Biomedical Microdevices

, Volume 13, Issue 3, pp 453–462

Exploiting osmosis for blood cell sorting


  • Vahidreza Parichehreh
    • Department of BioengineeringUniversity of Louisville
  • Rosendo Estrada
    • Department of BioengineeringUniversity of Louisville
  • Srikanth Suresh Kumar
    • Department of BioengineeringUniversity of Louisville
  • Kranthi Kumar Bhavanam
    • Department of BioengineeringUniversity of Louisville
  • Vinay Raj
    • Department of BioengineeringUniversity of Louisville
  • Ashok Raj
    • Division of Hematology/Oncology, Department of Pediatrics, School of MedicineUniversity of Louisville
    • Department of BioengineeringUniversity of Louisville

DOI: 10.1007/s10544-011-9513-y

Cite this article as:
Parichehreh, V., Estrada, R., Kumar, S.S. et al. Biomed Microdevices (2011) 13: 453. doi:10.1007/s10544-011-9513-y


Blood is a valuable tissue containing cellular populations rich in information regarding the immediate immune and inflammatory status of the body. Blood leukocytes or white blood cells (WBCs) provide an ideal sample to monitor systemic changes and understand molecular signaling mechanisms in disease processes. Blood samples need to be processed to deplete contaminating erythrocytes or red blood cells (RBCs) and sorted into different WBC sub-populations prior to analysis. This is typically accomplished using immuno-affinity protocols which result in undesirable activation. An alternative is size based sorting which by itself is unsuitable for WBCs sorting due to size overlap between different sub-populations. To overcome this limitation, we investigated the possibility of using controlled osmotic exposure to deplete and/or create a differential size increase between WBC populations. Using a new microfluidic cell docking platform, the response of RBCs and WBCs to deionized (DI) water was evaluated. Time lapse microscopy confirms depletion of RBCs within 15 s and creation of > 3 μm size difference between lymphocytes, monocytes and granulocytes. A flow through microfluidic device was also used to expose different WBCs to DI water for 30, 60 and 90 s to quantify cell loss and activation. Results confirm preservation of ∼ 100% of monocytes, granulocytes and loss of ∼ 30% of lymphocytes (mostly CD3+/CD4+) with minimal activation. These results indicate feasibility of this approach for monocyte, granulocyte and lymphocyte (sub-populations) isolation based on size.


MicrofluidicsCell sortingBlood cells

1 Introduction

Blood is a living tissue that perfuses the entire body and is the primary mediator of immune homeostasis. Local and systemic changes following injury and disease are almost immediately reflected as changes in molecular and cellular constituents of blood. Cellular populations in blood of particular interest for high-throughput gene and protein expression studies are leukocytes or white blood cells (WBCs) (Calvano et al. 2005; Cobb et al. 2005). Prior to analysis of WBCs, it is often beneficial to deplete contaminating erythrocytes or red blood cells (RBCs) which make up ∼ 99% of the cells in blood (Feezor et al. 2004). Further, separation of WBCs into individual populations can enhance the quality of analysis by minimizing heterogeneity within the sample and reducing noise in molecular expression data (Laudanski et al. 2006). An important yet often overlooked consideration during the isolation process is minimizing cell loss and artificial activation due to the isolation process (Kuijpers et al. 1991; Lundahl et al. 1995; Macey et al. 1995). This process is extremely critical to ensure generation of high fidelity information from subsequent genomics and proteomic analysis and will minimize generation of artifactual data as a consequence of isolation process induced events.

Currently, the most effective techniques for isolation of WBC sub-populations use immuno-specificity, (i.e.) antibodies specific to antigens or cell surface phenotype markers to accomplish separation (Neurauter et al. 2007; Sleasman et al. 1997; Ujam et al. 2003). While such approaches are highly specific and can be integrated with magnetic bead based separation, immuno-affinity columns and density gradient separation, the use of antibodies and the binding event itself is a source of unnecessary signaling and downstream signal transduction. For example, anti- cluster of differentiation (CD) 66b used to phenotype neutrophils and esonophils (sub-populations of granulocytes) is a key mediator in adhesion, activation and downstream signaling (Yoon et al. 2007). Therefore techniques that exploit physical properties of cells rather than chemical or biological targets to accomplish separation hold strong appeal for blood cell sorting.

Size-based sorting is one particular option that has been explored extensively in recent years. Three groups in particular have exploited flow phenomenon that develop in curvilinear micro-channels channels with a rectangular cross-section to separate cells and particles based on size (Kuntaegowdanahalli et al. 2009; Di Carlo et al. 2008; Russom et al. 2009a). This phenomenon is based on trapping cells/particles in equilibrium positions due to inertial lift forces generated in channels with rectangular cross-sections at Reynolds Numbers (Re) between 1–10 and Dean’s vortices that develop perpendicular to the direction of fluid flow when the channel is arranged in a curved or spiral fashion. Other alternatives to inertial sorting like deterministic lateral displacement (Green et al. 2009; Huang et al. 2004), acoustic size-based sorting (Kapishnikov et al. 2006), cross-flow filtration (Chen et al. 2008) and size-based filtration (Ji et al. 2008; Murthy et al. 2006; Sethu et al. 2006a) can also benefit from this technique to enable sorting of blood cells. With inertial sorting techniques, the smallest size difference necessary for reliable discrimination of particles/cells is 3 μm (Russom et al. 2009b) whereas with the deterministic lateral displacement the minimum size difference necessary for sorting is ∼ 2.5 μm (Huang et al. 2004). Other techniques do not report the smallest size difference necessary to accomplish sorting.

This manuscript describes efforts taken to determine if controlled exposure to hypotonic solution like deionized (DI) water can be used to deplete certain blood cell populations (particularly RBCs) and create a suitable size difference between other WBC populations to facilitate separation into unique sub-populations. To accomplish this, a cell docking module was designed and fabricated to allow extra-cellular conditions to be changed almost instantaneously while the response of cells can be monitored and characterized using an inverted microscope. Computational fluid dynamics (CFD) modeling was initially performed to determine if trapped cells remained within the grooves and mass transfer was almost instantaneous. The model was then experimentally validated and used to study the response of different blood cells (RBCs and WBCs) to DI water to characterize the size increase and eventual lysis as a function of time. Results indicate that specific exposure times allow complete depletion of RBCs and sorting of WBCs. Cell lysis and activation were also evaluated using a flow-through device to subject isolated WBCs to 30, 60 and 90 s exposure to DI water. This device essentially performs the same function as the docking device but does not allow for visualization of the cells. Since sample continuously flows through the channels, larger sample volumes can be processed and sufficient number of cells can be obtained for flow cytometry. Preservation of ∼ 100% of granulocytes and monocytes was observed along with ∼ 70% recovery of lymphocytes. Activation studies using early activation markers reveal that lymphocytes and monocytes are not activated even after 90 s of exposure to DI water whereas there was minimal (statistically insignificant) activation of granulocytes.

2 Materials and methods

2.1 Blood samples

4 mL of blood was withdrawn from healthy volunteers by venipuncture as per protocols approved by the University of Louisville Institutional Regulatory Board (IRB) and collected in green top vacctuainers (Fisher Scientific, Florence, KY) with heparin as anticoagulant. Samples were maintained on ice until use. All data is represented as mean values for a sample size of n = 5 ± SEM where SEM is the standard error of the mean and can be calculated by dividing the standard deviation by the square root of the sample size.

2.2 Cell docking device fabrication

The two layer device (Fig. 1) was fabricated using standard soft-lithography procedures. First, a layout of the channels and the grooves were created separately using AutoCAD (Autodesk, San Rafael, CA), a layout software. The layout as printed as a bright field mask on a transparency at 18,000 dots per inch (dpi) using a high resolution printer (Fineline Imaging, Colorado Springs, CO). The masks were then used to define features on silicon wafers coated with SU-8 50 (Microchem, Newton, MA) using ultraviolet (UV) light exposure. Finally, the features on the silicon wafers were developed and used as negative replica molds for fabrication of the channel structure and the grooves. The structures were molded using (poly)dimethyl siloxane (PDMS) (Dow Corning, Midland, MI) by mixing the pre-polymer with the cross-linking agent in a ratio of 10:1 and baking in an 80°C oven for 3 h. Following molding the channels and grooves were cut out from the silicon wafer, access holes were punched and irreversibly bonded after treatment with oxygen plasma in a plasma asher (Nordson March Instruments, Amherst, OH). Inlet and outlet tubing were then press fitted prior to use.
Fig. 1

A schematic of the cell docking device, layout of the channel with two sets of grooves at the bottom and 10× image of the grooves obtained using an inverted microscope and image of two cells captured within the grooves

2.3 Controlled exposure device

This device (Fig. 2) was fabricated as discussed above. The design of this device ensures introduction of cell sample and DI water via inlets 1 and 2. The sample in DI water traverses a channel that is 168 cm long, 500 μm wide and 128 μm tall. Samples were introduced into the device at a ratio of 10:1 (DI water: Cell Sample) and allowed to mix via double herringbone structures at the bottom of the channel to induce chaotic mixing. The 30, 60 and 90 s exposure was accomplished using DI water at a flow rates of 400, 200 and 100 μL/min and cell sample flow rates of 40, 20 and 10 μL/min. Samples were returned to isotonic conditions at the outlet using 2× Phosphate buffered Saline (PBS) which was flowed in at the same rate as that of the DI water to ensure return to isotonic conditions.
Fig. 2

Image of device used to subject isolated white blood cells to DI water for predetermined durations (30, 60 and 90 s). Whole blood (inlet 1) and mix with DI water (inlet 2) in a 30:1 ratio and are collected at the outlet following return to isotonic conditions via mixing with 2× PBS (inlet 3)

2.4 Fluid flow modeling

To predict wall shear stress as a function of channel height and flow rate, we used ANSYS 12.0 Academic Teaching Advanced FLOTRON module (ANSYS, Canonsburg, PA). The channel and grooves were modeled in 2D and the fluid flow was modeled as steady state and incompressible flow. The fluid was assigned physical properties of water: density (ρ) = 1000 kg/m3 and dynamic viscosity (μ) = 0.001 Pa s. The following boundary conditions were applied: (a) input fluid flow rate at inlet = 100 μL/min, (b) outlet pressure = 0 Pa and (c) rigid walls with no-slip for both the channels and grooves. The fluid domain was meshed using 2D quadrilateral element, FLUID141. Then the model was solved using ANSYS finite element software. This approach is based on a segregated sequential solver algorithm that uses a matrix system derived from the finite element discretization of the governing equation for each degree of freedom. At each simulation, the elements showing high velocity gradients were refined until convergence with predicted flow velocities was achieved and then the fluid flow stream lines and flow velocities were determined within the channel and the grooves (Fig. 3).
Fig. 3

CFD modeling results. As can be seen from the images, velocity of fluid in the main channel is ∼ 8 mm/s whereas at the bottom of the grooves, recirculation occurs at significantly lower velocity of ∼ 0.014 mm/s. This ensures that cells trapped within the grooves do not escape the grooves but the recirculation helps in accomplishing rapid mass transfer. It should be noted that cells did however experience slight lateral movement within the grooves

2.5 Experimental procedure for immobilization and analysis of cells

The device was primed with 1 × phosphate buffered saline (PBS) to eliminate air and trapped bubbles within the device prior to use. Cells were suspended in 1 × PBS with 0.4% trypan blue solution to aid visualization and introduced into the device. Cell concentration was adjusted to ∼ 1 × 105 cells/ml to provide sufficient cellular populations for analysis. Cells were allowed to sediment within the device for ∼ 2 min and then the cells not immobilized within the grooves were removed using 1 × PBS with 0.4% trypan blue. A 10 mL syringe was filled with DI water and loaded onto a syringe pump (Harvard Apparatus, Holliston, MA) which was set to deliver fluids at ∼ 100 μL/min. The flow of DI water was initiated and a stop clock was started as soon as a visible color change was observed within the grooves (DI water replacing the 1 × PBS with trypan blue). Images were taken every 3 s and analyzed using Metamorph software (Molecular Devices, Sunnyvale, CA). Briefly, the measurements of cell size were accomplished manually. Once the image was captured and the diameter of each individual cell was estimated by measuring the length of the cell from end to end. This was accomplished 10 different times from different directions and the average values were used to determine the cell diameter.

2.6 Separation of peripheral blood mononuclear cells (PBMCs) and polymorphonuclear cells (PNMs)

PBMC and PNM separation from whole blood was accomplished using standard density gradient separation. Briefly, 4 mL of blood was mixed with 4 mL of 1 × phosphate buffered saline (PBS) and carefully layered on top of 10 mL of Ficol Paque (GE Healthcare, Waukesha, WI) in a 50 mL tube. Samples were then balanced and centrifuged for 30 min at 450 g. Following this procedure, PBMCs were fractionated as a narrow band below the plasma and above the Ficol whereas the PNMs and RBCs pellet to the bottom below the Ficol. Each fraction was isolated and the PBMCs were washed once with 1 × PBS whereas the PNM and RBC mixture was mixed with 30 mL of ammonium chloride lysis buffer for 5 min to eliminate RBCs. The samples were then again centrifuged at 250 g for 5 min and resuspended in 1 × PBS and maintained at room temperature until needed.

2.7 Cell counts

Sample cell concentration was estimated prior to introduction into the controlled exposure device and again counted after exposure to DI water and return to isotonic conditions at the outlet. Cells were counted using a hemocytometer. The difference between the inlet and outlet counts represents the number of cells lost due to DI water exposure. Cell Counts of different sub-populations was estimated using flow cytometry using a combination of light scatter and phenotype markers.

2.8 Activation studies

Activation studies were performed using flow cytometry. Samples were first suspended in a flow cytometry buffer consisting of 1 × PBS and 1% bovine serum albumin (BSA). Antibodies used were conjugated to fluorescently labeled tags such as Fluorescein isothiocyanate (FITC), phycoerythrin (PE), Peridinin Chlorophyll Protein Complex (PerCP) and Allophycocyanin (APC) and evaluated using a four color flow cytometer (BD FACS Calibur, BD Biosciences, Nashville, TN). Antibodies specific to cell surface markers CD3 and CD4 for lymphocytes, CD14 for monocytes and CD66b for granulocytes were used to phenotype white blood cells into major sub-populations. Once cells were phenotyped, they were evaluated for activation using early activation markers identified previously (Sethu et al. 2006b). Expression of CD25 and CD69 were evaluated for lymphocytes and expression of CD18 and CD29 were evaluated for monocytes and granulocytes.

3 Results

3.1 Device design and modeling

To enable characterization of changes in cell size in response to changing extra-cellular conditions a device was designed to first immobilize cells within grooves (50 μm × 50 μm) in a microfluidic channel (Fig. 1). This device is compatible with an inverted microscope where images of cells at different time points can be acquired using a charge coupled device (CCD) camera and analyzed using image processing software. The aspect ratio of the grooves within the microchannel was designed to be ∼ 1:1. CFD modeling was used to determine the effect of various inlet flow rates on cells trapped at the bottom of the grooves. CFD results indicate that the fluid flowing in the main channel at a flow rate of 100 μL/min, corresponding to a main channel velocity of 8 mm/s does not directly affect cells trapped at the bottom of the channel. Rather, the fluid flow in the main channel causes recirculation at lower velocities (~ 0.014 mm/s) within the bottom half of each groove (Fig. 2). These results confirm theoretically, that cells trapped at the bottom of the grooves do not get carried away by the fluid in the main channel and mass transfer is accomplished instantaneously and effectively due to a combination of diffusion mixing and recirculation effects at the bottom of the channel. These modeling results were confirmed experimentally using cells suspended in trypan blue and introduced into the device and flushing with 1 × PBS. Cells remained immobilized within the grooves and mass transfer was accomplished in ∼ 2 s following entry into the device as visualized by replacement of the blue color of trypan blue in the grooves. This ∼ 2 s equilibration time was taken into account for all subsequent experiments.

3.2 RBC characterization

Blood was processed as described in the Methods Section and separated into PBMCs and PNMs. First, the swelling and lysis of RBCs were evaluated by introducing RBCs suspended in trypan blue containing 1× PBS into the device. Following 2 s of DI water perfusion into the device, RBCs begin to swell, almost instantaneously transition from a biconcave to spherical shape and finally lyse due to buildup of osmotic pressure. Analysis of images captured every 2 s using the CCD camera show that RBCs reach a maximum size of ∼ 10.6 ± 0.092 μm and begin to lyse at ∼ 10 s and are completely lysed by 15 s (Fig. 4(a)).
Fig. 4

(a) RBCs trapped within the grooves swell, increase in size and begin lysis at ∼ 10 s. Complete lysis is accomplished within 15 s. Lysis of RBCs results in ghost cells (cells w/ compromised membranes), (b) Lymphocytes immobilized within the grooves increase in size to ∼ 11 μm and begin lysis at ∼ 30 s. Nearly 30% of the lymphocytes are lysed whereas the remaining ∼ 70% remain at 11 μm in size without lysis, (c) Monocytes studies using this device are initially ∼ 10.7 ± 0.291 μm in diameter and attain a final diameter of ∼ 17 ± 0.210 μm following 100 s of exposure without lysis. Also seen in the image are lymphocytes in the PBMC fraction, (d) Granulocytes, initially 8.03 ± 0.262 μm in diameter increase in size to a final size of ∼ 14.8 ± 0.176 μm following 100 s of exposure

3.3 PBMC characterization

PBMCs or the mononuclear cell fraction is a mixture of primarily monocytes and lymphocytes. In isotonic conditions monocytes are typically ∼ 10.7 ± 0.291 μm in diameter and lymphocytes are ∼ 7.6 ± 0.156 μm in diameter allowing visual discrimination between the two cell types. PBMC samples with trypan blue were introduced into the device and evaluated in a similar fashion to the RBCs. Image analysis was performed to track cells identified as lymphocytes and monocytes based on initial size (Fig. 4(b) and (c)). Monocytes, initially ∼10.7 ± 0.291 μm in diameter swell and reach a size of 17 ± 0.210 μm following 100 s of exposure. Lymphocytes on the other hand increase from their initial size of ∼ 7.6 ± 0.156 μm to 11 ± 0.071 μm within 30 s. At this time lysis of lymphocytes begins and a ∼ 30% of the lymphocytes begin to lyse and are completely lysed by 40 s. The remaining 70% of the lymphocytes remain at a size of 11 ± 0.071 μm until 100 s without any change in size.

3.4 PNM characterization

PNMs collectively refer to granulocytes including neutrophils, basophils, eosinophils and other cells including mast cells. These cells are characterized by multiple nuclei and highly granular cytoplasm which is a source of high side scatter in flow cytometry. PNMs obtained following lysis of contaminating RBCs was mixed with trypan blue, introduced into the device and characterized in a similar fashion to RBCs and PBMCs. Image analysis of granulocytes in isotonic conditions measure ∼ 8.03 ± 0.262 μm in diameter and following exposure to DI water gradually increase in size to ∼ 14.8 ± 0.176 μm in diameter and remain intact even 100 s in DI water (Fig. 4(d)).

3.5 Combined analysis

Comparative analysis of all blood cell populations was accomplished by plotting cell size increase as a function of time for all cells (Fig. 5). Complete lysis of RBCs is accomplished at 15 s. At this point the size of lymphocytes, monocytes and granulocytes are ∼ 8.8 ± 0.139, 11.3 ± 0.225 and 9.5 ± 0.253 μm respectively which is not a sufficient size difference for sorting. At the 100 s time point, 70% of the lymphocytes remain and are ∼ 11 ± 0.071 μm in diameter whereas the monocytes are ∼ 17 ± 0.210 μm and granulocytes are ∼ 14.8 ± 0.176 μm in diameter. Therefore a 100 s exposure provides the best opportunity to separate WBCs into monocytes, granulocytes and a sub-population of lymphocytes.
Fig. 5

Graph tracking cell size increase over time. The plot represents the swelling behavior of Monocytes, Lymphocytes, Granulocytes and RBCs. RBCs lyse within 15 s whereas 30% of the lymphocytes lyse within 40 s. The shaded region represents the remaining 60% lymphocytes. A sufficient size difference (~ 3 μm) between the 3 WBC population is achieved following 90 s of exposure to DI water

3.6 Cell recovery

PBMC and PNM fractions were counted following isolation and after 30, 60 and 90 s of exposure to DI water at a ratio of 10:1 (DI water: sample) in the controlled exposure device. Results indicate that ∼ 30% of lymphocytes are lost for exposure times of 60 and 90 s whereas all other cell populations remain minimally changed (Table 1).
Table 1

Cell recovery following exposure to DI water (sample size n = 5)

% of cells recovered


30 s

60 s

90 s













































3.7 Cell activation

Following 30, 60 and 90 s of exposure to DI water in the controlled exposure device, cells were phenotyped using light scatter and phenotype markers. Lymphocytes were characterized into three major populations CD3+/CD4+ (T helper 1), CD3+/CD4 (T helper 2) and CD3/CD4 (B cells and NK cells) whereas Monocytes and Granulocytes were characterized using CD14 and CD66b respectively. Expression of early activation markers CD25 and CD69 was evaluated on the 3 lymphocyte populations (Fig. 6) and results indicate that there is no activation. Similarly, the expression of early activation markers CD18 and CD29 was profiled on CD14+ monocytes and CD66b+ granulocytes and results indicate that there is no activation of monocytes whereas there is minimal activation of granulocytes for both markers in comparison to controls (Fig. 7). Data represented in Figs. 6 and 7 is for one representative sample. Similar results were obtained for the other four samples that were evaluated using flow cytometry.
Fig. 6

Activation status of Lymphocytes following 30, 60 and 90 s of exposure to DI water determined using early activation markers CD25 and CD69. Lymphocytes were phenotyped into three subpopulations as CD3+/CD4+, CD3+/CD4 and CD3/CD4. Results indicate no DI water induced activation in comparison to controls
Fig. 7

Activation status of Monocytes (CD14+) and Granulocytes (CD66b+) following 30, 60 and 90 s of exposure to DI water using early activation markers CD18 and CD29. Results show no monocytes activation and minimal granulocyte activation in comparison to controls

4 Discussion

Blood cell sorting without the use of cell specific antibodies is important to minimize unnecessary signaling in cell samples prior to molecular expression analysis. Inertial microfluidic sorters that exploit flow phenomenon in curvilinear channels with rectangular cross-section hold great promise to accomplish antibody-free sorting based on size. Without appropriate sample pre-processing, blood cells cannot be sorted into WBC subpopulations using inertial microfluidic sorters due to abundance of unwanted RBCs and size overlap between different WBC populations. Other size based sorting techniques like lateral deterministic displacement, size-based filtration, size-based acoustic sorting and cross-flow filtration can also benefit from cell size amplification. Subjecting cells to hypotonic solutions like deionized water results in fluid transport across the cell membrane and a buildup in pressure within the cells. The process of cell swelling and ultimately lysis depends on the cells membrane properties including presence or absence of water channels (aquaporins), cytoplasm to nucleus ratio and intracellular ratios of F-actin to G-actin. This process can be potentially exploited to selectively deplete certain cells while also creating a sufficient size difference between remaining cell populations to provide a suitable starting sample for size-based sorting.

To determine the feasibility of using an osmosis based approach for sample pre-processing, the response of blood cells to hypotonic solutions like DI water needs to be characterized. A new microfluidic cell docking device was designed and constructed for this purpose. Fluid flow in channels with grooves that have aspect ratio ∼ 1:1, 50 μm (width): 50 μm (height) does not affect the bottom of the grooves directly but creates recirculation patterns at the bottom of the grooves which is extremely important in ensuring mass transfer is rapid and the conditions within the wells match conditions in the main channel. This provides a unique platform for visually characterizing the immediate and time dependent response of cells to changing extracellular stimulus.

Experiments performed with the cell docking device indicate that RBCs can be eliminated within 15 s of hypotonic exposure without loss of any WBC populations. At the 30 s time point, the lymphocytes begin to lyse and within 40 s of exposure, ~ 30% of the lymphocytes are lysed. The remaining 70% of the lymphocytes remain at a size of 11 μm in diameter till the conclusion of the experiment at 100 s. Lymphocytes consist of four unique sub-populations: T helper cells, cytolytic T cells, B cells and natural killer (NK) cells. We speculate that the remaining cells may represent a unique sub-population of lymphocytes. At the 40 s time point, the size difference is still unsuitable for separation of the three sub-populations. However, with increase in exposure time the granulocytes and monocytes differentially increase in size whereas the lymphocytes remain at 11 μm. At 60 s the size difference between the remaining lymphocytes and the larger cells is > 3 μm and which provides an opportunity to sort the remaining lymphocytes. A ∼ 3 μm size difference between granulocytes and monocytes appears only following 90–100 s of exposure. Therefore, a 90 s exposure provides the best opportunity to eliminate RBCs and sort monocytes, granulocytes and a sub-population of lymphocytes based on size.

To confirm this technique does not induce WBC activation, activation studies were performed using commonly used early activation markers. These studies confirm that that most cells were not activated and the granulocytes showed minimal activation which was found to be statistically insignificant. However, there was loss of lymphocytes following > 30 s of exposure to DI water. These cells were predominantly CD3+/CD4+ lymphocytes. Despite loss of this sub-population of lymphocytes, this technique can be used to isolate other lymphocyte subpopulations as well as granulocytes and monocytes without loss and minimal activation.

5 Conclusion

Size-based sorting is an attractive alternative to immuno-affinity based approaches. However, prior to application for blood cell sorting RBCs need to be eliminated and sufficient size difference need to created between different WBC populations. Using the microfluidic docking device, we have demonstrated that osmosis that ensues following DI water exposure can be exploited to first deplete RBCs and create sufficient size differences between different WBC populations for subsequent sorting using size-based sorting techniques.


The authors would like to thank Tim Andrews, phlebotomist, Division of Pediatric Hematology/Oncology for help with obtaining blood samples and helpful discussions. This work was supported through a proof-of-concept grant (POCG) through the Office of Technology Transfer and the Vice President of Research at the University of Louisville and by the National Science Foundation under Grant No. 0814194.

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© Springer Science+Business Media, LLC 2011