Highly Efficient Assay of Circulating Tumor Cells by Selective

Aug 10, 2012 - In Vitro Diagnostics Lab, Bio Research Center, Samsung Advanced Institute of Technology, Samsung Electronics Co., Ltd., San No. 14-1 ...
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Highly Efficient Assay of Circulating Tumor Cells by Selective Sedimentation with a Density Gradient Medium and Microfiltration from Whole Blood Jong-Myeon Park,*,†,‡ June-Young Lee,† Jeong-Gun Lee,† Hyoyoung Jeong,† Jin-Mi Oh,† Yeon Jeong Kim,† Donghyun Park,† Minseok S. Kim,† Hun Joo Lee,†,§ Jin Ho Oh,∥ Soo Suk Lee,† Won-Yong Lee,*,‡ and Nam Huh† †

In Vitro Diagnostics Lab, Bio Research Center, Samsung Advanced Institute of Technology, Samsung Electronics Co., Ltd., San No. 14-1, Nongseo-dong, Giheung-gu, Yongin-si, Gyeonggi-do, Republic of Korea ‡ Department of Chemistry and Center for Bioactive Molecular Hybrids, Yonsei University, Seoul 120-749, Republic of Korea § Interdisciplinary Program of Integrated Biotechnology, Sogang University, Shinsu-dong, Mapo-gu, Seoul, Republic of Korea ∥ Samsung Biomedical Research Institute, Samsung Advanced Institute of Technology, Samsung Electronics Co., Ltd., San 14, Nongseo-dong, Giheung-gu, Yongin-si, Gyeonggi-do, Republic of Korea S Supporting Information *

ABSTRACT: Isolation of circulating tumor cells (CTCs) by size exclusion can yield poor purity and low recovery rates, due to large variations in size of CTCs, which may overlap with leukocytes and render size-based filtration methods unreliable. This report presents a very sensitive, selective, fast, and novel method for isolation and detection of CTCs. Our assay platform consists of three steps: (i) capturing CTCs with antiEpCAM conjugated microbeads, (ii) removal of unwanted hematologic cells (e.g., leukocytes, erythrocytes, etc.) by selective sedimentation of CTCs within a density gradient medium, and (iii) simple microfiltration to collect these cells. To demonstrate the efficacy of this assay, MCF-7 breast cancer cells (average diameter, 24 μm) and DMS-79 small cell lung cancer cells (average diameter, 10 μm) were used to model CTCs. We investigated the relative sedimentation rates for various cells and/or particles, such as CTCs conjugated with different types of microbeads, leukocytes, and erythrocytes, in order to maximize differences in the physical properties. We observed that greater than 99% of leukocytes in whole blood were effectively removed at an optimal centrifugal force, due to differences in their sedimentation rates, yielding a much purer sample compared to other filter-based methods. We also investigated not only the effect of filtration conditions on recovery rates and sample purity but also the sensitivity of our assay platform. Our results showed a near perfect recovery rate (∼99%) for MCF-7 cells and very high recovery rate (∼89%) for DMS-79 cells, with minimal amounts of leukocytes present.

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One attractive separation method is cell size-based direct isolation of CTCs using membrane filters. Recently, various types of membrane filters, such as track-etched polycarbonate membrane filters,13 multilayered (or single) microfabricated parylene membrane filters, and microcavity arrays have been developed.14,15 In addition, size filtration devices that utilize lateral flow have also been developed.16−18 An alternative to filtration methods is density gradient centrifugation using Ficoll-Hypaque, which can separate mononuclear cells, including CTCs, with a density of less than 1.077 g/mL from other cells in whole blood. Density gradient centrifugation is a method that can be used to enrich any type of CTCs, because the performance of the density gradient medium is independent from the presence of specific markers. Similarly, OncoQuick is a

etastasis is responsible for over 90% of cancer-related deaths.1 Circulating tumor cells (CTCs) are disseminated from primary tumors or metastatic sites and then enter into the bloodstream. The presence of CTCs in the peripheral blood has been regarded as an important indicator of potential for metastatic disease.2−5 However, an assay that can isolate and count CTCs from human whole blood is technically challenging, due to the extremely low numbers if CTCs in the peripheral blood, which has been estimated to be as low as 1 in 109 blood cells. To overcome the hurdle of low concentrations of CTCs in peripheral blood, a variety of techniques have been developed to separate CTCs from whole blood. Many of these methods rely on differences between CTCs and other cells in either (a) physical characteristics, such as density, cell size, and electrical properties, or (b) biological characteristics, such as expression of protein markers, cancerspecific antigen−antibody interactions, and combinations of these two characteristics.6−12 © 2012 American Chemical Society

Received: May 2, 2012 Accepted: August 10, 2012 Published: August 10, 2012 7400

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Figure 1. Schematic illustration of the concept for CTC separation. (A) Distribution of size and density in hematopoietic cells, CTCs, and sizedensity amplification microbeads (SDABs) conjugated CTCs. (B) Selective sedimentation of CTC-SDABs in DGM from other blood cells. (C) Recovery and detection of the CTC-SDABs by the filtration through a filter device. The dimensions of filter device were (length × width × thickness) 30 mm × 15 mm × 2 mm, with slot widths of 10, 12, or 14 μm.

of CTCs and subsequent filtration. As shown in Figure 1A, physical properties of CTCs are very similar to those of leukocytes in density and size, and thus, small CTCs (less than 15 μm in diameter) cannot be clearly distinguished from leukocytes by size exclusion alone, while relatively large CTCs (larger than 15−20 μm in diameter) can be discriminated from leukocytes by size.27 To model the variety in size of CTCs, DMS-79 small cell lung cancer cells (mean diameter = 10 μm) and MCF-7 human breast cancer cells (mean diameter = 24 μm) were used.28 Anti-EpCAM capturing antibodies were conjugated to the surface of size-density amplification beads (SDABs), whose diameter and density are 2.8 μm and 1.3 g/ mL, respectively. The size and density of CTCs could simultaneously be amplified by incubating them with SDABs. As shown in Figure 1B,C, our approach consists of amplification of the sedimentation rate for CTCs by SDABs and selective sedimentation of SDAB-bound CTCs (CTCSDABs) from whole blood, by controlling the gravity force with a density gradient medium (DGM). CTC-SDABs are selectively precipitated at the bottom of a centrifuge container, while most of the leukocytes and erythrocytes remain in suspension at an optimum g force and time, in the presence of DGM, due to differences in sedimentation rates between CTCSDABs and unwanted cells. In order to recover and detect only the desired cells, the bottom-precipitated fractions of CTCSDABs is filtered through a microfilter device, after removal of the unwanted fraction. In this article, we detail the performance of our novel selective sedimentation assay, in terms of its improved recovery rate and purity, compared to previously available methods to isolate CTCs.

separation method, based on a density gradient medium, which has a unique porous barrier that separates layers with different densities but prevents any cross-contamination.19−22 Although these isolation technologies offer many advantages, they still suffer from several shortcomings. First, the size-based direct filtration method is based on the assumption that CTCs are larger than leukocytes.23 Unfortunately, to the best of our knowledge, there have been no studies that show all CTCs are indeed larger than leukocytes. Recently, in prostate cancer patients, the sizes of CTCs have been reported to range from 8 to 16 μm in diameter.24 Consequently, direct filtered CTCs can be contaminated by leukocytes with a relatively larger size and, at the same time, CTCs that are small in size may pass through the filter. Therefore, a size-based filtration method to isolate CTCs can lead to low purity and recovery rate. Second, the lateral flow applied to the filtration device is relatively slow, resulting in slow sample processing and limited overall sample throughput. Besides, application of a large volume of blood sample to direct microfilters may result in failed isolation, due to clogging of microfilters by a large number of unwanted cells.18 Lastly, because of the similarity in density between CTCs and monocytes, CTC fractions are still mixed with a number of unwanted monocytes after density gradient centrifugation with Ficoll. In addition, during the migration of cells to the plasma layer, while centrifuging, CTCs can be easily lost.2 It is well-known that the presence of leukocytes in CTC analysis can result in false positive enumeration of CTCs and may also causes severe interferences with CTC-specific molecular analysis.25,26 Here, we report a highly sensitive and selective novel CTC assay platform based on simultaneous size-density amplification 7401

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EXPERIMENTAL SECTION Preparation of SDAB. SDAB was prepared with monoclonal antihuman EpCAM/TROP1 antibody (R&D Systems, MN) using standard carbodiimide chemistry in a single reaction. Briefly, the magnetic protein G (pG) modified microbeads (Invitrogen, CA) were washed with 1 mL of NaOAc buffer at pH 5.6 three times and mixed with 100 μg of monoclonal antihuman EpCAM/TROP1 antibody. The reaction container was kept in rotation at 12 rpm for over 1.5 h at room temperature on a rotator (PTR-60, Grant-bio, Shepreth, U.K.). After the beads were collected, DMP (SigmaAldrich, MO) was added and mixed at room temperature for 1 h. The reaction was stopped by the addition of 50 mM ethanolamine. The resulting beads are then washed three times with PBS buffer at pH 7.4, followed by blocking with 5% BSA for 2 h at room temperature. Finally, the beads were resuspended and stored in 1% BSA. CTC Selective Sedimentation by SDAB. CTC recovery was investigated using both a breast cancer (MCF-7) and small cell lung cancer (DMS-79) cell line. All cell lines were obtained from ATCC (Rockville, MD) and cultured in accordance with their recommendations. For binding cells and SDABs, the actual number of cells in the cell suspension was determined by the cell counting method, as previously described.14 A total of 100 cells were introduced into a tube with 1 mL of whole blood sample, followed by the addition of 40 μL (2.0 × 109/mL) of SDABs. The entire tube was incubated for 1.5 h at room temperature, while constantly being rotated at 12 rpm. Then, 1 mL of preincubated blood samples were carefully layered onto 2 mL of Ficoll-Hypaque gradient medium. The blood-cell mixture was centrifuged with a centrifuge (Eppendorf 5810 R, Hamburg, Germany). CTC Isolation and Characterization. After centrifugation at a given g force, 2 mL of the upper layer (leukocytes with erythrocyte-concentrated fraction) was carefully aspirated and discarded. About 1 mL of the residual sample was resuspended with a pipet and transferred to the filtration device which was fabricated by silicon-on-glass (SOG) technology to make an accurate and precise gap between filter slots. The liquid in the blood samples were withdrawn through the filter, using a syringe pump (KDS LEGATO 110, KD Scientific Inc., New Hope, PA), at a specific flow rate. After this initial filtration, cells trapped in the microfilter device were washed with 2 mL of PBS buffer at a flow rate of 100 μL/min. Prior to any immunostaining, cell fixation was conducted with 4% paraformaldehyde through the microfilter for 20 min at a flow rate of 50 μL/min. Cells were then permeabilized in 0.01% Triton X-100 (Sigma-Aldrich, MO) for 10 min at a flow rate of 50 μL/min. To identify CTCs and leukocytes, the mixture of DAPI, anticytokeratin-PE antibody, and anti-CD45-FITC antibody was flowed through the filter at 7.5 μL/min for 60 min. Finally, the microfilter was washed with PBS at a flow rate of 50 μL/min for 10 min. Tumor cells were imaged by a fluorescence microscope (IX81, Olympus Corp., Japan), integrated with a computer-operated motorized stage (TANGO, Märzhäuser Wetzlar GmbH & Co. KG, Germany) (see Figure S-1 in the Supporting Information).

physical properties of erythrocytes and leukocytes. The density and diameter of erythrocytes have been reported to be 1.1− 1.15 g/mL and 6.6−7.5 μm, respectively, while leukocytes have a density of 1.07−1.09 g/mL and a wide range of diameters, 8− 20 μm. The density of CTCs is on par with that of leukocytes. As for their diameter, since most CTCs are derived from epithelial cancers, it had been initially assumed that they will be larger than leukocytes; however, it has been since discovered that there is significant variation in their size range.29 This creates the difficulty of having much of the size range of leukocytes overlapping with that of CTCs. Therefore, CTC isolation, relying solely on size-based filtration, may exclude a significant portion of the CTC population.24 The sedimentation rate of a particle in suspension is proportional to the centrifugal force applied to it. At a fixed centrifugal force and liquid viscosity, the sedimentation rate is proportional to the size of the particle and the difference in density between a particle and its surrounding solution. The equation for the sedimentation of a sphere in a centrifugal field is v=

d 2(ρp − ρl ) 18η

×g

(1)

where ν is the sedimentation rate, d is the diameter of particle, ρp is the particle density, ρl is the density of the DGM, η is the viscosity of the DGM, and g is the centrifugal force. To calculate the sedimentation rate of cells, such as leukocytes, erythrocytes, and CTC-SDABs, at a given g force from eq 1, we assumed that CTCs have diameters between 10 and 24 μm and a uniform density of 1.07 g/mL, respectively. We also assumed that cell surfaces are fully covered with SDABs. For leukocytes and erythrocytes, we assumed that leukocytes have a 20 μm diameter with a density of 1.09 g/mL and erythrocytes have a 7.5 μm diameter and a density of 1.15 g/mL. As is shown in Figure 2A, the density of material used to create SDAB has a great effect on the sedimentation rate of CTC-SDABs. CTCs conjugated to SDAB with a density of 1.05 g/mL cannot settle out of a blood suspension, because CTCSDABs are smaller in density than DGM (d = 1.077 g/mL). However, sedimentation rates increased dramatically, when the density of SDAB increases from 1.3 to 1.6 g/mL. Besides, the size of SDAB is also critical in determining the sedimentation rate of CTC-SDABs. Our calculation showed that the sedimentation rate of CTC-SDABs increases when the diameter of SDAB is increased from 1 to 6 μm, since both the diameter and the density of SDAB-CTC correlates positively with the diameter of SDAB, given that the density of SDAB is greater than that of DGM. Thus, to maximize the difference in sedimentation rates between CTC-SDABs and leukocytes, SDAB should have a high density and large size. However, as large microbeads (diameter, 6 μm) showed poor binding efficiency between microbeads and tumor cells,27 the optimal diameter of SDAB to be used in our selective sedimentation assay was decided as 2.8 μm. We speculate that one reason for this effect may be physical damages to tumor cells during incubation, in blood, with relatively large microparticles. Such issues, including mechanical forces damaging cells, good chance of encounter of rare cells with SDABs, and shear forces acting on CTCs, are commonly shared among all microbead-based techniques, including CellSearch. Thus, further optimization of this assay, including, but not

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RESULTS AND DISCUSSION Estimation of Sedimentation Rate and Selection of SDAB. In order to estimate the sedimentation rate required to selectively isolate CTCs, it is first necessary to consider the 7402

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expression level may vary significantly. To address this issue, we first investigated how much of the cell surface has to be covered by SDABs in order to separate CTC-SDABs from blood cells. The results showed that the sedimentation rate was different between CTC-SDABs and blood cells if greater than 10% of the surface of a CTC is covered by SDAB. This equates to about 4 beads for a CTC with a diameter of 10 μm (see Figure S-2 in the Supporting Information). Consistent to our findings, Nagrath et al. previously reported that the capture efficiency of their system did not differ significantly among cells with differing EpCAM expression level, even between SKBr-3 cells with >500 000 antigens per cell and T-24 cells with ∼2 000 antigens per cell.30 Although we are not able to estimate exactly how many antigens per cell are required to cover 10% of the CTC surface with SDAB, it may be feasible to discriminate low EpCAM expression from the absence of EpCAM. At this moment, however, our assay is limited to capturing and detecting EpCAM-positive CTCs. Regarding this issue, development of new CTC-specific markers is needed to fully realize the clinical utility of an immunoaffinity-based CTC isolation technique, including our platform, which does not have to be limited to be used with only anti-EpCAM antibody. Optimization of the g Force and Detection of CTCs. To verify that our approach is feasible, 1 mL of whole blood was spiked with 100 MCF-7 cells and 40 μL of SDABs was added. The samples after incubation were layered onto the 2 mL of DGM preloaded container. Various centrifugal forces ranging from 50g to 800g were applied for 2 min. Because of the difference in the sedimentation rate, most erythrocytes and leukocytes were found in the uppermost layer, while the lowermost layer contained CTC-SDABs precipitated to the bottom of the container after centrifugation at less than 200g. However, at centrifugal forces greater than 200g, more erythrocytes began to be found in the lowermost layer, reaching to a complete lack of layer separation at centrifugal forces greater than 400g (see Figure S-3 in the Supporting Information). In order to recover and evaluate the fraction of the settled tumor cells bound with SDABs, ∼2 mL of the upper layer, containing leukocytes and erythrocytes, was removed, and the bottommost 1 mL of remaining blood sample was obtained as a CTC fraction. The CTC fraction was directly injected into an inlet hole by applying negative pressure. Subsequently, reagents for washing and staining were serially injected into the micro filter chip for optical inspection via fluorescence microscopy. To identify and count the entrapped cells, whole cellular images were obtained. Recovered cells that were DAPI-positive, CKpositive, and CD45-negative were identified as tumor cells, and cells that were DAPI-positive and CD45-positive were identified as leukocytes. In our experience with fluorescence microscopy, detection of CTCs was straightforward, as cells that were DAPI-positive, CK-positive, and CD45-negative could usually be identified easily. However, because of the presence of beads, the samples could not be used for detailed cellular or subcellular morphological analysis. One option in the future, for better image analysis, is to employ an optically transparent microbead, since our assay does not specifically require magnetic beads. Indeed, our preliminary data with melamine resin beads (d = 1.51 g/mL) as SDAB offered improved structural details in immunofluorescence analysis (data not shown). The second possibility is to remove the microbeads, especially for subcellular image analysis, as reported previously.31 However,

Figure 2. Calculated sedimentation rate: (A) comparison of sedimentation rates for CTC-SDABs (solid lines) and hematopoietic cells (dashed lines) and (B) ratio of the sedimentation rate between CTC-SDABs and leukocytes. The term “d” in the figure denotes the density of the SDAB.

limited to, bead binding time, SDAB size, and SDAB material might be necessary to apply this assay to clinical samples. Figure 2B shows the ratio for calculated sedimentation rates between CTC-SDABs and leukocytes. The sedimentation rate of CTCs (diameter, 10−24 μm) conjugated with 2.8 μm diameter SDABs of density ranging from 1.3 to 1.6 g/mL were 5−25 times larger than that of leukocytes. Although our assay has nothing to do with a magnet, protein G-modified magnetic microbeads, as the beads to be used in our assay, were made for the following nonmagnetic physical properties: (i) appropriate diameter (2.8 μm) and density (1.3−1.8 g/mL), (ii) convenience for conjugating antibodies with the protein Gcoated beads, and (iii) improved activity of antibodies due to proper orientation established by interaction with protein G. In addition to the size and density of SDABs, the number of SDABs bound to a CTC is another critical element in determining sedimentation rates. Although we initially assumed that CTC surfaces are completely covered with SDABs, this may not be the case for clinical samples, where the EpCAM 7403

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Recent studies presumed that these triple positive cells are likely leukocytes, thereby not counting them as CTCs.32−35 Therefore, the optimal centrifuge conditions for selective sedimentation were determined to be 100g for 2 min with a suspension height of 3−4 cm, considering both the recovery rate for CTCs and the final removal rate of leukocytes. The numbers of erythrocytes and leukocytes were initially measured as 3.6 × 109 and 9.5 × 106 cells in the blood samples. After sedimentation, 4.3 × 107 erythrocytes (n = 3, ±5.9%) and 3.9 × 102 (n = 3, ±11.7%) leukocytes were found in the CTC fraction. As a result, the removal efficiency of erythrocytes and leukocytes after the selective sedimentation step were 99.81% and 99.996%, respectively. After filtration, the recovery rate for CTC was 103 ± 6% (n = 4) and the total removal efficiency was determined to be 99.9995% based on the leukocyte starting from 9.5 × 106 and finally observed as less than 50 ± 7 cells (n = 4). To rule out the distinct possibility that triple positive cells were recovered, due to cross-reactivity with anti-EpCAM antibody, we examined if these cells would sediment without SDAB. Whole blood, without spiked tumor cells and SDABs, was layered onto Ficoll-Hypaque gradient medium. After centrifugation at 400g for 20 min at 20 °C, each fraction was filtered through our microfilter device with a 10 μm slot width. A large number of triple-positive leukocytes could be observed in fractions of the lower layer (erythrocyte fraction). On the other hand, CK-positive cells could not be observed in the upper layer (leukocyte fraction) (see the Supporting Information, Figure S-4). From this result, it is clear that triple positive cells are not monocytes but should rather be considered as leukocytes that have relatively high density (>1.077 g/mL) and large size. Furthermore, we were not able to detect leukocytes bound to SDABs, suggesting negligible nonspecific binding between SDABs with anti-EpCAM and leukocytes (data not shown). Although CTC purification based on immunomagnetic beads is currently the leading technology in the clinical setting,14,33,36 one of the main problems with this approach is that CTCs are still outnumbered by around 1000−10000 of leukocytes.26 The contamination of CTCs with leukocytes needs to be addressed by further assays, by utilizing cell-type specific antibody staining and molecular analysis. Compared to previous studies reporting relatively high numbers of leukocyte contamination, our approach shows a marked improvement in removal of leukocytes from CTCs, offering future promise of detailed molecular analysis of CTCs. Assessment of Flow Rate in CTC Detection. As shown in Figure 4, in order to demonstrate the effect that flow rate may have on tumor cell recovery rate and leukocyte removal efficiency, the CTC fractions obtained by selective sedimentation assay were passed through the microfilter device with a slot width of 10 μm at flow rates ranging from 50 to 2000 μL/min, after being subjected to the optimal g force of 100g for 2 min. Results show that recovery rates were decreased from 107 ± 4.6% (n = 3) at 100 μL/min to 84 ± 13.3% (n = 3) at 1000 μL/ min. The standard deviation for recovery rates was significantly increased at flow rates greater than 500 μL/min, while the number of leukocytes remained constant at various flow rates, with a high purity of less than 50 cells at all flow rates tested. In conclusion, a change in the flow rate did not have a significant effect on improvement of purity. In fact, most leukocytes were already removed by the selective sedimentation assay, prior to introduction into the microfilter and, hence, the

for the purpose of enumeration, there was no significant problem to identify marker protein (CK) used to designate a filtered cell as a tumor cell. To confirm the relationship of centrifugal forces to recovery rates of MCF7 cells in tumor cell-spiked blood samples and removal rates of leukocytes by selective sedimentation, g forces in the range of 10−800g were applied to the tumor cell-spiked blood samples. Centrifugation time and filter slot width were fixed at 2 min and 10 μm, respectively. The control samples were not treated with selective sedimentation. Recovery rate was defined as percentage of recovered cancer cells, found optically on the filter, divided by the total number of cancer cells input. As shown in Figure 3A, the recovery rates were very

Figure 3. Recovery rates at different g forces and fluorescence images of tumor cells (MCF-7) captured on the microfilter. The average number of input cell is 101 (n = 5, ±10). (A) Recovery rate (shown as gray bars) of MCF-7 cells and the number of leukocytes (shown as solid line) captured on the slot microfilter at different g forces ranging from 10g to 800g at a fixed centrifugation time of 2 min. Error bars show standard deviations for each condition (n = 3). (B) Three-color immunofluorescence staining method based on DAPI nuclear staining, cytokeratins 7/8, and CD45, which was employed to identify and enumerate MCF-7 cells and leukocytes, nonspecifically trapped on the slot microfilter.

good when the g force applied was in the range of 50−100g and the number of leukocytes was less than 100 cells. However, poor recovery rates were observed in cases where the centrifugal force was less than 20g or greater than 800g. Furthermore, the number of leukocytes, optically identified in the filter, dramatically increased at g forces greater than 200g. As mentioned before, tumor cells were identified by DAPIpositive, CK-positive, and CD45-negative expression, when visualized by fluorescence microscopy. However, in some cases, a large number of triple positive cells (DAPI-positive, CKpositive, and CD45-positive) were observed at g forces greater than 200g. Numerous recent studies reported triple positive cells from healthy volunteers and cancer patients in the CTC fraction obtained by the anti-EpCAM based CellSearch system or the size-based isolation method, such as ScreenCell filtration and the parylene membrane filter-based portable microdevice. 7404

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CTC capturing platform for small-sized CTCs, small cell lung cancer DMS-79 cells, which express EpCAM,37 were used as a model for small-sized CTCs. As shown in Figure 5A, control DMS-79 cells and DMS-79 cells with attached SDABs show a distribution in diameter from 7 to 19 μm (n = 100, mean; 10 μm diameter, ±1.8) and from 11 to 24 μm (n = 100, mean; 15 μm diameter, ±2.2), respectively. The recovery rates for MCF7 and DMS-79 cells were compared at filter slot widths of 10, 12, and 14 μm. Unlike MCF-7 cells, which greatly overexpress EpCAM, to enhance the binding efficiency between DMS-79 cells and SDABs, plasma was removed from whole blood, prior to incubation with antibody-conjugated beads. As shown in Figure 5B, size amplified DMS-79 cells showed recovery rates of 89 ± 8% (n = 3) at a slot width of 10 μm. As the slot width was increased from 10 to 14 μm, the recovery rate for DMS-79 cells decreased significantly to 49 ± 6% (n = 3), whereas the recovery rate for MCF-7 cells remained constant at approximately 100 ± 5%. The number of leukocytes detected on the microfilter did not exceed 50 cells at the most narrow slot width, where the highest number of leukocytes is expected. Many groups who use size-based direct filtration for CTC isolation and/or detection have adopted a filter pore size of 5− 9 μm in diameter. However, the results from these efforts tend to be less pure, as the small-sized filter pores recovered not only CTCs but also approximately 103−104 relatively large-sized leukocytes.14,27,38 Thus, filter-based CTC detection methods have a trade-off relationship between recovery rates and sample purity. In contrast, our platform showed that small CTCs, as well as large CTCs, can be efficiently recovered and detected in a microfilter (slot width of 10 μm) with high purity because selective sedimentation, prior to filtration, had already removed a great number of leukocytes. Assessment of Sensitivity in Recovery Rates. To validate the efficiency of our CTC-capture platform for cancer cells in blood, a series of blind spiking studies were performed using MCF-7 breast cancer cells. A series of MCF-7 cell-spiked blood samples were prepared by introducing 5 (±1.6), 10 (±3.3), 25 (±4.4), 50 (±6.3), or 100 (±8.5) MCF-7 cells to 1 mL of whole blood. The number of input cells was controlled as before, by serial dilution and verification of this dilution

Figure 4. Effect of flow rate on MCF-7 cell recovery rate (gray bars) and the number of captured leukocytes (■). Error bars show the standard deviations at each condition.

washing effect was marginal. It could be argued that purity is determined by selective sedimentation and does not depend on microfluidic conditions. Therefore, flow rate should not be determined by the level of CTC purity. On the other hand, the effect of flow rates on the recovery rate could be seen, as a greater flow rate resulted in larger variations in recover rate, as evidenced by their standard deviations. Therefore, the optimal flow rate was determined to be 100 μL/min, since this flow rate maximizes the recovery rate, while minimizing variation within the recovery rates. Relationship of Recovery Rate between Size of CTCs and Filter Slot Width. To evaluate the recovery rate of our

Figure 6. Sensitivity of the present CTC detection method at various tumor cell concentrations. A known number of MCF-7 cells ranging from 5 to 100 were spiked into 1 mL of whole blood and followed by an isolation procedure. The plot was obtained from an average of three separate repeated experiments. The dot line represents the spiked cell number, and the solid line represents the recovered number of cells. Error bars show standard deviations at each condition.

Figure 5. (A) Size distribution of original DMS-79 cells (black line) and DMS-79 cells with SDABs (red line). (B) Recovery rates of tumor cells, as a function of slot width of the microfilter. MCF-7 and DMS-79 cells were spiked into whole blood at 100 cells/mL. Error bars show standard deviations at each condition. 7405

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through five independent manual counts through visualization with an optical microscope. Assessment of sensitivity showed that our platform was capable of detecting MCF-7 cancer cells at concentrations as low as 5 tumor cells per 1 mL of blood. The recovery rate was found to be the 99 ± 4% (n = 15) when 5−100 MCF-7 cancer cells were present in 1 mL of blood. As shown in Figure 6, a linear regression model showed good correlation between the number of observed cells and the number of expected cells, i.e., the number of input cells (adjusted R2 = 0.987, P < 0.05).

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CONCLUSIONS Currently available methods for isolation and/or detection of CTCs do not meet the requirements necessary (high recovery rate and purity) to enable highly sensitive downstream analysis, such as molecular characterization, to further study the biological characteristics of CTCs, which may be important in improving our understanding of metastasis and disease progression. In this work, we have demonstrated a highly efficient assay for detection of CTCs by selective sedimentation and subsequent filtration. To overcome the similar size and density between CTCs and leukocytes, antibody-immobilized size-density amplification microbeads (SDABs) were used to maximize discriminating physical features (size and density) between CTC-conjugated SDABs (CTC-SDABs) and leukocytes. Furthermore, we found that selective sedimentation of CTCs minimized the number of leukocytes that resulted in the final isolated CTC sample, offering a distinct advantage over most previously reported discrimination approaches. The recovery rate and purity of our platform was on par or better than currently available methods for isolating CTCs. The methods were also reproducible, showing good sensitivity with both a different number of cells present in a blood sample and differently sized cells. With further development, the techniques highlighted in this article will allow for efficient and pure isolation of CTCs from blood, allowing for improved understanding of the biological characteristics of CTCs and metastasis and may ultimately play an important role in patient treatment.

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ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Jong-Myeon Park: e-mail, [email protected]; phone, +82 31 280 1864; fax, +82 31 280 6816. Won-Yong Lee: e-mail, [email protected]; phone, +82 2 2123 2649; fax, +82 2 364 7050. Notes

The authors declare no competing financial interest.

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REFERENCES

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