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Quantitative Analysis of Drug-Induced Complement-Mediated Cytotoxic Effect on Single Tumor Cells Using Atomic Force Microscopy and Fluorescence Microscopy Mi Li, Lianqing Liu*, Member, IEEE, Ning Xi*, Fellow, IEEE, Yuechao Wang, Xiubin Xiao, and Weijing Zhang

Abstract—In the antibody-based targeted therapies of B-cell lymphomas, complement-mediated cytotoxicity (CMC) is an important mechanism. CMC is activated after the binding of drugs (monoclonal antibodies) to tumor cells. The activation of CMC ultimately leads to the lysis of tumor cells. However, it remains poorly understood how CMC alters the morphology and mechanics of single tumor cells at the nanoscale. In recent years, nanoscopic observations of cellular behaviors with the use of atomic force microscopy (AFM) have contributed much to the field of cell biology. In this work, by combining AFM with fluorescence microscopy, the detailed changes in cellular ultra-microstructures and mechanical properties during the process of CMC were quantitatively investigated on single tumor cells. AFM imaging distinctly showed that the CMC effect could lead to the formation of nano holes on the tumor cells. Quantitative analysis of AFM images indicated that cell surface became lower and rougher after the CMC process. The cellular mechanics measurements showed that during the process of CMC cells firstly softened and finally stiffened, which was validated by dynamically monitoring the mechanical changes of single living cells during CMC. The experimental results provide novel insights into the antibody-dependent CMC. Index Terms—Antibody, atomic force microscopy, cell, complement- mediated cytotoxicity, lymphoma, mechanical properties.

Manuscript received March 20, 2014; revised July 29, 2014; accepted November 11, 2014. Date of publication November 20, 2014; date of current version February 27, 2015. This work was supported in part by the National Natural Science Foundation of China (61175103, 61327014 and 61433017), the Research Fund of the State Key Laboratory of Robotics (No. 2014-Z07) and CAS FEA International Partnership Program for Creative Research Teams . Asterisk indicates corresponding authors. M. Li is with the State Key Laboratory of Robotics, Shenyang Institute of Automation, Chinese Academy of Sciences, Shenyang 110016, China, and also with the University of Chinese Academy of Sciences, Beijing 100049, China (e-mail: [email protected]). *L. Liu is with the State Key Laboratory of Robotics, Shenyang Institute of Automation, Chinese Academy of Sciences, Shenyang 110016, China (e-mail: [email protected]). *N. Xi is with the State Key Laboratory of Robotics, Shenyang Institute of Automation, Chinese Academy of Sciences, Shenyang 110016, China, and also with the Department of Mechanical and Biomedical Engineering, City University of Hong Kong, Hong Kong, China (e-mail: [email protected]). Y. Wang is with the State Key Laboratory of Robotics, Shenyang Institute of Automation, Chinese Academy of Sciences, Shenyang 110016, China (e-mail: [email protected]). X. Xiao and W. Zhang are with the Department of Lymphoma, Affiliated Hospital of Military Medical Academy of Sciences, Beijing 100071, China (e-mail: [email protected]; [email protected]). Color versions of one or more of the figures in this paper are available online at http://ieeexplore.ieee.org. Digital Object Identifier 10.1109/TNB.2014.2370759

I. INTRODUCTION

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N RECENT years, monoclonal antibody (mAb)-based targeted therapy has rapidly developed into an important therapeutic strategy in the clinic for treating various diseases, e.g., cancer [1], cardiovascular disease [2], rheumatoid arthritis [3], autoimmunity, and inflammatory disease [4]. Rituximab, approved by U.S. Food and Drug Administration (FDA) in 1997, is the first mAb for treating B-cell lymphomas. Rituximab has achieved unprecedented success in the clinical practice. Rituximab therapy is now the standard care in the treatment of both aggressive (in combination with chemotherapy) and indolent (single agent or in combination with chemotherapy) B-cell lymphomas [5]. The explanation for this success appears to lie with the inherent properties of its target, CD20, which is highly expressed on B cell lymphomas and allows rituximab to recruit immune cells to attack tumor cells with unusual efficiency [6]. The in vitro mechanisms by which rituximab kills the tumor cells have been clarified for years: direct signaling, complement-mediated cytotoxicity (CMC), and antibody-dependent cellular cytotoxicity (ADCC) all appear to play a role in rituximab efficacy [7]. For CMC, the binding of rituximab to the CD20 antigen on the tumor cells can activate C1 complex of complement and subsequently triggers the classical complement pathway. The activation of complement system can lead to the generation of membrane attack complex (MAC) on the cell surface which can lead to the cell lysis. Though the signaling pathway of the CMC effect has been elucidated, it remains poorly understood how CMC alters the tumor cells at the nanoscale. Traditional methods to investigate CMC effect are based on the optical microscopy [8], [9]. Due to the diffraction limit of 200 nm, optical microscopy can not reveal the nanoscopic cellular behaviors. Besides, the results of traditional ensemble experiments reflect the averaged behaviors of large populations of cells, masking the behaviors of individual cells. Evidence has proved that cell heterogeneity occurs even for the monoclonal cells that have been cultured under identical conditions [10]. Investigating the biochemical and biophysical behaviors at the single-cell level can bring breakthrough in uncovering functional diversity between cells and in deciphering cell states and circuits [11], which can improve our understanding of diseases and promote the development of new drugs.

1536-1241 © 2014 IEEE. Personal use is permitted, but republication/redistribution requires IEEE permission. See http://www.ieee.org/publications_standards/publications/rights/index.html for more information.

LI et al.: QUANTITATIVE ANALYSIS OF DRUG-INDUCED COMPLEMENT-MEDIATED CYTOTOXIC EFFECT ON SINGLE TUMOR CELLS

In recent years, nanoscopic observations of cellular behaviors with the used of atomic force microscopy (AFM) have produced a lot of new knowledge to cell biologist to unravel the fundamental mechanisms guiding cellular processes [12]. AFM can dynamically probe the nanostructures of single cells in aqueous conditions, e.g., dynamics of the spore germination of single microbial cells [13], [14], organization of cytoskeletons [15], [16], and drug-induced changes of cellular ultra-microstructures [17], [18]. Besides nanoscopic imaging, AFM can measure the cell mechanical properties via indenting technique [19]. In the force curve mode, AFM tip first approaches and then retracts from the cell surface. During the approach-retract cycle, force curves that reflect the cell mechanical properties are recorded. Cell mechanical properties play an important role for the accurate performing of cellular physiological functions. The deviations of cell mechanical properties are often associated with pathological changes (e.g., occurrence of diseases) [20]. AFM single-cell indenting researches have indicated that cancer cells are significantly softer than normal cells [21], and cell mechanical properties can reflect the aggressive ability of the different types of cancer cells [22], [23]. Hence cell mechanical properties have been considered as an effective biomarker for indicating the cell activities. Investigating the cell mechanical properties of single cells can provide better understanding about the pathological processes of diseases. Here, with the use of AFM and fluorescence microscopy, we quantitatively investigated the nanoscale changes of cellular ultra-microstructures and mechanical properties on single lymphoma cells during the process of CMC. II. MATERIALS AND METHODS A. Cell Lines and Reagents Human B-cell Burkitt's lymphoma Raji cell line was used for the study. Raji cells were obtained from Affiliated Hospital of Military Medical Academy of Sciences (Beijing, China). Clinical-used rituximab solution (10 mg/mL) was also obtained from Affiliated Hospital of Military Medical Academy of Sciences (Beijing, China). Native human serum was purchased from Ruizekang Technology Co., Ltd (Beijing, China). Propidium iodide (PI) dye was purchased from Ruitaibio Technology Co., Ltd (Beijing, China). Raji cells were cultured in RPMI-1640 medium (Thermo Scientific Hyclone, Logan, UT, USA) containing 10% fetal bovine serum and 1% penicillin-streptomycin at 37 ( ). 24-well sterile tissue culture plates were purchased from Jet Bio-Filtration Products Co., Ltd (Guangzhou, China). Phosphate buffered saline (PBS) were purchased from Thermo Scientific Hyclone Co., Ltd (Logan, UT, USA). B. Fluorescence Microscopy Four wells of the 24-well plates were used for CMC fluorescence experiments. For simplicity, the four wells of the 24-well plate were named A, B, C, and D, respectively. 1) All of the four wells were added with 1 mL Raji cell suspension. 2) 1 mL RPMI-1640 medium was added to well A, 1 mL human serum was added to well B, were added to well C, were added to well D. Well A, B, C were the control groups. Well D was the CMC group. 3) Put the 24-well plate into the cell

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incubator and incubated for 2 h at 37 ( ). 4) After the incubation, the solution in the four wells were transferred to four centrifuge tubes respectively and centrifuged at 1000 rpm for 5 min. 5) After remove the supernatant of the four centrifuge tubes, 250 PI solution was added to the four centrifuge tubes and incubated for 5 min. 6) After the incubation, cells of the four centrifuge tubes were dropped onto the glass slides which were coated with poly-L-lysine. 7) Put the glass slides onto the stage of an inverted fluorescence microscope (Ti, Nikon, Tokyo, Japan) for fluorescence observation. Green light was used for excitation. For convincing comparison, the fluorescence imaging parameters (exposure time, gain) were the same for the cells from the four wells. According to the previous literatures, for CMC experiments, the stimulation concentration of rituximab was , the amount of human serum was and the co-incubation time was about 2 h [8], [9]. , Hence the concentration of rituximab used here was 20 the ratio of cell suspension to human serum was 1:1, and the co-incubation time was 2 h. C. Atomic Force Microscopy AFM experiments were performed using a Bioscope Catalyst AFM (Bruker, Santa Barbara, CA, USA) which was set on an inverted fluorescence microscope (Ti, Nikon, Tokyo, Japan). The type of the probe was MLCT (Bruker, Santa Barbara, CA, USA) and the cantilever with nominal spring constant 0.01 N/m was used. The exact spring constant was calibrated by thermal noise method [24]. For AFM imaging, cells of the four wells from 24-well tissue culture plates after 2 h incubation were dropped onto the glass slides which were coated by poly-L-lysine and then chemically fixed by 4% paraformaldehyde for 30 min. The poly-L-lysine is positively charged and cells are negatively charged. Hence the poly-L-lysine-coated glass slides can adsorb the cells via electrostatic adsorption. AFM imaging was performed in PBS and in air. The imaging mode was contact mode. The scanning rate was 0.5 Hz. The scanning line was 256 and the sampling point for each scanning line was 256. Both height image and deflection image were recorded. First large-size scanning was performed to obtain the AFM images of ensemble cells. Then small-size scanning was performed to obtain the AFM images of the local areas on single cells by modulating the parameters (x offset, y offset, scanning size) of the AFM manipulation software which can automatically controlled the tip move to the surface of single cells. For each of the four situations (three control groups and one CMC group), 15 cells were randomly selected to obtain AFM images for quantitative analysis. In order to analyze the cell mechanical properties during the process of CMC, force curves were obtained on Raji cells before and after the treatment of rituximab and human serum. The process of AFM indenting experiments was the following. 1) Raji cells (before or after the treatment of rituximab and human serum) were PI-stained and then dropped onto the poly-L-lysine-coated glass slides. 2) The glass slides were then put in a Petri dish containing PBS and the Petri dish was placed onto the AFM stage. 3) After setting the parameters of AFM manipulation software, the AFM tip was controlled to enter into the liquid in the Petri dish. 4) Force curves were obtained on the bare area of the substrate to calibrate the deflection sensitivity

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of the cantilever and then thermal tune method was used to calibrate the spring constant of the cantilever. 5) Under the guidance of optical/fluorescence microscopy, AFM tip was moved to the Raji cells to obtain force curves on the central area of the cells. For the cells before the treatment (cells did not exhibit fluorescence), six cells were randomly selected to obtain force curves. For the cells after the treatment (that is, cells were incubated with rituximab and human serum for 2 h), some cells exhibited fluorescence which indicated that these cells were with CMC effect, and some cells did not exhibit fluorescence which indicated that these cells were without CMC effect. For each situations (with CMC effect and without CMC effect after the treatment), six cells were randomly selected to obtain force curves. Force curves were obtained on different points on the cell surface at the same loading rate (4 ). The maximum loading force was 10 nN. In order to monitor the mechanical changes of single tumor cells during the process of CMC, successive force curves were obtained on four cells. The total measurement time was 150 min. About 20 force curves were obtained on the central area of the cell each 10 min. At the time of 60 min, rituximab and human serum was added to activate the CMC effect.

Fig. 1. PI staining of Raji cells from control groups. (A), B), (C) were the bright field images. (D), (E), (F) were the corresponding fluorescence images. (A), (D) were from the Raji cells after 2 h of incubation in 1640 medium. (B), (E) were from the Raji cells after 2 h of incubation in 1640 medium containing 50% human serum. (C), (F) were from the Raji cells after 2 h of incubation in 1640 medium containing rituximab.

D. Data Analysis For the conical tips used in the study, the Young's modulus of cells was computed by applying Sneddon-modified Hertz model [19]: (1) where is the Poisson ratio of cell (often 0.5), is the indentation depth, is the half-opening angle of the AFM tip, is the cell Young's modulus, and is the applied loading force. The original force curves were exported as text files which were then imported by using Matlab (Ver.7.6.0) software for data processing and analysis.

Fig. 2. PI staining of Raji cells after 2 h of incubation with human serum and rituximab. (A), (B) were bright field image (A) and corresponding fluorescence image (B). (C), (D) were the higher-resolution bright field image (C) and corresponding fluorescence image (D).

III. RESULTS AND DISCUSSION We firstly examined the CMC effect on Raji cells by fluorescence experiments. PI dye is a commonly used agent for discriminating dead and viable cells [8], [9]. PI molecules can infiltrate into the cells which have damaged plasma membranes, but cannot infiltrate into the cells which have intact plasma membranes. Hence we used PI dye to detect the cell lysis in this study. Fig. 1 shows the PI staining results of Raji cells from three control groups. Three control groups were designed, including 1640 medium, 1640 medium containing 50% human serum, 1640 medium containing rituximab. Fig. 1(A-C) were the bright field images and Fig. 1(D-F) were the corresponding fluorescence images. Fig. 1(A, D) were the results of Raji cells cultured in 1640 medium. Fig. 1(B, E) were the results of Raji cells cultured in 1640 medium containing 50% human serum. Fig. 1(C, F) were the results of Raji cells cultured in 1640 medium containing rituximab. We can see that the cells from the three control groups did not exhibit fluorescence. The only one cell that exhibited fluorescence [Fig. 1(D)] may due to the mechanical damage during the experimental procedure, such as centrifugation and micropipette stirring. Rituximab can bind the CD20 antigen on the surface of Raji cells which can then activate the

classical complement pathway. Complement is a complex proteolytic cascade comprised of over 30 proteins that act to lyse target cells through assembly of MAC [25]. The activation of CMC on Raji cells requires the existence of rituximab and complement. While in the cases of the three control groups, this requirement was not met and thus CMC effect was not activated. Hence cells cultured in the control groups were viable and did not exhibit fluorescence. Rituximab can directly induce the apoptosis of Raji cells after binding to the CD20 antigen, but this requires more than 24 h [8], [26]. Here, cells were cultured only for 2 h and thus the apoptosis directly induced by rituximab was not observed. Fig. 2 shows the PI staining results of Raji cells cultured in 1640 medium containing human serum and rituximab for 2 h. In this case, the requirement of CMC was met. Fig. 2(A), (B) were the bright field image and corresponding fluorescence image. Fig. 2(C), (D) were the higher-resolution bright field image and corresponding fluorescence image. There were many cells which exhibited fluorescence. Human serum contains complement for CMC. When Raji cells were incubated with human serum and rituximab, ritximab bound to the CD20 antigen on

LI et al.: QUANTITATIVE ANALYSIS OF DRUG-INDUCED COMPLEMENT-MEDIATED CYTOTOXIC EFFECT ON SINGLE TUMOR CELLS

Fig. 3. AFM images of Raji cells from three control groups. (A)-(D) were the AFM deflection images of Raji cells cultured in 1640 medium. (E)-(H) were the AFM deflection images of Raji cells cultured in 1640 medium containing 50% human serum. (I)-(L) were the AFM deflection images of Raji cells cultured in 1640 medium containing rituximab. For each situation (1640 medium, 1640 medium containing 50% human serum, 1640 medium containing rituximab), representative AFM images of cells were shown. (A), (C), (E), (G), (I), (K) were the AFM images of whole cells. (B), (D), (F), (H), (J), (L) were the AFM images of local areas denoted by the squares in (A), (C), (E), (G), (I), (K).

the surface of Raji cells and then the Fc domains of rituximab could bind the complement to trigger the CMC effect whose final product was the MAC in the cell membrane. MAC creates 10 nm pores in the cell membrane that facilitate free passage of water and solutes into and out of the cell [27]. PI molecules can infiltrate into the cells via the MAC pores to stain the cell nucleus and thus cells exhibited bright fluorescence. From the bright field image, we can see that viable cells [denoted by the white arrows in Fig. 2(C)] were round and bright, while the lysis cells [denoted by the blue arrows in Fig. 2(C)] were irregular and dark. The special optical traits of lysis cells are caused by the CMC effect. After the formation of MAC pores in the cell membrane, small molecules (e.g., water, ion) entered into the cell freely, which caused that cell swelled and eventually burst [28]. The results of Figs. 1 and 2 qualitatively showed that the incubation of Raji cells with human serum and rituximab can activate the CMC effect. However, due to the 200 nm resolution limit of optical microscopy, it cannot resolve the detailed situations of CMC effect at the nanoscale. We then used AFM to image the ultra-microstructures of Raji cells from three control groups and from CMC group to reveal the detailed morphological changes of Raji cells during the process of CMC. Fig. 3 shows the representative AFM images of Raji cells from the three control groups. For each of the three control groups, we obtained AFM images on 15 cells and only the AFM images of two cells were shown in Fig. 3. Fig. 3(A-D) corresponded to the Raji cells cultured in 1640 medium. From the AFM images of whole cells [Fig. 3(A), (C)], we can see that the cells exhibited a round shape. From the AFM images of local areas [Fig. 3(B), (D)], we can see that the cell surface was intact and rough. The roughness topography of Raji cells may be related to the structure of B lymphocyte. Raji cell originates from the malignant B lymphocyte. We know microvilli are abundant on the surface of B lymphocytes [29]. The existence of microvilli on the cell surface facilitates B lymphocytes to perform some functions, such as cell adhesion [30]. The microvilli can thus cause the roughness topography of B

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Fig. 4. AFM images of Raji cells incubated with human serum and rituximab. (A)-(F) corresponded to the initial stage of CMC where holes began forming on the cell surface. (A) AFM deflection image of large size. AFM deflection image (B) of a cell (denoted by the blue square I in A) and local area (C). AFM deflection image (D) of another cell (denoted by the blue square II in A). AFM height image (E) and deflection image (F) of local area denoted by the square in (D). (G)-(L) corresponded to the stage of CMC where cytoplasm escaped. Deflection image (G) of a cell [denoted by the red square I in (A)]. AFM height image (H) and deflection image (I) of local area denoted by the square in G. Deflection image (J) of another cell [denoted by the red square II in (A)] and local area (K). Height image (L) of local area. (M)-(R) corresponded to the stage of CMC where cells had collapsed. AFM height images (M), (O), (Q) and deflection images (N), (P), (R).

lymphocytes. Fig. 3(E)–(H) corresponded to the Raji cells cultured in 1640 medium containing 50% human serum. From the AFM images of whole cells [Fig. 3(E), (G)], we can see that the cells were not plump as the cells cultured in 1640 medium. From the AFM images of local areas [Fig. 3(F), (H)], we can see that the cell surface was still intact but looked smoother than the cells cultured in 1640 medium. This may because of the influence of human serum. The addition of human serum into the cell culture medium changed the growth environment of Raji cells, which may cause the morphological changes of Raji cells. Fig. 3(I)–(L) corresponded to the Raji cells cultured in 1640 medium containing rituximab. From the AFM images of whole cells [Fig. 3(I), (K)], we cannot see distinct difference compared to the cells cultured in 1640 medium. The cell surface was still intact. While from the AFM images of local areas [Fig. 3(J), (L)], we can see that some protrusions occurred on the cell surface (denoted by the arrows). The binding of rituximab to the CD20 antigen on the Raji cells can trigger the apoptotic signaling pathway, which may cause the morphological changes of cell surface, as reported previously [31]. Fig. 4 shows the representative AFM images of Raji cells cultured in 1640 medium containing human serum and rituximab. In this case, CMC effect can be activated, which can lyse the cells via generating MAC pores on the cell surface. Fig. 4(A)-(F) corresponded to the initial stage of CMC effect where holes began forming on the cell surface. We first performed large size scanning to visualize many cells and then perform small size scanning on interesting cells. From the AFM images, we can clearly see that some holes occurred on the cell surface [denoted by the arrows in Fig. 4(C), (E)]. From the AFM images, we can measure the diameter of holes by applying the AFM offline software (Nanoscope Analysis, Bruker, Santa Barbara, CA, USA). The size of holes was summarized in Table I. From Table I(A), we can see that the size of the holes at the initial stage was about 150~500 nm. At this stage, we did not see the outflow of cytoplasm. Fig. 4(G)-(L) corresponded to the

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TABLE I SIZES OF HOLES ON THE SURFACE OF RAJI CELLS AT DIFFERENT STAGES OF CMC EFFECT. A. HOLE FORMING. B. CYTOPLASM ESCAPE. C. CELL MEMBRANE DEBRIS

stage where holes on the cell surface became larger and cytoplasm began escaping from the cell. At this stage, many more holes formed [denoted by the arrows in Fig. 4(H), (K), (L)]. From Table I(B), we can see that the size of the holes at this stage was about 400~700 nm, significantly larger than that at the initial stage. Besides, we can distinctly see that some portion of cytoplasm had escaped from the cell [denoted by the red arrows in Fig. 4(G), (J)]. Fig. 4(M)-(R) corresponded to the CMC stage where cytoplasm had totally escaped outside the cell and only cell membrane debris left. Fig. 4(M), (O), (Q) were the AFM height images and Fig. 4(N), (P), (R) were the corresponding deflection images. At this stage, the holes continued to became larger. From Table I(C), we can see that the size of holes were in the range of 600~2000 nm. Besides, at this stage, because of the outflow of cytoplasm, the cells collapsed and we can only see the cell membrane debris. Compared the AFM imaging results of Figs. 3 and 4, we can see that Raji cells had many holes on the cell surface after 2 h incubation with human serum and rituximab. For Raji cells from the control groups where the CMC effect did not occur, there were no holes on the cell surface. We have statistically analyzed the cells that had holes from CMC group and from control groups. The results showed that more than 40% of cells generated holes when incubated with human serum and rituximab, while less than 1% of cells exhibited holes from the control groups. These results demonstrated that the observed cellular ultra-structures (holes, cytoplasm) in Fig. 4 were related to the CMC effect. There are three complement activation pathways: classical pathway, lectin pathway and alternative pathway [32]. All of the three pathways have the same terminal product, MAC, which can cause the cell lysis. The binding of rituximab to the CD20 antigen on the surface of Raji cells can induce the formation of MAC via classical pathway. MAC is composed of complement proteins C5b, C6, C7, C8, and C9 [33]. Electron microscopy images of MACs isolated from erythrocytes have indicated that MAC exhibits a hollow cylinder structure [34].

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From the optical images (Fig. 2), we cannot see the detailed situations on the cell surface during the process of CMC due to the essential 200 nm resolution limit. AFM can resolve the changes of ultra-microstructures (such as holes, cytoplasm) of single cells during the process of CMC, as shown in Fig. 4. The occurrence of holes was due to the CMC effect on Raji cells. The sizes of holes on the cell membrane observed here (150~2000 nm) were evidently larger than the size of MAC ( 10 nm). When the MAC assembled in the cell membrane, the initial size of MAC pore was about 10 nm. Then due to the shear effect of the flushing fluid (water molecules can freely enter into the cell membrane through the MAC pores), the pores can reasonably became larger. In fact, researchers have observed large sizes of pore-like structures (30~60 nm) [33]. Hence the holes on the cell surface observed here should be closely related to the MAC pores. As the CMC effect proceeded, the size of holes became larger, as shown in Fig. 4 and Table I. Researches of MAC have shown that besides directly inducing cell lysis, MAC can also influence the inflammation, immune cell chemotaxis, phagocytosis, and up-regulation of antibody production [35]. However, the dynamics of how MAC alters the cells is still poorly understood. Here we used AFM to image the dynamic changes of cellular ultra-microstructures during the process of CMC, directly revealing the distinct cellular morphological changes at the different stages of CMC effect. Though MAC pore structures can be observed by electron microscopy [36], the flaw of electron microscopy is that the sample preparation is complex and it requires the samples fixed and dried. AFM has easy sample preparation and can work in liquids which allow us to investigate the biological activities on living cells, providing novel knowledge for understanding the cell biology. From the AFM height images, we can extract the morphological information of cells, such as cell height, and surface roughness [37]. For each of the four situations (three control groups and one CMC group), 15 cells were selected. For each cell, AFM images of whole cell and local area (1 1 ) were obtained. Due to the fact that the surface roughness computed from the AFM images can be influenced by the curvature of the cell, we obtained small size AFM images (1 1 ) on the cell surface to compute the roughness in order to eliminate the disturbance of cell curvature. The AFM images of local areas were obtained at the central area on the cell surface. The cell height was calculated from the AFM image of whole cell and the cell roughness was calculated from the AFM image of local area. Fig. 5 shows extracting the cell height and surface roughness of one cell. Fig. 5(A) was a large-size scan AFM image. From the line profile [Fig. 5(B)], we can calculate the cell height ( 5.8 ). Fig. 5(C) was the 1 1 local area image obtained on the central area of the cell [denoted by the red square in Fig. 5(A)]. The corresponding roughness curve of Fig. 5(C) was shown in Fig. 5(D), indicating that the roughness of Fig. 5(C) was 11.83 nm. For all cells, the process of calculating the cell height and surface roughness was the same as described in Fig. 5. Information of cell height and surface roughness for the four situations were summarized in Table II. From Table II(A), (B), (C), we cannot see the significant difference of cell height and cell roughness between the three control groups. The cell height of cells from

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TABLE II HEIGHT AND SURFACE ROUGHNESS OF RAJI CELLS CULTURED IN CONTROL GROUPS AND CMC GROUP. (A) 1640 MEDIUM (CONTROL GROUP). (B) 1640 MEDIUM CONTAINING 50% HUMAN SERUM (CONTROL GROUP). (C) 1640 MEDIUM CONTAINING RITUXIMAB (CONTROL GROUP). D. 1640 MEDIUM CONTAINING 50% HUMAN SERUM AND RITUXIMAB (CMC GROUP)

Fig. 5. Extracting the cell height and surface roughness from AFM height images. (A) AFM height image of large-size scan. (B) Line profile of one cell taken in the height image (A) along the red line. (C) AFM height image of 1 1 local area of one cell (denoted by the red square in A). (D) Roughness curve computed from the AFM height image (C).

control groups was in the range of 3~6 and cell surface ) was in the range of 5~30 nm. While for roughness (1 1 the cells from the CMC group [Table II(D)], cell height was in the range of 1~5 and cell surface roughness (1 1 ) was in the range of 10~40 nm. The results in Table II showed that cell height decreased and cell surface roughness increased during CMC effect. The formation of MAC pores and the subsequently formed larger size holes can lead to the outflow of cytoplasm (as observed in Fig. 4), which caused the decrease of cell height. The generated holes on the cell membrane can make cell surface rougher. From AFM high-resolution images of cell topography (Fig. 4), we can visually see the detailed situations occurred at the nanoscale on single cells. From the information extracted from the AFM images (Tables I and II), we can quantitatively characterize the changes of cell surfaces during the biological activities. Plomp et al. [13] have applied AFM to investigate the dynamic structural changes of single germinating bacterial spores, revealing the germination-induced alterations in spore coat architecture and topology as well as the disassembly of outer spore coat rodlet structures. Kirmse [15] have studied the detailed morphology (such as lamellipodia, filopodia) of melanoma cells which were cultured on different substrates (anisotropic collagen matrix and isotropic collagen matrix), and the results showed the distinct cellular morphological difference for different substrates. AFM can quantitatively reveal the nanoscopic situations on single living cells, which is of great significance in clinical medicine. We know traditional methods for evaluating drugs take long time (often several weeks). For AFM, we can immediately monitor the detailed morphological changes of single cells after the stimulation of drug molecules, and the detected morphological changes of cell surface (such as cell surface roughness) may be used for evaluating the drug efficacy. AFM detection is fast. Hence with

AFM we may evaluate the drug efficacies for each patient to examine whether the patients can benefit from the drugs. This is of important significance in the era of personalized medicine. It should be noted that AFM images obtained here were on chemically fixed cells. In the future, we would like to investigate the cellular morphological changes directly on living cells during the process of CMC effect. We know almost all of the current knowledge about cell biology has come from ensemble experiments [38]. More and more evidence have indicated that the results from ensemble experiments cannot reflect the real situation and the incidents of small subpopulations may be masked [39]. Consequently, quantitatively investigating the cellular behaviors at the single-cell level will undoubtedly have great impact on life sciences and medicine. AFM is a powerful analytical tool that can image native samples with nanometer resolution and quantitatively obtain multiple properties of biosystems (including molecules, cells, and tissues), thus offering new possibilities for understanding the mysteries of life [40]. By AFM indenting technique, we can obtain the mechanical properties (e.g., Young's modulus) of single cells. Researches of using AFM to measure the mechanical properties of single cells have shown that cell mechanical properties are a new reliable indicator of cell states. Investigating the cell mechanics will potentially have great impact on improving the diagnosis, treatment and prognosis of critical diseases, such as cancer [41]. In order to understand the CMC effect on Raji cells from the view of cell mechanics, we studied the changes of cellular mechanical properties during the process of CMC. Fig. 6 shows the process

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Fig. 6. Under the guidance of fluorescence, AFM tip was moved to the target cells. (A) Bright field image, (B) fluorescence image, and (C) overlay image before moving AFM tip to the target cell (exhibiting fluorescence). (D) Bright field image, (E) fluorescence image, and (F) overlay image after moving AFM tip to the target cell to obtain force curves. The inset in (F) was the upright optical image of AFM cantilever.

of obtaining force curves on single cells which had CMC effect under the guidance of fluorescence. Raji cells were cultured with human serum and rituximab for 2 h. After the incubation, cells were PI-stained and then dropped onto the glass slides. The glass slides were then placed in a Petri dish in PBS. From the bright field image [Fig. 6(A)], we can see that there were three cells. From the fluorescence image [Fig. 6(B)], we can see that one cell exhibited fluorescence. The fluorescence demonstrated that the cell was with CMC effect. Then the tip was moved to the fluorescence-shining cell to measure its mechanical properties. Because the direction of approaching process of our used AFM was not vertical, we should move the AFM tip to an adequate position and then controlled the tip to automatically approach the cell. The position can be obtained by performing an approaching process. Before approaching, we recorded the original horizontal position of AFM tip. After the approaching, we recorded the tip position at this moment. If the tip was just on the cell, then the original position was the adequate position . If the tip was not on the cell, then we can obtain the difference between and the position of target cell. Then we can calibrate the original position by subtracting the difference and thus obtain the adequate position . Fig. 6(D), (E), (F) were the images after moving the AFM tip onto the fluorescence cell and obtaining force curves on cell surface. From the overlay image [Fig. 6(F)], we can clearly see that the fluorescence cell was beneath the AFM cantilever. In order to confirm the position of AFM tip on the cantilever, we obtained the optical image of AFM tip by using an upright optical microscope (KH7700, HIROX company, Japan), as shown in the inset in Fig. 6(F). From the upright optical image of AFM tip, we can see that the tip was close to the end of the cantilever, which demonstrated that the cells were under the tip. According to the process of moving AFM tip to target cells in Fig. 6, we obtained force curves on single Raji cells for three situations (before treatment, without CMC after treatment, with CMC after treatment). Fig. 7 shows the process of computing the Young's modulus of cells from the obtained force curves for the three situations. Fig. 7(A)-(F) corresponded to the Raji cells cultured in 1640 medium. At this case, due to the lack of CMC effect, Raji cells did not exhibit fluorescence. Under the guidance of optical microscopy, AFM tip was moved to the cells

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Fig. 7. Measuring the mechanical properties of Raji cells for three situations (before treatment, without CMC after treatment, with CMC after treatment). (A)-(F) corresponded to the Raji cells before treatment. (G)-(L) corresponded to the Raji cells after treatment (without CMC). (M)-(R) corresponded to the Raji cells after treatment (with CMC). (A), (B), (C), (G), (H), (I) were the bright field images. (M), (N), (O) were the overlay images of bright field images and fluorescence images. (D), (J), (P) Typical force curves. (E), (K), (Q) Indentation curves. (F), (L), (R) Histogram of Young's modulus computed from the indentation curves in (E), (K), (Q). The black arrow denoted the abrupt peak.

[Fig. 7(A-C)] to obtain force curves. Fig. 7(D) was a representative force curve. The blue curve corresponded to the approach curve and the red curve corresponded to the retract curve. After converting the approach curve into indentation curve according to the contact point, we can obtain many Young's modulus by applying Hertz model and the histogram of Young's modulus was shown in Fig. 7(F). Though Hertz model does not consider electrostatic forces, adhesion or friction between contact surfaces, it is the mostly widely used model in practice [19], [42]. From the Gaussian fitting, we can see that the mean value was 2.9 kPa. After putting the 2.9 kPa into the (1), we can obtain the Hertz fitting indentation curve, as shown in Fig. 7(E). We can see that the experimental indentation curve was consistent with the Hertz fitting indentation curve, proving the effectiveness of Hertz model in characterizing the indenting process between AFM tip and cell. From the results of Fig. 2, we can see that some cells exhibited fluorescence after the incubation with human serum and rituximab and some cells did not exhibit fluorescence. For the cells which did not exhibit fluorescence, we measured their mechanical properties, as shown in Fig. 7(G)-(L). Under the guidance of optical microscopy, AFM tip was moved to the cells [Fig. 7(G)-(I)] to obtain force curves. Fig. 7(K) shows the contrast of experimental indentation curve and Hertz fitting indentation curve. They were also consistent with each other. Fig. 7(M)-(R) corresponded to the Raji cells with CMC effect after incubation in 1640 medium containing 50% human serum and rituximab. Under the guidance of fluorescence [Fig. 7(M)-(O)], AFM tip was moved to the target cells to obtain force curves. Fig. 7(P) was a typical force curve. From the force curve, we can see that there was an abrupt peak in the approach curve (denoted by the black arrow). This peak represented that the tip penetrated the cell membrane [43], [44]. For the cells with CMC effect, MAC pores formed and these pores can become larger as the CMC progressed, as observed in Fig. 4. Due to the existence of these pores on the cell membrane, the tip can easily penetrate the cell membrane from these pores. This caused the abrupt peaks in the approach curves. Fig. 7(R) was the histogram of Young's modulus computed from the indentation curve [Fig. 7(Q)] converted from the approach curve [Fig. 7(P)]. Fig. 7(Q) shows the contrast of experimental indentation curve and Hertz fitting indentation curve.

LI et al.: QUANTITATIVE ANALYSIS OF DRUG-INDUCED COMPLEMENT-MEDIATED CYTOTOXIC EFFECT ON SINGLE TUMOR CELLS

For each cell, about 50 force curves were obtained at the central area on the cell surface. Each force curve was processed according to the procedure described in Fig. 7. For three situations (before treatment, without CMC after treatment, with CMC after treatment), the Young's modulus of Raji cells were computed and the results were shown in Fig. 8. From Fig. 8, we can see that the Young's modulus of Raji cells before the treatment of human serum and rituximab was in the range of 1.5~3 kPa. After the treatment of human serum and rituximab, the Young's modulus of Raji cells without CMC effect was in the range of 0.5~2.5 kPa, and the Young's modulus of Raji cells with CMC effect was in the range of 2~9 kPa. The results in Fig. 8 clearly showed that the Young's modulus of Raji cells during the process of CMC effect decreased firstly and then increased remarkably. For the cells without CMC effect after the treatment of human serum and rituximab were softer than the Raji cells without treatment. This may be caused by the direct apoptotic effect of rituximab. In previous studies, we have shown that the treatment of rituximab could cause the Raji cells become softer [31]. The binding of rituximab to the CD20 antigen on the surface of Raji cells can activate the apoptotic signaling pathway, which could thus induce various changes of cells, such as the changes of mechanical properties. Researches have shown that the apoptotic cells can become softer than their normal counterparts [17], [45]. Hence when culturing Raji cells in 1640 medium containing human serum and rituximab, the apoptotic effect of rituximab can thus cause the decrease of Young's modulus of Raji cells. It should be noted that the direct apoptotic effect of rituximab on cells can be detected by fluorescence after more than 24 h [8], [26]. Here the stimulation time was only 2 h, meaning that cell membrane was still intact (as observed in Fig. 3) and cells did not exhibit fluorescence. For the cells with CMC effect after the treatment, the cell Young's modulus was significantly larger than the cells without CMC effect. This may be related to the CMC effect on Raji cells. The CMC effect can generate MAC pores on the cell membrane and this finally caused the cell lysis. This biological process can reasonably cause the changes of mechanical properties. Lam et al. [46] have investigated the changes of mechanical properties of leukemia cells after the treatment of anti-cancer chemotherapy drugs and the results indicated that dead cells (PI positive) caused by chemotherapy drugs were prominently stiffer than live cells. Here we can see that we obtained similar results: the cells with CMC effect (PI positive) became stiffer than live cells (PI negative). The mechanical properties of cells were dependent on the cytoskeletons [47]. Hence the CMC effect may cause the changes of cytoskeleton. The changes of cytoskeleton may then lead to the increase of cell Young's modulus. It is noticed that some cells (denoted by the black arrow in Fig. 8) exhibited a much larger Young's modulus, about four times larger than the live cells. This may because the tip probed the cell nucleus. The obtained force curves showed that the tip can easily penetrate the membrane of cells with CMC effect [denoted by the black arrow in Fig. 7(P)]. Researchers have shown that the stiffness of cellular (chondrocytes, endothelial cells) was 3~10 times stiffer than cytoplasmic stiffness [48]. A recent study using AFM needle penetration technique showed that the Young's modulus of nucleus from bladder cancer cells was 7~10 kPa and the Young's modulus of cytoplasm from

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Fig. 8. The Young's modulus of Raji cells changed during the process of CMC.

Fig. 9. Dynamic changes of cellular Young's modulus of four Raji cells during the process of CMC by successively recording force curves on the four cells every 10 min. (A) Cell 1, (B) Cell 2, (C) Cell 3, and (D) Cell 4. Rituximab and human serum was added at the time of 60 min.

bladder cancer cells was 4~6 kPa [49]. Here we can see that our results (the Young's modulus of nucleus of lymphoma cells was 4 times larger than that of cytoplasm) were comparable to the previous researches. The results of Fig. 8 were obtained on different cells. The intrinsic heterogeneity between cells may cause the different cellular mechanical responses. In order to observe the mechanical changes of single cells during the process of CMC effect, successive force curves were obtained on the same cells in 150 min. The dynamic changes of four cells during the process of CMC effect were shown in Fig. 9. Before the adding of rituximab and human serum, the cellular Young's modulus basically kept stable in 60 min. For cell 1, the cellular Young's modulus began to decrease 30 min later after the addition of rituximab and human serum. At the time of 150 min, the cellular Young's modulus (2.2 kPa) was significantly smaller than the cellular Young's modulus at the time of 60 min (3.4 kPa). For cell 2, the cellular Young's modulus began to decrease 30 min after the addition of rituximab and human serum and finally increased to 10.3 kPa at the time of 150 min. For cell 3 and cell 4, we can also see that the cellular Young's modulus increased after the addition of rituximab and human serum. Especially, the Young's

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modulus of cell 2 and cell 4 at the time of 150 min (denoted by the black arrows in Fig. 9) was much larger than the Young's modulus before stimulation. This may because that the tip penetrated the cell membrane and probed the cell nuclei. For cell 1, we cannot see the increase of cellular Young's modulus after stimulation. This may due to the cellular heterogeneity. However, on the whole, the cellular Young's modulus first decreased and finally increased after the addition of rituximab and human serum, which was consistent with the results in Fig. 8. The tip radius of the probe used here was about 20 nm and hence the tip measured the local mechanical properties of cells. In order to better represent the cellular mechanic properties, we obtained force curves at many different points on the central area of the cells. Studies have shown that the measured cellular Young's modulus was related to the loading rate [20]. In order to make the obtained data comparative, force curves were obtained at the same loading rate (4 ). Studies have also shown that other factors can influence the measured cellular Young's modulus, including the depth of indentation, the substrate used for cell attachment, and the experimental solution [50], [51]. Here the range of indentation used for Young's modulus calculation was 1000 nm. We can see that in this range the Hertz fitting curve was consistent with the experimental indentation curve, as shown in Fig. 7(E), (K), (Q). In addition, the substrate and the experimental solution was the same for all cells measured in this study. By doing these, we can greatly eliminate the artificial errors. In fact, before the disturbance of rituximab and human serum, there were no significant changes of cellular Young's modulus of tumor cells. Combining Fig. 8 with Fig. 9, it can be concluded that tumor cells firstly softened and finally stiffened during the process of CMC effect. So far, progress in the treatment of cancer has been slow. A key challenge is that cancer therapies are effective only in some patients and oftentimes only for a limited period of time [52]. Cancer is a highly heterogeneous disease that is determined by several factors, such as the genetic alterations in tumor cells, the tumor micro-environments and their interactions with tumor cells, and the cancer cell plasticity [53], [54]. As we enter the era of personalized medicine, identifying patient subpopulations that would benefit from specific targeted therapeutics has been increasingly important in the field of medical oncology [53]. Major advancements in basic science (e.g., the personal genome sequencing [55], new methods in biological informatics [56]) have created the opportunity for significant progress in clinical medicine [57]. However, just knowing the cell genomics is insufficient for completely understanding cancers, because genetically identical cells can have different phenotypes [10]. In recent years, using AFM to investigate the cell behaviors has attracted the attention of researchers from the field of life sciences [40]. AFM has been proved as a powerful tool for characterizing the cellular and molecular physiological activities on single cells. Traditional biochemical experiments based on optical microscopy can not reveal the detailed events at the nanoscale, causing that how various molecules localize, assemble and interact on the surface of living cells is poorly understood [58]. With AFM, we can directly observe the spontaneous or drug-induced real-time changes of cellular ultra-microstructures [13], [14], [18] and even the real-time motion of molecules on the native cell membrane [59], greatly improving our under-

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standing of the dynamics of cellular and molecular activities on live cells. Here we observed the changes of cellular ultra-microstructures during the process of CMC effect and the results distinctly showed that nano holes formed during the process. Recent researches have demonstrated the effectiveness of cell mechanical properties in indicating the cell states. From the results in Figs. 8 and 9, we can see that the Young's modulus of tumor cells changed as the CMC effect progressed, meaning that we can discern the different stages in the CMC process by simply monitoring the cellular mechanical properties after the activation of complement system. The important significance impact is that cell mechanical property is a label-free biomarker and thus avoids the serious flaw of fluorescence staining (fluorescence staining can inevitably alter the natural activities of cells [60]). By investigating the cell mechanical properties during the biological activities, we can thus develop novel methods for diagnosing the diseases and real-timely monitoring the effect of drugs for drug screening. However, the mechanisms that cause the changes of cellular mechanical properties during the CMC effect are not known and further researches are needed. Besides indenting technique, AFM cantilever can be used as a nanomechanical sensor to detect the cell mechanics [61], opening the new possibilities for rapidly detecting the cell resistance to drugs with low concentrations of cells. Overall, AFM provides a powerful platform for cell biology and the further applied of AFM in life sciences will bring many novel breakthroughs which can help us to better understand the mysteries of life. IV. CONCLUSION In this work, we used AFM and fluorescence microscopy to quantitatively study the morphological and mechanical changes during the process of CMC effect on single lymphoma cells. The AFM cellular ultra-microstructue imaging showed that MAC pore-like holes formed during the CMC process and the size of holes became larger as the CMC effect progressed. The quantitative analysis of AFM images showed that cell height decreased and cell surface roughness increased in the CMC process. Both of the AFM cell indenting experiments on different cells and on single cells indicated that, after the treatment of rituximab and human serum, the cellular Young's modulus decreased firstly at the stage where cells were without CMC effect (PI negative), and then the cellular Young's modulus increased largely at the stage where cells were with CMC effect (PI positive). These experimental results can improve our understanding of the CMC effect during the immune therapy of cancers. REFERENCES [1] A. M. Scott, J. D. Wolchok, and L. J. Old, “Antibody therapy of cancer,” Nat. Rev. Cancer, vol. 12, pp. 278–287, 2012. [2] A. L. Catapano and N. Papadopoulos, “The safety of therapeutic monoclonal antibodies: Implications for cardiovascular disease and targeting the PCSK9 pathway,” Atherosclerosis, vol. 228, pp. 18–28, 2013. [3] L. Bossaller and A. Rothe, “Monoclonal antibody treatments for rheumatoid arthritis,” Expert Opin. Biol. Ther., vol. 13, pp. 1257–1272, 2013. [4] A. C. Chan and P. J. Carter, “Therapeutic antibodies for autoimmunity and inflammation,” Nat. Rev. Immunol., vol. 10, pp. 301–316, 2010. [5] D. G. Maloney, “Anti-CD20 antibody therapy for B-cell lymphomas,” N. Engl. J. Med., vol. 366, pp. 2008–2016, 2012. [6] S. A. Beers, C. H. T. Chan, R. R. French, M. S. Cragg, and M. J. Glennie, “CD20 as a target for therapeutic type I and II monoclonal antibodies,” Semin. Hematol., vol. 47, pp. 107–114, 2010.

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Mi Li received the B. S. degree in software engineering in 2006 and M.S. degree in pattern recognition and intelligent systems in 2008 from Huazhong University of Science and Technology, Wuhan, China. He is currently working toward the Ph.D. degree from Shenyang Institute of Automation Chinese Academy of Sciences, Shenyang, China. His research interests include AFM-based imaging and mechanical analysis of biological systems.

Lianqing Liu received the B.S. degree in industry automation from Zhengzhou University, Zhengzhou, China, in 2002 and the Ph.D. degree in pattern recognition and intelligent systems from Shenyang Institute of Automation Chinese Academy of Sciences, Shenyang, China, in 2009. He is currently a Professor with the State Key Laboratory of Robotics, Shenyang Institute of Automation Chinese Academy of Sciences, Shenyang, China. His research interests include micro/nano system, nanodevice fabrication, nanobiotechnology, and biosensors.

Ning Xi received the D.Sc. degree in system science and mathematics from Washington University, St. Louis, MO, USA, in 1993 and the B.S. degree in electrical engineering from Beijing University of Aeronautics and Astronautics, Beijing, China. Currently, he is the John D. Ryder Professor of Electrical and Computer Engineering at Michigan State University, East Lansing, MI, USA, and a Professor of Mechanical and Biomedical Engineering, City University of Hong Kong, Hong Kong, China. His research interests include robotics, manufacturing automation, nanosensors, micro/nano manufacturing, and intelligent control and systems.

Yuechao Wang received the M.S. degree in pattern recognition and intelligent systems from Shenyang Institute of Automation, Chinese Academy of Sciences, Shenyang, China, in 1997 and the Ph.D. degree in mechatronic engineering from Harbin Institute of Technology, Harbin, China, in 1999. Since 1987, he has been with the Shenyang Institute of Automation, Chinese Academy of Sciences, where he is currently a Professor. His current research interests include robot control, multirobot systems, and micro-nano manipulations.

Xiubin Xiao received the M.M. degree in blood in 2002 and D.M. degree in immunology in 2009 from Academy of Military Medical Sciences, Beijing, China. She is currently an Associate Chief Physician with the Department of Lymphoma, Academy of Military Medical Sciences. Her current research interests include diagnosis and treatment of lymphoma.

Weijing Zhang received the B. M. degree in immunology in 1983 from the First Military Medical University, Guangzhou, China, and the M.M. degree in immunology in 1994 from Academy of Military Medical Sciences, Beijing, China. He is currently a Chief Physician with the Department of Lymphoma, Academy of Military Medical Sciences. His current research interests include diagnosis and treatment of lymphoma.