class of flow cytometers has been developed which allows the detection of particles of down to 100 nm in size. The developers of these cytometers state that the unique optical design of the apparatus eliminates the unwanted light thus giving the best signal to noise ratio. Due to these adjustments, the cytometers are supposedly representing the highest light scatter sensitivity and resolution available making it an attractive tool in MP research. Additionally, they also offer the volumetric-based absolute count ability of

(1)

**Enumeration of MP by flow cytometry**

2010]:

Lewis, 2010].

**Role of instrumentation**

enabling cells to be distinguished by size and granularity.

Fig. 2. Tracking of CD61 expression to follow platelets and PMP elimination in a gastric patient plasma sample subjected to differential centrifugation. A- initial plasma; B- plasma subjected to centrifugation at 15,000xg; C- plasma subjected to centrifugation at 50,000xg

samples as well as high fluorescence sensitivity making their use even more practical. One of such product is manufactured by Apogee Flow Systems and allows detection of particles less than 150 nm using 3 light scatter detectors. However, for appropriate MP detection an even lower detection limit is required as compared to microspheres [Chandler, et al. 2011]. For smaller particles, like exosomes (< 100 nm in diameter) some other approaches can be introduced to allow their analysis by flow cytometry. Caby et al. in a very elegant study have presented detection and characterization of peripheral blood exosomes by flow cytometry analysis using CD63-coated latex beads, which can be easily detected by standard instrumentation [Caby et al., 2005]. Other approaches introduce electrical detection systems to detect beads with attached MP/exosomes. Those are based on the work of Wallace Coulter, who demonstrated that electrical charge can be used to detect, size and count particles in solution. Advances in nanotechnologies have drawn many research groups to develop Coulter counters on chip-based platforms [Kim & Ligler 2010]. Holmes et al. has demonstrated a microfabricated flow cytometer for rapid analysis of microspheres using impedance for particle detection [Holmes et al., 2007]. Such cytometers could measure impedance at high (10 MHz) and low (0.5 MHz) frequencies to distinguish mixed bead populations [Kim & Ligler, 2010]. These beads may be coated with antibodies capturing MP/exosomes of different origin.

#### **2.4 Multicolor flow cytometry analysis of MP**

Multicolor flow cytometry analysis of MP, using monoclonal antibodies, opened a new way of extensive investigation and characterization of MP. Multicolor analysis is used to detect the cellular origin of MP based on their phenotype. Gelderman and Simak have developed a three-color flow cytometric assay for immunophenotyping MP that are present in plasma [Gelderman & Simak, 2008]. The assay has been used to study MP present in plasma of healthy donors and in patients with paroxysmal nocturnal hemoglobinuria, sickle cell anemia, and in patients with acute ischemic stroke [Simak et al., 2002, Simak et al., 2004, Simak et al., 2002a]. With the use of monoclonal antibodies conjugated to different fluorochromes, a combination of three or even more antigens can be analyzed on a single particle. In addition, annexin V conjugated to a fluorochrome can be used to detect PS on

Application of Flow Cytometry in the Studies of Microparticles 211

Preanalytical stages like blood sampling site (cubital vein, central venous caterer), needle diameter, discharge of the first portion of blood, collection (Vacutainer, tube, etc), anticoagulant and transport circumstances are extremely important for PMP quality and quantity. Blood should be anticoagulated with 3.8% sodium citrate (9:1 v/v, BD Vacutainer blood collection tubes are recommended) as versenate salts lead to platelet aggregation. Changes in temperature, like overheating or cooling should be avoided. Also, shaking of blood may induce PMP release from platelets as well as other blood cells. The delay between sampling and further processing should be as short as possible (less than 1 hour), as storage results in increased number of PMP [Simak & Gelderman, 2006,

The protocols for obtaining PMP differ between laboratories and their standardization is a subject of debate. The first step in standardization was undertaken by the Scientific Standardization Committee of the International Society on Thrombosis and Haemostasis. The Committee recommends double centrifugation step to ensure removal of platelets from platelet poor plasma (PPP) and washing PPP samples to remove non specific particles, however these procedures may results in the loss of MP [Ayers et al., 2011]. Differential centrifugation-protocol results in isolation of PMP that in many cases contain exosomes. To avoid the exosomes contamination Grant et al. proposed filtration of PPP through a 0.1 μm pore size filter [Grant et al., 2011]. MP may be also isolated from PPP by capturing them using immobilized annexin V which binds to PS present on PMP and other MP. In this assay, MP are captured with biotinylated annexin V, then incubated with streptavidincoated plates, and after being washed used for further experiments [Mallat et al.,1999].

Flow cytometry allows the analysis of large numbers of PMP using tiny volume (about 5 μl) of plasma [Simak & Gelderman, 2006, Michelson et al., 2000]. It is of importance to use double filtered (0.2 μm) sheath fluid (e.g. phosphate buffered saline - PBS) for flow cytometry analysis of PMP. Generally, an accepted background "noise" consists of 25-50

**Size.** Standard beads of different sizes (diameter) may, to some extend, be used for PMP sizing, e.g. monodisperse fluorescent Megamix beads (BioCytex). However, one has to be aware that size related data derived from beads and that derived from biological particles

It is generally accepted that PMP are larger than 100 nm, but at the same time it should be noted that PMP are very heterogeneous size-wise. The upper limit of PMP size is about 1.5 μm. Additionally, distinguishing between PMP aggregates and platelet-PMP aggregates by flow cytometry may cause problems [Simak & Gelderman, 2006], because of their size overlap. Better results in platelet/PMP sizing can be achieved by using impedance-based flow cytometry instead of light-scatter based one. For this purpose electron microscopy or

**Number.** App. 75% of laboratories use flow cytometry to enumerate PMP in clinical samples [Lacroix et al., 2010]. However, a variety of preanalytic and analytic variables may lead to variations in PMP values in healthy individuals ranging from 100 to 4000 x 103 per μl [Robert at al., 2009]. Although, standardization in PMP count is still inadequate some steps

Kim et al., 2002].

**Flow cytometry in PMP analysis**

events per second when filtered PBS is run.

are not fully comparable [Lacroix et al., 2010].

AFM analysis are highly recommended [Yuana et al., 2011].

MP. Some authors limit the analysis of MP to only PS positive ones, however, it has been shown that only a limited population of MP in blood binds annexin V. Although immunophenotyping of MP seems to be a straightforward procedure, here are some methodological requirements in MP staining with monoclonal antibodies. First of all, monoclonal antibodies should be directly conjugated to fluorochomes (indirect staining is not recommended), and fluorochromes used should be as bright as possible (FITC – fluorescein isothiocyanate and PerCP – peridinin-chlorophyll-protein complex conjugated antibodies are not recommended). The titration of antibodies using MP prepared from their parental cells *in vitro* as well as from plasma, should be a rule. Gelderman and Simak recommend the use of two clones against different epitopes of an antigen to confirm specificity of detection [Gelderman & Simak, 2008]. In regard to identification of cellular origin, they suggest using glycophorin A (CD235a) and the leukocyte common antigen (CD45) for detection of red blood cell and leukocyte derived MP, respectively. Plateletderived MP are detected using monoclonal antibodies against GPIIb (CD41) or GPIIIa (CD61). Monoclonal antibodies to CD14, CD66b, CD4, CD8 and CD20 are used to detect MP from monocytes, granulocytes, T helper, T suppressor and B lymphocytes, respectively. For endothelial-derived MP the use of anti-PECAM (CD31), anti-CD34, anti-E selectin (CD62E) and anti-Endoglin (CD105) monoclonal antibodies is recommended [Gelderman & Simak 2008]. And finally, relevant isotype controls to detect the nonspecific staining should be used in parallel. It is of importance to keep in mind that the presence of specified antigen on MP does not clearly identify their cellular origin. In some cases MP may absorb a soluble antigen circulating in the plasma that is derived from another cell type. A good example is absorption of prostate antigen (PSA) by human monocytes [Faldon et al., 1996]. Thus MP derived from such monocytes might be positive for PSA.

#### **3. Biology of platelet derived microvesicles (PMP)**

PMP are the most abundant MP population in blood stream constituting approximately 70- 90% of circulating MP [Horstman et al., 1999]. First demonstration of "platelet dust" was done in 1967 by Wolf [Wolf, 1967]. Nowadays, it is known that PMP may be released by activated as well as resting platelets, both in circulation and *in vitro* experiments. A body of experimental and clinical data has shown the association between PMP and diseases.

#### **3.1 Characteristics of PMP**

#### **3.1.1 Methods of PMP generation and their phenotype analysis by flow cytometry**

Blood platelets activated by a variety of stimuli undergo shape change and degranulation. During this process platelets secrete thin walled vesicles called microparticles or microvesicles (PMP) and smaller exosomes. Under physiological conditions PMP are released after activation with agonists such as epinephrine, adenosine diphosphate, thrombin or collagen [Matzdorff et al., 1998], and also by exposure to complement protein C5b-9 or high shear stress (higher than physiological), [Holme et al., 1997]. *In vitro*, PMP may be generated by stimulation of platelets with physiological factors like thrombin [George, 1982], thrombin and collagen [Baj-Krzyworzeka et al., 2002], or nonphysiological agonist like calcium ionophore [Forlow et al., 2000]. PMP shedding was also described in stored and apoptotic platelets [Jy et al., 1995].

MP. Some authors limit the analysis of MP to only PS positive ones, however, it has been shown that only a limited population of MP in blood binds annexin V. Although immunophenotyping of MP seems to be a straightforward procedure, here are some methodological requirements in MP staining with monoclonal antibodies. First of all, monoclonal antibodies should be directly conjugated to fluorochomes (indirect staining is not recommended), and fluorochromes used should be as bright as possible (FITC – fluorescein isothiocyanate and PerCP – peridinin-chlorophyll-protein complex conjugated antibodies are not recommended). The titration of antibodies using MP prepared from their parental cells *in vitro* as well as from plasma, should be a rule. Gelderman and Simak recommend the use of two clones against different epitopes of an antigen to confirm specificity of detection [Gelderman & Simak, 2008]. In regard to identification of cellular origin, they suggest using glycophorin A (CD235a) and the leukocyte common antigen (CD45) for detection of red blood cell and leukocyte derived MP, respectively. Plateletderived MP are detected using monoclonal antibodies against GPIIb (CD41) or GPIIIa (CD61). Monoclonal antibodies to CD14, CD66b, CD4, CD8 and CD20 are used to detect MP from monocytes, granulocytes, T helper, T suppressor and B lymphocytes, respectively. For endothelial-derived MP the use of anti-PECAM (CD31), anti-CD34, anti-E selectin (CD62E) and anti-Endoglin (CD105) monoclonal antibodies is recommended [Gelderman & Simak 2008]. And finally, relevant isotype controls to detect the nonspecific staining should be used in parallel. It is of importance to keep in mind that the presence of specified antigen on MP does not clearly identify their cellular origin. In some cases MP may absorb a soluble antigen circulating in the plasma that is derived from another cell type. A good example is absorption of prostate antigen (PSA) by human monocytes [Faldon et al., 1996]. Thus MP

PMP are the most abundant MP population in blood stream constituting approximately 70- 90% of circulating MP [Horstman et al., 1999]. First demonstration of "platelet dust" was done in 1967 by Wolf [Wolf, 1967]. Nowadays, it is known that PMP may be released by activated as well as resting platelets, both in circulation and *in vitro* experiments. A body of

experimental and clinical data has shown the association between PMP and diseases.

**3.1.1 Methods of PMP generation and their phenotype analysis by flow cytometry**

Blood platelets activated by a variety of stimuli undergo shape change and degranulation. During this process platelets secrete thin walled vesicles called microparticles or microvesicles (PMP) and smaller exosomes. Under physiological conditions PMP are released after activation with agonists such as epinephrine, adenosine diphosphate, thrombin or collagen [Matzdorff et al., 1998], and also by exposure to complement protein C5b-9 or high shear stress (higher than physiological), [Holme et al., 1997]. *In vitro*, PMP may be generated by stimulation of platelets with physiological factors like thrombin [George, 1982], thrombin and collagen [Baj-Krzyworzeka et al., 2002], or nonphysiological agonist like calcium ionophore [Forlow et al., 2000]. PMP shedding was also described in

derived from such monocytes might be positive for PSA.

**3.1 Characteristics of PMP**

stored and apoptotic platelets [Jy et al., 1995].

**3. Biology of platelet derived microvesicles (PMP)**

Preanalytical stages like blood sampling site (cubital vein, central venous caterer), needle diameter, discharge of the first portion of blood, collection (Vacutainer, tube, etc), anticoagulant and transport circumstances are extremely important for PMP quality and quantity. Blood should be anticoagulated with 3.8% sodium citrate (9:1 v/v, BD Vacutainer blood collection tubes are recommended) as versenate salts lead to platelet aggregation. Changes in temperature, like overheating or cooling should be avoided. Also, shaking of blood may induce PMP release from platelets as well as other blood cells. The delay between sampling and further processing should be as short as possible (less than 1 hour), as storage results in increased number of PMP [Simak & Gelderman, 2006, Kim et al., 2002].

The protocols for obtaining PMP differ between laboratories and their standardization is a subject of debate. The first step in standardization was undertaken by the Scientific Standardization Committee of the International Society on Thrombosis and Haemostasis. The Committee recommends double centrifugation step to ensure removal of platelets from platelet poor plasma (PPP) and washing PPP samples to remove non specific particles, however these procedures may results in the loss of MP [Ayers et al., 2011]. Differential centrifugation-protocol results in isolation of PMP that in many cases contain exosomes. To avoid the exosomes contamination Grant et al. proposed filtration of PPP through a 0.1 μm pore size filter [Grant et al., 2011]. MP may be also isolated from PPP by capturing them using immobilized annexin V which binds to PS present on PMP and other MP. In this assay, MP are captured with biotinylated annexin V, then incubated with streptavidincoated plates, and after being washed used for further experiments [Mallat et al.,1999].

#### **Flow cytometry in PMP analysis**

Flow cytometry allows the analysis of large numbers of PMP using tiny volume (about 5 μl) of plasma [Simak & Gelderman, 2006, Michelson et al., 2000]. It is of importance to use double filtered (0.2 μm) sheath fluid (e.g. phosphate buffered saline - PBS) for flow cytometry analysis of PMP. Generally, an accepted background "noise" consists of 25-50 events per second when filtered PBS is run.

**Size.** Standard beads of different sizes (diameter) may, to some extend, be used for PMP sizing, e.g. monodisperse fluorescent Megamix beads (BioCytex). However, one has to be aware that size related data derived from beads and that derived from biological particles are not fully comparable [Lacroix et al., 2010].

It is generally accepted that PMP are larger than 100 nm, but at the same time it should be noted that PMP are very heterogeneous size-wise. The upper limit of PMP size is about 1.5 μm. Additionally, distinguishing between PMP aggregates and platelet-PMP aggregates by flow cytometry may cause problems [Simak & Gelderman, 2006], because of their size overlap. Better results in platelet/PMP sizing can be achieved by using impedance-based flow cytometry instead of light-scatter based one. For this purpose electron microscopy or AFM analysis are highly recommended [Yuana et al., 2011].

**Number.** App. 75% of laboratories use flow cytometry to enumerate PMP in clinical samples [Lacroix et al., 2010]. However, a variety of preanalytic and analytic variables may lead to variations in PMP values in healthy individuals ranging from 100 to 4000 x 103 per μl [Robert at al., 2009]. Although, standardization in PMP count is still inadequate some steps

Application of Flow Cytometry in the Studies of Microparticles 213

are of megakaryocyte origin [Flaumenthaft et al., 2009, 2010]. They recommended multicolor flow cytometric analysis (CD41, CD62P and LAMP-1) to distinguish PMP from megacaryocyte-derived MP [Flaumenthaft et al., 2009, 2010]. Flow cytometry analysis of PMP needs standardization comprising consensual panel of antibodies (clones, concentration) and beads specification. It appears that labeling low amounts of PMP results in higher variability of MFI (Mean Fluorescence Intensity) in comparison to labeling higher amounts of PMP [Orozco & Lewis, 2010]. Moreover, it should be noted that differences in *in vitro* research protocols for PMP generation result in variable PMP subpopulations which differ in size, marker expression, protein content and thrombogenic potential [Baj-

Interactions of cells with PMP may result in modulation of biological function in the

• First, PMP may provide interactions between cells without the need for direct cell-to-

• Third, PMP may act as "transfer vehicles" which may deliver/exchange protein, lipids, mRNA or even pathogens to target cells. For instance, PMP may change cell activity by lipid transfer [Barry et al., 1997] or may deliver CCL23, CXCL4,7, FGF and TGF [Dean et al., 2009], thus transferring chemoattractant capabilities to various cells [Baj-Krzyworzeka

Changes in circulating PMP level have been described in patient with many disease states.

The PMP number is decreased in preterm neonates compared with adults [Rajasekhar et al., 1994]. Patients with Castaman's defect and Scott syndrome, whose platelets are defective, also have a defect in generation of PMP [Castaman et al., 1996, Weiss et al., 1994], which is associated with a bleeding tendency. On the other hand, an increased number of PMP suggests a potential prognostic marker for arherosclerotic vascular disease [Boulanger et al., 2006, Michelsen et al., 2008], transient ischemic attacks, cardiopulmonary bypass and

For instance, PMP activate and stimulate neutrophils to secrete proteases, may induce the transformation of peripheral blood monocytes into endothelial progenitor cells [Prokopi et al., 2009], and may facilitate the interactions between leukocytes and endothelial cells [Forlow et al., 2000, Barry et al., 1998]. PMP may also interact with CD34+ cells and increase their bone marrow homing [Janowska-Wieczorek et al., 2001]. • Second, PMP can bind to target cells and fuse with their membranes, resulting in the acquisition of new surface antigens and thus new biological properties and activities of the target cells. For example, the chemokine receptor, CXCR4 which is present on PMP, may be transferred to various cells and make them susceptible to infection by X4-HIV [Rozmysłowicz et al., 2003]. PMP may also transfer GPIIb/IIIa to neutrophils (allowing its interaction with CD18) and neutrophil activation [Salanova et al., 2007] or transfer

Krzyworzeka et al., 2002, Dean et al., 2009, Sandberg et al., 1985].

CD40L which may activate B cells [Sprague et al., 2008].

**3.2 PMP in health and disease (counting and procoagulant activity)**

following ways:

cell contact.

et al., 2002, Fig. 4.]

**Hemostasis and thrombosis**

**3.1.2 Role of PMP in modulation of biological activity of human cells**

have been already undertaken to uniform this procedure. Three Scientific and Standardization Subcommittees of the International Society on Thrombosis and Haemostasis (ISTH) have initiated a project aimed at standardizing an enumeration of cellular MP by flow cytometry. The main objective was to establish the resolution and the level of background noise of the instruments, and to define reproducibility of PMP count in plasma using flow cytometers manufactured by different vendors. The study demonstrated that different systems were heterogeneous with respect to FSC resolution and background noise. In 2010 the Scientific and Standardization Committee of the International Society on Thrombosis and Haemostasis proposed procedures to be followed for obtaining good reproducibility in PMP count by flow cytometry [Lacroix et al. 2010]. The advised strategy is based on the use of size-calibrated fluorescent beads (0.5 μm and 0.9 μm) in a fixed ratio (Megamix) to gate PMP in a defined size-restricted window [Robert et al. 2009]. However, the procedure seems to be more adequate for Beckman-Coulter instruments rather than Becton-Dickinson's, as the former measure the forward-scattered light at a relatively wide angle (1-19º) compared to the latter (1-8º) [Lacroix et al. 2010].

**Phenotype.** Flow cytometry is used as a method of choice for PMP phenotyping. Similarly to platelets, almost all PMP, express glycoprotein complex IIb-IIIa (CD41/CD61) [Heijnen et al., 1999, George, 1982], CD42a [Simak and Gelderman, 2006, Matsumoto et al., 2004]. Caution is recommended when staining for glycoprotein Ib (CD42b) as only 40-50% of PMP express this marker. Usually, large PMP stain better than the small ones [Zdebska et al., 1998]. Other markers were also detected on PMP, such as: CD31, PAR1, CXCR4, CD154, PF4, as well as ligands for annexin V or lactadherin [Ayers et al., 2011, Baj-Krzyworzeka et al., 2002, Fig 3.]. It has been suggested that CD41+ and CD42+ PMP represent two different populations, thus it is important to use both of these markers for better assessment. PMP released by activated platelets express also platelet activation markers such as CD62P and activated complex GPIIb-IIIA [Michelson, 2000]. Flaumenhaft et al. showed that cultured megakariocytes shed MP positive for CD41 and concluded that a part of blood CD41+ MP

Fig. 3. Expression of selected surface proteins on PMP obtained from platelets stimulated by calcium ionophore A23187. Staining with an isotype control is shown as a red line. GPAglycophorin A (negative marker)

have been already undertaken to uniform this procedure. Three Scientific and Standardization Subcommittees of the International Society on Thrombosis and Haemostasis (ISTH) have initiated a project aimed at standardizing an enumeration of cellular MP by flow cytometry. The main objective was to establish the resolution and the level of background noise of the instruments, and to define reproducibility of PMP count in plasma using flow cytometers manufactured by different vendors. The study demonstrated that different systems were heterogeneous with respect to FSC resolution and background noise. In 2010 the Scientific and Standardization Committee of the International Society on Thrombosis and Haemostasis proposed procedures to be followed for obtaining good reproducibility in PMP count by flow cytometry [Lacroix et al. 2010]. The advised strategy is based on the use of size-calibrated fluorescent beads (0.5 μm and 0.9 μm) in a fixed ratio (Megamix) to gate PMP in a defined size-restricted window [Robert et al. 2009]. However, the procedure seems to be more adequate for Beckman-Coulter instruments rather than Becton-Dickinson's, as the former measure the forward-scattered light at a relatively wide angle (1-19º) compared to

**Phenotype.** Flow cytometry is used as a method of choice for PMP phenotyping. Similarly to platelets, almost all PMP, express glycoprotein complex IIb-IIIa (CD41/CD61) [Heijnen et al., 1999, George, 1982], CD42a [Simak and Gelderman, 2006, Matsumoto et al., 2004]. Caution is recommended when staining for glycoprotein Ib (CD42b) as only 40-50% of PMP express this marker. Usually, large PMP stain better than the small ones [Zdebska et al., 1998]. Other markers were also detected on PMP, such as: CD31, PAR1, CXCR4, CD154, PF4, as well as ligands for annexin V or lactadherin [Ayers et al., 2011, Baj-Krzyworzeka et al., 2002, Fig 3.]. It has been suggested that CD41+ and CD42+ PMP represent two different populations, thus it is important to use both of these markers for better assessment. PMP released by activated platelets express also platelet activation markers such as CD62P and activated complex GPIIb-IIIA [Michelson, 2000]. Flaumenhaft et al. showed that cultured megakariocytes shed MP positive for CD41 and concluded that a part of blood CD41+ MP

the latter (1-8º) [Lacroix et al. 2010].

SSC-H

Counts

300

100

100

0

> 101

101

FSC-H

FSC-H

102

102 103

103

glycophorin A (negative marker)

104 Counts

Counts

104 100

300

0

300

100

0

> 101

101

FSC-H

FSC-H

102

102 103

> 103

104

Fig. 3. Expression of selected surface proteins on PMP obtained from platelets stimulated by calcium ionophore A23187. Staining with an isotype control is shown as a red line. GPA-

Counts

Counts

104 100

**PAR1 CD154 PF4 GPA**

100

0

300

0

300

101

101 FSC-H

FSC-H

102

102 103

**CXCR4 CD41 CD62**

103 104

104 Counts

Counts

100

100

0

300

0

300

101

101

FSC-H

FSC-H

102

102 103

103 104

104 are of megakaryocyte origin [Flaumenthaft et al., 2009, 2010]. They recommended multicolor flow cytometric analysis (CD41, CD62P and LAMP-1) to distinguish PMP from megacaryocyte-derived MP [Flaumenthaft et al., 2009, 2010]. Flow cytometry analysis of PMP needs standardization comprising consensual panel of antibodies (clones, concentration) and beads specification. It appears that labeling low amounts of PMP results in higher variability of MFI (Mean Fluorescence Intensity) in comparison to labeling higher amounts of PMP [Orozco & Lewis, 2010]. Moreover, it should be noted that differences in *in vitro* research protocols for PMP generation result in variable PMP subpopulations which differ in size, marker expression, protein content and thrombogenic potential [Baj-Krzyworzeka et al., 2002, Dean et al., 2009, Sandberg et al., 1985].

#### **3.1.2 Role of PMP in modulation of biological activity of human cells**

Interactions of cells with PMP may result in modulation of biological function in the following ways:

• First, PMP may provide interactions between cells without the need for direct cell-tocell contact.

For instance, PMP activate and stimulate neutrophils to secrete proteases, may induce the transformation of peripheral blood monocytes into endothelial progenitor cells [Prokopi et al., 2009], and may facilitate the interactions between leukocytes and endothelial cells [Forlow et al., 2000, Barry et al., 1998]. PMP may also interact with CD34+ cells and increase their bone marrow homing [Janowska-Wieczorek et al., 2001].

#### **3.2 PMP in health and disease (counting and procoagulant activity)**

Changes in circulating PMP level have been described in patient with many disease states.

#### **Hemostasis and thrombosis**

The PMP number is decreased in preterm neonates compared with adults [Rajasekhar et al., 1994]. Patients with Castaman's defect and Scott syndrome, whose platelets are defective, also have a defect in generation of PMP [Castaman et al., 1996, Weiss et al., 1994], which is associated with a bleeding tendency. On the other hand, an increased number of PMP suggests a potential prognostic marker for arherosclerotic vascular disease [Boulanger et al., 2006, Michelsen et al., 2008], transient ischemic attacks, cardiopulmonary bypass and

Application of Flow Cytometry in the Studies of Microparticles 215

stimulated with fMLP (formyl-methionyl-leucylphenylanin), calcium ionophore, IL-8 or PMA release small vesicles very quickly (in the matter of minutes) [Hess et al., 1999, Mesri et al., 1999]. Analysis of these MP by flow cytometry showed expression of complement receptor 1 (CR1, CD35), CD66b, HLA class I, LFA-1/CD11a, Mac-1/CD11b, CD62L, CD46, CD55, CD16 and CD59 [Gasser et al., 2003]. However, PMN-MP did not express detectable amount of others PMNs markers like CD32 and CD87 [Gasser et al., 2003]. In general, PMN-MP may be distinguished from other non-PMN-MP by expression of CD66b and annexin V

MP released by monocytes can be defined by CD14, CD18 and TF expression [Aharon et al., 2008]. Moreover, these microvesicles contain caspase-1 and may deliver a cell death signal to e.g. vascular smooth muscle cells [Sarkar et al., 2009]. MP generated by the human monocytic cell line U937 were positive for the co-stimulatory molecules CD80 and CD86, whereas the expression of the adhesion molecules CD11a, CD11c (the DC marker and complement receptor 4), HLA-DR and the scavenger receptor CD36 were rather low. Also, expression of HLA class II molecules such as HLA-DR, HLA-DP and HLA-DQ was lower on MdM (monocyte-derived macrophages)-derived MP than MP of mature or immature DC [Kolowos et al., 2005]. Kolowos et al observed that MP derived from LPSstimulated MdM showed high expression of the activation marker CD71 in comparison to MP of untreated or UV-B irradiated MdM. Stimulation of another monocytic cell line (THP-1) by starvation or by endotoxin and calcium ionophore A23187 resulted in the release of MP which expressed exosomal marker Tsg 101, monocyte markers (CD18, CD14) and active tissue factor (TF) [Aharon et al., 2008]. The number of monocyte-derived MP was elevated in meningococcal sepsis and in patients with acute coronary syndromes

MP derived from B lymphocytes (defined by the expression of CD19) and DC have the capacity to present antigens to induce antigen-specific T-cell responses [Raposo et al., 1996, Zitvogel et al., 1998]. Dendritic cell-derived MP showed expression of HLA class I and II, as well as costimulatory molecules CD80 and CD86 [Wieckowski & Whiteside, 2006] and may present antigens to T cells [Montecalvo et al., 2008]. DC- derived exosomes were proposed to be a short range mechanism to spread alloantigen during T cell

Exosomes from activated T cells can mediate "activation-induced cell death" in a cellautonomous manner, defined by the nature of the initial T cell activation events and can play central roles in both central and peripheral deletion events involved in tolerance and homeostasis [Prado et al., 2010]. On the other hand thymic cell-derived MP express several proteins that are known to be involved in leukocyte rolling on endothelial surfaces, as well

Tumor derived microvesicles (TMV), also called oncosomes, are released by tumor cells during their activation (Fig. 5) by different stimuli, hypoxia, irradiation, exposure to proteins from an activated complement cascade and exposure to shear stress [Ratajczak et

as in transendothelial leukocyte migration [Turiak et al., 2011].

**5. Tumor derived microvesicles – Biology and function**

binding.

[Nieuwland et al., 2000].

allorecognition.

al., 2006].

Fig. 4. Chemotactic activity of human blood monocytes to PMP analyzed by flow cytometry. The dot-plots show: monocytes defined in PBMC population by FSC and SSC parameters – region R1 (left); spontaneous chemotaxis of monocytes to culture medium without serum (middle) and chemotaxsis of monocytes to PMP (right)

thrombocytopenias (heparin-induced, thrombotic and idiopathic thrombocytopenic purpura) [Sheriadan et al., 1986, Warkentin et al., 1994]. PMP are strongly procoagulant as they contain the anionic phospholipid PS. Procoagulant PMP are present *in vivo* in blood of patients with activated coagulation and fibrinolysis including patients suffering from Disseminated Intravascular Coagulation (DIC), however, such PMP are also found in healthy donors but in low amounts only [Holme et al., 1994]. An elevated number of PMP was described in patients with diabetes mellitus, which causes the development and progression of artherosclerosis in these patients [Nomura et al., 1995].

#### **Cancer**

Although there is no set standard for monitoring PMP levels, some studies have shown that their number is elevated in the plasma of cancer patients as compared to the normal samples suggesting that the tumor itself may be responsible for stimulating their release presumably to enhance its survival. For example, elevated numbers of PMP were detected in plasma of patients with gastric cancer or in urine of patients with bladder cancer [Kim et al., 2003]. Microvesicles derived from activated platelets was also reported to induce metastasis and angiogenesis in lung cancer [Janowska-Wieczorek et al., 2005].

#### **Inflammatory diseases**

Boilard et al. demonstrated that platelets are crucial for the development of arthritis. They showed that PMP may facilitate arthritis progression [Boilard et al., 2010]. Other authors reported increased level of PMP in septic patients [Nieuwland et al., 2000, Mostefai et al., 2008]. Suprisingly, early elevated level of PMP and endothelial MP may predict a more favorable outcome in severe sepsis [Soriano et al., 2005].

#### **4. Blood leukocyte derived MP**

Major part (~80%) of MP present/detected in serum/blood is released by platelets, while the remaining ~20% constitute MP shed by erythrocytes, leukocytes and endothelial cells [Ratajczak et al., 2006].

Leukocyte-derived MP circulate in the blood stream under normal conditions and are rapidly up-regulated by inflammatory stimuli, e.g. polymorphonuclear leukocytes (PMNs)

0 50 100 150 200 250 FSC-H

Fig. 4. Chemotactic activity of human blood monocytes to PMP analyzed by flow cytometry. The dot-plots show: monocytes defined in PBMC population by FSC and SSC parameters – region R1 (left); spontaneous chemotaxis of monocytes to culture medium without serum

thrombocytopenias (heparin-induced, thrombotic and idiopathic thrombocytopenic purpura) [Sheriadan et al., 1986, Warkentin et al., 1994]. PMP are strongly procoagulant as they contain the anionic phospholipid PS. Procoagulant PMP are present *in vivo* in blood of patients with activated coagulation and fibrinolysis including patients suffering from Disseminated Intravascular Coagulation (DIC), however, such PMP are also found in healthy donors but in low amounts only [Holme et al., 1994]. An elevated number of PMP was described in patients with diabetes mellitus, which causes the development and