Chemotherapy exposure increases leukemia cell stiffness

Alterations of biophysical properties of blood cells contribute to the pathophysiology of hematologic diseases.^1-3  While chemotherapy-induced cell death has been a mainstay of cancer treatment for decades and is well-studied biochemically, little is known about the mechanical effects chemotherapy may have on leukemia cells. Furthermore, since hyperleukocytosis accompanies some cases of acute leukemia, mechanical changes in leukemia cells due to chemotherapy could significantly alter the overall blood rheology.

In this work, we quantified the effect of standard induction chemotherapy on the stiffness of acute lymphoblastic leukemia (ALL) and acute myeloid leukemia (AML) cells using atomic force microscopy (AFM), a tool for imaging and characterization of materials at the nanometer scale.⁴  The high force sensitivity of AFM and its ability to measure properties of individual cells over long times makes the technique particularly appropriate for measuring dynamic changes in cell stiffness. We find that when exposed to chemotherapy, leukemia cell stiffness increased by nearly 2 orders of magnitude at a rate dependent on the type of chemotherapy employed.

Leukemia cells were isolated via centrifugation from the blood of patients with newly diagnosed acute leukemia noted to have peripheral blast cells. Leukemic cell lines were purchased commercially (ATCC, Manassas, VA). University of California, San Francisco (UCSF) and UC Berkeley institutional review boards approved all experimental procedures. Informed consent was obtained from each human subject, in accordance with the Declaration of Helsinki, before a blood sample was obtained. Dexamethasone and daunorubicin (Sigma-Aldrich, St Louis, MO) are mainstay induction chemotherapeutic agents for ALL and AML, respectively. Accordingly, lymphoid and myeloid leukemic cells were exposed to typical treatment doses of 1 μM dexamethasone and 1 μM daunorubicin respectively.^5,6  Positive staining with either 0.9 μM propidium iodide (PI) or 1 μM Sytox Green (Invitrogen, Carlsbad, CA), markers for loss of cell membrane integrity, indicated cell death or late apoptosis. To identify early apoptosis, we used cresyl violet conjugated to Aspartate-Glutamate-Valine-Aspartate (Immunochemistry Technologies, Bloomington, MN), an indicator for caspase 3 and 7 activity.

Force microscopy measurements were obtained on a modified commercial AFM. Details involving the use of AFM for biological applications, the modifications made to our system, and the analytic methodology used to calculate cell stiffness are given in Document S1 (available on the Blood website; see the Supplemental Materials link at the top of the online article) and were described previously.⁷  Briefly, a Bioscope AFM (Veeco, Santa Barbara, CA) mounted atop an epifluorescence microscope (Carl Zeiss, Thornwood, NY) held the fluid-cell–mounted cantilever (Veeco). Gold-coated silicon nitride cantilevers with a spring constant between 0.009 and 0.019 N/m, calibrated by the thermal noise method,⁸  were used in all experiments. Cells were mechanically immobilized in microfabricated wells⁷  within a 37°C perfusion chamber for AFM indentation. The desired chemotherapeutic agents, along with cell death markers, were then added. The AFM cantilever and nearby cells could be visualized simultaneously by fluorescence microscopy to determine whether the cells were alive, early apoptotic, or late apoptotic/dead.

Using standard photolithographic techniques,⁹  polydimethylsiloxane (PDMS) microfluidic channels were molded from a SU-8 photoresist master on a silicon wafer. Microchannels were designed to geometrically emulate a microvasculature network branching into 5-μm wide by 12-μm tall, capillary-sized channels. ALL cells were incubated with dexamethasone for 6 hours, yielding approximately 10% cell death by trypan blue exclusion, and then passed through the channels driven by a physiologic pressure difference of approximately 1 kPa typically observed across capillary beds.¹⁰  Microchannel obstruction by live and dead cells was visualized with brightfield and epifluorescence microcopy (Carl Zeiss).

All reported cell stiffness values represent the average of 5 consecutive AFM measurements. Stiffness levels in the different cell populations (n ≥ 15 cells unless otherwise specified) were compared with analyses of variance using 2-tailed significance tests. Errors are reported as standard errors of the mean.

In the presence of chemotherapeutic agents, dexamethasone for ALL cells and daunorubicin for AML cells, leukemia cells exhibited a 14- to 91-fold increase in stiffness as they underwent cell death (Figure 1A-C; see Table S1 for a summary of patient data). The average stiffness of both lymphoid and myeloid leukemia cells held at 37°C and exposed to chemotherapy (mean: 4.7 kPa) was significantly higher than the average stiffness of untreated control populations (mean: 0.2 kPa, P < .05, Figure 1D-E). This observation was not, however, isolated to chemotherapy-induced cell death, as Fas-induced apoptotic cells and the rare dead cells in control populations were also noted to be significantly stiffer than live, untreated cells (data not shown). Although cell death is often coupled with a decrease in cell volume,^11,12  we found that cell shrinkage occurs after chemotherapy-induced cell stiffening (Document S1 and Figure S1). This suggests that the cause of stiffness increases is not simply increased density due to decreased volume.

Results from microchannel experiments using primary leukemia cells exposed to chemotherapy show that dead cells are more likely to obstruct capillary-sized channels and cause cell aggregation than live cells (Figure 1F-G, Video S1). The fraction of cells initiating microchannel obstruction relative to those traversing the microchannel system was approximately 7 times higher for dead cells than live cells.

To determine the relationship between increasing cell stiffness and chemotherapy exposure time, serial single-cell stiffness measurements were taken over several hours after exposure. During stiffness measurements on each cell (n = 15-20), the apoptotic state was tracked using dual fluorescent labeling with cresyl violet-DEVD and Sytox Green. Figure 2A shows the change in stiffness for a single leukemic cell taken from a patient newly diagnosed with M5 AML after it was exposed to daunorubicin. Cell stiffness began increasing within an hour of exposure and increased most significantly after apoptosis was detected, peaking near the point of cell death. Interestingly, cell stiffness began to increase before peak caspase activation in apoptosis. Accordingly, for the same patient sample, the average apparent stiffness for populations of control cells, early apoptotic cells, and late apoptotic/dead cells exhibit significant increases with progression through the stages of chemotherapy-induced cell death (Figure 2B, P < .05).

From serial single-cell stiffness measurements on several patient samples and cell lines, different chemotherapeutic agents were found to change cell stiffness at different rates (Figure 2C). In general, the stiffness of myeloid leukemia cells exposed to daunorubicin rapidly increased before the onset of cell death and then stabilized. Lymphoid leukemia cells exposed to dexamethasone exhibited a more gradual increase in cell stiffness as cell death occurred. Control cells from each sample remained PI negative throughout the experiments, and their stiffness remained close to the initial stiffness. To determine whether the difference in stiffening rate is due to the chemotherapeutic agent or to the cell type, populations of HL60 (myeloid) and Jurkat (lymphoid) cells were exposed to daunorubicin and dexamethasone separately, and cell deformability was tracked in 4 separate experiments. For both HL60 and Jurkat cell lines, cells exposed to daunorubicin exhibited an increase in cell stiffness and onset of cell death significantly earlier than cells exposed to dexamethasone, indicating chemotherapy type dominates the kinetics of cell stiffness, not leukemia cell type.

Previous research has shown that chemotherapy-induced cell death is associated with reorganization of actin in leukemic cell lines.^13-17  To test whether the actin cytoskeleton was involved in the observed chemotherapy-induced stiffness increase, the stiffness of single HL60 cells exposed to daunorubicin was tracked, as 2 μM cytochalasin D, an inhibitor of actin polymerization, was added (Figure 2D). Cells exposed simultaneously to both cytochalasin D and daunorubicin exhibited almost no increase in cell stiffness as cell death progressed. When cytochalasin D was added 45 minutes after daunorubicin exposure, as cell stiffness was already increasing, cell stiffness ceased to increase within 15 minutes and declined to 40% of its maximum stiffness after 70 minutes. The average stiffness of dead HL60 and Jurkat cell populations exposed simultaneously to daunorubicin and cytochalasin D was found to be significantly less than the stiffness of cells exposed to only daunorubicin (P < .05, Figure 2E). These decreases in stiffness due to cytochalasin D suggest that the stiffness increase with chemotherapy-induced cell death is at least partly due to dynamic changes in the actin cytoskeleton.

Our results reveal that chemotherapy-induced cell death increases the stiffness of leukemia cells, which may influence vascular flow in the microcirculation. This observed link between cell death and increasing cell stiffness may have implications for patients with acute leukemia and hyperleukocytosis. In some cases, clinical deterioration due to leukostasis paradoxically does not occur until after chemotherapy has been initiated,^18-21  and alteration of leukemic cells' biophysical properties by chemotherapeutic agents has been hypothesized as a possible link.^19,20,22  Further research is needed to fully characterize the impact of decreased deformability of dying cells as well as the role of other factors like leukemia cell–endothelial interactions and transmigration on the pathophysiology of leukostasis in acute leukemia. With the capability to investigate these biophysical phenomena at the single-cell level, new research platforms like AFM and microfluidic assays may provide valuable insight into this and many other hematologic problems.

Contribution: W.A.L. and M.J.R. performed experiment design and execution and data analysis, and wrote the manuscript; and D.A.F. performed experiment design and wrote the manuscript.

Conflict-of-interest disclosure: The authors declare no competing financial interests.

Correspondence: Daniel A. Fletcher, Department of Bioengineering, UC Berkeley, 481 Evans Hall, no. 1762, Berkeley, CA 94720-1762; e-mail: fletch@berkeley.edu

The authors wish to thank Kevin Shannon, Benjamin Braun, Michelle Hermiston, Todd Sulchek, Sanjay Kumar, Sapun Parekh, Joshua Shaevitz, and Martijn van Duijn for discussions and careful reading of the manuscript. Microfabrication was performed in the UC Berkeley Microlab. Several patient samples were obtained from the UCSF Hematopoietic Tissue Cell Bank courtesy of Mignon Loh. This work was supported by a National Institutes of Health (NIH) National Research Service Award (NRSA) and the Hammond Research Fellowship of the National Childhood Cancer Foundation/Children's Oncology Group (W.A.L.), a National Science Foundation (NSF) Graduate Research Fellowship (M.J.R.), and an NSF Faculty Early Career Development (CAREER) Award and support from the University of California Cancer Research Coordinating Committee and Lawrence Livermore National Laboratory (D.A.F.).