Cell Cycle Dependent Mechanisms Underlie Vincristine-Induced … · Anisha Kothari1,Walter N. Hittelman2, and Timothy C. Chambers1 Abstract Microtubule-targeting agents (MTA), such

Therapeutics, Targets, and Chemical Biology

Cell Cycle–Dependent Mechanisms UnderlieVincristine-Induced Death of Primary AcuteLymphoblastic Leukemia CellsAnisha Kothari1, Walter N. Hittelman2, and Timothy C. Chambers1

Abstract

Microtubule-targeting agents (MTA), such as the taxanes andvinca alkaloids, are used to treat a variety of cancers due to theirability to perturb microtubule dynamics. In cell culture, MTAsexert their anticancer effects primarily by causing mitotic arrestand cell death. However, accumulating indirect evidence suggeststhat MTAs may exert their cytotoxicity in human tumors byinterfering with interphase microtubules. In this study, we soughtto develop and characterize an experimental system in which totest the hypothesis thatMTAs induce cell death during interphase.Primary adult acute lymphoblastic leukemia (ALL) cells treatedwith vincristine only weakly exhibited colocalization betweenmitotic and apoptoticmarkers andmajor characteristics ofmitoticdeath, such as an increase in cells with 4NDNA content before theappearance of cells with <2N DNA content, suggesting a mixed

response. Therefore, we separated ALL cells into distinct phasesof the cell cycle by centrifugal elutriation, labeled cells with5-ethynyl-20-deoxyuridine (EdU), and then treated each popu-lation with vincristine. Cells isolated during G1 underwent celldeath without evidence of EdU uptake, indicating that thecytotoxic effects of vincristine took place during G1. Conversely,cells isolated during S or G2–M phases underwent death fol-lowing mitotic arrest. Thus, vincristine induces distinct deathprograms in primary ALL cells depending on cell-cycle phase,and cells in G1 are particularly susceptible to perturbationof interphase microtubules. Primary ALL cells may thereforeprovide a powerful model system in which to study themultimodal mechanisms underlying MTA-induced cell death.Cancer Res; 76(12); 3553–61. �2016 AACR.

IntroductionMicrotubule targeting agents (MTA) are clinically important

chemotherapeutic agents used in the treatment of various cancers(1). MTAs act by interfering with microtubule dynamics, whichplays a major role in mitotic spindle formation. MTAs such as thevinca alkaloids, including vincristine, cause microtubule depo-lymerization, whereas taxanes stabilize microtubules preventingtheir depolymerization (2). Treatment of cultured cells withMTAsoften leads to prolonged activation of the mitotic spindle check-point, leading to mitotic arrest and eventually cell death, provid-ing a rationale for their use as antitumor agents (3–5). Cells maydie in mitosis or exit mitosis without proper chromosome seg-regation or cytokinesis, referred to as mitotic slippage. Afterslippage, cells may die in interphase, or arrest and survive, orresume cycling (6, 7). Whether cells die in mitosis or undergomitotic slippage has been postulated to depend on the balance oftwo pathways, one controlling cyclin B1 levels and the othercaspase activation (8). We have reported that cyclin-dependent

kinase 1 (Cdk1)/cyclin B1-mediated Bcl-xL/Bcl-2 phosphoryla-tion acts as a functional link coupling mitotic arrest and celldeath (9), and that cell fate after mitotic arrest is dictated by thedegree of Cdk1/cyclin B1–mediated phosphorylation of anti-apoptotic Bcl-2 proteins (10).

Implicit in conventional theories of MTA action is that theyinduce mitotic arrest prior to cell death. This viewpoint hasrecently been challenged, on the basis of largely indirect evi-dence but nonetheless compelling lines of reasoning (11, 12).The major points underlying this paradigm shift include thefollowing: (i) all of the mechanistic data derive from cellculture and xenograft models, which may not recapitulatepatient tumor conditions; (ii) there is a poor relationshipbetween the status of the spindle assembly checkpoint andcellular sensitivity to MTAs (6); (iii) MTAs are effective clini-cally despite the slow doubling times and lowmitotic indices ofsome human tumors (11–13); and (iv) mitosis-specific inhi-bitors, such as those targeting Aurora kinases and polo-likekinases, are effective in vitro but have shown poor activity inclinical trials (11–13). Such collective evidence has led to thesuggestion that the therapeutic efficacy of MTAs in the clinicalsetting may be attributed to interference with interphase micro-tubule functions (11, 12). This viewpoint has itself been ques-tioned, resulting in an intense current debate (14–17). Forexample, it has been suggested that the slow doubling timeof tumors likely reflects the balance of proliferation and death,and thus cycling cells may be more abundant than predictedbased solely on doubling time (14). In addition, it has beencontended that since drugs such as taxol are concentrated andretained in tumor cells for lengthy periods, their effects may beexerted on quiescent cells that reenter the cycle after a pro-longed delay (13, 18, 19).

1Department of Biochemistry and Molecular Biology, University ofArkansas for Medical Sciences, Little Rock, Arkansas. 2Departmentof Experimental Therapeutics, University of Texas MDAnderson Can-cer Center, Houston, Texas.

Note: Supplementary data for this article are available at Cancer ResearchOnline (http://cancerres.aacrjournals.org/).

CorrespondingAuthor: Timothy C. Chambers, Department of Biochemistry andMolecular Biology, University of Arkansas for Medical Sciences, 4301 W. Mark-ham St., Little Rock, AR 72205. Phone: 501-686-5755; Fax: 501-686-8169; E-mail:[email protected]

doi: 10.1158/0008-5472.CAN-15-2104

�2016 American Association for Cancer Research.

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Rapid induction of interphase death by vinca alkaloids hasbeen reported in a limited number of cell lines, but generally onlywith relatively high drug concentrations or in combination withother drugs (20–23). A studyof breast cancer cell lines treatedwithmicrotubule destabilizers reported arrest in G1 and G2 phases,suggesting the drugs may perturb cells in interphase, but thesubsequent fate of the treated cells was not examined (24).Because MTAs typically manifest their effects in mitosis in mostcultured cell lines, a major obstacle to progress in this area is thelack of appropriate laboratorymodels ofMTA-induced interphasedeath.

To develop and characterize a cell system to test the hypothesisthat MTAs could also induce interphase death, we assessed theextent of mitotic arrest in several different cell types after MTAtreatment. Among these were primary adult acute lymphoblasticleukemia (ALL) cells derived from several individual adultpatients, expanded, and established in culture using a definedserum-free medium (25, 26). In the current study, to determinewhether cells in different cell-cycle contexts were differentiallyaffected by vincristine, we used centrifugal elutriation to isolatepopulations of ALL cells in different cell-cycle phases. We showthat vincristine can induce cell death directly in G1 phase cells,whereas cells beyond G1 phase transit to M-phase before under-going cell death. Thus, vincristine induces distinct cell-cycle–dictated pathways of cell death in primary ALL cells. To ourknowledge, these results are the first to definitively demonstrateinduction of interphase death by relatively low and therapeuti-cally relevant concentrations of an MTA used alone, and, as such,provide a powerful model system to study clinically applicabledeath mechanisms induced by drugs in this class and to providehypotheses to explore in vivo.

Materials and MethodsFor materials, cell extraction and immunoblotting, cell-cycle

analysis, analysis of microtubule content, and Statistical analysis,see Supplementary Materials.

Cell cultureKB3 cells (HeLa subline) were maintained in DMEM, and

RS4;11 ALL cells were maintained in RPMI1640 medium, eachsupplemented with 10% bovine growth serum, 2 mmol/LL-glutamine, 50 U/mL penicillin, and 50 mg/mL streptomycin.ALL cell cultures were maintained in suspension as describedpreviously (25) in Iscove's modified Dulbecco's medium(IMDM) containing serum-free supplement (10 mg/mL choles-terol, 6 mg/mL human serum albumin, 0.5 mg/mL amphoter-icin, 1 mg/mL insulin, 200 mg/mL human apo-transferrin,50 mmol/L 2-mercaptoethanol, 2 mmol/L glutamine, and50 U/mL penicillin). Cells were maintained at 37�C and 5%CO2. The ALL cultures utilized, namely ALL-2 and ALL-5,correspond to their original designations of CM and PH cells,respectively (25). Doubling time was determined by monitor-ing viable cell count daily for up to 6 days. Authentication ofthe cell lines and ALL cultures was established via short tandemrepeat (STR) profiling in September, 2014, by Genetica DNALaboratories (LabCorp Specialty Testing Group). The STR pro-files of KB3 and RS4;11 cell lines matched that of referenceprofiles available in the ATCC database, and, as expected, thetwo primary ALL cell cultures each gave unique profiles that didnot match any in the repository.

Cell viability assayCell viability was determined using 3-(4,5-dimethylthiazol-2-

yl)-2,5-diphenyltetrazolium bromide (MTT) as described previ-ously (27). Cells (2,000 per well for KB3 and 30,000 per well forothers)were seeded in96-well plates, and vincristinewas added ina fixed final concentration of 0.1% DMSO. After incubating for atime period equivalent to twice their doubling time (see Table 1),MTT reagent (50 mg/10 mL/well) was added and incubated over-night at 37�C. The next day, 0.1mLof 10%SDS in 0.01mol/LHClwas added, and after overnight incubation, absorbance readingswere taken at 540 nm.

Centrifugal elutriationCentrifugal elutriation was performed using a Beckman JE-6

elutriator rotor as described previously (28). Cells (3.5–4 � 108)were suspended in 20-mL elutriation buffer (Hank's bufferedsalt solution containing 1.6 g/L 2-naphthol-6,8-disulfonic aciddipotassium salt and 2% calf serum), passed through a 25Gneedle twice, and introduced into the chamber at a flow rateof 0.8–1.1 mL/minute with a rotor speed of 3,300 rpm. Rotorspeed was reduced in decrements of 80 rpm to collect a total of20 fractions. Aliquots of individual fractions were fixed andsubjected to propidium iodide staining and flow cytometry.Specific fractions were pooled to enrich for cells in defined cell-cycle phases. Cells with the smallest diameter (earliest in the cellcycle) eluted first. Pooled fractions were centrifuged and cellsresuspended in fresh medium at 1–1.5 � 106 cells/mL. Celldiameter was measured using a Beckman Z2 Coulter Counterand data analyzed with Z2 AccuComp software.

EdU incorporation assay5-ethynyl-20-deoxyuridine (EdU) incorporation assay was per-

formed according to manufacturer's instructions (Invitrogen)using the Click-iT EdU Imaging Kit (cat. no. C10086), with somemodifications. Cells isolated after centrifugal elutriation wereincubated with 10 mmol/L EdU and either 0.1% DMSO (vehicle)or 100 nmol/L vincristine and harvested at specific time intervalsby washing in PBS and fixing with 70% cold ethanol for 15minutes. Cells were then washed with wash solution (3%BSA inPBS), incubatedwithpermeabilizationbuffer (0.5%TritonX-100 in PBS) for 20minutes, washed again, suspended in 200 mLof Click-iT reaction cocktail (with either Alexa Fluor 488 or 647),and incubated for 30minutes in the dark. Cells were thenwashed,resuspended, and 20 mL propidium iodide (BD Pharmingen) wasadded prior to analysis by flow cytometry.

Cell proliferation assayTo monitor individual cell divisions and to determine whether

ALL cells harbor a quiescent population, the Cell ProliferationDye eFluor 450 kit (eBioscience) was used. A total of 1� 106 ALL

Table 1. Doubling time and vincristine sensitivity of KB3, RS4;11, and primaryALL cells

Doubling time (d) IC50 VCR (nmol/L)

KB3 1.0 � 0.1 2.10 � 0.09RS4;11 1.5 � 0.3 1.05 � 0.24ALL-2 2.5 � 0.5 3.90 � 0.07ALL-5 2.8 � 0.6 0.60 � 0.07

NOTE: The values for doubling time represent mean � SD (n ¼ 6), and IC50

values were derived from data in Supplementary Fig. S1.Abbreviation: VCR, vincristine.

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cells were washed with PBS thrice and resuspended in PBS. Anequal volume of 20 mmol/L eFluor 450 was added and incubatedat room temperature for 20 minutes according to manufacturer'sinstructions. Labeling was stopped by adding 4 and 0.04 volumesof completemedia and 20%human serum albumin, respectively,and incubatingon ice for 5minutes. Cellswerewashed three timeswith complete media, resuspended in medium, and returned toculture. Cell aliquots were removed at daily intervals, washed andresuspended in PBS, and propidium iodide was added. Fluores-cence wasmeasured using a BD LSRFortessa Flow Cytometer withdata analysis using ModFit LT 4 (VSH) software. Nonviable(propidium iodide–positive) cells were excluded from analysis.

ResultsVincristine induces cell death without significant prior mitoticarrest in primary ALL cultures

In initial experiments, we compared the effects of vincristine onmitotic arrest in exponentially growing populations of KB3(HeLa) and RS4;11 (ALL) cell lines and in two primary ALL cellcultures, namely ALL-2 andALL-5, derived as described previously(25) and recently characterized with respect to Bcl-2 dependence(26). DNA content analysis showed that KB3 cells underwent

mitotic arrest (determined by 4N DNA content) and subsequentapoptosis (determined by <2N DNA content) in response to100 nmol/L vincristine (Fig. 1A), similar to the results we hadreported previously using vinblastine (29, 30). RS4;11 cells gave asimilar response, although the extent of M-phase arrest was lessthan observed for KB3 cells (Fig. 1A). In contrast, when weexamined the primary ALL cell cultures, cell death occurredwithout evidence of a prolonged mitotic arrest (Fig. 1A). Quan-titation of the results confirmed that vincristine induced time-dependent death in all cell types as determined by <2N DNAcontent (Fig. 1B, top), but significant mitotic arrest was onlyobserved in KB3 and RS4;11 cells (Fig. 1B, bottom). This pro-found difference in response was not due to major differences invincristine sensitivity, as IC50 values for vincristine of the ALLcultures were similar to those of the KB3 and RS4;11 cell lines(Supplementary Fig. S1; Table 1). Two other MTAs were tested inALL-5 cells: docetaxel, a microtubule polymerizing agent, andvinblastine, amicrotubule depolymerizing agent (SupplementaryFig. S2). Both drugs induced significant levels of apoptosis with-out evidence of a prolonged or marked mitotic accumulation,suggesting that they can also induce death from interphase. Thelack of a robust mitotic arrest in response to vincristine in the ALLcells could not be attributed to defects in the spindle assembly

Figure 1.Vincristine induces cell death without prior robust mitotic arrest in primary ALL cultures. A, flow cytometric analysis for cell-cycle distribution of KB3, RS4;11,ALL-2, and ALL-5 cells, untreated or treated with 100 nmol/L vincristine for the times indicated. Histograms are representative of three independent experiments.B, quantitation of data from A showing the proportion of cells with <2N (sub-G1) or 4N (G2–M) DNA content with respect to time of vincristine treatment.Results are expressed as mean � SD (n ¼ 3). � , P � 0.05 (Student t test). C and D, kinetics of PARP cleavage and relationship to the mitotic marker MPM2.Extracts were made from untreated or vincristine-treated (100 nmol/L) KB3, ALL-2, or ALL-5 (C) or RS4;11 (D) cells at the times indicated and subjected toimmunoblotting for PARP or MPM2. Intact and cleaved species of PARP are shown. GAPDH was used as a loading control.

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checkpoint, because robust mitotic arrest was observed in cellstreated with the Aurora A kinase inhibitor, MLN8237 (Supple-mentary Fig. S3). To further examine the kinetics of cell death andrelationship tomitotic arrest, immunoblotting was performed forPARP, which is cleaved during apoptosis, and with MPM2 anti-body, which recognizesmitotic phosphoproteins (Fig. 1C andD).PARP cleavage in KB3 cells was coincident with strong MPM2staining, consistent with Fig. 1A and with the occurrence ofmitotic death, whereas PARP cleavage in the primary ALL cultureswas incomplete at 24hours andoccurred in concert with relativelyweak MPM2 staining (Fig. 1C). In RS4;11 cells, PARP cleavagepreceded a relatively strong MPM2 signal (Fig. 1D), suggestingthat some cells may die via nonmitotic death. Preliminary experi-ments have indicated that a range of vincristine concentrations,from 10 to 1,000 nmol/L, induces death of ALL cells withoutsignificant mitotic arrest, as determined by examination ofDNA content by propidium iodide staining and flow cytometry,suggesting that interphase death in these cells is a general responseto microtubule inhibition and not a concentration-dependentresponse.

Distinct responses of primary ALL cells to vincristine indifferent phases of the cell cycle

The results of Fig. 1 suggested two possibilities; either that theALL cellswere dying in response to vincristine after a very transientmitotic arrest, such that mitotic markers were only weakly repre-senteddue to their rapid reversal, or that theywere dying primarilyin interphase without transit first to mitosis. To distinguish thesepossibilities, isolation or synchronization of cells at different cell-cycle phases was needed. However, their relatively long doublingtimes (Table 1) precluded chemical methods, as the cells wereunable to survive lengthy treatments with agents such as thymi-dine or hydroxyurea. Therefore, a physical method of cell syn-chronization, centrifugal elutriation, which separates cells basedon their size, which, in turn, increases with advance through the

cell cycle, was tested. Flow cytometric analysis of side-scatter, anindicator of cell size, and of DNA content after propidium iodidestaining, showed a good correlation between the two parameters(Supplementary Fig. S4). Initial elutriation experiments indeedestablished the effectiveness of the technique in isolation of cellsin different phases of the cell cycle. As shown in SupplementaryFig. S5, pools of early fractions were highly enriched (95.4%) inG1 (or G0) phase cells with 2N DNA content, intermediatefractions contained larger diameter cells in G1 and S phases, andlater fractions were enriched (61.2%) in cells with 4N DNAcontent, representing G2–M phases. Measurement of average celldiameter in individual fractions confirmed the relationshipbetween cell diameter and cell-cycle phase. For example, specificvalues in a typical experiment ranged from 8.9 mm for earlyfractions representing early G1 phase cells, to 11.7 mm for latefractions representing G2–M phase cells.

To determine the response to vincristine of cells in differentphases of the cell cycle, elutriated ALL-2 cells in either G1 or G2–Mphases were treated with vehicle (0.1% DMSO) or 100 nmol/Lvincristine, harvested at specific times, andDNA content analyzed(Fig. 2).When treatedwith vehicle (Fig. 2A, top row), cells initiallyin G1 transited into S- and G2–M phases over time, with 20.1%exhibiting 4N DNA at 24 hours, with a return to approximatelynormal cell-cycle proportions by 48 hours. Over this 48-hourperiod, there was essentially no evidence of cell death, with only2.3% of cells with <2N DNA at 48 hours, indicating that the cellssuffered no ill-consequences from elutriation and subsequentculture. When cells initially in G1 were treated with vincristine(Fig. 2A, second row), a smaller percentage of the total populationtransited into S and G2–M phases compared with vehicle-treatedcells, and between 24 and 48 hours there was a large increasein cells with <2N DNA content indicative of cell death. Similarresults were obtained with ALL-5 cells (Supplementary Fig. S6A).We next analyzed cells enriched for those with 4N DNA content(Fig. 2A, third and fourth rows). When treated with vehicle, the

Figure 2.Distinct responses of primary ALL cells to vincristine in different phases of the cell cycle. A, ALL-2 cells separated by centrifugal elutriation and enrichedfor cells in either G1 (2N DNA, top two rows) or G2–M (4N DNA, bottom two rows) were treated with 0.1% DMSO or 100 nmol/L vincristine (VCR) forthe times indicated. Cells were fixed and stained with propidium iodide and analyzed by flow cytometry. The histograms are representative of threeindependent experiments. Inset, average 4N DNA content and selective <2N DNA content, expressed as percentage of total cells analyzed. B and C,ALL-2 or ALL-5 cells, as indicated, in G1 phase (B) or G2–M phase (C) were treated with 100 nmol/L vincristine (VCR) for the times indicated andextracts subjected to immunoblotting for PARP or MPM2. Intact and cleaved species of PARP are shown. Untreated or vincristine-treated KB3 cells (left twolanes) served as positive control. GAPDH was used as a loading control.

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cells completed the cell cycle and divided, mainly returning to G1

phase at 16–24hours,with little evidenceof cell deathby48hours(Fig. 2A, third row). In contrast, when treated with vincristine,cells underwent M-phase arrest, with 86.4% cells having 4NDNAcontent within 8 hours of vincristine treatment, and by 48 hours,50% of cells had <2N DNA content (Fig. 2A, fourth row). Similarresults were obtained with ALL-5 cells, which are representedgraphically in Supplementary Fig. S6B.

Elutriated ALL-2 or ALL-5 cells initially in G1 (Fig. 2B) or G2–M(Fig. 2C) were treated with 100 nmol/L vincristine for indicatedtime periods and analyzed by immunoblotting for PARP andMPM2. G1 cells exhibited vincristine-induced PARP cleavage aftera delay of 24 hours and in the absence of the mitotic markerMPM2 (Fig. 2B). In contrast, G2–M ALL cells exhibited muchmore rapid vincristine-induced PARP cleavage coincident withincreased MPM2 staining (Fig. 2C). Together, the results of Fig. 2suggest that vincristine induces distinct pathways of cell death inprimary ALL cells, dependent on position in the cell cycle whenthe drug is encountered. The fact that themajority (typically 70%–

75%) of ALL cells are in G1 phase with only a small proportion(9%–12%) in G2–M phases (Fig. 1A) likely explains why mitoticdeath signals are not prominent when asynchronous cells wereexamined (Fig. 1), but become readily detectable when G2–M-enriched cells were used (Fig. 2). As a control for these experi-ments, elutriated cells were treated with 0.1% DMSO up to 48hours, and extracts subjected to immunoblotting for PARP andMPM2. As shown in Supplementary Fig. S7, PARP remained intactthroughout, consistent with maintenance of cell viability, andMPM2 staining was largely lacking, consistent with an absence ofcells undergoing mitotic arrest.

Vincristine causes microtubule depolymerizationBecause vincristine appeared to promote death in G1 phase in

primary ALL cells, it was particularly important to confirm thatmicrotubules were targeted by the drug. ALL cells were thereforetreated with vincristine, or with taxol or CaCl2 to act as positiveand negative controls, respectively, for tubulin polymerization,and depolymerized and polymerized tubulin were separated andanalyzed, as described inMaterials andMethods. As shown in Fig.3, tubulin was present in control cells in both depolymerized(soluble) and polymerized forms. Vincristine caused an increasein depolymerized tubulin and a corresponding decrease in poly-merized tubulin in both asynchronous (Fig. 3A) and G1 phase(Fig. 3B) ALL cells. The known depolymerizing agent CaCl2 (31)gave very similar results; conversely, the microtubule stabilizing

agent taxol (32) caused an increase in polymerized tubulin and acorresponding decrease in depolymerized tubulin. Quantitationof tubulin expression, performed as described in Materials andMethods, confirmed these observations (see Fig. 3 legend). Totaltubulin levels were unaffected. These results confirm that micro-tubules are a target of vincristine action in ALL cells.

ALL cells in G1 phase can undergo death directly in response tovincristine without cell-cycle transit

To more rigorously test whether primary ALL cells in G1 phasedie directly in response to vincristine without cell-cycle transit,ALL-5 cells in G1 phase were incubated with 5-ethynyl-20-deox-yuridine (EdU) and either vehicle or 100 nmol/L vincristine andharvested at specific intervals up to 48 hours. After fluorescentlylabeling the EdU incorporated into DNA, cells were stained withpropidium iodide and subjected to two-parameter flow cytome-try. The resulting cytograms are shown inFig. 4A, andquantitationof the proportion of EdU-positive versus EdU-negative cells thatwere either alive (>2NDNA)or dead (<2NDNA) shown in Fig. 4BandC, respectively. Cells initially gated to the left side of the lowerright quadrant of the cytogram, reflecting a DNA content (x-axis)of 2N and an absence of EdU labeling (y-axis; Fig. 4A). Whentreated with vehicle (Fig. 4A, top row), the cells became EdUlabeled and increased their DNA content, reflecting transitthrough S-phase and beyond. By 48 hours, 43% of live cells wereEdU labeled (Fig. 4B), and only a minor population had <2NDNA content (Fig. 4A and C). When treated with vincristine(Fig. 4A, bottom row), a majority of cells in G1 phase shiftedfrom 2N DNA content to <2N DNA content without becomingEdU labeled, representing 43.5% of total cells by 48 hours, andonly a minor proportion (2.4%) of dead cells were EdU positive(Fig. 4C). These results indicate that ALL cells treated with vin-cristine can die directly from G1 without further cell-cycle transit.Essentially identical results were obtained in two independentexperiments with ALL-5 cells and the same pattern was obtainedwith ALL-2 cells (Supplementary Fig. S8).

Validation of EdU labeling through use of a mixed populationTo rule out the possibility that the presence of vincristine

affected the efficiency of EdU labeling, ALL-5 cells were subjectedto centrifugal elutriation, and intermediate fractions pooled suchthat bothG1 (82.4%) and S (17.6%) phases were represented (seeSupplementary Fig. S5B). Cells were incubated with EdU and 100nmol/L vincristine or 0.1%DMSO, harvested at intervals up to 48

Figure 3.Vincristine depolymerizes microtubules in both asynchronous and G1-phase ALL cells. Asynchronous (A) or G1-phase (B) ALL-5 cells were treated with 0.1 %DMSO (Ctrl) or 100nmol/L vincristine (VCR) for 1 hour andharvested. Cellswere also treatedwith 100nmol/L taxol (TAX) for 1 hour or 2mmol/LCaCl2 for 5minutes toserve as polymerization and depolymerization controls, respectively. Soluble tubulin (S) and polymerized tubulin (P) were separated as described inMaterials and Methods and subjected to immunoblotting for a-tubulin. Total tubulin, present in cell homogenates prior to centrifugation, was used as aloading control. The relative proportions of a-tubulin present in soluble and pellet fractions, determined as described in Materials and Methods and given as%S/%P, is as follows. A, Ctrl, 42/58; TAX, 21/79; CaCl2, 50/50; VCR, 56/44. B, Ctrl, 60/40; TAX, 51/49; CaCl2, 75/25; VCR, 76/24.






hours and subjected to flow cytometry (Fig. 5). On treatment withvehicle (Fig. 5A, top row), by 48 hours, about half the populationof cells had become EdU labeled (Fig. 5B) with only a lowincidence of dead cells (Fig. 5C). When treated with vincristine(Fig. 5A, bottom row), by 48hours, dead cells were approximatelyevenly divided between EdU-positive and EdU-negative, withabout 21% of the total appearing in the top left quadrant, thatis, dead and EdU-positive (Fig. 5A and C). This corresponds wellwith the initial S-phasepopulation (17%of total cells), and showsthat vincristine does not interfere with EdU labeling. Essentially,the same pattern was obtained with ALL-2 cells. These data alsoemphasize and reinforce the conclusion that G1 cells treated withvincristine are highly susceptible to death without further cell-cycle transit.

All cells within the primary ALL cultures are activelydividing

We considered the possibility that ALL cells with 2NDNA may be comprised of two subpopulations, namelyquiescent (or non-cycling, G0) and cycling (G1), and thatG0 cells may be the population most susceptible to vincris-tine-induced death. To determine whether quiescent cellswere present, a cell proliferation assay using the violet fluo-rescent dye, eFluor 450, was performed, as described inMaterials and Methods. The dye binds to primary amines inproteins, and the label is expected to dilute with each cell

division but be maintained at original levels in nondividingcells. ALL-5 cells were labeled with the dye and after washing,fluorescence was determined at daily intervals. As shown inSupplementary Fig. S9, at day zero, a single parent peak offluorescence was observed, and over 3 days the fluorescentintensity reduced to approximately one-half with no residualsignal at the day zero position. These results indicate that ALLcells are in an actively dividing state with no evidence of aquiescent population, and are also consistent with a doublingtime of 2.8 days determined independently (Table 1). Similarresults were obtained when carboxyfluorescein succinimidylester (CFSE), another proliferation marker similar to eFluor450, was used. These results are also consistent with a pre-vious study that inferred that all cells within the ALL cultureswere in a proliferative state (25).

DiscussionIn this study, we provide evidence that primary ALL cell cultures

treated with vincristine undergo two distinct pathways of celldeath. Using centrifugal elutriation to separate cells into differentcell-cycle phases together with EdU labeling to monitor subse-quent S-phase progression, we showed that cells in G1 phase candie directly, whereas cells in later phases transit to M-phase firstand then die. A model summarizing these findings is presentedin Fig. 6. We propose that primary ALL cells in G1 phase are

Figure 4.Primary ALL cells in G1 phase canundergo death directly without transitto S-phase. ALL-5 cells in G1 phaseobtained by centrifugal elutriationwere incubated with EdU and either0.1% DMSO or 100 nmol/L vincristine(VCR) for the times indicated. Cellswere harvested, fixed, and stained forEdU incorporation andwith propidiumiodide (PI) and analyzed by flowcytometry, with propidium iodidestaining represented on the x-axis andEdU staining on the y-axis. B and C,graphical representation of data fromA. B, distribution between EdU-negative andEdU-positive for live cells(>2N DNA). C, distribution betweenEdU-negative and EdU-positive fordead cells (<2N DNA), expressed aspercentage of total cells. Results areexpressed as mean � SD (n ¼ 3).

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susceptible to death directly, whereas cells that have passeda putative "microtubule sensitivity checkpoint" in late G1/earlyS-phase continue to cycle and undergo M-phase arrest and celldeath. Docetaxel and vinblastine appear to act similarly (Sup-plementary Fig. S2), suggesting that this model applies to MTAsin general. Further studies will be needed to elucidate thecellular and molecular basis for G1 phase sensitivity to MTAs,and the mechanisms that allow many cell lines to bypass suchsusceptibility.

When G1 phase cells were subjected to EdU labeling,only about half the population migrated out of their startingposition, and many remained EdU-negative with 2N DNA(Fig. 4A). However, when G1 phase cells were subjected tothe cell proliferation dye assay in the absence of drug treat-ment, all the cells showed the expected dye dilution within twodays of culture, indicating they had progressed through the cellcycle. Thus, isolated G1 cells appear to be fully cell-cyclecompetent. The reason for the apparent inability of manyG1-phase cells to progress into subsequent cell-cycle phasesduring the EdU assay may therefore be related to the specificconditions of the assay. Exhaustion of the pool of EdU doesnot seem to be a factor, because cells would still be expected toenter S-phase and show an increase in DNA content from 2N to4N. As this fraction of the population remained EdU-negative

Figure 6.Proposed model for mode of action of vincristine in ALL cells. Cells in G1

phase are susceptible to death directly, whereas cells that have passed aputative "microtubule sensitivity checkpoint" in late G1/early S-phasecontinue to cycle and die following mitotic arrest.

Figure 5.Vincristine does not affect EdUlabeling of ALL cells. A, ALL-5 cells,separated by centrifugal elutriationand representing cells in both G1

(82.4%) and S (17.6%) phases (seeSupplementary Fig. S5B), wereincubated with EdU and either 0.1%DMSOor 100 nmol/L vincristine (VCR)for the time indicated, stained forincorporation of EdU and withpropidium iodide (PI), and subjectedto flow cytometry, as in Fig. 4. B andC,graphical representation of data fromA. B, distribution between EdU-negative and EdU-positive for livecells (>2N DNA). C, distributionbetween EdU-negative and EdU-positive for dead cells (<2N DNA),expressed as percentage oftotal cells. Results are expressed asmean � SD (n ¼ 3).






with 2N DNA content, it appears they failed to enter S-phase,perhaps because EdU incorporation limits DNA replication incertain cells. Further work will be required to clarify the basisfor this observation.

Previous studies have provided indirect evidence for inter-phase death or cell-cycle–independent death in response tovinca alkaloids (20–24). In most instances, significant levels ofinterphase death were only observed in conjunction with otheragents, such as that seen in ML-1 cells cotreated with vinblas-tine plus inhibitors of either ERK or CDKs (21, 23). In a morerecent study of a panel of leukemia cell lines, one cell line,OC1-AML1, underwent very rapid death, with 100% of thecells affected within 4 hours after being treated with vincristinealone (22). In another study, rapid cell death was observed inU2OS osteosarcoma cells treated with vinorelbine (20). How-ever, in both cases, relatively high drug concentrations, in thelow- to mid-micromolar range, were used, and it remainsunclear whether such acute effects would have been observedat lower concentrations. Importantly, in these and relatedstudies, evidence for interphase death has often been inferredbased on the rapid appearance of death signals such as cleavedPARP or activated caspase-3 in asynchronous cells. We alsonoted PARP cleavage at early time points in vincristine-treatedasynchronous ALL cell cultures (Fig. 1C), but examination ofcells enriched in different cell-cycle phases revealed that thosein G2 and/or M-phase were the source of these signals (Fig.2C). In contrast, vincristine-induced cell death in G1-phasecells was considerably delayed (Fig. 2B). Thus, the earlyappearance of death signals in cells treated with an MTA doesnot necessarily imply death originated in interphase cells, ascells advancing to and entering M-phase can be rapidly sus-ceptible, especially if death ensues without a prolonged mitot-ic arrest. The use of cells enriched in different phases asexploited here eliminates interpretational problems that canresult from the study of asynchronous populations undergoingmixed responses.

Our data imply that microtubules play some essential role inG1 phase in primary ALL cells. Although the exact mechanismof interphase cell death induced by MTAs is still unclear, otherstudies have implicated rapid events such as the induction ofthe proapoptotic protein Noxa in CLL cells (33) or the activa-tion of JNK inML-1 cells (21). However, in preliminary studies,we have failed to observe any acute effects on Bcl-2 familyprotein expression or JNK pathway activation (unpublishedobservations). Recently, interference with intracellular proteintrafficking has been suggested as a possible mechanism toexplain the interphase effects of MTAs. One study showed thatin prostate cancer cells, taxanes act, at least in part, by inhibit-ing nuclear transport of the androgen receptor (AR), and henceblock AR-mediated signaling (34). It was also recently reportedthat MTAs interfere with the trafficking of DNA repair proteinson interphase microtubules, providing a mechanistic explana-tion for the long-standing observation that MTAs and DNA-damaging agents act synergistically (35). We observed that theinduction of apoptosis in primary ALL cells in G1 phase was notinitiated until 24 hours after vincristine treatment (Fig. 2B).Thus, the response is delayed rather than acute, and as such ismore compatible with a mechanism involving interferencewith protein trafficking than one involving rapid induction ofa death stimulus.

The results of this study have several important implica-tions. First, MTAs are well established as an effective therapyfor many cancer types, many of which are relatively slow-growing with low mitotic indices (12, 36). Our finding thatvincristine also targets cells in G1 phase provides a plausibleexplanation for these clinical observations and a potentialsolution to this paradox. Second, the results suggest thatprimary ALL cell cultures, and perhaps other primary or non-immortalized tumor cell types, represent unique tools touncover novel mechanisms of MTA action with clinical rele-vance. Third, our observation that relatively low and clinicallyrelevant concentrations of vincristine can kill cells in G1 phaseadds to a growing body of evidence that microtubules con-tribute to fundamental cellular processes outside of mitosis(37). Finally, understanding drug mechanism is criticallyimportant in rational drug design and discovery, in the testingand development of novel drug combinations, and in patientstratification. Conversely, a lack of understanding of clinicalmechanisms can lead to the development of ineffective drugs(12), resulting in poor utilization of available resources. As weenter an era where customized therapies are under develop-ment and may replace standardized regimens, studies designedto elucidate the molecular mechanisms of action of antitumoragents are even more vital.

Disclosure of Potential Conflicts of InterestNo potential conflicts of interest were disclosed.

Authors' ContributionsConception and design: A. Kothari, W.N. Hittelman, T.C. ChambersDevelopment of methodology: A. Kothari, W.N. Hittelman, T.C. ChambersAcquisition of data (provided animals, acquired and managed patients,provided facilities, etc.): A. Kothari, W.N. HittelmanAnalysis and interpretation of data (e.g., statistical analysis, biostatistics,computational analysis): A. Kothari, W.N. Hittelman, T.C. ChambersWriting, review, and/or revision of the manuscript: A. Kothari, W.N.Hittelman, T.C. ChambersAdministrative, technical, or material support (i.e., reporting or organizingdata, constructing databases): A. Kothari, T.C. ChambersStudy supervision: W.N. Hittelman, T.C. Chambers

AcknowledgmentsThe authors thank Dr. Fred Falkenburg for providing the ALL cell cultures,

AndreaHarris andDebopamGhosh for their assistancewithoptimizationof theEdU assay, Dr. Gang Lee for advice on the cell proliferation assay, Dr. GwenChilds for advice on centrifugal elutriation, Sarah Hardin and Katie Hart fortechnical assistance, Dr. Joshua Eichhorn for critical review of the manuscript,and Megan Reed for help with the figures.

Grant SupportThis workwas supported byNIH grant CA-109821 from theNational Cancer

Institute (T.C. Chambers) and in part by internal funds from UAMS College ofMedicine Research Council, UAMS Department of Biochemistry andMolecularBiology, Arkansas Biosciences Institute, and the Center for Microbial Patho-genesis and Host Inflammatory Responses grant P20GM103625 through theNIH National Institute of General Medical Sciences Centers of BiomedicalResearch Excellence.

The costs of publication of this article were defrayed in part by thepayment of page charges. This article must therefore be hereby markedadvertisement in accordance with 18 U.S.C. Section 1734 solely to indicatethis fact.

Received July 31, 2015; revised February 25, 2016; accepted April 3, 2016;published OnlineFirst May 6, 2016.

Kothari et al.





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2016;76:3553-3561. Published OnlineFirst May 6, 2016.Cancer Res Anisha Kothari, Walter N. Hittelman and Timothy C. Chambers Death of Primary Acute Lymphoblastic Leukemia Cells

Dependent Mechanisms Underlie Vincristine-Induced−Cell Cycle

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Cell Cycle Dependent Mechanisms Underlie Vincristine-Induced … · Anisha Kothari1,Walter N. Hittelman2, and Timothy C. Chambers1 Abstract Microtubule-targeting agents (MTA), such