molecular determinants of apoptosis induced by the cytotoxic ribonuclease … · mechanism of...

[CANCER RESEARCH 60, 1983–1994, April 1, 2000]

Molecular Determinants of Apoptosis Induced by the Cytotoxic RibonucleaseOnconase: Evidence for Cytotoxic Mechanisms Different from Inhibitionof Protein Synthesis1

Mihail S. Iordanov, Olga P. Ryabinina, John Wong, Thanh-Hoai Dinh, Dianne L. Newton, Susanna M. Rybak, andBruce E. Magun2

Department of Cell and Developmental Biology, Oregon Health Sciences University, Portland, Oregon 97201 [M. S. I., O. P. R., J. W., T-H. D., B. E. M.], and IntramuralResearch Support Program, Science Applications International Corporation Frederick [D. L. N.] and Laboratory of Drug Discovery, Research and Development, DevelopmentalTherapeutics Program, Division of Cancer Treatment and Diagnosis [S. M. R.], National Cancer Institute-Frederick Cancer Research and Development Center, Frederick,Maryland 21702

ABSTRACT

Cytotoxic endoribonucleases (RNases) possess a potential for use incancer therapy. However, the molecular determinants of RNase-inducedcell death are not well understood. In this work, we identify such deter-minants of the cytotoxicity induced by onconase, an amphibian cytotoxicRNase. Onconase displayed a remarkable specificity for tRNAin vivo,leaving rRNA and mRNA apparently undamaged. Onconase-treated cellsdisplayed apoptosis-associated cell blebbing, nuclear pyknosis and frag-mentation (karyorrhexis), DNA fragmentation, and activation of caspase-3-like activity. The cytotoxic action of onconase correlated with inhibitionof protein synthesis; however, we present evidence for the existence of amechanism of onconase-induced apoptosis that is independent of inhibi-tion of protein synthesis. The caspase inhibitor benzyloxycarbonyl-Val-Ala-Asp(OMe) fluoromethyl ketone (zVADfmk), at concentrations thatcompletely prevent apoptosis and caspase activation induced by ligation ofthe death receptor Fas, had only a partial protective effect on onconase-induced cell death. The proapoptotic activity of the p53 tumor suppressorprotein and the Fas ligand/Fas/Fas-associating protein with death domain(FADD)/caspase-8 proapoptotic cascade were not required for onconase-induced apoptosis. Procaspases-9, -3, and -7 were processed in onconase-treated cells, suggesting the involvement of the mitochondrial apoptoticmachinery in onconase-induced apoptosis. However, the onconase-induced activation of the caspase-9/caspase-3 cascade correlated withatypically little release of cytochromec from mitochondria. In turn, thelow levels of cytochromec released from mitochondria correlated with alack of detectable translocation of proapoptotic Bax from the cytosol ontomitochondria in response to onconase. This suggests the possibility ofinvolvement of a different, potentially Bax- and cytochromec-independentmechanism of caspase-9 activation in onconase-treated cells. As one pos-sible mechanism, we demonstrate that procaspase-9 is released frommitochondria in onconase-treated cells. A detailed understanding of themolecular determinants of the cytotoxic action of onconase could providemeans of positive or negative therapeutic modulation of the activity of thispotent anticancer agent.

INTRODUCTION

Programmed cell death or apoptosis is an integral part of embryonicdevelopment and of homeostasis in higher eukaryotic organisms (forreviews, see Refs. 1–3). With the exception of the specific case ofgranzyme B-mediated cell death utilized by cytotoxic T-lymphocytes,two autonomous but interacting pathways of apoptosis induction havebeen identified in mammalian cells: (a) apoptosis induced in responseto engagement by proapoptotic cytokines of specialized cell surface

death receptors (exemplified, for instance, by the death receptorFas/APO1/CD95 and its ligand, FasL3); and (b) apoptosis triggered bychanges in the homeostasis of mitochondria (for recent reviews, seeRefs. 2 and 3). For each of the two apoptotic pathways, a triggeringevent converges through specific signal transduction pathways ontoinitiation and execution machinery that is comprised of cascades ofproteolytic enzymes known as caspases (for reviews, see Refs. 4–6).Before an apoptosis-triggering event, caspases exist as dormant proen-zymes. The most apical “initiation” caspase that participates in thedeath receptor-dependent pathway is caspase-8 (7, 8). Death receptor-triggered autocatalytic processing of procaspase-8 releases the activecaspase-8 heterotetrameric enzyme, which, in turn, can directly proc-ess and activate the “executioner” caspase-3 (9, 10). Similarly, themost apical initiation caspase that participates in mitochondria-depen-dent cell death is caspase-9. Its autocatalytic processing (11) andactivation involve the participation of several coactivator molecules,one of which is cytosolic cytochromec (12). This protein, which isnormally loosely attached to the outer surface of the inner mitochon-drial membrane (for a review, see Ref. 13), is released from themitochondria through mechanisms that are still debated into thecytosol in response to apoptotic stimuli (for a review, see Ref. 3). Theproapoptotic protein Bax induces apoptosis by triggering the releaseof cytochromec from mitochondria (14–20). Activated caspase-9 candirectly process caspase-3 and its closest relative, caspase-7 (11). Theactivation of caspase-8/caspase-3 or caspase-9/caspase-3 cascadesensures the irreversibility of the apoptotic process. Ultimately, acti-vation of executioner caspases leads to the proteolytic cleavage ofnumerous cellular substrates, the activation of apoptosis-specificnucleases (21, 22) that lead to internucleosomal chromatin cleavage,and the development of a typical apoptotic morphology characterizedby nuclear pyknosis and fragmentation and by the blebbing of the cellplasma membranes. In the final step of apoptosis, the cell corpses areengulfed and destroyed by neighboring phagocytic cells.

Based on results obtained in recent knockout models, the apoptoticprogram in response to DNA damage appears to involve the mito-chondrial apoptotic pathway mediated by caspase-9 (23–25). How-ever, in certain Fas-expressing cells, anticancer drugs and radiationactivate the transcriptional induction of the gene for FasL that leads toincreased FasL expression. FasL further engages Fas and triggersFas-dependent, caspase-8-mediated cell death (26–28). Furthermore,the absence of a functional wild-type p53 tumor suppressor proteingenerally impedes the induction of apoptosis by DNA-damagingagents such as genotoxic drugs and ionizing or UV radiation (forreviews see Refs. 29 and 30), thus establishing an important role for

Received 9/30/99; accepted 2/2/00.The costs of publication of this article were defrayed in part by the payment of page

charges. This article must therefore be hereby markedadvertisementin accordance with18 U.S.C. Section 1734 solely to indicate this fact.

1 Supported by USPHS Grants CA-39360 and ES-08456 (to B. E. M.) and by an N. L.Tartar Research Fund Fellowship (to M. S. I.).

2 To whom requests for reprints should be addressed, at the Department of Cell andDevelopmental Biology, Oregon Health Sciences University, 3181 S.W. Sam JacksonPark Road, Mail Code L215, Portland, OR 97201. Phone: (503) 494-7811; Fax: (503)494-4253; E-mail: [email protected].

3 The abbreviations used are: FasL, Fas ligand; zVADfmk, benzyloxycarbonyl-Val-Ala-Asp(OMe) fluoromethyl ketone; polyIz polyC, polyinosinic z polycytidylic acid;DEVDase, caspase-3 or another caspase with caspase-3-like proteolytic specificity;PARP, poly(ADP)ribose polymerase; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling; TBS, Tris-buffered saline; FADD, Fas-associatingprotein with death domain.

1983

on May 5, 2020. © 2000 American Association for Cancer Research. cancerres.aacrjournals.org Downloaded from

http://cancerres.aacrjournals.org/

p53 in the DNA damage-induced apoptosis. However, the relation-ships between p53 and either of the apoptotic pathways is poorlyunderstood. Despite the incompletely understood mechanisms of ap-optosis induced by DNA-damaging chemicals and ionizing radiation,these agents constitute the predominant part of the available antican-cer therapy.

An entirely novel approach to induce cytotoxicity in target cancercells is based on the ability of the amphibian endoribonuclease(RNase) onconase (P-30 protein) to kill rapidly proliferating cells(with a certain preference for cancer cells) both in tissue culture andin the mouse (31–35). Onconase, originally isolated from oocytes ofRana pipiens(36), is a member of a growing family of extracellularcytotoxic RNases (for a review, see Ref. 37). The cytotoxic propertiesof onconase ultimately depend on its ability to enter target cells and onits RNA-hydrolyzing capacity (33). Recently, onconase, in combina-tion with doxorubicin, has been found to dramatically increase the lifespan of nude mice bearing human breast carcinoma cells (38). Inter-estingly, pancreatic RNase A, which is strongly homologous to on-conase, is not cytotoxic (39). The cellular protein RI, which is aninhibitor of ribonucleases, appears to possess a much greater affinityfor RNase A than for onconase (40), thus providing one possibleexplanation for the different cytotoxic abilities of the two RNases. Anonconase-based immunotoxin with increased tumor cell specificity isapproved for clinical trials.4 In the present work, we undertook toelucidate the intracellular mechanisms responsible for the cytotoxiceffects of onconase. We used a lipofection-mediated delivery ofenzyme into mammalian cells and asked whether it could inducecharacteristic features of apoptosis in these cells. We demonstrate herethat onconase treatment caused degradation of cellular tRNA but leftrRNA and mRNA apparently undamaged. Onconase induced a char-acteristic apoptotic pattern of cell death involving chromatin degra-dation, nuclear pyknosis and fragmentation, cell membrane blebbing,and activation of caspases-9, -3, and -7. The proapoptotic effects ofonconase did not require the presence of wild-type p53 and of theFasL/Fas/FADD/caspase-8 proapoptotic cascade. Although onconase-induced cytotoxicity correlated with onconase-induced inhibition ofprotein synthesis, we present evidence for the existence of an onco-nase-triggered apoptotic mechanism that is likely to be independent ofthe inhibition of protein synthesis. Finally, we demonstrate that on-conase-induced cell death and caspase-9 activation are accompaniedby unusually little release of cytochromec from mitochondria and alack of detectable translocation of Bax from the cytosol onto themitochondria, suggesting that Bax- and cytochromec-independentmechanisms of caspase-9 activation might be involved in mediatingonconase cytotoxicity.

MATERIALS AND METHODS

Chemicals. The source, storage, methods of application, and modes ofaction of emetine and cycloheximide have been described (41). The caspaseinhibitor zVADfmk was from Calbiochem, and it was stored as a 50 mM

(10003) stock solution in (H3C)2SO at220°C. Whenever zVADfmk pretreat-ment was applied, the corresponding control cell received 1ml of (H3C)2SOper milliliter of medium. Lipofectin Reagent was from Life Technologies, Inc.PolyI z polyC was from Calbiochem. All commonly used chemicals, digitonin(D-5628), and acridine orange were from Sigma Chemical Company. Allradiochemicals were from DuPont New England Nuclear Research Products.

Cell Culture. HeLa tk2 cells were maintained in DMEM supplementedwith 10% calf serum (HyClone, Logan, UT). All experiments presented herewere performed with logarithmically growing cultures that had not reachedmore than 50% confluence. For this purpose, cells were plated 18–24 h beforetreatment at a density of 2.63 106, 5 3 105, or 2 3 105 cells/plate (or well)

in 10-cm plates, 6-well plates, or 12-well plates, respectively. The p530/0

fibroblast cell line was derived from p530/0 mouse primary embryonic fibro-blasts (a generous gift from Dr. Markus Grompe) through spontaneous im-mortalization. The absence of p53 in these cells at both allele and protein levelswas verified by PCR and Western blot analyses, respectively, and it will bedescribed elsewhere.5

Onconase Preparation.Native onconase was purified fromRana pipiensoocytes (Nasco, Fort Atkinson, WI) following the published protocol (36) asdescribed previously (42).

Lipofectin-mediated Delivery of Onconase.Delivery of onconase withLipofectin was performed in DMEM containing 0.5% calf serum. Lipofectin/onconase mixes were prepared following the procedure outlined for the deliv-ery of diphtheria toxin anda-sarcin in Ref. 41. Before the application of theLipofectin/onconase mixes, the cells were washed once with serum-freeDMEM. Four h after the addition of the Lipofectin/onconase mix, the cellswere fed calf serum to a final concentration of 10%.

Antibodies. Antibodies against PARP (H-250, sc-7150, and A-20, sc-1562), cytochromec (H-104, sc-7159), Bax (N-20, sc-493HRP), rabbit IgG(sc-2004), and mouse IgG (sc-2005) were from Santa Cruz Biotechnology.Antibodies against caspase-9 (#66571A), caspase-8 (#66231A), caspase-7(#66871A), caspase-3 (#65906E), and caspase-1 (#66441A) were from PharM-ingen. The activating (CH11, #05-201) and blocking (ZB4, #05-338) antibod-ies against human Fas were from Upstate Biotechnology.

Measurement of Protein Synthesis via [3H]Leucine Incorporation. In-corporation of [3H]leucine was performed as described previously (40), exceptthat the cells were not subjected to leucine deprivation before onconasetreatment and [3H]leucine pulse labeling. TwomCi of [3H]leucine/ml mediumwere used for labeling.

Detection of Apoptosis.For detection of apoptosis using TUNEL, cellswere plated at low density on Thermanox slides (Nunc) and treated as indi-cated in the text, and apoptotic cells were detected using theIn SituCell DeathDetection kit from Boehringer Mannheim following the instructions of themanufacturer. For detection of apoptosis by morphological criteria, cells wereplated on 6-well plates and treated with onconase as described in the text or inthe figure legends. At the indicated times after the treatment, the cells werewashed twice with PBS to remove dead cells. Cells that remained attached tothe plates were fixed in a solution containing 95% ethanol and 5% acetic acidfor 15 minutes at RT. The cells were then rehydrated with several washes ina sodium phosphate buffer [10 mM (pH 7.9)] for 1 h. The cells were stained byapplying a 10mg/ml acridine orange solution in sodium phosphate buffer (pH7.9) for 1 h, followed by extensive washing with sodium phosphate buffer (pH7.9). Visualization of the incorporated acridine orange into nuclear DNA wasachieved in fluorescent microscopy using an ARC Lamp UV source (LudlElectronic Products Ltd., Hawthorne, NY) and an IM35 Zeiss microscope.Phase-contrast viewing using the same microscope was done with the in-builtvisual light source. Photographs of cell were taken with a CONTAX 167MTcamera (Kyocera Co., Tokyo, Japan) and Kodak 100 Elite Chrome film(Eastman-Kodak). For DNA fragmentation analysis, cells were treated, har-vested, and lysed, and nuclei were sedimented as described below for prepa-ration of nuclear extracts. The nuclear pellets were resuspended in 50 mM

Tris-HCl (pH 7.8), 10 mM EDTA, and 0.5% (w/v) sodiumN-lauroyl sarcosi-nate. DNase-free RNase A (Sigma) was added (f.c. 0.5 mg/ml) for 20 min at50°C. Proteinase K (Life Technologies, Inc.) was added (f.c. 0.5 mg/ml) for anadditional 50 min at 50°C. The samples were then resolved in 2% nondena-turing agarose gels. DNA was visualized using ethidium bromide and UVtransillumination.

Measurement of Cell Death/Survival. For the 24-h death assay, a modi-fication of the technique of Goillotet al. (43) was used. Briefly, HeLa cellswere plated in 12-well plates. Eighteen to 24 h later, the cells were treated asdescribed in the text or in the figure legends. Four h after the treatment, thecells were supplemented with calf serum to 10%. Twenty-four h after thetreatments, the cells were washed twice with PBS to remove dead cells. Cellsthat remained attached to the plates were simultaneously fixed and stained for10 min by the addition of a solution containing 0.05% crystal violet, 20% (v/v)ethanol, 0.37% (v/v) formaldehyde, 80% (v/v) H2O. The wells were thenrinsed extensively with water and dried. Crystal violet was extracted for 1 h

4 D. L. Newton and S. M. Rybak, unpublished results. 5 M. S. Iordanov and B. E. Magun, manuscript in preparation.

1984

ONCONASE-INDUCED APOPTOSIS



from the cells by addition of 2 ml methanol/well and vigorous shaking. Theabsorbance of the extract was measured at 570 nm. After subtracting thebackground staining from plates containing only growth medium but not cellsand processed as described above, the absorbance of control cells was taken toindicate a 100% survival, and the percentage cell survival of the treated cellswas calculated accordingly. Percentage cell death was calculated as follows:100 2 percent survival.

Preparation of RNA. Total RNA containing the fraction of tRNA wasprepared as follows. Cells were treated as described in the text or in the figurelegends. At the indicated times after treatment, the cells were washed twice inice-cold PBS, scraped in ice-cold PBS, and centrifuged at 16,0003 g for 1 minat 4°C. The cellular pellet was resuspended in 375ml of a solution containing10 mM Tris-HCl (pH 7.0), 150 mM NaCl, and 1 mM EDTA; thereafter, 26mlof 10% (v/v) NP40 were added to lyse the cells for 2 min on ice. Nuclei weresedimented at 16,0003 g for 30 s at 4°C, and 375ml of a solution containing20 mM Tris-HCl (pH 7.8), 350 mM NaCl, 20 mM EDTA, and 1% (v/v) SDSwere added to the supernatant. One extraction with 750ml of phenol/chloro-form/isoamyl alcohol (25;24;1) and one extraction with chloroform/isoamylalcohol (24;1) were performed and RNA from the aqueous phase was precip-itated with 2 volumes of ethanol. RNA precipitate was dissolved in RNase-freewater, and an equal volume of a 23RNA-loading solution [50% (v/v)formamide, 6% (v/v) formaldehyde, 20% (v/v) glycerol, 2 mM sodium phos-phate buffer (pH 7.0), and 10mg/ml ethidium bromide without bromphenolblue] was added. The RNA samples were denatured by heating for 10 min at85°C.

Northern Blot Analysis. RNA samples were resolved electrophoreticallyin either 1% or 4% denaturing agarose gels (44) and transferred onto Hy-bond-N membrane (Amersham Life Science). Hybridization with radioactivelylabeled DNA probes was done using the ExpressHyb hybridization solution(Clontech) following the manufacturer’s instructions. The hybridization probefor tRNALys was the oligonucleotide 59-CTGAGATTAAGAGTCTCAT-GCTCTACCGACTGAGCTAGCC-39(Life Technologies, Inc.). The hybrid-ization probe for cyclophilin is described elsewhere (44).

Reverse Transcription of rRNA by Primer Extension. Probing for dam-age to 28S rRNA by reverse transcriptase-mediated primer extension was doneas described previously (40). The following two primers (45) were used (a)59-CCCACAGATGGTAGCT-39; and (b) 59-CGACATCGAAGGATCA-39.

Preparation of Nuclear and Cytosolic Extracts for Western Blot Anal-ysis. The following protocol applies to the experiments presented in Figs. 4,6B, 7A, and 7B.Cells were treated as described in the text and in the figurelegends. At the indicated times, both attached and detached cells were col-lected by scraping directly in the medium, sedimented at 16,0003 g for 1 minat 4°C, washed by resuspending in ice-cold PBS, and resedimented. Cellularpellets were resuspended in 100ml of a solution containing 10 mM HEPES-KOH (pH 7.9), 60 mM KCl, 1 mM EDTA, 0.5% (v/v) NP40, 1 mM DTT, andComplete Protease Inhibitor mixture (Roche Molecular Biochemicals, Indian-apolis, IN) and lysed for 5 min on ice. Nuclei were sedimented at 7353 g for5 min and resuspended in a solution containing 250 mM Tris-HCl (pH 7.8), 60mM KCl, 1 mM DTT, Complete Protease Inhibitor mixture, whereas thepost-nuclear supernatants were designated “cytosolic extracts” after removal ofcellular debris at 16,0003 g for 10 min at 4°C. For extraction of nuclearproteins, the salt concentration was elevated to 500 mM KCl, and the proteinswere extracted at 4°C for 30 min with vigorous agitation, followed by removalof nuclear envelopes at 16,0003 g for 10 min at 4°C.

Western Blot Analysis. Nuclear or cytosolic protein extracts were re-solved by SDS-PAGE and transferred onto polyvinylidene difluoride mem-brane (Millipore). After the transfer, the nonspecific antibody-binding activi-ties of the membranes were blocked in Tris-buffered saline (TBS, 10 mM

Tris-HCl, (pH 8.0), 150 mM NaCl) containing 0.3% (v/v) Tween-20 and 10%(w/v) nonfat dry milk for 30 min at room temperature. Incubations withappropriate dilutions of primary and secondary antibodies were done for 60and 45 min, respectively, in TBS containing 0.3% Tween-20 and 5% nonfatdry milk, followed by extensive washes with TBS containing 0.3% Tween-20.Detection of immunolabeled proteins was done using the Renaissance chemi-luminescence kit (DuPont New England Nuclear Research Products) andHyperfilm enhanced chemiluminescence film (Amersham).

Preparation of Mitochondria. For the detection of cytochromec pre-sented in Fig. 7C, mitochondria were prepared and analyzed as described inBossy-Wetzelet al. (46). For the combined analysis of Bax and cytochromec

shown in Fig. 8A, the procedure described in Saikumaret al. (19) was applied,with modifications. Control and appropriately treated cells were harvested byscraping directly in the medium, sedimented, washed once in ice-cold PBS andonce in isotonic sucrose buffer [SU buffer-250 mM sucrose, 10 mM HEPES, 10mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA (pH 7.1)] and thenpermeabilized for 1 min at room temperature in SU buffer containing 0.025%digitonin and Complete Protease Inhibitor mixture. The efficiency of perme-abilization was nearly 100% as measured by trypan blue exclusion. Aftersedimenting the cellular pellet, the supernatant was designated “cytosolicfraction.” The pellet of permeabilized cells was resuspended in SU buffercontaining 0.5% Triton X-100 and Complete Protease Inhibitor mixture andkept for 10 min on ice, after which the Triton X-100-soluble and -insolublefractions were separated by centrifugation. All three fractions were resolved in15% SDS-PAGE, transferred onto polyvinylidene difluoride membrane, andprobed with appropriate antibodies as shown in Fig. 8A.

RESULTS

tRNA Is a Specific Target for Onconasein Vivo. Preliminarystudies have demonstrated that the cationic lipid vehicle Lipofectinfacilitated the delivery of onconase into a variety of cell lines, includ-ing HeLa cells (measured by cytotoxicity, see below). To determinethe relevantin vivo target(s) for onconase, we treated HeLa cells withLipofectin and onconase (1mg/ml, a concentration that causes;75%cell killing within 24 h of treatment, see below). Immediately after,2 h after, or 4 h after the treatment with onconase, the cells wereharvested, and total cellular RNA was prepared and resolved electro-phoretically in either 1% or 4% denaturing agarose gels. Hydrolysis of28S or 18S rRNA was undetectable by ethidium bromide visualizationin the 1% gel (Fig. 1,Lanes 4–6). We also applied a highly sensitivemethod (see “Materials and Methods”) based on the lesion-inducedarrest of a reverse transcriptase-driven primer extension to detectsingle-nucleotide damages to domains V and VI of 28S rRNA (41,45). These domains are directly involved in translational elongationand are the most common targets for fungal antibiotics and polypep-tide ribotoxins (see Ref. 41 and references therein). Using two primersthat allow the scanning of an approximately 1-kb region within thesedomains, we failed to detect any sites that were susceptible to cleav-age by onconase (data not shown). We therefore concluded that rRNAis an unlikely candidate for an onconase targetin vivo.

During the course of the former experiments, we noticed that a bandof RNA with a mobility greater than the 28S and 18S rRNAs dis-played a graded decrease in ethidium bromide staining after treatmentwith onconase (Fig. 1,Lanes 4–6). The same band was furtherresolved in a 4% gel into three distinct RNA species, of which onlythe species with the highest mobility was significantly affected byonconase treatment (Fig. 1,Lanes 7–9). Based on the apparent mo-bilities of these three bands, we concluded that they represent the 5.8SrRNA, the 5S rRNA, and the population of tRNAs, respectively. Toconfirm that the band that decreased in abundance after onconasetreatment of cells corresponded to tRNA, we performed Northern blotanalysis of the same RNA preparations using a radioactively labeled40-mer DNA oligonucleotide probe complementary to the humantRNALys. Indeed, at 2 and 4 h after exposure of cells to onconase, therelative abundance of tRNALys was decreased to 68% and 55% of thecontrol cells, respectively (Fig. 1,Lanes 1–3). Furthermore, probingof the same membrane with a probe specific for cyclophilin mRNArevealed a single hybridizing band and an absence of degradationproducts corresponding to these mRNAs (Fig. 1,Lanes 10–12). Theabundance and integrity of three other nonrelated mRNAs (c-jun,c-fos, and surviving mRNAs) was assessed; these mRNAs also dis-played a single hybridizing band and an absence of degradationproducts (data not shown).

1985




Onconase-induced Cytotoxicity.Although onconase is a cyto-toxin with intrinsic internalization capacity, its translocation acrossthe cellular membrane is the rate-limiting step of cytotoxic action(33). We avoided this potential complication by Lipofectin-mediateddelivery of onconase into all cell types used in this work. To deter-mine the ability of Lipofectin-delivered onconase to kill HeLa cells,the cells were treated with Lipofectin alone or with onconase (0.01,0.1, or 1mg/ml) and Lipofectin, and the cell survival was measured24 h later using crystal violet staining as described in “Materials andMethods.” Fig. 2 demonstrates that onconase caused cell death thatcorrelated strongly with the logarithm of onconase concentration; thehalf-maximal killing concentration (IC50) for onconase in HeLa cellswas;0.05mg/ml (Fig. 2; Table 1, column C). In contrast, onconaseadded without Lipofectin to the medium of cells at concentrations ofup to 30mg/ml was not cytotoxic (data not shown).

A potential target for the action of a ribonuclease is the process ofprotein synthesis. We therefore determined, in parallel to the deter-mination of the cell killing, the effect of onconase on translation. Cellswere pulse-labeled with [3H]leucine between 2.5 and 3 h after onco-nase treatment. As for the killing, onconase caused an inhibition ofincorporation of [3H]leucine that correlated with the logarithm ofonconase concentration; the IC50 for inhibition of protein synthesiswas ;0.1 mg/ml (Fig. 2; Table 1, column A). Correlation analysis(percentage of inhibition of protein synthesisversuspercentage ofkilling) revealed that at IC50 for inhibition of protein synthesis,onconase kills 656 5% of the cells in 24 h (Table 1, column E).

Can the cytotoxic potential of onconase result entirely from theinhibition of protein synthesis? We reasoned that if this were the case,a similar correlation between cytotoxicity and protein synthesisshould be observed for other inhibitors of protein synthesis. To testthis hypothesis, we performed the same correlation analysis (transla-tion versussurvival) for emetine, a potent and irreversible inhibitor ofribosomal translocation (see Ref. 41 and references therein). Weobserved reproducibly that at similar levels of inhibition of proteinsynthesis, onconase is significantly more cytotoxic. For instance, atIC50 for inhibition of protein synthesis (;0.04mg/ml; Table 1, col-

umn A), emetine kills merely 156 5% of the cells in 24 h (Table 1,column E). Although not as pronounced, similar results were obtainedusing another antibiotic inhibitor of protein synthesis, cycloheximide(Table 1, column E). These results are consistent with the notion thata part of the cytotoxic action of onconase results from onconase-triggered death mechanisms other than inhibition of protein synthesis(see below).

Onconase Triggers an Apoptotic Cell Death Program.To iden-tify and characterize the mode of cell death that is triggered by

Fig. 1. Differential damage to tRNA, rRNA, or mRNA by onconaseinvivo. HeLa cells logarithmically growing in 10-cm plates were treatedwith 1 mg/ml onconase in Lipofectin (12.5mg/ml)-containing DMEM/0.5% serum for the indicated times. (Hereafter, “onconase treatment” willalways indicate treatment in the presence of 12.5mg/ml Lipofectin.) Noapparent loss of cells was detectable at any of these time points. Forpreparation of RNA, the cells were washed twice with ice-cold PBS andprocessed as described in “Materials and Methods.” Equal volumes ofsamples were resolved in either 1% or 4% denaturing agarose gels, andRNA was visualized by means of ethidium bromide contained in the RNAsample and UV transillumination. The 1% gels were transferred onto amembrane and probed in Northern blot analysis with the indicated hy-bridization probes for tRNALys and cyclophilin as described in “Materialsand Methods.”

Fig. 2. Inhibition of protein synthesis and cytotoxicity induced by onconase. HeLa cellsgrowing in 12-well plates were treated with 0, 0.01, 0.1, or 1mg/ml onconase. Two h and30 min after the treatment, half of the plates were labeled with 2mCi/ml [3H]leucine foranother 30 min, after which time [3H]leucine incorporation was determined as describedpreviously (41). The incorporation of [3H]leucine in the control cells (hereafter, “controlcells” will always indicate Lipofectin-treated cells, unless otherwise specified) is shownas 100%.Error bars represent SD from experimental points in triplicates. Twenty-four hafter the treatment, the other half of the plates was used to determine cell survival usingcrystal violet staining as described in “Materials and Methods.” The survival of the cellsin the control cells is shown as 100%.Error bars represent SD from experimental pointsin triplicates.

1986




onconase treatment, we sought to determine whether onconase-treatedcells display characteristic features of apoptosis. Twenty-four h afterthe onconase treatment, HeLa cells displayed a characteristic internu-cleosomal chromatin cleavage (Fig. 3A, Lane 2). Furthermore, treat-ment of HeLa cells with onconase induced a distinct pattern of nuclearpyknosis and fragmentation (karyorrhexis) as viewed by fluorescencemicroscopy after acridine orange staining (Fig. 3B). Treatment withonconase also induced a characteristic blebbing of the cellular mem-brane. Membrane blebbing in HeLa cells was detectable as early as4 h after treatment and continued to be evident at 16 h after the

treatment (data not shown for HeLa cells, but see Fig. 3Cfor p530/0

mouse fibroblasts). We next investigated whether the mode of onco-nase-induced cell death involves the activation of an apoptosis-specific DEVDase activity. To this end, we tested for the appearancein the nuclear fractions of HeLa cells of specific cleavage products ofthe enzyme PARP, a target for caspase-3 (47, 48). Fig. 4A (top)demonstrates that onconase treatment caused a dose-dependent dis-appearance of theMr 116,000 full-size PARP and a concomitantaccumulation of theMr 89,000 cleavage product. In conclusion, theability of onconase to induce internucleosomal chromatin fragmenta-

Table 1 Correlation between onconase-, emetine-, or cycloheximide-induced inhibition of protein synthesis and cell death in the absence or presence of zVADfmk

HeLa cells growing in 12-well plates were treated with 0, 0.01, 0.1, or 1mg/ml onconase, either in the absence or in the presence of 50mM zVADfmk added 1 h before onconase.Two h and 30 min after the onconase treatment, half of the plates were labeled with 2mCi/ml [3H]leucine for another 30 min, after which time [3H]leucine incorporation was determinedas described in Fig 2. Twenty-four h after the treatment, the other half of the plates was used to determine cell survival using crystal violet staining as described in “Materials andMethods.” Both protein synthesis and cell killing were plotted as functions of the concentration of onconase and IC50 values (6SD from experimental points in triplicates) for eithereffect were calculated using DeltaGraph Professional software from Deltapoint, Inc. (columns A–D). Plotting percentage (of control) cell killingversuspercentage (of control)translation allowed the calculation of percentage killing at IC50 for inhibition of translation (columns E and F). Identical experimental set-up and calculations were performed for emetineusing 0, 0.03, 0.1, or 0.3mg/ml emetine and for cycloheximide using 0, 0.001, 0.01, 0.1, 1, 10, or 100mg/ml cycloheximide. Not shown: emetine and cycloheximide have been appliedboth without and with Lipofectin (to match the experimental conditions used for onconase) with identical outcome.

IC50 translation (mg/ml) IC50 killing (mg/ml) Killing at 50% inhibition of translation

2 1zVADfmk 2 1zVADfmk 2 1zVADfmk

A B C D E F

Onconase 0.1a 0.1a 0.05a 1.0a 65%b 35%b

Emetine 0.04a 0.04a 0.4a 1.8a 15%b 15%b

CHXc 0.1a 0.2a 0.6a 40a 35%b 25%b

a SD ¶ 10%.b SD ¶ 5%.c CHX, cycloheximide.

Fig. 3. Hallmarks of apoptosis triggered by onconase.A, HeLa cellswere treated, where indicated, with 1mg/ml onconase. Twenty-four hlater, genomic DNA was prepared from the nuclei as described in“Materials and Methods.” Equal volumes of DNA samples were re-solved in nondenaturing 2% agarose gels, and DNA was visualized bymeans of ethidium bromide contained in the gel and UV transillumi-nation. Chromatin fragments resulting from internucleosomal cleavageare indicated byarrowheads. B, HeLa cells were treated, where indi-cated, with 1mg/ml onconase for 20 h, after which time the cells werestained with acridine orange, studied by fluorescence microscopy, andphotographed as described in “Materials and Methods.” A representa-tive case of nuclear pyknosis and fragmentation (karyorrhexis) isshown (inset). Magnification,364 (3256 for theinset). C, logarith-mically growing p530/0 cells were treated, where indicated, with 1mg/ml onconase for 16 h, after which time phase-contrast microscopyof living cells was used to detect membrane blebbing (inset). Magni-fication, 364 (3256 for theinset).

1987




tion, karyorrhexis, membrane blebbing, and activation of caspasessupported the notion that onconase actively engages the cellularapoptotic machinery and triggers apoptotic death.

Onconase and Emetine Trigger Apoptosis with Different Ki-netics. We have previously reported that emetine (at a concentrationof 100 mg/ml) inhibits protein synthesis in mammalian cells com-pletely ($98%), instantaneously (within the first minute after addi-tion), and irreversibly (41). Therefore, emetine-induced apoptosis(Fig. 4B) represents, in its purest form, the mode of apoptosis trig-gered by inhibition of protein synthesis. When studied using TUNEL,emetine-treated HeLa cells did not display TUNEL-positive (apo-ptotic) cells at 6 or 8 h after treatment; TUNEL-positive cells weredetected only at 10 h after treatment (Fig. 5A). In contrast, a substan-tial portion of onconase-treated cells were TUNEL-positive as early as6 h after treatment (Fig. 5A). The observed early TUNEL positivity ofonconase-treated cells was not caused by a deoxyribonuclease(DNase) activity (or contamination) of the onconase preparation be-cause we failed to detect such activity (or contamination) under fourdifferent experimental conditionsin vitro (Fig. 5B). Because theability of onconase to inhibit protein synthesis has neither the potencynor the immediacy so characteristic for emetine, the results shown inFig. 5Aare consistent with the notion that the early onconase-inducedapoptosis (e.g., at 6 h after treatment) does not result entirely frominhibition of translation (see “Discussion”).

Onconase-induced Apoptosis Does Not Require p53.In general,the absence of a functional wild-type p53 protein impedes the induc-tion of apoptosis by certain DNA-damaging agents such as genotoxicdrugs and ionizing or UV radiation (for reviews, see Refs. 29 and 30).Although the wild-typep53 gene is present in HeLa cells (49), thelevels of p53 protein are kept very low in these cells due to its rapiddegradation through the ubiquitin/proteasome pathway (50, 51). Toaddress the possible requirement for a functional p53 protein inonconase-induced apoptosis, we used a mouse fibroblast cell linederived from p530/0 embryonic fibroblasts. These p530/0 cells areresistant to both UV radiation- and doxorubicin-induced cell death.5

However, when treated with onconase, the p530/0 cells displayedmassive cell blebbing (Fig. 3C), nuclear pyknosis, DEVD-specificcaspase activity, DNA fragmentation, and massive cell death (Fig. 3Cand data not shown). Importantly, we did not observe significantdifferences between the responsiveness of p530/0 and p531/1 mousefibroblasts to onconase (data not shown). These experiments conclu-sively demonstrated that the presence of a functional p53 protein isnot required for onconase-induced apoptosis.

Partial Resistance of Onconase-induced Apoptosis andCaspase Activity to Inhibition by the Caspase Inhibitor zVAD-fmk. When treated with the anti-Fas agonistic antibody CH11,HeLa cells undergo a massive apoptosis within a 24-h period aftertreatment (Fig. 6A). This cell death is accompanied by a markedincrease in DEVDase activity as measured by the specific cleavageof the p116 PARP, generating the two fragments p89 and p27 (Fig.6B, compareLanes 1and3). Both cell death and caspase activityinduced by CH11 could be completely prevented by pretreatmentof cells with the nonspecific caspase inhibitor zVADfmk (Fig. 6,AandB, compareLanes 3and4). Because the CH11 dose used forthese experiments (0.5mg/ml) was approximately 10 times higherthan the one required for maximum killing and PARP cleavage(data not shown), a conclusion can be safely reached that zVAD-fmk (at the concentration used) possesses a complete ability toprevent Fas-mediated apoptosis and caspase activity. However,when applied to pretreat cells before onconase treatment, the effectof zVADfmk was markedly different. zVADfmk was not able tocompletely prevent cell death but caused a shift to the right in theIC50 of onconase by more than 1 log (Table 1, compare columns Cand D). As expected, this IC50 shift was not caused by an effect ofzVADfmk on the ability of onconase to inhibit protein synthesis(Table 1, compare columns A and B). Rather, the IC50 shiftcorrelated well with a partial inhibition in onconase-induced DEV-Dase activity (Fig. 4A, compare top and bottom). Correlationanalysis revealed that at IC50 for inhibition of protein synthesis,onconase kills only 35% of the zVADfmk-pretreated cells, com-pared with 65% cell killing in the absence of zVADfmk (Table 1,compare columns E and F).

Does zVADfmk interfere with the part of onconase-inducedcytotoxicity that results from inhibition of protein synthesis? Toaddress this question, we investigated the effect of zVADfmk onemetine-induced cell death. First, we observed that a concentrationof emetine that caused;70% killing (0.3 mg/ml; data not shown)also caused DEVDase activity, as measured by PARP cleavage(Fig. 4B, top, Lane 4). This DEVDase activity was due to specificcaspase activation because zVADfmk almost completely abolishedthe cleavage of PARP in response to emetine (Fig. 4B, bottom,Lane 4). However, in contrast to the effect of zVADfmk ononconase-induced cytotoxicity, the caspase inhibitor caused only amodest (4.5-fold) increase in the killing IC50 for emetine (Table 1,columns C and D) and failed to significantly affect the cell killinginduced by emetine at IC50 for translation (Table 1, columns E andF). Taken together, the results presented in Figs. 2 and 4 and Table

Fig. 4. Induction of DEVDase activity in onconase- or emetine-treated cells in theabsence or the presence of zVADfmk. Cells were treated in parallel and identically tothose in Table 1, except that they were grown in 6-cm tissue culture plates. Twenty-fourh after the treatments, the cells were harvested, and nuclear proteins were extracted (see“Materials and Methods”). Twentymg protein/lane were resolved in 7.5% SDS-PAGEgels, transferred onto polyvinylidene difluoride membrane, and probed in an immunoblotassay with an anti-PARP antibody (see “Materials and Methods”).

1988




1 are consistent with the following two notions: (a) cell deathcaused by classical inhibitors of protein synthesis (emetine andcycloheximide) requires a significant inhibition of the cellulartranslation machinery, and, despite the appearance of activecaspases, this cell death can proceed in the absence of caspaseactivity as well; and (b) onconase-induced cell death appears toinvolve a component that is probably independent of inhibition ofprotein synthesis and that is zVADfmk sensitive (see Fig. 9 and“Discussion”).

Onconase-induced Apoptosis Is Not Mediated by Engaging theFasL/Fas Cell Death Cascade.Recent studies have demonstratedthat apoptosis in Fas-expressing cells subjected to DNA-damagingagents is triggered in part through drug- or radiation-induced activa-tion of FasL production and cell surface presentation, followedby autocrine ligation of Fas (26–28). The results obtained usingzVADfmk suggested that the mechanisms of onconase-induced celldeath are different from those used by Fas ligation. However, thehypothesis could not be ruled out without directly determiningwhether onconase-induced apoptosis could be mediated in part byonconase-stimulated production of FasL and autocrine ligation of Fas.To test this hypothesis, we used an antagonistic anti-Fas monoclonalantibody, ZB4, that binds to Fas and prevents Fas agonists (such asFasL or CH11) from engaging the receptor and triggering apoptosis(52). Fig. 6C shows that preincubation of HeLa cells with ZB4

abrogated the cell death induced by CH11, whereas Fig. 6D demon-strates that the same pretreatment completely failed to inhibit onco-nase-induced apoptosis. We therefore concluded that onconase-in-duced apoptosis does not utilize the FasL/Fas death signalingpathway.

Procaspases-9, -3, and -7, But Not Procaspase-1 or -8, AreProcessed in Onconase-treated Cells.To determine the caspasesthat are activated by onconase, we prepared cytosolic extracts fromHeLa cells that were treated with Lipofectin alone or with Lipo-fectin and onconase for 0, 24, or 48 h and studied the processingof procaspase-1, -8, -9, -3, and -7. The use of caspase-1 as anegative control was determined by the fact that this caspase doesnot seem to be involved in mediating apoptosis (53–55). The(auto)processing of caspase-8 and -9 was detected by the decreasedintensity in Western blot analysis of the respective band corre-sponding to the full-size procaspases. As expected, 24 h afteronconase treatment, no significant caspase-1 cleavage was ob-served (Fig. 7A, compareLanes 1and 2 with Lanes 3and 4).Similarly, both 24 h (data not shown) and 48 h after onconasetreatment, the majority of caspase-8 was in the procaspase form(Fig. 7B, Lanes 3and 4). In contrast, there was a massive proc-essing of procaspase-8 in cells treated for 48 h with CH11 (Fig. 7B,Lanes 1and 2). Procaspase-9, however, displayed onconase-in-duced processing that was detectable both 24 and 48 h after the

Fig. 5. Early apoptotic changes in cells treated with onconase, butnot with emetine.A, cells plated at low density on Thermanox slideswere treated with either emetine (100mg/ml) or onconase (1mg/ml),both in the presence of Lipofectin (12.5mg/ml). At the indicated timesafter treatment, apoptotic cells were detected using TUNEL.B, absenceof DNase activity in the preparation of onconase used. Onemg ofsupercoiled pcDNA3 plasmid was incubated for 1 h at 37°C in theabsence (Lane 1) or presence (Lanes 2–5) of 10mg/ml onconase in fourdifferent buffer conditions optimized for various restriction endonucle-ases (restriction digest buffers 1–4 from Life Technologies, Inc.) andthen resolved electrophoretically in an 0.8% agarose gel.Lane 6dem-onstrates the linearization of the same plasmid by theEcoRI restrictase.

1989




treatment (Fig. 7A, compareLanes 1–3with Lane 4and Fig. 7B,compareLanes 3 and 4). Procaspase-9 was also processed inresponse to treatment of cells with CH11 (Fig. 7B, Lanes 1and2).Active caspase-9 can directly process two DEVDases, caspase-3and its closest relative, caspase-7 (11); therefore, if the processingof procaspase-9 observed in onconase-treated cells leads to thegeneration of active caspase-9, then a subsequent processing of

caspase-3 and -7 should be expected. Indeed, cells treated withonconase displayed processing of the procaspase forms and accu-mulation of the characteristic COOH-terminalMr 17,000 frag-ments that result from the specific processing of theMr 32,000procaspase-3 and theMr 35,000 procaspase-7, respectively (Fig.7A, Lane 4; data not shown for caspase-7). This proteolytic proc-essing of procaspase-3 and -7 is likely to generate the active

Fig. 6. Effects of the caspase inhibitor zVADfmk and of theFas-blocking antibody ZB4.A, HeLa cells growing in 12-wellplates were treated, where indicated, with the anti-Fas agonisticantibody CH11 (0.5mg/ml), in either the absence or presence,where indicated, of 50mM zVADfmk added 1 h before CH11.Twenty-four h after the treatment, cell survival was determinedusing crystal violet staining as described in “Materials and Meth-ods.” The survival of the cells in the control cells is shown as 100%.Error bars represent SD from experimental points in triplicates.B,HeLa cells were treated as described inA, except that the cells werein 6-cm plates. Twenty-four h later, PARP cleavage was studied asdescribed in Fig. 4, except that in addition to the p89 proteolyticfragment, the p27 proteolytic fragment was also detected.C andD,HeLa cells grown in 12-well plates were either not pretreated orpretreated, where indicated, for 1 h with the ZB4 antibody (1mg/ml). Cells were then either left untreated or treated with CH11(0.5 mg/ml), with Lipofectin alone or with onconase (1mg/ml) andLipofectin; all pretreatments and treatments were performed in sucha way that the final concentration of ZB4 remained unchangedthroughout the duration of the experiment. Twenty-four h later, thecell survival was measured using crystal violet staining as describedin “Materials and Methods.”Error bars, SD from experimentalpoints in triplicates. The presence of Lipofectin affected neither theability of CH11 to kill HeLa cells nor the ability of ZB4 to preventCH11-induced cell killing (data not shown).

Fig. 7. Processing of procaspase-9 and -3, but not of procaspase-1and -8, and lack of massive loss of mitochondrial cytochromec inonconase-treated HeLa cells.A and B, HeLa cells were treated withLipofectin without or with onconase (1mg/ml), for 0 or 24 h (A) or for48 h (B), after which time the cells were harvested, and cytoplasmicextracts were made as described in “Materials and Methods.” Tenpercent SDS-PAGE gels were used for the immunoblot detection ofprocaspase-1, -8, and -9, and 15% SDS-PAGE gels were used for thedetection of procaspase-3. For the detection of procaspase-1, -8, and -3,10 mg of total protein were used per lane, and for the detection ofprocaspase-9, 50mg of total protein were used per lane.C, cells weretreated, where indicated, with either CH11 (0.5mg/ml) or onconase (1mg/ml). Twenty-four h after the treatment, cytosolic and mitochondrialfractions were prepared as described in “Materials and Methods.”Twenty mg of total protein were resolved in 15% SDS-PAGE gels,transferred onto a polyvinylidene difluoride membrane, and probed inan immunoblotting assay with an anti-cytochromec antibody as de-scribed in “Materials and Methods.”

1990




caspase-3 and -7 because DEVDase activities were detected bymeans of PARP cleavage in onconase-treated cells (Fig. 4A). Thus,procaspase-9, -3, and -7, but not procaspase-1 and -8, appeared tobe activated by onconase.

Lack of Massive Release of Cytochromec from Mitochondriainto the Cytosol and of Translocation of Bax from the Cytosolonto Mitochondria in Response to Onconase.Release of cyto-chromec from mitochondria is a key event for initiating a caspase-9/caspase-3 cascade and for amplifying the effectiveness of Fas liga-tion through a caspase-8-dependent activation of caspase-9 (56–58).Most notably, cytochromec is a cofactor for caspase-9 activation (12).We therefore investigated whether cytochromec (normally entirelysequestered in the mitochondria) is released into the cytosol ononconase treatment. Twenty-four h after treatment of HeLa cells withonconase, there was a detectable amount of cytochromec in thecytosolic fraction compared to the absence of detectable cytochromec in the Lipofectin-treated cells (Fig. 7C,top, compareLanes 3and4).Consistently, however, significantly less cytochromec was releasedfrom mitochondria into the cytosol in response to onconase than inresponse to Fas ligation (Fig. 7C,top, compareLanes 2and 4) andother proapoptotic stimuli including actinomycin D or UV radiation(both not shown; all agents were applied at doses that triggered similarlevels of cell death). To find a possible explanation for the low levelof cytochromec release from mitochondria in onconase-treated cells,we studied the subcellular distribution of the proapoptotic protein Baxin control and onconase-treated cells. Recent studies have demon-strated that the transition of Bax from a soluble (cytosolic) form to amitochondrial membrane-inserted form is sufficient and possibly re-quired for the release of cytochromec from apoptotic mitochondria,

both in in vitro reconstitution systems andin vivo (14–20). In searchof a positive control, we investigated the subcellular distribution ofBax in cells induced to undergo apoptosis in response to treatmentwith polyI z polyC, a mimic of double-stranded RNA. In HeLa cells,polyI z polyC treatment induces classical features of apoptosis (e.g.,caspase activation).6 Importantly, polyI z polyC (under treatmentconditions described in Fig. 8A), in HeLa cells, causes inhibition ofprotein synthesis with kinetics and amplitude similar to those inducedby onconase and triggers a similar degree of cell death within 24 hafter treatment.6 For instance, in the experiment shown in Fig. 8A,polyI z polyC caused;70% cell death, and onconase caused.65%cell death 24 h after the treatment. Fig. 8 shows that in the controlcells, the majority of Bax protein was found in the cytosol, with aminor fraction found attached to sedimentable structures that weresoluble in Triton X-100 (Fig. 8A,top, compareLanes 1and4). Six hafter polyI z polyC treatment, there was a dramatic redistribution ofBax: the protein levels were almost undetectable in the cytosol andwere greatly increased in the Triton X-100-soluble organellar fraction(Fig. 8A,top, compareLanes 2and5). In contrast, in onconase-treatedcells, the levels of Bax protein were substantially decreased in boththe cytosolic and the organellar fractions, and there were no indica-tions of a subcellular redistribution of Bax (Fig. 8A, top, compareLanes 3and6). The disappearance of Bax from onconase-treated cellswas almost complete 24 h after the treatment (data not shown). Thedifferences in the subcellular location of Bax in polyIz polyC- andonconase-treated cells correlated with even more dramatic differences

6 M. S. Iordanov and B. E. Magun, unpublished results.

Fig. 8. Lack of detectable Bax translocation in onconase-treatedcells. A, HeLa cells were treated, where indicated, with Lipofectinalone (control), with Lipofectin in the presence of 1mg/ml onconase,or with Lipofectin in the presence of 3mg/ml polyI z polyC. Six hafter the treatment, the cells were harvested and permeabilized withdigitonin, and cytosolic and organellar fractions were prepared asdescribed in “Materials and Methods.” The abundance and distribu-tion of Bax and cytochromec were assessed in immunoblot assay.Molecular weight markers are shown to theleft. Nonspecific bandsare indicated (p).Co, control cells.pIC, polyI z polyC-treated cells.On, onconase-treated cells.B, cells were treated with onconase andprocessed as described inA. To avoid overloadingLanes 1and2,only 20% of the cytosolic fraction was loaded into these lanes. Toallow appropriate comparison ofLanes 1and2 with Lanes 5and6,the immunoblot signal (procaspase-9 and -8) inLanes 1and2 shouldtherefore be scaled up fivefold.Co, control cells.On, onconase-treated cells.

1991




in the location of cytochromec. In the control cells, the majority ofcytochromec was found in the Triton X-100-insoluble organellarfraction (Fig. 8A,bottom, compareLanes 1and7). After polyI z polyCtreatment, the Triton X-100-insoluble organellar fraction was substan-tially depleted of cytochromec, and the levels of cytochromec weredramatically increased in the cytosol (Fig. 8A,bottom, compareLanes2 and8). In contrast, in onconase-treated cells, there was no detectablerelease of cytochromec 6 h after the treatment (Fig. 8A,bottom,compareLanes 3and9). The results shown in Figs. 7 and 8Araisedthe possibility that a yet-to-be-identified cytochromec-independentmechanism for caspase-9 activation is triggered in cells in response toonconase. Recently, it was discovered that a substantial portion ofprocaspase-9 is sequestered in mitochondria (59, 60). On apoptoticstimulation, mitochondrial procaspase-9 was found to translocate tothe cytosol and undergo a subsequent proteolytic activation there (59,60). Therefore, using the approach described in Fig. 8A, we investi-gated whether procaspase-9 undergoes translocation to the cytosolfrom an organellar (presumably mitochondrial) storage site before itsproteolytic activation in response to onconase treatment (Fig. 7,A andB). Indeed, the organellar fraction of onconase-treated HeLa cellsdisplayed decreased levels of procaspase-9 6 h after the treatment(Fig. 8B, top panel, compareLanes 5and 6), concomitant with adetectable increase in the levels of soluble cytosolic procaspase-9(Fig. 8B,top panel, compareLanes 1and2). In contrast, procaspase-8was found to be entirely cytosolic (Fig. 8B,bottom panel,Lanes 1and2) and, as described above (Fig. 7,A andB), unchanged in responseto onconase treatment.

DISCUSSION

Onconase is a prototypic member of a growing family of endori-bonucleases that are found in the bodily fluids of vertebrates fromamphibians to mammals and that seem to have emerged as part ofevolutionary ancient antiviral and antiparasitic defense systems (37).The most interesting property of onconase is its potential for use as anonmutagenic alternative for (or a supplement to) the conventionalDNA-damaging therapy of cancer. Two major directions of researchcould contribute to the realization of this potential: (a) increasing thetarget cell specificity and efficiency of delivery of onconase; and (b)understanding the mechanisms of onconase-induced cytotoxicity. Inthis work, we have begun to address the question of the intracellularmechanisms used by onconase to kill target cells. The most importantfinding we report here is that on internalization, onconase elicits anapoptotic death signal that seems to be, in part, independent ofinhibition of protein synthesis.

Apoptotic Mode of Cell Death Induced by Onconase.Adversetoxins are known to trigger cytolysis via nonapoptotic mechanisms.One such example is the drug capsaicin, the pungent ingredient inchili peppers, that has a therapeutic potential for targeted killing ofprimary afferent neurons (61, 62). However, a nonapoptotic mode ofcell death could hamper the potential application of onconase in thetherapy of cancer because nonapoptotic cell death could producesevere inflammatory and immune complications in patients. Althoughpreviously reported as apoptotic (63, 64), we present, for the firsttime, a detailed characterization of the mode of cell death induced byonconase, demonstrating that onconase-treated cells display classicalhallmarks of apoptosis such as chromatin fragmentation, nuclearpyknosis and karyorrhexis, plasma membrane blebbing, and activa-tion of caspases.

The Nature of the Onconase-induced Death Signal(s): PossibleInvolvement of tRNA? Previous studies have demonstrated that thecytotoxicity of onconase invariably depends on its catalytic capacityas a ribonuclease (33, 40). The data presented in this work prompt a

speculation that the major determinant of onconase-induced cytotox-icity is a death signal that is generated in response to induced RNAhydrolysis. This signal is generated even when concentrations ofonconase are applied that are otherwise insufficient for severe inhi-bition of translation. The only RNA population significantly affectedby onconase in HeLa and mouse fibroblast cells appeared to be tRNA(Fig. 1; data not shown for fibroblasts). This is in agreement with ourprevious findings that onconase preferentially degrades tRNAin vitroin reticulocyte lysates and when injected intoXenopus laevisoocytes(34). Therefore, the most likely candidate for an apoptosis-signalingintermediate is onconase-damaged tRNA. Although we cannot ex-clude the existence of certain onconase-sensitive mRNAs, we con-sider this possibility unlikely. The coding region of cyclophilinmRNA contains 67 potential sites for hydrolysis by onconase (59-UpG-39), yet apparent hydrolysis of this highly abundant mRNA wasundetectable in onconase-treated cells (Fig. 1). Curiously, onconaseeasily cleaves rRNA and diverse mRNAs (including cyclophilinmRNA) in vitro (data not shown). This finding demonstrates thatintracellular conditions determine the specificity of onconasein vivo.Among the possible candidates for such specific conditions, we con-sider the proper subcellular localization of both target RNAs andonconase and/or specific postinternalization modification(s) of onco-nase. These possibilities are currently being explored.

The Complex Nature of Onconase-induced Cell Death.A hy-pothetical model of the cytotoxic pathways triggered by onconase ispresented in Fig. 9 and is discussed below. One of the first effects ofonconase on target cells is the degradation of cellular tRNA. Aninevitable effect of sustained tRNA degradation is inhibition of pro-tein synthesis. Prolonged inhibition of translation, in turn, will inducecell death that results, in part, from caspase-mediated apoptosis (see,for instance, the example with emetine, Figs. 4B and 5A). However,three lines of evidence suggest that onconase-induced apoptosis doesnot entirely result from inhibition of protein synthesis. First, whereas

Fig. 9. A hypothetical model of onconase-induced cell death. One of the first effectsof onconase on target cells is the degradation of cellular tRNA. tRNA degradation causesinhibition of protein synthesis, which, in turn, triggers cell death through apoptoticpathways involving activation of caspases. However, the results presented in Fig. 2, Fig.4, Fig. 5A, and Table 1 strongly support the existence of an apoptotic pathway foronconase that is independent of inhibition of protein synthesis (downstream of a hypo-thetical proapoptotic eventX). This pathway is partially sensitive to zVADfmk, whereasthe pathway dependent on protein synthesis inhibition is likely to be zVADfmk-resistantand probably involves secondary necrosis. In contrast, cell death induced by ligation ofFas does not involve inhibition of protein synthesis in HeLa cells (see “Discussion”) andis entirely zVADfmk sensitive. The zVADfmk-resistant steps are indicated byitalic. Thehypothetical connections between tRNA degradation and protein synthesis inhibition, onone hand, and apoptosis, on the other hand, suggested by our results but yet to bedemonstrated are marked with a question mark (?).

1992




antibiotic inhibitors of translation (as exemplified in Fig. 5A byemetine) induce apoptotic cell death with relatively slow kinetics,onconase triggered a markedly early apoptosis (as measured by theappearance of TUNEL-positive cells; Fig. 5A). Second, emetine andcycloheximide required significantly higher levels of inhibition oftranslation to induce cell death than onconase (Table 1; data notshown for cycloheximide). Third, zVADfmk, although almost entirelyabolishing DEVDase activity induced by emetine (Fig. 4B), could notsignificantly protect cells from emetine-induced cell death (Table 1,compare columns E and F). It is likely that in the presence ofzVADfmk, the emetine-treated cells die from secondary necrosis, aform of cell death that has been recently postulated for cells in whichcertain aspects of apoptosis are impaired or proceed in an atypical way(65). Onconase-induced DEVDase activity was, overall, similarlysusceptible to zVADfmk (Fig. 4), but the inhibitor increased thekilling IC50 for onconase by more than 1 log (Table 1, comparecolumns C and D) and efficiently reduced the cytotoxic potential ofonconase at IC50 for translation (Table 1, compare columns E and F).In contrast to both onconase and antibiotic inhibitors of proteinsynthesis, cell death and DEVDase activation in response to Fasligation are completely inhibited by zVADfmk (Fig. 6,A andB). Themost obvious mechanistic difference between Fas- and onconase-induced cell death is the inability of Fas ligation to inhibit proteinsynthesis in HeLa cells (data not shown). Our results are thereforeconsistent with the notion that in addition to the mode of cell deaththat is dependent on protein synthesis inhibition (and shared by anyinhibitor of translation in HeLa cells), onconase triggers a comple-mentary proapoptotic mechanism that is zVADfmk sensitive. Thiscomplementary proapoptotic mechanism activates caspase-9 and -3/-7but is distinct from the classical Fas-induced pathway (FasL/Fas3FADD3 caspase-83 Bid3 Bax3 cytochromec3 caspase-93caspase-3). Furthermore, our results (Figs. 6C and 8) also raised thepossibility of a Bax- and cytochromec-independent mechanism ofcaspase-9 activation in cells treated with onconase. Alternatively, it isconceivable that feedback self-amplifying caspase “loops” are in-volved in executing the onconase-induced cell death program. Sub-sequent to its processing and activation by caspase-9, caspase-3 canprocess and activate additional molecules of caspase-9, thus leading tothe self-amplification of the caspase cascade (11). We have detectedcaspase-3/-7 (DEVDase) activity as early as 1–2 h after onconaseapplication (data not shown). At these time points (and up to 6 h, seeFig. 8A), we were unable to detect a measurable release of cytochromec. However, it is possible that low levels of cytochromec release andactive caspase-9 (below the sensitivity of the assays used here) lead tothe initial increase in DEVDase activity and that these DEVDases, inturn, process more procaspase-9, thereby amplifying the caspase-9/caspase-3(-7) cascade without further involvement of Bax or cyto-chromec.

Onconase in Comparison with Other Cytotoxic Enzymes Usedfor Designing Anticancer Immunotoxins. Experimental evidencefrom different laboratories shows that the mechanisms of cytotoxicactions of natural toxins used to design novel therapeutic agents withanticancer properties (such as ricin A chain andPseudomonasexo-toxin A) are more complex than mere of inhibition of protein synthe-sis, a property which these agents have in common (see Ref. 41 andreferences therein). For instance, Keppler-Hafkemeyeret al.(66) haveinvestigated the mode of cell death triggered by a genetically engi-neered immunotoxin containingPseudomonasexotoxin A as a cyto-toxic agent.Pseudomonasexotoxin A causes inhibition of proteinsynthesis through inactivation of translation elongation factor EF-2(see Ref. 41 and references therein). Similar to our findings usingonconase, Keppler-Hafkemeyeret al. have found that the antibody-Pseudomonasexotoxin A fusion protein also triggered apoptosis and

activated caspases (66). However, zVADfmk has been inefficient inpreventingPseudomonasexotoxin A-induced cell death, despite theablation of apoptotic morphology (66). In this respect,Pseudomonasexotoxin A displayed a behavior more reminiscent of emetine (asdescribed in this work) than of onconase (see Table 1). Therefore, inlight of the possible use of both cytotoxic enzymes for immunotoxindesign, future parallel studies are required to investigate whetheronconase andPseudomonasexotoxin A use the same or differentcytotoxic mechanisms.

Lack of p53 Dependence of the Onconase-induced ApoptoticProgram: Possible Implications in Therapy. We demonstrate inthis work that tRNA damage might be a physiologically relevant deathsignal in mammalian cells. Similar to DNA damage, this signal isultimately communicated to caspases involved in inducing an apo-ptotic phenotype. However, unlike DNA damage, the ability of RNAdamage to induce cell death is not affected by the presence or theabsence of a functional p53 protein (Fig. 3Cand text). The vastmajority of human tumors have inactivated the function of p53, eitherthrough acquired or inherited mutation in one allele, followed by lossof heterozygosity or by altering the expression or function of criticalp53 regulators, such as the human homologue ofmdm-2.Increasedresistance of some tumors with inactivated p53 to conventional DNA-damaging therapy has been reported (67, 68). Therefore, both the p53independence and the nonmutagenicity of RNA damage makeRNA an attractive target for developing novel proapoptotic anticancerstrategies.

ACKNOWLEDGMENTS

We thank D. Pribnow for expert advice and help with the RNA damagestudies. We also thank J. A. Pearson and J. L. Magun for technical assistance.

REFERENCES

1. Green, D. R. Apoptotic pathways: the roads to ruin. Cell,94: 695–698, 1998.2. Ashkenazi, A., and Dixit, V. M. Death receptors: signaling and modulation. Science

(Washington DC),281: 1305–1308, 1998.3. Green, D. R., and Reed, J. C. Mitochondria and apoptosis. Science (Washington DC),

281: 1309–1312, 1998.4. Cohen, G. M. Caspases: the executioners of apoptosis. Biochem. J.,326: 1–16, 1997.5. Cryns, V., and Yuan, J. Proteases to die for. Genes Dev.,12: 1551–1570, 1998.6. Thornberry, N. A., and Lazebnik, Y. Caspases: enemies within. Science (Washington

DC), 281: 1312–1316, 1998.7. Muzio, M., Chinnaiyan, A. M., Kischkel, F. C., O’Rourke, K., Shevchenko, A., Ni,

J., Scaffidi, C., Bretz, J. D., Zhang, M., Gentz, R., Mann, M., Krammer, P. H., Peter,M. E., and Dixit, V. M. FLICE, a novel FADD-homologous ICE/CED-3-like prote-ase, is recruited to the CD95 (Fas/APO-1) death-inducing signaling complex. Cell,85: 817–827, 1996.

8. Srinivasula, S. M., Ahmad, M., Fernandes-Alnemri, T., Litwack, G., and Alnemri,E. S. Molecular ordering of the Fas-apoptotic pathway: the Fas/APO-1 protease Mch5is a CrmA-inhibitable protease that activates multiple Ced-3/ICE-like cysteine pro-teases. Proc. Natl. Acad. Sci. USA,93: 14486–14491, 1996.

9. Stennicke, H. R., Jurgensmeier, J. M., Shin, H., Deveraux, Q., Wolf, B. B., Yang, X.,Zhou, Q., Ellerby, H. M., Ellerby, L. M., Bredesen, D., Green, D. R., Reed, J. C.,Froelich, C. J., and Salvesen, G. S. Pro-caspase-3 is a major physiologic target ofcaspase-8. J. Biol. Chem.,273: 27084–27090, 1998.

10. Martin, D. A., Siegel, R. M., Zheng, L., and Lenardo, M. J. Membrane oligomeriza-tion and cleavage activates the caspase-8 (FLICE/MACHa1) death signal. J. Biol.Chem.,273: 4345–4349, 1998.

11. Srinivasula, S. M., Ahmad, M., Fernandes-Alnemri, T., and Alnemri, E. S. Autoac-tivation of procaspase-9 by Apaf-1-mediated oligomerization. Mol. Cell,1: 949–957,1998.

12. Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S. M., Ahmad, M., Alnemri, E. S.,and Wang, X. Cytochromec and dATP-dependent formation of Apaf-1/caspase-9complex initiates an apoptotic protease cascade. Cell,91: 479–489, 1997.

13. Gonzales, D. H., and Neupert, W. Biogenesis of mitochondrial c-type cytochromes.J. Bioenerg. Biomembr.,22: 753–768, 1990.

14. Rosse, T., Olivier, R., Monney, L., Rager, M., Conus, S., Fellay, I., Jansen, B., andBorner, C. Bcl-2 prolongs cell survival after Bax-induced release of cytochromec.Nature (Lond.),391: 496–499, 1998.

15. Pastorino, J. G., Chen, S. T., Tafani, M., Snyder, J. W., and Farber, J. L. Theoverexpression of Bax produces cell death upon induction of the mitochondrialpermeability transition. J. Biol. Chem.,273: 7770–7775, 1998.

1993




16. Jurgensmeier, J. M., Xie, Z., Deveraux, Q., Ellerby, L., Bredesen, D., and Reed, J. C.Bax directly induces release of cytochromec from isolated mitochondria. Proc. Natl.Acad. Sci. USA,95: 4997–5002, 1998.

17. Eskes, R., Antonsson, B., Osen-Sand, A., Montessuit, S., Richter, C., Sadoul, R.,Mazzei, G., Nichols, A., and Martinou, J. C. Bax-induced cytochromec release frommitochondria is independent of the permeability transition pore but highly dependenton Mg21 ions. J. Cell Biol.,143: 217–224, 1998.

18. Narita, M., Shimizu, S., Ito, T., Chittenden, T., Lutz, R. J., Matsuda, H., andTsujimoto, Y. Bax interacts with the permeability transition pore to induce perme-ability transition and cytochromec release in isolated mitochondria. Proc. Natl. Acad.Sci. USA,95: 14681–14686, 1998.

19. Saikumar, P., Dong, Z., Patel, Y., Hall, K., Hopfer, U., Weinberg, J. M., andVenkatachalam, M. A. Role of hypoxia-induced Bax translocation and cytochromecrelease in reoxygenation injury. Oncogene,17: 3401–3415, 1998.

20. Desagher, S., Osen-Sand, A., Nichols, A., Eskes, R., Montessuit, S., Lauper, S.,Maundrell, K., Antonsson, B., and Martinou, J. C. Bid-induced conformationalchange of Bax is responsible for mitochondrial cytochromec release during apopto-sis. J. Cell Biol.,144: 891–901, 1999.

21. Enari, M., Sakahira, H., Yokoyama, H., Okawa, K., Iwamatsu, A., and Nagata, S. A.caspase-activated DNase that degrades DNA during apoptosis, and its inhibitorICAD. Nature (Lond.),391: 43–50, 1998.

22. Shiokawa, D., and Tanuma, S. Molecular cloning and expression of a cDNA encodingan apoptotic endonuclease DNase-g. Biochem. J.,332: 713–720, 1998.

23. Hakem, R., Hakem, A., Duncan, G. S., Henderson, J. T., Woo, M., Soengas, M. S.,Elia, A., de la Pompa, J. L., Kagi, D., Khoo, W., Potter, J., Yoshida, R., Kaufman,S. A., Lowe, S. W., Penninger, J. M., and Mak, T. W. Differential requirement forcaspase 9 in apoptotic pathwaysin vivo. Cell, 94: 339–352, 1998.

24. Kuida, K., Haydar, T. F., Kuan, C. Y., Gu, Y., Taya, C., Karasuyama, H., Su, M. S.,Rakic, P., and Flavell, R. A. Reduced apoptosis and cytochromec-mediated caspaseactivation in mice lacking caspase 9. Cell,94: 325–337, 1998.

25. Yoshida, H., Kong, Y. Y., Yoshida, R., Elia, A. J., Hakem, A., Hakem, R., Penninger,J. M., and Mak, T. W. Apaf1 is required for mitochondrial pathways of apoptosis andbrain development. Cell,94: 739–750, 1998.

26. Kasibhatla, S., Brunner, T., Genestier, L., Echeverri, F., Mahboubi, A., and Green,D. R. DNA damaging agents induce expression of Fas ligand and subsequentapoptosis in T lymphocytes via the activation of NF-k B and AP-1. Mol. Cell,1:543–551, 1998.

27. Friesen, C., Herr, I., Krammer, P. H., and Debatin, K. M. Involvement of the CD95(APO-1/FAS) receptor/ligand system in drug-induced apoptosis in leukemia cells.Nat. Med.,2: 574–577, 1996.

28. Herr, I., Wilhelm, D., Bohler, T., Angel, P., and Debatin, K. M. Activation of CD95(APO-1/Fas) signaling by ceramide mediates cancer therapy-induced apoptosis.EMBO J.,16: 6200–6208, 1997.

29. Hansen, R., and Oren, M. p53: from inductive signal to cellular effect. Curr. Opin.Genet. Dev.,7: 46–51, 1997.

30. Levine, A. J. p53, the cellular gatekeeper for growth and division. Cell,88: 323–331,1997.

31. Darzynkiewicz, Z., Carter, S. P., Mikulski, S. M., Ardelt, W. J., and Shogen, K.Cytostatic and cytotoxic effects of Pannon (P-30 Protein), a novel anticancer agent.Cell Tissue Kinet.,21: 169–182, 1988.

32. Mikulski, S. M., Ardelt, W., Shogen, K., Bernstein, E. H., and Menduke, H. Strikingincrease of survival of mice bearing M109 Madison carcinoma treated with a novelprotein from amphibian embryos. J. Natl. Cancer Inst.,82: 151–153, 1990.

33. Wu, Y., Mikulski, S. M., Ardelt, W., Rybak, S. M., and Youle, R. J. A cytotoxicribonuclease. Study of the mechanism of onconase cytotoxicity. J. Biol. Chem.,268:10686–10693, 1993.

34. Lin, J. J., Newton, D. L., Mikulski, S. M., Kung, H. F., Youle, R. J., and Rybak, S. M.Characterization of the mechanism of cellular and cell free protein synthesis inhibi-tion by an anti-tumor ribonuclease. Biochem. Biophys. Res. Commun.,204: 156–162, 1994.

35. Rybak, S. M., Pearson, J. W., Fogler, W. E., Volker, K., Spence, S. E., Newton, D. L.,Mikulski, S. M., Ardelt, W., Riggs, C. W., Kung, H. F., and Longo, D. L. Enhance-ment of vincristine cytotoxicity in drug-resistant cells by simultaneous treatment withonconase, an antitumor ribonuclease. J. Natl. Cancer Inst.,88: 747–753, 1996.

36. Ardelt, W., Mikulski, S. M., and Shogen, K. Amino acid sequence of an anti-tumorprotein from Rana pipiensoocytes and early embryos. Homology to pancreaticribonucleases. J. Biol. Chem.,266: 245–251, 1991.

37. Schein, C. H. From housekeeper to microsurgeon: the diagnostic and therapeuticpotential of ribonucleases. Nat. Biotechnol.,15: 529–536, 1997.

38. Mikulski, S. M., Newton, D. L., Wiltrout, R. H., and Rybak, S. M. Onconase anddoxorubicin synergy in prolonging survival of MDA-MB-231 human breast cancer-bearing nude mice. Proc. Am. Assoc. Cancer Res.40: 491, 1999.

39. Rybak, S. M., Saxena, S. K., Ackerman, E. J., and Youle, R. J. Cytotoxic potential ofribonuclease and ribonuclease hybrid proteins. J. Biol. Chem.,266: 21202–21207,1991.

40. Leland, P. A., Schultz, L. W., Kim, B. M., and Raines, R. T. Ribonuclease A variantswith potent cytotoxic activity. Proc. Natl. Acad. Sci. USA,95: 10407–10412, 1998.

41. Iordanov, M. S., Pribnow, D., Magun, J. L., Dinh, T. H., Pearson, J. A., Chen, S. L.,and Magun, B. E. Ribotoxic stress response: activation of the stress-activated proteinkinase JNK1 by inhibitors of the peptidyl transferase reaction and by sequence-specific RNA damage to thea-sarcin/ricin loop in the 28S rRNA. Mol. Cell Biol.,17:3373–3381, 1997.

42. Newton, D. L., Xue, Y., Boque, L., Wlodawer, A., Kung, H. F., and Rybak, S. M.Expression and characterization of a cytotoxic human-frog chimeric ribonuclease:potential for cancer therapy. Protein Eng.,10: 463–470, 1997.

43. Goillot, E., Raingeaud, J., Ranger, A., Tepper, R. I., Davis, R. J., Harlow, E., andSanchez, I. Mitogen-activated protein kinase-mediated Fas apoptotic signaling path-way. Proc. Natl. Acad. Sci. USA,94: 3302–3307, 1997.

44. Pribnow, D., Muldoon, L. L., Fajardo, M., Theodor, L., Chen, L. Y., and Magun, B. E.Endothelin induces transcription of fos/jun family genes: a prominent role for calciumion. Mol. Endocrinol.,6: 1003–1012, 1992.

45. Iordanov, M. S., Pribnow, D., Magun, J. L., Dinh, T. H., Pearson, J. A., and Magun,B. E. Ultraviolet radiation triggers the ribotoxic stress response in mammalian cells.J. Biol. Chem.,273: 15794–15803, 1998.

46. Bossy-Wetzel, E., Newmeyer, D. D., and Green, D. R. Mitochondrial cytochromecrelease in apoptosis occurs upstream of DEVD-specific caspase activation and inde-pendently of mitochondrial transmembrane depolarization. EMBO J.,17: 37–49,1998.

47. Lazebnik, Y. A., Kaufmann, S. H., Desnoyers, S., Poirier, G. G., and Earnshaw, W. C.Cleavage of poly(ADP-ribose) polymerase by a proteinase with properties like ICE.Nature (Lond.),371: 346–347, 1994.

48. Tewari, M., Quan, L. T., O’Rourke, K., Desnoyers, S., Zeng, Z., Beidler, D. R.,Poirier, G. G., Salvesen, G. S., and Dixit, V. M. Yama/CPP32b, a mammalianhomolog of CED-3, is a CrmA-inhibitable protease that cleaves the death substratepoly(ADP-ribose) polymerase. Cell,81: 801–809, 1995.

49. Yaginuma, Y., and Westphal, H. Analysis of thep53 gene in human uterine carci-noma cell lines. Cancer Res.,51: 6506–6509, 1991.

50. Hoppe-Seyler, F., and Butz, K. Repression of endogenous p53 transactivation func-tion in HeLa cervical carcinoma cells by human papillomavirus type 16 E6, humanmdm-2, and mutant p53. J. Virol.,67: 3111–3117, 1993.

51. Huibregtse, J. M., and Beaudenon, S. L. Mechanism of HPV E6 proteins in cellulartransformation. Semin. Cancer Biol.,7: 317–326, 1996.

52. Oka, M., Hirazawa, K., Yamamoto, K., Iizuka, N., Hazama, S., Suzuki, T., andKobayashi, N. Induction of Fas-mediated apoptosis on circulating lymphocytes bysurgical stress. Ann. Surg.,223: 434–440, 1996.

53. Li, P., Allen, H., Banerjee, S., Franklin, S., Herzog, L., Johnston, C., McDowell, J.,Paskind, M., Rodman, L., Salfeld, J., Towne, E., Tracey, D., Wardwell, S., Wei, F-Y.Wong, W., Kamen R., and Seshadri, T. Mice deficient in IL-1b-converting enzymeare defective in production of mature IL-1b and resistant to endotoxic shock. Cell,80:401–411, 1995.

54. Smith, D. J., McGuire, M. J., Tocci, M. J., and Thiele, D. L. IL-1b convertase (ICE)does not play a requisite role in apoptosis induced in T lymphoblasts by Fas-dependent or Fas-independent CTL effector mechanisms. J. Immunol.,158:163–170,1997.

55. Li, P., Allen, H., Banerjee, S., and Seshadri, T. Characterization of mice deficient ininterleukin-1b converting enzyme. J. Cell. Biochem.,64: 27–32, 1997.

56. Kuwana, T., Smith, J. J., Muzio, M., Dixit, V., Newmeyer, D. D., and Kornbluth, S.Apoptosis induction by caspase-8 is amplified through the mitochondrial release ofcytochromec. J. Biol. Chem.,273: 16589–16594, 1998.

57. Li, H., Zhu, H., Xu, C. J., and Yuan, J. Cleavage of BID by caspase 8 mediates themitochondrial damage in the Fas pathway of apoptosis. Cell,94: 491–501, 1998.

58. Luo, X., Budihardjo, I., Zou, H., Slaughter, C., and Wang, X. Bid, a Bcl2 interactingprotein, mediates cytochromec release from mitochondria in response to activation ofcell surface death receptors. Cell,94: 481–490, 1998.

59. Susin, S. A., Lorenzo, H. K., Zamzami, N., Marzo, I., Brenner, C., Larochette, N.,Prevost, M. C., Alzari, P. M., and Kroemer, G. Mitochondrial release of caspase-2 and-9 during the apoptotic process. J. Exp. Med.,189: 381–394, 1999.

60. Krajewski, S., Krajewska, M., Ellerby, L. M., Welsh, K., Xie, Z., Deveraux, Q. L.,Salvesen, G. S., Bredesen, D. E., Rosenthal, R. E., Fiskum, G., and Reed, J. C.Release of caspase-9 from mitochondria during neuronal apoptosis and cerebralischemia. Proc. Natl. Acad. Sci. USA,96: 5752–5757, 1999.

61. Maggi, C. A. Therapeutic potential of capsaicin-like molecules: studies in animalsand humans. Life Sci.,51: 1777–1781, 1992.

62. Caterina, M. J., Schumacher, M. A., Tominaga, M., Rosen, T. A., Levine, J. D., andJulius, D. The capsaicin receptor: a heat-activated ion channel in the pain pathway.Nature (Lond.),389: 816–824, 1997.

63. Mikulski, S. M., Viera, A., Deptala, A., and Darzynkiewicz, Z. Enhancedin vitrocytotoxicity and cytostasis of the combination of onconase with a proteasome inhib-itor. Int. J. Oncol.,13: 633–644, 1998.

64. Deptala, A., Halicka, H. D., Ardelt, B., Ardelt, W., Mikulski, S. M., Shogen, K., andDarzynkiewicz, Z. Potentiation of tumor necrosis factor induced apoptosis by onco-nase. Int. J. Oncol.,13: 11–16, 1998.

65. Kroemer, G., Dallaporta, B., and Resche-Rigon, M. The mitochondrial death/liferegulator in apoptosis and necrosis. Annu. Rev. Physiol.,60: 619–642, 1998.

66. Keppler-Hafkemeyer, A., Brinkmann, U., and Pastan, I. Role of caspases in immu-notoxin-induced apoptosis of cancer cells. Biochemistry,37: 16934–16942, 1998.

67. Benhattar, J., Cerottini, J. P., Saraga, E., Metthez, G., and Givel, J. C. p53 mutationsas a possible predictor of response to chemotherapy in metastatic colorectal carcino-mas. Int. J. Cancer,69: 190–192, 1996.

68. Iwadate, Y., Fujimoto, S., Tagawa, M., Namba, H., Sueyoshi, K., Hirose, M., andSakiyama, S. Association ofp53 gene mutation with decreased chemosensitivity inhuman malignant gliomas. Int. J. Cancer,69: 236–240, 1996.

1994




2000;60:1983-1994. Cancer Res Mihail S. Iordanov, Olga P. Ryabinina, John Wong, et al. Different from Inhibition of Protein SynthesisRibonuclease Onconase: Evidence for Cytotoxic Mechanisms Molecular Determinants of Apoptosis Induced by the Cytotoxic

Updated version

http://cancerres.aacrjournals.org/content/60/7/1983

Access the most recent version of this article at:

Cited articles

http://cancerres.aacrjournals.org/content/60/7/1983.full#ref-list-1

This article cites 65 articles, 27 of which you can access for free at:

Citing articles

http://cancerres.aacrjournals.org/content/60/7/1983.full#related-urls

This article has been cited by 14 HighWire-hosted articles. Access the articles at:

E-mail alerts related to this article or journal.Sign up to receive free email-alerts

Subscriptions

Reprints and

[email protected] at

To order reprints of this article or to subscribe to the journal, contact the AACR Publications

Permissions

Rightslink site. Click on "Request Permissions" which will take you to the Copyright Clearance Center's (CCC)

.http://cancerres.aacrjournals.org/content/60/7/1983To request permission to re-use all or part of this article, use this link



http://cancerres.aacrjournals.org/content/60/7/1983.full#ref-list-1

http://cancerres.aacrjournals.org/content/60/7/1983.full#related-urls

http://cancerres.aacrjournals.org/cgi/alerts

mailto:[email protected]



molecular determinants of apoptosis induced by the cytotoxic ribonuclease … · mechanism of...

Documents