Research Laboratory of Children's Hospital, University of Ulm

Chairman: Prof. Dr. Klaus-Michael Debatin

Activation of apoptosis pathwaysby different classes of anticancer drugs

Dissertation for the applying for aDoctor Degree of Medicine (Dr. med.)

Faculty of Medicine, University of Ulm

Presented by Jiahao Liu

Born in Hubei, P. R. China

2001

Amtierender Dekan: Prof. Dr. R. Marre

1. Berichterstatter: Prof. Dr. K. M. Debatin

2. Berichterstatter: Prof. Dr. Dr. Dr. A. Grünert

Tag der Promotion: 26. 10. 2001

To my family:

Chen Longgui &Liu Chang

1

Contents

Contents 1

Abbreviations 4

1. Introduction1.1. Apoptosis: definitions and mechanisms 6

1.1.1. Cell biology of apoptosis 6

1.1.2. Execution of programmed cell death by caspases 7

1.1.3. Two main pathways of apoptosis 8

1.2. Cytotoxic anticancer drugs and apoptosis 9

1.3. Aims and summary of the project 11

2. Materials and Methods

2.1. Materials 14

2.1.1. Reagents and equipment for cell culture 14

2.1.2. Reagents and equipment for flow cytometric analysis 14

2.1.3. Reagents and equipment for western blot 15

2.1.4. Anticancer drugs 16

2.1.5. Antibodies 17

2.2. Methods 19

2.2.1. Cell culture 19

2.2.2. Cell preservation and reconstitution 19

2.2.3. Cell stimulation 19

2.2.4. Inhibitor studies 20

2.2.5. Flow cytometry 20

2.2.5.1. Analysis of annexin V and PI positive cells 20

2.2.5.2. Quantification of DNA fragmentation 21

2.2.5.3. Analysis of mitochondrial membrane potential (∆Ψm) 21

2.2.5.4. Quantification of cytoplasmic cytochrome c 22

2.2.6. Cytosolic and mitochondrial extracts preparation 22

2

2.2.7. Western blot analysis 23

2.2.7.1. Cell lysis 23

2.2.7.2. Quantification of protein 24

2.2.7.3. Electrophoreses 24

2.2.7.4. Immunoblotting 25

2.2.7.5. Detection 26

2.2.7.6. Reprobing membranes 26

2.2.8. Statistical analysis 27

3. Results

3.1. Induction of apoptosis by anticancer drugs 28

3.1.1. Apoptosis induced by treatment of Jurkat cells with etoposide,

cytarabine, 4-hydroxy-cyclophosphamide, doxorubicin and

methotrexate 28

3.1.1.1. Dose and time kinetics for anticancer drug treatment 28

3.1.1.2. Comparison of different apoptotic signs induced by

different drugs 30

3.1.1.3. Annexin V single positive cells appear through

treatment with early anticancer drugs, but not with

late anticancer drugs 34

3.1.2. Comparison of apoptosis induced by anticancer drugs with

apoptosis induced by death receptor signaling and γ-radiation 36

3.1.2.1. Agonistic anti-CD95 antibody-induced apoptosis 36

3.1.2.2. γ-radiation-induced apoptosis 39

3.2. Expression of death receptor associated molecules induced by

anticancer drugs 39

3.3. Caspases activation by anticancer drugs 41

3.3.1. Caspase-8, caspase-3 activation and PARP cleavage 41

3.3.2. Caspase inhibitor zVAD-fmk blocks drug-induced apoptosis 43

3.4. Disturbance of mitochondrial function induced by anticancer drugs 46

3.4.1. Alterations of mitochondrial transmembrane potential (∆Ψm) 46

3.4.1.1. Time course of drug-induced ∆Ψm loss 46

3

3.4.1.2. Caspase inhibitor abrogated anti-CD95-induced ∆Ψm

loss but not drug-mediated ∆Ψm loss 49

3.4.2. Cytochrome c release induced by anticancer drugs 52

3.4.3. Anticancer drugs cleave Bid and Bcl-2 52

4. Discussion4.1. Different drugs induce apoptosis in a different time and dose

fashion and represent different feature of apoptosis 55

4.2. CD95-associated signaling molecules and anticancer drugs 57

4.3. The central role of caspase in drug-induced apoptosis 59

4.4. Disturbance of mitochondrial function induced by anticancer drugs 62

5. Summary 67

6. References 69

7. Acknowledgements 81

4

Abbreviations

PARP poly (ADP-ribose) polymerase

PBS phosphate-buffered saline

PAGE polyacrylamide gel electrophroresis

zVAD-fmk benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl ketone

ROS reactive oxygen species

∆Ψm mitochondrial transmembrane potential

DiOC6(3) 3,3'-dihexyloxacarbocyanine iodide

FACS fluorescence-activated cell-sorting

PT permeability transition

AIF Apoptosis-inducing factor

BetA betulinic acid

BA bongkrekic acid

BSA bovine serum albumin

mAb monoclonal antibody

CD95L CD95 ligand

Eto etoposide

Ara-c cytarabine

Doxo doxorubicin

4-HCP 4-hydroxy-cyclophosphamide

MTX methotrexate

Cyt. C cytochrome c

DISC death-inducing signaling complex

FADD Fas-associated death domain protein

Apaf-1 apoptosis protease-activating factor 1

VDAC voltage-dependent anion channel(s)

FBS fetal bovine serum

FITC fluorescein isothiocyanate-conjugated

PI propidium iodide

Gy gray

5

PS phosphatidylserine

DTT dithiothreitol

RIP receptor interacting protein

RT room temperature

SDS Sodium dodecyl sulfate

TNF Tumor necrosis factor

Triton X-100 octylphenol-polyethyleneglycol ether

EDTA ethylene diamintetraacetic acid

6

1. Introduction

Cells die by two primary processes: A) necrosis, in which the release of intracellular

proteases and lysozymes induce an inflammatory response, or B) apoptosis, also known as

programmed cell death, where the cell remnants quietly disappear as they are

phagocytosed by surrounding cells.

1.1. Apoptosis: definitions and mechanisms

1.1.1. Cell biology of apoptosis

The major physiological mechanism of cell removal is apoptosis - a Greek descriptive term

for falling leaves or petals. Apoptosis describes the process by which cells are 'silently'

removed under normal conditions when they reach the end of their life span, are damaged,

or superfluous. It is a general tissue phenomenon necessary for development and

homeostasis: elimination of redundant cells during embrogenesis, cell atrophy upon

endocrine withdrawal or loss of essential growth factors or cytokines, tissue remodelling

and repair, and removal of cells that have sustained genotoxic damage. Its conserved

features reflect its similarly evolutionarily conserved genetic characteristics, from

nematode worm to man. Apoptosis is strictly a morphological description and other

morphologies of developmental programmed cell death exist (Kerr et al., 1972; Majno et

al., 1995; Häcker, 2000; Chinnaiyan et al., 1997).

Apoptotic cells exhibit a characteristic pattern of changes, including cytoplasmic

shrinkage, active membrane blebbing, chromatin condensation and, typically,

fragmentation into membrane-enclosed vesicles, apoptotic body (Ucker et al., 1992;

Kawabat et al., 1999; Wyllie, 1999; Mills et al., 1999). This readily visible transformation

is accompanied by a number of biochemical changes. Changes at the cell surface include

the externalization of phosphatidylserine and other alterations that promote recognition by

phagocytes. Intracellular changes include the degradation of the chromosomal DNA into

high-molecular-weight and oligonucleosomal fragments, as well as cleavage of a specific

7

subset of cellular polypeptides (Ellis et al., 1991; Rotello et al., 1994; Franc et al., 1996;

Savill, 1996).

The apoptotic process may be set in motion by: A) genes responding to DNA damage; B)

death signals received at the cell membrane (Fas ligand), or C) proteolytic enzymes

entering directly into the cell (granzymes). The final events, evidenced by the changes in

cell structure and disassembly, are the work of specific proteases (caspases) (Evans, 1993).

1.1.2. Execution of programmed cell death by caspases

Caspases are currently considered as the central executioners of many, if not all, apoptotic

pathways (Chinnaiyan et al., 1997; Alnemri, 1997; Kroemer et al., 1998; Budihardjo et al.,

1998). Many of the proteolytic cleavages during apoptosis result from the action of a

unique family of cysteine-dependent proteases called caspases. The various members of

this protease family differ in primary structure and substrated specificity but share several

carboxyl side of aspartate residues. First, each caspase cleaves at the carboxyl side of

aspartate residues. Second, each active caspase is a synthesized as a zymogen that contains

an N-terminal prodomain, a large subunit and a small subunit. Finally, proteolytic cleavage

to liberate each caspase involves sequential cleavages at two or more small subunits from

one another and from the prodomain. The fact that these activating cleavages occur at sites

that could be cleaved by caspases led to the concept that caspase activation might involve

either a proteolytic cascade or an autoactivation process (Earnshaw et al., 1999; Nicholson,

1999; Walker et al., 1994; Salvesen et al., 1997).

Of the twelve known human caspases, six (caspases-3, -6, -7, -8, -9, and -10) are definitely

involved in apoptosis in various model systems. One current classification scheme divides

these apoptotic caspases into two classes, effector (or 'downstream') caspases, which are

responsible for most of the cleavages that disassemble the cell, and initiator (or 'upstream')

caspases, which initiate the proteolytic cascade (Depraetere et al., 1998; Thornberry et al,

1998).

Caspase-3, -6, and -7 are the major effector caspases characterized to date. Once activated,

these enzymes are capable of cleaving the vast majority of polypeptides that undergo

8

proteolysis in apoptotic cells (Earnshaw et al., 1999; Tewari et al., 1995; Sakahira et al.,

1998; Sahara et al., 1999). Interestingly, overexpression of these caspases in mammalian

cells is relatively non-toxic, suggesting that these precursors have limited capacity for

autoactivation. Instead, effector caspases are usually activated by other proteases.

Caspase-8 and -9 are the major initiator caspases identified to date. Zymogen forms of

these enzymes display low but detectable protease activity. This activity increases when

the prodomains of these zymogens interact with certain binding partners. Upon activation,

caspase-8 and -9 acquire the ability to cleave and activate caspases (Juo et al., 1998;

Nagata, 1997).

An increasing number of proteins have been found to be cleaved by caspases, and for some

of them an apoptotic function has been proposed. Among different substrates are enzymes

involved in genome function, such as the DNA repair enzyme poly (adenosine

diphosphate-ribose) polymerase (PARP) and DNA-dependent protein kinase (DNA-PK),

or regulators of the cell cycle, including retinoblastoma protein, the p53 regulator MDM-2,

MEKK, and protein kinase C-δ. Substrates of the nucleus and cytoskeleton include lamins,

Gas2, gelsolin, and fodrin. Furthermore, it has been found that DNA cleavage is triggered

upon caspase-mediated degradation of the inhibitory subunit of a novel endonuclease,

designated caspase-activated DNase.

Current knowledge indicates that individual caspase have distinct substrate specificities,

inhibitor profiles, and abilities to process each other. These findings suggest that caspases

form a hierarchical network which, similar to the complement system, may function as an

amplifier for a given apoptotic stimulus (Garcia-Calvo et al., 1999; Nicholson, 1999; Los

et al., 1999).

1.1.3. Two main pathways of apoptosis

One of the best-defined apoptotic pathways is mediated by the death receptor CD95 (APO-

1/Fas). Triggering of CD95 by its natural ligand or agonistic antibodies induces the

formation of DISC that consists of the adapter protein FADD and FLICE/caspase-8.

Complex formation is initiated through homophilic interaction of the death domains

9

present in the intracellular part of both CD95 and FADD. FADD, in addition, contains a

second interaction region called the DED, which couples to caspase-8 as the most proximal

element in the caspase cascade. Further downstream, caspse-8 presumably triggers the

proteolytic activation of other caspases and cleavage of cellular substrates (Krammer,

1999; Schulze-Osthoff et al., 1998).

Another apoptotic pathway is the mitochondrial pathway. It has been shown that

mitochondria play an important role in regulation of apoptosis. An early event in this

process is the loss of the mitochondrial transmembrane potential ∆Ψm, which induce the

release of molecules contained in the intermembrane space of the mitochondria to the

cytosol. Among the released molecules is cytochrome c that, on entry in the cytosol,

induces oligomerization of Apaf-1 (caspase recruitment domain) in the presence of ATP.

In turn, oligomerized Apaf-1 binds to cytosolic procaspase-9 in a so-called apoptosome

complex and induces processing and activates the downstream caspase cascade. Other

molecules released from the mitochondria include several procaspases and the flavoprotein

AIF (apoptosis inducing factor) that translocates to the nucleus and triggers caspase-

independent nuclear changes. Mitochondrial apoptosis signal is regulated by molecules of

the Bcl-2 family, which have been shown to control mitochondrial membrane integrity by

interaction with the mitochondrial membranes. The antiapoptotic properties of Bcl-2 and

related proteins have been related to their ability to prevent these mitochondrial events,

whereas the targeting of BH3 domain-only proteins of the Bcl-2 family such as Bid, Bim

or Bad from various parts of the cell to the mitochondria was shown to activate the death

process by inducing the mitochondrial release of proapoptotic molecules (Li et al., 1997;

Stennicke et al., 1999).

1.2. Cytotoxic anticancer drugs and apoptosis

Anticancer drugs have been shown to target diverse cellular functions in mediating cell

death in chemosensitive tumors. Cytotoxic drugs which are currently used for the

treatment of malignant diseases such as etoposide, cytarabine, cyclophosphamide,

doxorubicin and methotrexate are thought to exert their effects through inhibition of

topoisomerase II (etoposide), DNA-polymerase (cytarabine), antagonization of folic acid

10

(methotrexate), inhibition of DNA-crosslinking (cyclophosphamide) and DNA-

intercalation (doxorubicin).

Although the primary intracellular targets of drug action are rather distinct, it has become

evident that drug-induced cytotoxicity ultimately converges on a common pathway,

causing apoptosis (Debatin, 1999; Kaufmann et al., 2000; Mkin et al., 2000). Cells exposed

to anticancer drugs display apoptosis alterations, such as cell shrinkage, chromatin

condensation, and internucelosomal DNA fragmentation. A close link between apoptosis

and the mechanism of drug action has been demonstrated by the involvement of similar

genetic components. Overexpression of Bcl-2 proteins can confer drug resistance in

transfected tumor cells. A number of investigations exposed a critical role of the tumor

suppressor p53 in apoptosis after drug treatment. p53 requires an upstream DNA damage

signal to allow it to function, and the clues as to how this comes about were provided by

the observations that p53 is induced by DNA-damaging agents, including γ-irradiation and

chemotherapeutic agents. After exposure of the cell to DNA damaged, p53 protein levels

are rapidly upregulated by a post-transcriptional mechanism involving both stabilization

and possible modification of a latent form of p53. DNA strand breaks are sufficient

stimulus for this p53 response and it has been suggested that a single double-strand DNA

break per cell is sufficient. The outcome of the activation of p53 is either apoptosis or a

cell cycle arrest. But it is clear that the response to activation of p53 is tissue specific; the

cellular outcome will also depend upon the balance between pro-apoptotic signaling from

p53, and its downstream events, and anti-apoptotic survival signaling provided by various

molevules. Finally, it has been recently shown that drug-induced cytotoxicity involves

proteases of the caspase family, because specific inhibitors of caspases prevented cell

death after treatment with different anticancer agents.

A number of studies have raised the possibility that anticancer drugs trigger apoptosis by

inducing the synthesis of CD95-L, which then bind to CD95 receptor and activates the

death receptor pathway in an autocrine or paracrine fashion. Drug-induced increases in

CD95 mRNA, CD95-L and upregulation of CD95 was observed after treatment of

different tumors with cytotoxic drugs such as doxorubicin, cisplatin, methotrexate,

cytarabine and etoposide at therapeutic concentrations. Treatment of leukemias or solid

tumors, including neuroblastoma, hepatoblastoma, medulloblastoma, colon carcinoma and

breast cancer cells with cytotoxic drugs induces CD95-L expression and mediates

11

autocrine suicide or paracrine cell death following binding to its receptor. Blockade of

CD95-L/receptor interaction using antagonistic antibodies to the receptor markedly

reduced drug-triggered apoptosis. Thus, production of CD95-L and crosslinking of its

cognate receptor are probably involved in drug-mediated cell death. Moreover, CD95

expression was unregulated upon treatment with cytotoxic drugs, which increases

sensitivity of physiological apoptotic signals (Friesen et al., 1996; Fulda et al., 1997a, b).

Alterations of mitochondrial functions such as permeability transition (PT) have been

found to play a major role in the apoptotic process induced by anticancer drugs (Costantini

et al., 2000). Exposure of many cultured hematological cell lines to a cytotoxic anticancer

drug can cause mitochondrial dysfunction that include loss of mitochondrial membrane

potential (∆Ψm), release of cytochrome c and AIF from the mitochondrial intermembrane

space to the cytosol, and the generation of reactive oxygen species. Anticancer drugs also

destroy the balance of between proapoptotic and antiapoptotic members of Bcl-2 family,

which reduce the stabilization role of the mitochondrial membrane by anti-apoptotic Bcl-2-

like proteins.

1.3. Aims and summary of the project

Over the past 20 years, anticancer combination therapy using protocols based on clinical

experience and empirical data has achieved long term remission and cure in 70-80% of

patients with pediatric acute lymphoblastic leukemia. However, the most prevalent of

malignancies have proved to be more or less resistant to anticancer drugs. Dose escalation

using high-dose chemotherapy may have resulted in a modest improvement in responses

but has not constituted a breakthrough. The dose intensity of most chemotherapeutic

regimens is limited primarily by the degree of toxicity encountered. Acute toxicities

common to many of the anticancer drugs include myelosuppression, nausea and vomiting,

alopecia, orointestinal mucositis, liver function test abnormalities, allergic or cutaneous

reactions, and local ulceration from subcutaneous drug extravasation.

The primary role of the pediatric oncologist is to orchestrate the administration of complex

combination chemotherapy regiments to children. Special care must be taken because the

anticancer drugs used in these regimens have the lowest therapeutic index of any class of

12

drugs and predictably produce significant, even life-threatening toxic reactions at

therapeutic doses. However, allowing significant dose reductions or delays in therapy to

attenuate these toxicities may compromise dose-intensity and place the patient at an

increased risk for disease recurrence. The cancer chemotherapist must carefully balance the

risks of toxicities from therapy against the risk of tumor recurrence from inadequate

treatment. Unfortunately, the development and clinical usage of cancer chemotherapy

remains largely empiric, and the mechanisms of action of most anticancer drugs are

nonselective targeting vital macromolecules (e.g. nuclei acids) or metabolic pathways.

To ensure that these drugs are used safely and effectively, an understanding of the

mechanism of drug action and time kinetics is crucial. Although extensive studies of the

biochemical and molecular pharmacology of drug-target cell interaction have been

performed, the precise molecular requirement by which anticancer drugs initiate apoptosis

pathways are poorly defined.

It is known that in empirical medicine, these different anticancer drugs have different

characteristic concerning specific anti-tumor or anti-leukemic efficacy and side effects on

normal tissue. While solid tumors are often treated with cisplatin, treatment of leukemia is

based on the use of anthracyclines and antimetabolites. Many drugs also have unique

toxicities affecting normal tissues, such as cardiotoxicity associated with anthracyclines,

hemorrhagic cystitis associated with cyclophosphamide and ifosfamide, peripheral

neuropathy from vincristine, and coagulopathy from L-asparaginase

We therefore hypothesized that the different clinical used anticancer drugs might induce

apoptosis in a drug specific manner. Thus the clinical observed differences could be

reflected by different activation of apoptosis signaling pathways.

We therefore investigated induction of apoptosis, activation of caspases and involvement

of mitochondrial signaling in the well defined Jurkat cell culture system by five

conventional used anticancer drugs: etoposide, cytarabine, 4-hydroxy-cyclosphamide,

doxorubicin, and methotrexate, in order to identify drug specific activation of distinct

apoptosis pathways.

13

In the studies presented, we found some differences in apoptosis induced by these five

anticancer drugs: (1) the anticancer drug-induced apoptosis appears in different time

kinetics, etoposide and cytarabine were found to be early acting drugs, while 4-hydroxy-

cyclophosphamide, doxorubicin, and methotrexate were late acting drugs. (2)

Interestingly, higher doses of cytarabine induce less apoptosis, whereas lower doses of

cytarabine induce more apoptosis. (3) Etoposide strongly induced caspases activation,

compared to cytarabine, 4-hydroxy-cyclophosphamide, doxorubicin and methotrexate.

Besides this difference, we also found that all drugs induced apoptosis in a similar manner.

Both activation of mitochondrial signaling and caspase activation were essential for

execution of programmed cell death (PCD) induced by anticancer drugs.

14

2. Materials and Methods

2. 1. Materials

2.1.1. Reagents and equipment for cell culture

Human Leukemia T-cell line Jurkat American Type Culture Collection

Human Leukemia T-cell line H9 American Type Culture Collection

Human neuroblastoma cell line SHEP American Type Culture Collection

RPMI 1640 medium Life Technologies, Paisley, Scotland

Penicillin-Streptomycin Life Technologies, Paisley, Scotland

L-Glutamine Life Technologies, Paisley, Scotland

Fetal Calf Serum (FCS) Biochrom KG, Berlin, Germany

HEPES- Buffer (1 M) Biochrom, Berlin, Germany

Trypsin/EDTA Biochrom, Berlin, Germany

Trypan Blue Solution (0.4%) Sigma-Aldrich, England

Safety Cabinet Heraeus, Germany

CO2 Incubator WTC binder, Germany

Inver Microscope Leika, Portugal

Tissue Culture Flask Becton Dickinson Labware, England

Tissue Culture Plate Becton Dickinson Labware, USA

Sterile Syringe Becton Dickinson Labware, Germany

Sterile Pipette Becton Dickinson Labware, USA

Pipetter Bilson, France

Sterile Filter Schleicher & Schnell, Germany

2.1.2. Reagents and equipment for flow cytometric analysis

PBS Biochrom KG, Berlin, Germany

HANKS' Life Technologies, Paisley, Scotland

Annexin V FITC Boehringer Mannheim, Germany

15

Steofundin B/Braun, Germany

Propidium Iodide Sigma-Aldrich Chemie, Germany

Triton X-100 Sigma-Aldrich Chemie, Germany

Trinatriumcitrate Dihydrate Sigma-Aldrich Chemie, Switzerland

Paraformaldehyde (PFA) Merck, Darmstadt, Germany

β-Mercaptoethernol Merck, Darmstadt, Germany

Protein A Sigma-Aldrich Chemie, USA

z-VAD.fmk Enzyme Systems, San Diego, USA

(Z-Val-Ala-Asp (Ome)-FMK)

Ethanol Merck, Darmstadt, Germany

3,3'-Dihexyloxycarbocyanine Iodide Mo Bi Tec, Netherlands

(DiOC6 (3))

Dimethyd Sulfoxide (DMSO) Serva Electrophoresis, Germany

Bovine Serum Albumin (BSA) Boehringer Ingelheim, Germany

Sodium Azide Sigma, USA

Optimized Sheath Fluid Becton Dickinson, Belgium

Flow Cytometry (FACSCalibur) Becton Dickinson, Heiderberg,

Germany

Thermobath Sink Elvo Labortechnik, Germany

Vortex Scientific Industries, USA

Centrifuge (Varifuge 3.0R) Heraeus, Germany

Centrifuge Tube (Polypropylene Conical Tube) Becton Dickinson Labware, France

2.1.3. Reagents and equipment for western blot

Tris Base Sigma, USA

Glycin Carl Roth, Karlsruhe, Germany

Glycerol J.T. Baker, Holland

Skim Milk Powder Merck, Darmstadt, Germany

Bromophenol Blue Sigma, USA

Sodium Salt (SDS) Sigma-Aldrich Chemie, Germany

Methanol Merck, Darmstadt, Germany

Tween 20 Gerbu Biotecknik, Germany

Protease Inhibitor Cocktail Sigma, USA

16

EDTA Gerbu Biotechnik, Germany

EGTA Sigma-Aldrich Chemie, Germany

Dithiothreitol (DTT) Sigma, USA

BCA Protein Assay Reagent Pierce, USA

KCl Merck, Darmstadt, Germany

MgCl2 Merck, Darmstadt, Germany

HEPES Carl Roth, Karlsruhe, Germany

Sucrose Merck, Darmstadt, Germany

Eppendorf Centrifuge 5417R Eppendorf-Netheler-Hinz, Germany

Mini Centrifuge MS Laborgerät, Heidelberg, Germany

-200C refrigerator Liebher, Italy

-800C refrigerator Heraeus, Germany

Electrophoresis Cell (Criterion Cell) Bio-Rad, USA

Semi Dry Transfer Cell Bio-Rad, USA

Precast Gel Bio-Rad, USA

Electrophoresis Power Supply EP300 Pharmmcia Biotech, Sweden

Thermomixer Eppendorf-Netheler-Hiny, Germany

Hyperfilm Amersham Pharmacia Biotech,

England

Nitrocellulose Membrane (Hybond ECL) Amersham Pharmacia Biotech,

England

Gel-Blotting-Paper Merck Eurolab, Germany

Molecular Weight Marker (Rainbow) Amersham Pharmacia Biotech,

England

Western Blotting Regents (ECL) Amersham Pharmacia Biotech,

England

X-ray Film Processor (Hyper Processor) Amersham Life Science. England

UV/Visible Spectrophotometer Phamacia Biotech, England

(Ultrspec 2000)

Digital pH Meter (210A) Orion Research, Boston, USA

Platform Shaker (Ploymax 1040) Heidolph, Germany

2.1.4. Anticancer drugs

17

Etoposide Sigma-Aldrich Chemie, Germany

Cytarabine (Ara-c) Pfizer, Germany

4-Hydroxy-cyclophosphamide Asta, Germany

Doxorubicin Pharmcia, Italy

Methotrexate Lederle, Germany

2.1.5. Antibodies

Anti-CD95 monoclonal antibody (IgG3)

Mouse IgG1,κ (MOPC-21)(M9269) Sigma-Aldrich, Germany

Mouse IgG2b (clone DAK-G09)(X 0944) DAKO, Denmark

(Isotype, Negative Control)

Goat F (ab') 2 Anti-Mouse-IgG2b-FITC Southern Biotechnology Associates

(GAM IgG 2b FITC) Birmingham, USA

Anti-mouse Cytochrome c PharMingen, USA

(65981A) (clone 7H8.2C12)

Anti-cytochrome oxidase subunit IV mAb Molecular Probes, Germany

(COX-IV) (A6431, clone 20E8-C12)

Anti-Caspase-8/Flice mAb Kindly Provided by Prof. Krammer,

(clone C15, hybridoma supernatant) DKFZ Heidelberg, Germany

Anti-Caspase-3/CPP32 mAb Transduction Laboratories, Lexington,

(C31720) KY

Anti-PARP mAb PharMingen, USA

(65196A, clone C2-10)

Anti-Fas Ligand/CD95L mAb (F37720) Transduction Laboratories

Anti-RIP mAb (R41220) Transduction Laboratories

Anti-FADD mAb (F36620) Transduction Laboratories

Anti-Human Bid (AF846) R&D Systems, England

Anti-human Bcl-2 mAb PharMingen, San Diego, CA

(65111A, clone Bcl-2/100)

Anti-β-Actin mAb Sigma, USA

(A-5441, clone AC-15)

Anti-Mouse IgG-HRP Santa Cruz Biotechnology, Germany

(sc-2005, HRP-conjugate)

18

Anti-Rabbit IgG-HRP Santa Cruz Biotechnology, Germany

(sc-2004, HRP-conjugate)

19

2.2. Methods

2.2.1. Cell culture

Human leukemia T-cell lines Jurkat, H9, and Neuroblastoma (Shep) cell line were

obtained from American Type Culture Collection (Rockville, MD) and cultured in 75-cm2

tissue culture flasks in RPMI-1640 medium containing 10% heat-inactivated fetal calf

serum (FCS), 100 U of penicillin per milliliter, 0.1mg streptomycin per milliliter, 2 mM

glutamine, and 10 mmol/L HEPES. Cells were grown at 370C in a 5% CO2 atmosphere and

maintained in log phase.

2.2.2. Cell preservation and reconstitution

When being in a best growing phase, cells were spun down and washed 3 time with PBS,

resuspended with FCS plus 10% DMSO. Aliquots of 1 x 106 cells were transferred into a

cryogenic vial and frozen at -800C. For long-term storage, the frozen cells were placed in

liquid nitrogen. For reconstitution, a face guard and protective gloves and clothing must

be worn whenever an vial is removed from liquid nitrogen, because the vial that has been

submerged in liquid nitrogen can explode upon removal if it has not been properly sealed

and the plastic fragments fly at high force in all directions creating a hazard. The frozen

cells were thawed by incubation of cryogenic vials in a covered water bath at 370C for 1

min and washed with prewarmed medium before resuspension with the corresponding

medium.

2.2.3. Cell stimulation

Anti-Apo-1 mAb 1mg/ml were kept as stock solution at -200C. Etoposide (Eto) was

dissolved in Dimethyd Sulfoxide (DMSO) at a concentration of 20 mg/ml, Methotrexate

(MTX) dissolved in 0.1 N NaOH at a concentration of 20mg/ml, doxorubicin (Doxo) and

cytarabine (Ara-C) dissolved in sterile distilled water at a concentration of 1mg/ml and

kept as stock solution at -200C. 4-hydroxy-cyclophosphamide (4-HCP) was dissolved in

sterile distilled water at a concentration of 1mg/ml and kept as stock solution at -800C.

Cyclophosphamide, which is used in patients, is not an appropriate stimulus in vitro,

20

because this compound must undergo hydroxylation at the 4-carbon position before

expressing cytotoxic activity; this reaction is catalyzed by hepatic microsomal enzymes.

So 4-hydroxy-cyclophosphamide must be used in vitro to get the same effect as the clinic

use.

Prior to stimulation, cells were incubated 24 hours in 75-cm2 cell culture flasks in same

medium and then seeded in 24-well plate for stimulation. Cells were stimulated with

agonist Anti-Apo-1 (anti-CD95) mAb or etoposide, cytarabine, 4-hydroxy-

cyclophosphamide, doxorubicin, and Methotrexate, or irradiated using a 137Cs source (2 x

415 Ci) at the doses and time points as indicated in the individual figure legends. Control

culture were treated with the appropriate amount of DMSO and NaOH, used as solvent for

some anticancer drugs and peptide inhibitor. Cells were harvested at the appropriate time

points, and then subjected to various processing procedures for the different purpose of

analysis.

2.2.4. Inhibitor studies

Benzyloxycarbonyl-Val-Ala-Asp fluoromethyl ketone (zVAD-fmk) was dissolved in

DMSO at a concentration of 20 mM and kept as solution at -200C.

For the inhibitor studies, cells cultured as described above were treated with 50 or 100 µM

zVAD-fmk prior to the addition of stimulus, or only using zVAD-fmk without stimulus as

control.

2.2.5. Flow cytometry

2.2.5.1. Analysis of annexin V and PI positive cells

A. Annexin V and PI staining

For double labeling procedures, after exposure to the apoptotic stimulus, 5 x 105 cells were

Harvested into a 5 ml test tube and washed with annexin V buffer (140 mM NaCl, 2.5 mM

CaCl2, 10 mM Hepes, pH 7.4) for 7 min, at 40C, 1300 rpm. And then the pellets were

21

resuspended with annexin V working solution (1 µl Annexin V-FITC and 19 µl buffer) and

incubated for 15 min, at 40C in the dark. The cells were washed again with buffer at 1300

rpm for 7 min, at 40C. Before the measurement, the cells were added 10 µl propidium

iodide (PI) work solution (20 µg/ml) 10 µl (the concentration of PI work solution is 1

mg/ml).

B. Flow cytometry

Labeled cells were suspended in 300 µl buffer. Flow cytometric analysis was performed

on a FACSCalibur flow cytometry with an excitation wavelength of 488 nm. Data

acquisition and analysis were performed by the CellQuest software (Becton Dickinson).

30.000 events were collected for each analysis. Cell debris was excluded by setting

appropriate light scatter gates. Photomultiplier voltages were adjusted to have the

unlabeled cell population fall in the first decade of fluorescence. Cells labeled with only -

FITC or PI were used to adjust the compensation. Annexin V-FITC were detected through

the FL1 channel, equipped with a 530-nm (20-nm band pass) filter. PI were detected

through the FL2 channel, equipped with a 575-nm (20-nm band pass) filter. The data were

acquired and analyzed with CELLQuest software (Becton Dickinson).

2.2.5.2. Quantification of DNA fragmentation

After stimulated, Aliquots 5 x 105 cells were washed with PBS without Ca++ at 1300 rpm

for 7 min, at 40C. The cell pellets were gently resuspended in 525 µl hypotonic lysis

solution (PI 25 µg in 0.1% trinatriumcitrate-dihydrate 250 µl plus 0.1% Triton X-100

250µl) and incubated at 4 0C overnight in the dark. This hypotonic lysis solution can break

the cell membrane and let PI stain the nuclear, because early during apoptosis the cell

membrane is still intact. Cells were analyzed for DNA content using flow cytometry

(FACSCalibur) by examining 10,000 events with excitation wavelength of 488 nm. The

emission wavelengths were detected through the FL3 channel, equipped with a 650 nm

(20-nm band pass) filter. The data were acquired and analyzed with CELLQuest software

(Becton Dickinson).

2.2.5.3. Analysis of mitochondria membrane potential (∆Ψm)

22

To evaluate ∆Ψm, the cationic lipophilic fluorochrome 3,3'-dihexyloxycarbocyanine iodide

(DiOC6(3)) was used. 5 x 105 cells were placed into a 5 ml test tube, added 3 ml PBS, and

centrifuged at 1300 rpm for 7 min, at 40C. The supernatants were removed by aspiration.

The cell pellets were mixed gently with 20 nM DiOC6 (3) and incubated for 15 min, at

370C in the dark. DiOC6(3) was prepared from a 40 µM stock solution in DMSO. This

solution was diluted with PBS, pH 7.4, to a 400 nM working solution. Cells were washed

once with PBS; cell suspensions were prepared for flow cytometry. The live cells were

gated and DiOC6(3) was detected through the FL1 channel. ∆Ψm low cells were those

displaying DiOC6(3) fluorescence less than the fluorescence of control cells in the absence

of the apoptotic stimulus. The data were acquired and analyzed with CELLQuest software

(Becton Dickinson).

2.2.5.4. Quantification of cytoplasmic cytochrome c

5 x 105 cells were placed into a 5 ml test tube, added 3 ml wash solution (HANKS' plus 1%

BSA and 0.1% sodium azide) and centrifuged at 1300 rpm for 7 min, at 40C. The pellets

were resuspended and fixed for 20 min, at 40C with 100 µl 4% paraformaldehyde (PFA),

which was freshly prepared. The cells were washed again as described above, resuspened

with 50 µl 0.2% saponin in PBS containing 5 µl Mouse IgG1, k (MOPC-21) for the

purpose of permeabilization and blocking nonspecific binding and incubated for 5 min, at

the room temperature. And then the cells were added with 20 µl first antibody, anti-

cytochrome c (1:20 dilution in wash solution) and incubated for 20 min, at 40C. As a

negative control, in parallel cells were added with only 20 µl mouse IgG2b (isotype Ab)

(1:20 dilution in wash solution) and incubated as the first antibody. The wash step was

repeated. The pellets were suspended with 20 µl second antibody, Goat F (ab')2 Anti-

Mouse-IgG2b-FITC (GAM IgG 2b TITC) (1:20 dilution in wash solution) for 20 min, at

40C. In parallel, cells were added with only second antibody. Wash step was repeated.

Pellets were resuspended with 100 µl 4% PFA for flow cytometric analysis. Cytochrome c

was detected through the FL1 channel with an excitation wavelength of 488 nm.

2.2.6. Cytosolic and mitochondrial extracts preparation

23

Approximately 2.5 x 107 cells were required for each preparation. The cell suspension was

transferred into a 50 ml centrifuge tube. The cells were pelleted by centrifugation at 600 x

g for 5 min, at 40C. The cell pellets were washed twice with 20 ml of cold PBS, pH 7.4. It

is important to remove all PBS form the cell pellet. The cell pellets were resuspended with

800 µl fractionation buffer (20 mM HEPES, pH7.5, 10 mM KCL, 1.5 mM MgCL2, 1 mM

EDTA, 1 mM EGTA, 1 mM dithiothreitol, 250 mM sucrose, and protease inhibitor

cocktail), and incubated for 20 min, on ice. During this time the tubes were tapped from

time to time in order to resuspend the cell pellet.

Cells were then disrupted by 25 strokes with Dounce homogenizer. To establish the

optimum conditions for cell homogenization, the trypan blue exclusion assay, which

discriminates between broken (stained) cells and intact cells (unstained), can be used. For

the trypan blue exclusion test, a 0.4% solution of trypan blue in PBS was diluted 1: 10 with

the cell suspension and was examined under the microscope. After dounce

homogenization, the cell homogenates were transferred to an Eppendorf tube and nuclei,

unbroken cells, and large debris were removed by centrifugation at 800 x g for 10 min, at

40C. Supernatants containing mitochondria were transferred to a new Eppendorf tube and

further centrifuged at 10,000 x g for 25 min, at 40C. The resulting supernatants were saved

as cytosolic extracts at -700C until further analysis. The mitochondrial pellets were lysed

with 100 µl of fractionation buffer. Samples were vortexed from time to time during the 20

min incubation period on ice. This was mitochondrial fraction and could be stored at -700C

until further analysis.

2.2.7. Western blot analysis

2.2.7.1. Cell lysis

After incubation with apoptosis-inducing stimuli for indicated length of time, aliquot of 1 x

107 cells were transferred to 50 ml centrifuge tube and centrifuged at 1300 rpm for 5 min,

at 40C. Pellets were suspended with 2 times cold PBS and centrifuged as described above.

Pellets were solubilized in 500 µl lysis buffer (20 mM HEPES, pH 7.4, 150 mM NaCl, 5

mM EDTA, pH 7.5, 10% Glycerol, 0.5% Triton X-100, and protease inhibitor cocktail)

and incubated for 30 min. Samples were vortexed form time to time during the 30 min

24

incubation period on ice. After lysis, samples were centrifuged at 14000 rpm for 15 min, at

40C. The resulting supernatants containing extraction of proteins were stored at -700C until

further analysis.

2.2.7.2. Quantification of protein

Quantification of protein was carried out with the BCA reagent. The reagent combines the

well-known reduction of Cu++ by protein in an alkaline medium with the highly sensitive

and selective colorimetric detection of the cuprous cation (Cu+) using a unique reagent

containing bicinchoninic acid. The purple-colored reaction product of this assay is formed

by the chelation of two molecules of BCA with one cuprous ion. This water soluble

complex was detected with spectrophotometer at 562 nm. Standard working curve was

obtained from the different dilution of BSA.

2.2.7.3. Electrophoreses

A. Loading and running samples

40 µg protein per well were mixed with 5x SDS loading buffer (50 mM Tris-Cl (pH 6.8),

1% SDS, 0.05% bromophenol blue, 5% glycerol, and 10% β-mercaptoethanol) and boiled

for 5 min to denature the proteins. Precast gel was mounted in the electrophoresis

apparatus and wells were washed immediately with deionized water. Running buffer (25

mM Tris, 250 mM glycine, 0.5% SDS) was added to the top and bottom reservoirs. Any

bubble was removed that become trapped at the bottom of the gel between the glass plates.

This is best done with a bent hypodermic needle attached to a syringe. Samples were

loaded into the bottom of the wells and molecule weight marker was loaded in the same

time.

The Electrophoresis Apparatus was connected to an electric power supply. 80 V was

applied to the gel and the gel was run until the bromophenol blue reaches the bottom of the

resolving gel (about 3 hours). The power supply was turned off.

B. Blotting

25

The graphite plates of Semi Dry Apparatus were rinsed with distilled water and any bead

of liquid was wiped off. Gloves were worn. Six pieces of Gel-Blotting Paper and one piece

of Nitrocellulose Membrane were cut to the exact size of the SDS-polyacrylamide gel. If

the paper or membrane was larger than the gel, there was a good chance that the

overhanging edges of the paper and the filter would touch, causing a short circuit that

would prevent the transfer of protein from the gel. One corner of the filter was marked

with a soft-lead pencil. The one piece of nitrocellulose membrane and the six piece of

paper were soaked in a shallow tray containing a small amount of transfer buffer (39 mM

glycine, 48 mM Tris base, 0.037% SDS, 20% methanol). The three sheets of paper and one

piece of the nitrocellulose membrane were placed on the bench (the bottom was the anode)

one on top of the other so that they were exactly aligned. The glass plates holding the SDS-

plyacrylamide gel were removed from the electrophoresis tank and the gel was placed

exactly on the top of the nitrocellulose membrane. The final three sheets of paper were

placed on the gel. Any air bubbles was squeezed out with a glass pipette. The electrical

leads were connected and 120 mA current was applied for a period of 145 min.

2.2.7.4. Immunoblotting

A. Block non-specific binding

The nitrocellulose membrane was taken out of the Semi Dry Transfer Apparatus, soaked

into blocking buffer (5% nonfat dried milk, 0.1% Tween 20 and PBS) and incubated for 1

hour at room temperature with gentle agitation on a platform shaker or incubated overnight

at 40C. This step was used for blocking unspecific bindings. After that, the membrane was

washed for 10 min with wash buffer (PBS-T) (0.1% Tween and PBS). The wash step was

repeated three times.

B. Binding of the primary antibody

Primary antibodies were suitable diluted in 10-15 ml with wash buffer: mouse anti-

cytochrome c monoclonal antibody 1:1000; mouse anti-cytochrome oxidase (subunit IV)

(COX IV) monoclonal antibody 1:1000; mouse anti-caspase-8/FLICE monoclonal

antibody C15 1:5 dilution of hybridoma supernatant; mouse anti-caspase-3/CPP32

monoclonal antibody 1:1000; mouse anti-PARP monoclonal antibody 1:2500; mouse anti-

26

Fas ligand/CD95L monoclonal antibody 1:1000; mouse ant-RIP monoclonal antibody

1:1000; mouse ant-FADD monoclonal antibody 1:250; rabbit anti-bid polyclonal antibody

1:1000; mouse ant-Bcl-2 monoclonal antibody 1:500; mouse anti-β-actin antibody 1:5000.

The membrane was incubated with the primary antibody as described above at room

temperature for 1 hour or overnight at 4 0C on a platform shaker with gentle agitation.

After that, the membrane was washed for three times with PBS-T as described above.

C. Binding of the secondary antibody

Second antibodies were suitable diluted in 15ml with wash buffer: The horseradish

peroxidase-conjugated anti-mouse IgG or horseradish peroxidase-conjugated anti-rabbit

IgG 1:5000. The membrane was incubated with the secondary antibody at room

temperature for 1 hour on a platform shaker. After that, the membrane was washed for

three times as described above.

2.2.7.5. Detection

The ECL detection reagents were take out of bottle, mixed at an equal volume of detection

solution 1 with detection solution 2 to give sufficient liquid to cover the membrane and

incubated for 1 min, at room temperature. The blots, protein side up, were placed in the

film cassette. All the wok should be done as quickly as possible. The film was developed

on X-ray Film Processor (Hyper Processor) after an appropriate length of exposure time.

Sometimes it may take one or several hours to see the faint cleaved fragments.

2.2.7.6. Reprobing membranes

For protein loading equivalent control, the membrane was stripped and reprobed. The

membrane was washed with 20 ml PBS-T for 30 min, at room temperature on a platform

shaker and then submerged in 20 ml stripping buffer (0.2 M NaOH) for 10 min, at room

temperature. The membrane washed with deionized water for 10 min and with PBS-T for 2

x 15 min. After that, the membrane was undergone blocking, immunoblotting and

detection procedure as described above. But if the reprobed protein was β-actin, the

blocking step can be skipped.

27

2.2.8. Statistical analysis

All experiments were performed in triplicate unless otherwise noted; results are expressed

as mean ± standard deviation.

28

3. Results

3.1. Induction of apoptosis by anticancer drugs

3.1.1. Apoptosis induced by treatment of Jurkat cells with

etoposide, cytarabine, 4-hydroxy-cyclophosphamide,

doxorubicin and methotrexate

3.1.1.1. Dose and time kinetics for anticancer drug treatment

To investigate whether there are difference of induction of apoptosis by different

anticancer drugs in dose and time, Jurkat cells were treated with various doses of

anticancer drugs for different time points. All conditions were performed in triplicate. The

cells were dual-stained with annexin V-FITC and propidium iodide (PI) and analyzed by

flow cytometry. Cells without stimuli were used as control.

These results demonstrate that there was a time-dependent fashion in annexin V/PI double

positive cells induced by etoposide, cytarabine (Ara-c), 4-hydroxy-cyclophosphamide,

doxorubicin, methotrexate as shown in Figure 1. Apoptosis in response to etoposide and

cytarabine was early, whereas apoptosis in response to 4-hydroxy-cyclophosphamide,

doxorubicin, methotrexate was significantly late. At 12 hours, the observed maximal

apoptosis for etoposide at the maximal concentration, 100 µg/ml, was about 50%, for

cytarabine at the concentration of 1-10 µg/ml was greater than 35%. In contrast, at same

time point, the observed maximal apoptosis for 4-hydroxy-cyclophosphamide,

doxorubicin, and methotrexate at their maximal concentration (4-hydroxy-

cyclophosphamide 5µg/ml, doxorubicin 1µg/ml, methotrexate 500µg/ml) was no more

than 15%. Until 36 hours, the observed apoptosis for 4-hydroxoy-cyclophosphamide at

maximal concentration was greater than 50%; until 24 hours, the observed apoptosis for

doxorubicin at maximal concentration was still less than 35%.

This data suggested that the effect of etoposide and cytarabine was earlier than 4-hydroxy-

cyclophosphamide, doxorubicin and methotrexate. Collectively, these results demonstrate

29

A.

B.

Figure 1. Different anticancer drugs induced apoptosis in a different time and dose fashion.

Jurkat cells were cultured in normal growth medium and then either left untreated (medium, as

control) or treated with etoposide (Eto), cytarabine (Ara-c), 4-hydroxy-cyclophosphamide (4-HCP),

doxorubicin (Doxo) and methotrexate (MTX) respectively at the indicated time points and doses.

Apoptosis was assessed by FACS analysis of annexin V and propidium iodide double staining.

0

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30

that etoposide and cytarabine were early acting drugs, whereas 4-hydroxy-

cyclophosphamide, doxorubicin and methotrexate were late acting drugs.

The cells underwent dose-dependent apoptosis in response to anticancer drugs, except for

cytarabine, The doses of etoposide, 10-100µg/ml induced a similar amount of apoptosis

(about 50%), at 12 hours. The maximum apoptosis induced by 4-hydroxy-

cyclophosphamide at maximum concentration 5µg/ml at 36 hours was nearly 55%. The

maximum apoptosis induced by doxorubicin at maximum concentration 1µg/ml at 36

hours was 70%. The amount of apoptosis induced by methotrexate at concentration of 1-

500µg/ml at 36 hours was not big different, from 25% to 32%.

Interestingly, we found an exceptional pattern of dose effect relationship for cytarabine.

Higher concentration of cytarabine induced less apoptosis, whereas lower concentration

induced more apoptosis. At 12 hours, e.g. a concentration of 1µg/ml, 5µg/ml or 10µg/ml

produced: about 35% apoptotic cells, whereas a concentration of 100µg/ml and 500µg/ml

produced less than 20% apoptotic cells. The data indicated different mechanisms of

apoptosis induction at low or high concentration.

3.1.1.2. Comparison of different apoptotic signs induced by different

drugs

Since we found differences in time of apoptosis induction, we further investigated that the

different drugs induced different signs of apoptosis. For this, cells were treated with the

drugs in one dose and different time points and apoptosis was detected with forward/side

scatter, annexin V/PI double staining and DNA fragmentation by flow cytometry.

Morphological, biochemical, and molecular changes that occur during apoptosis serve as

specific markers to identify apoptotic cells by cytometry. An early event of apoptosis is

dehydration, which leads to cell shrinkage. This change is reflected by an alteration in the

way the cells scatter the light of the laser beam in a flow cytometer. The intensity of light

scattered by apoptotic cells in a forward direction along the laser beam, which correlates

with cell size, is diminished. Late apoptotic cells or individual apoptotic bodies are

characterized by a low intensity of the forward scatter signal. Chromatin condensation,

31

A. B.

C. D.

Figure 2. Detection of early drug-induced apoptosis by different apoptosis assays. Jurkat cellswere cultured in normal growth medium and then either left untreated (medium, as control) ortreated with epotoside (Eto 30µg/ml) and cytarabine (Ara-c 30µg/ml) respectively at 37 0C in a CO2

incubator for the indicated time points. (A) Dead cells were observed according to the morphologicand granularity changes on flow cytometry. (B) Hypodiploid cells were detected by flowcytometry after the cell's nuclei were stained with propidium iodide; (C) Annexin V positive cellswere measured by flow cytometry after the cells were stained with annexin V. (D) PI positive cellswere assessed by flow cytometry after the cells were stained with propidium iodide.

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32

which often is followed by nuclear fragmentation, is another characteristic feature of

apoptosis. These changes increase the propensity of the cell to refract light, which may be

manifested by a transient increase in the intensity of light scattered at a 900 angle in the

direction of the laser beam (side scatter). Analysis of the forward and side light scatter

signals of the cells thus provide the means to identify apoptotic cells by flow cytometry on

the basis of their properties without measurement of fluorescence.

From the FSC versus SSC dot plot of FACSCarlibur, the population of apoptosis cells was

easily distinguished from the population of live cells. The population of apoptosis cells was

formed with a low forward scatter (FSC) and high side scatter (SSC) profile, which is a

characteristic of apoptotic cells. More apoptotic cells appeared in 12 hours for early drugs,

whereas apoptotic cells appeared in 24 hours for late drugs as shown in Figure 2A, 3A.

There was no difference in the apoptotic phenotype induced by different drugs in

forward/side scatter analysis.

DNA fragmentation is so characteristic an event of apoptosis that it is considered to be a

hallmark of this mode of cell death. Initially, DNA is cleaved at the sites of attachment of

chromatin loops to the nuclear matrix. Which results in discrete 50- to 300-kb size

fragments. Subsequently, although not in every cell type, DNA is cleaved at the

internucleosomal sections. As a result, the products of DNA cleavage are discontinuous,

nucleosomal and oligonucleosomal DNA fragments of approximately 180 bp and multiples

of this size. The analysis of the cellular DNA content of apoptotic cells from which

degraded DNA was extracted reveals them as cells with fractional DNA content,

represented by the sub-G1 peak on DNA content frequency histograms. This approach is

currently the most frequently used in flow cytometry to identify and quantify apoptotic

cells.

The DNA fragmentation can be measured by analysis of propidium iodide stained nuclei

on flow cytometry. The reduced DNA content of apoptotic nuclei resulted in a unequivocal

hypodiploid DNA (sub G1) peak, which was easily discriminable from the narrow peak of

cells with normal (diploid) DNA content in red fluorescence channels. The percentage of

apoptotic nuclei was quantified. This assay is more sensitive than forward/side scatter

assay. In the same time points (except for etoposide in 24 hours), the percentage of

apoptosis was detected by this assay was higher than that detected by forward/side scatter

33

A. B.

C. D.

Figure 3. Detection of late drug-induced apoptosis by different apoptosis assays. Jurkat cellswere cultured in normal growth medium and then either left untreated (medium, as control) ortreated with 4-hydroxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml) andmethotrexate (MTX 30µg/ml) respectively at 37 0C in a CO2 incubator for the indicated time points.(A) Dead cells were observed according to the morphologic and granularity changes on flowcytometry. (B) Hypodiploid cells were detected by flow cytometry after the cell's nuclei werestained with propidium iodide; (C) Annexin V positive cells were measured by flow cytometryafter the cells were stained with annexin V. (D) PI positive cells were assessed by flow cytometryafter the cells were stained with propidium iodide.

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34

as shown in Figure 2A, 2B, 3A, 3B. For etoposide in 24 hours, more than 85% cell death

(forward/side scatter), in this situation this assay is not sensitive. From the results of

doxorubicin in 24 hours, the percentage of dead cells was lower than 70% (forward/side

scatter). In this situation, this assay was also more sensitive than forward/side scatter.

In live cells the plasma membrane phospholipids, phosphatidylcholine, and spingomyelin

are exposed on the external leaflet of the lipid bilayer while phophatidylserine is almost

exclusively on the inner surface. The phospholipid asymmetry leading to exposure of

phosphatidylsereing on the outside of the plasma membrane occurs early during apoptosis.

This change is detected by annexin V-fluorescein isothiocyanate (FITC) conjugate, which

preferentially binds to negatively charged phospholipids such as phosphatidylserine. The

staining is done in combination with propidium iodide (PI), which is excluded from live

and early apoptotic cells but stains DNA and RAN in late apoptotic and necrotic cells, the

plasma membranes of which are disrupted. In this experiment system, annexin V staining

assay for detecting apoptosis was most sensitive, especially for etoposide as shown in Fig.

2C. 3C. At 12 hours, Annexin V positive cells-induced by etoposide was 80%, which was

two times greater than morphologic changes (FSC/SSC) at the same time point.

The duration of apoptosis is relatively short and variable depending on cell type, inducer of

apoptosis. Each of the methods presented above has its advantages and suffers limitations.

So combination of several apoptosis assay can provide a more definite identification of

apoptotic cells.

Among these four assays detecting apoptosis, a similar pattern of anticancer drug-induced

apoptosis was found that the number of apoptotic cells increased over the increasing time

period. The results further confirm that the etoposide and cytarabine effect was earlier than

4-hydroxy-cyclophosphamide, doxorubicin and methotrexate.

3.1.1.3. Annexin V single positive cells appear through treatment with

early anticancer drugs, but not with late anticancer drugs

Previous studies showed that after cells were stimulated by anticancer drugs and agonistic

anti-CD95-antibodies, through staining cells with annexin V-FITC and PI, it is possible to

35

A. B.

Figure 4. Difference of annexin V single positive for early and late drugs. Jurkat cells werecultured in normal growth medium and then either left untreated (medium, as control) or treatedwith anti-CD95 antibody (20ng/ml) plus protein A (2.5ng/ml), etoposide (Eto 30µg/ml), cytarabine(Ara-c 30µg/ml), 4-hydroxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml)and methotrexate (MTX 30µg/ml) respectively at 370C in a CO2 incubator for the indicated timepoints. (A) Annexin V positive cells in gate 1 were assessed by flow cytometry after the cells weredouble stained with annexin V and propidium iodide. (B) Flow cytometry profiles of forwardscatter versus side scatter and propidium iodide staining versus annexin V staining.

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Anti-CD95

R1

R290.18%

9.76%

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44.88%

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3.22%

39.51%

58.09%

Anti-CD95

3h

6h

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10.42%

R1R2

62.52%

37.46%

7.92%

2.04%

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8.67%

12h

48h

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2.35%

26.36%

39.89%

37.74%

R1

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45.03%

R1

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12.17%6h

12h

Eto Eto

Forward Scatter FL2 - PI

36

detect live, nonapoptotic cells (annexin V/PI double negative), early apoptotic cells

(annexin V single positive), and late apoptotic or necrotic cells (annexin V/PI double

positive) by flow cytometry. But in this experiment system, the annexin V single positive

profile of early and late drugs was different. After cells were stained with annexin V and

PI, several subpopulation were visualized in the quadrant profile. In the FSC/SSC dot plot,

the apoptotic cells were nearly same for anti-CD95 in 3h, 6h, compared with etoposide in 6

hours, 12 hours and 4-hydroxy-cyclophosphamide in 12 hours, 48 hours, whereas in the

annexin V versus PI quadrant profile, there was big difference between etoposide and 4-

hydroxy-cyclophosphamide in annexin V single positive. Etoposide induces more annexin

V single positive (upper left quadrant) like anti-CD95. Especially in the early time points

such as in 3 hours (anti-CD95) and in 6 hours (etoposide), annexin V single positive much

greater than annexin V/PI double positive (upper right quadrant). In contrast, 4-hydoxy-

cyclophosphamide induce much greater annexin V/PI double positive than single positive

in 12 hours and 48 hours as shown in Figure 4. This means that PS externalization is an

early event in apoptosis induced by anti-CD95 and etoposide, In contrast, in apoptosis

induced by late drugs, PS externalization occurs as the same time as disruption of

membrane integrity.

3.1.2. Comparison of apoptosis induced by anticancer drugs with

apoptosis induced by death receptor signaling and γ-radiation

3.1.2.1. Agonistic anti-CD95 antibody-induced apoptosis

One of the best-defined apoptotic pathways is mediated by the death receptor CD95 (Apo-

1/Fas). In order to compare anti-CD95 antibody-induced apoptosis with anticancer drug-

induced apoptosis, we investigated the agonist anti-CD95 antibody induce apoptosis using

the same approach.

Jurkat cells were treated with the agonist anti-CD95 Ig3 antibody plus protein A as shown

in Figure 5 and apoptosis was detected with forward/side scatter, annexin V/PI double

staining on flow cytometry FACSCalibur. Agonist anti-CD95 antibody induce apoptosis

was more rapidly and efficiently than anticancer drugs. In 6 hours more than 40% cells

underwent apoptotic morphologic changes. In this situation, the pattern of membrane

37

A.

B.

C.

Figure 5. Detection of anti-CD95-induced apoptosis by different apoptosis assays. Jurkat cellswere cultured in normal growth medium and then either left untreated (medium, as control) ortreated with anti-CD95 antibody (20ng/ml) plus protein A (2.5ng/ml) at 370C in a CO2 incubatorfor the indicated time points. (A) Dead cells were observed according to the morphologic andgranularity changes on flow cytometry. (B) Annexin V positive cells were measured by flowcytometry after the cells were stained with annexin V. (C) PI positive cells were assessed by flowcytometry after the cells were stained with propidium iodide.

0

20

40

60

80

100

0 3 6 9

Time (hours)

Medium

Apo20ng

0

20

40

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100

0 3 6 9

Time (hours)

Medium

Apo20ng

0

20

40

60

80

100

0 3 6 9

Time (hours)

Medium

Apo20ng

38

A. B.

C. D.

Figure 6. Detection of γ-radiation-induced apoptosis by different apoptosis assays.Jurkat cells were cultured in normal growth medium and then either left untreated ( ) or treatedwith γ-radation 20 Gy ( ), γ-radation 10 Gy ( ), γ-radation 5 Gy ( ). After irradiation, the cellswere incubated at 370C in a CO2 incubator for the indicated time points. (A) Dead cells wereobserved according to the morphologic and granularity changes on flow cytometry. (B)Hypodiploid cells were detected by flow cytometry after the cell's nuclei were stained withpropidium iodide; (C) Annexin V positive cells were measured by flow cytometry after the cellswere stained with annexin V-FITC. (D) PI positive cells were assessed by flow cytometry after thecells were stained with propidium iodide.

0

20

40

60

80

0 12 24 36 48

Time (hours)

60

20

40

60

80

0 12 24 36 48

Time (hours)

6

0

20

40

60

80

0 12 24 36 48

Time (hours)

60

20

40

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80

0 12 24 36 48

Time (hours)

6

39

integrity loss was as the same as that of morphologic changes. In these three assays

detecting apoptosis, annexin V positive was most sensitive. In 6 hours nearly 100% cells

were annexin V positive. It suggested that in anti-CD95 antibody-induced apoptosis, PS

externalization was a very early event.

3.1.2.2. γ-radiation-induced apoptosis

γ-radiation is a common procedure to treat cancer patients in the clinic. In the present

study, we included the γ-radiation-induced apoptosis to compare with anticancer drug-

induced apoptosis.

Jurkat cells were treated with γ-radiation with the dose used in the clinic patients. After

irradiation, the cells were incubated at 370C and 5% CO2 for different time points as shown

in Figure 6. γ-radiation-induced apoptosis was time- and dose-dependent. These apoptotic

features were like those induced by late drugs. PS externalization (annexin V positive)

appears at the similar time point as morphologic changes, DNA fragmentation and

membrane integrity loss. In other words, PS externalization was not an early event in γ-

radiation-induced apoptosis.

3.2. Expression of death receptor associated

molecules induced by anticancer drugs

Previous studies found that the CD95 system, which is known as a key regulator of the

immune system, also mediated drug-induced apoptosis of leukemia. To investigate whether

anticancer drug treatment of Jurkat cells stimulate expression of death receptor associated

molecules, we detected CD95 ligand (Apo-1/ Fas ligand), FADD (Fas-associated death

domain protein) and RIP (receptor interacting protein) by western blot.

Jurkat cells were either left untreated or treated with etoposide, cytarabine, 4-hydroxy-

cyclophosphamide, doxorubicin, methotrexate and anti-CD95 antibody plus protein A

respectively as described in Figure 7. At the indicated time points, cells were collected.

Protein extracts were prepared and fractionated by SDS-PAGE.

40

Figure 7. Expression of death receptor associated molecules induced by anticancer drugs andanti-CD95 antibody. Jurkat cells were cultured in normal growth medium and then either leftuntreated (medium, as control) or treated with etoposide (Eto 30µg/ml), cytarabine (Ara-c30µg/ml), 4-hydroxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml),methotrexate (MTX 30µg/ml) and anti-CD95 antibody (20ng/ml) plus protein A (2.5ng/ml)respectively at 370C in a CO2 incubator for the indicated time points. Total cellular protein 40µgper lane was separated by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to nitrocellulose, and probed with antibodies that recognize mouse anti-Fasligand monoclonal antibody, mouse anti-FADD monoclonal antibody and mouse anti-RIPmonoclonal antibody. Arrows indicate the full-length molecules or specific cleavage fragments.Equal protein loading was confirmed by reprobing the stripped blots with mouse anti-β-actinmonoclonal antibody. The results shown are representative of at least three independentexperiments

0 12 24 48 12 24 48 12 24 48

M 4-HCP Doxo MTX

0 6 12 24 6 12 24 3 6 9

M Eto Ara-c Anti-CD95

Time (hours)

Fas Ligand

FADD

β-actin

RIP

Cleaved fragment

β-actin

41

CD95L, 45kDa, and FADD, 24kDa, and RIP, 74kDa, expression were detected by specific

antibody. The results showed anticancer drugs, similar to anti-CD95, upregulate CD95L

and FADD expression. For early drugs, at 6 hours CD95L and FADD increased markedly,

for anti-CD95 antibody, the increase of the expression appeared as early as 3 hours, for late

drugs, the increase appeared at 12 hours. Cleavage of RIP was observed in Figure 7.

Among the stimuli, anti-CD95 antibody and etoposide most efficiently cleave RIP in this

system. These findings suggest CD95 system take part in anticancer drug-inducing

apoptosis in Jurkat cells, at least in part.

3.3. Caspases activation by anticancer drugs

3.3.1. Caspase-8, caspase-3 activation and PARP cleavage

Caspases comprise a family of different cysteine protease that are synthesized as inactive

zymogens and converted to an active complex composed of several heterodimeric subunits.

To investigate differences in caspase activation by different anticancer drugs, we analyzed

activation of caspase-8, caspase-3 and cleavage of caspase substrate PARP. We monitored

the processing of procaspases in immunoblot analyses using antibodies specific to

individual proteases. Jurkat cells were either left untreated or treated with etoposide,

cytarabine, 4-hydroxy-cyclophosphamide, doxorubicin, methotrexate and anti-CD95

antibody plus protein A respectively as described in Figure 8. At the indicated time points,

cells were collected. Protein extracts were prepared and fractionated by SDS-PAGE.

We first investigated the processing of caspase-8, the most proximal caspase during CD-

95-mediated apoptosis. Caspase-8 is synthesized as an inactive precursor of 55 kDa, which

was detected a double protein band, representing the isoforms porcaspase-8a and

porcaspase-8b, and, following formation of a 43-kDa, 41-kDa intermediate cleavage

product, processed to a p18 heterodimer.

Treatment of cells with anticancer drugs resulted in the conversion of the inactive 32-kDa

caspase-3 precursor to the proteolytically cleaved p17 subunit, indicating that caspase-3

was activated during drug-induced apoptosis. In a detailed time-response assay, we further

42

Figure 8. Caspase-8,caspase-3 and PARP are cleaved with different kinetics by early and latedrugs. Jurkat cells were cultured in normal growth medium and then either left untreated (M, ascontrol) or treated with etoposide (Eto 30µg/ml), cytarabine (Ara-c 30µg/ml), 4-hydroxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml), methotrexate (MTX 30µg/ml)and anti-CD95 antibody (20ng/ml) plus protein A (2.5ng/ml) respectively for the indicated timepoints. Total cellular protein 40µg per lane was separated by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to nitrocellulose, and probed with therespective mAb. Arrows indicated the position of full length caspase-8 which exist as two isoformsand it's cleavage fragment p43, p41 and p18, full length caspase-3 and it's cleavage fragment p17,full length PARP and it's leavage fragment p85. Equal protein loading was confirmed by reprobingthe stripped blots with mouse anti-β-actin monoclonal antibody.

Time (hours) 0 6 12 24 6 12 24 3 6 9 0 12 24 48 12 24 48 12 24 48

M Eto Ara-c Anti-CD95 M 4-HCP Doxo MTX

β-actin

PARP p85

β-actin

p17

Caspase-3

β-actin

p18

p43p41

Caspase-8

43

measured the cleavage of poly (ADP-ribose) polymerase (PARP), an enzyme involved in

DNA repair, which is specifically cleaved by caspase-3 during apoptosis. Figure 8

demonstrates that PARP, a 116-kDa protein, was cleaved into the characteristic 85-kDa

fragment in the course of treatment of anticancer drugs and anti-CD95 antibody.

The cleavage pattern of the individual caspase and PARP did not differ between the

anticancer drugs and anti-CD95 antibody stimulation. However, corresponding to the

different kinetics of apoptosis, Anti-CD95 antibody-induced caspase activation was more

rapid and efficient than etoposide and cytarabine; 4-hydoxy-cyclosphamide-, doxorubicin-

and methotrexate-induced caspase activation occurs latest. For etoposide and anti-CD95

antibody-induced caspase-8 cleavage, the active fragment p18 was clearly presented on

blot, but for cytarabine and late drugs, although the p43 and p41 intermediate cleavage

products were clear on the blot, the p18 was not appear on the blot. The reason could be

that the time points are not appropriate for these drugs. For early drug-induced caspase-3

activation, the cleaved fragment p17 was clearly presented on the blot. For late drug-

induced caspase-3 activation, the faint band of p17 also can be seen on the blot. Taken

together, these data showed all anticancer drugs activated caspases .

3.3.2. The caspase inhibitor zVAD-fmk blocks drug-induced

apoptosis

To determine whether anticancer drug-induced apoptosis was dependent upon caspases

activation, cells were pretreated with zVAD-fmk, abroad peptide caspase inhibitor, and

then exposed to the stimuli, etoposide, cytarabine, 4-hydroxy-cyclophosphamide,

doxorubicin, methotrexate or anti-CD95 antibody, respectively, in the indicated dose and

time points as shown in figure 9, 10. In the parallel experiment , cells were treated only

with stimuli. Apoptosis of the cells was determined by forward/side scatter, annexin V/PI

double staining and quantification of DNA fragmentation on flow cytemetery

FACSCalibur. Similar to anti-CD95, early drug (etoposide, cytarabine)-induced apoptosis

was completely inhibited by zVAD-fmk. Late drug (4-hydroxy-cyclophosphamide,

doxorubicin, methotrexate)- induced apoptosis was completely inhibited by zVAD in early

time point; whereas in late time point only partially inhibited. The pan-caspase inhibitor

zVAD-fmk blocked not only externalization of phosphatidylserine (PS) and DNA

44

A. B.

Figure 9. zVAD-fmk inhibits early drug-induced apoptosis. (A) Jurkat cells were cultured innormal growth medium and then either left untreated (M, medium) or treated with etoposide (Eto30µg/ml) and cytarabine (Ara-c 30µg/ml) in the presence or absence of zVAD-fmk (100µM)respectively at 37 0C in a CO2 incubator for the indicated time points. Dead cells, DNAfragmentation, annexin V and PI positive cells were assessed by flow cytometry as described inMaterials and Methods. Data were the mean o f triplicates with standard deviation. Similar resultswere obtained in 3 separated experiments. (B) Representative profiles of DNA fragmentationinduced by anticancer early drugs in the presence or absence of zVAD-fmk (100µM) for theindicated time points

0

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6 12 24Time (hours)

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6 12 24Time (hours)

6h 12h 24h

Medium

Eto

Eto +zVAD

Ara-c

Ara-c +zVAD

M1

2.56%

M1

11.69%

M1

3.81%

M1

5.34%

M1

3.96%

M1

5.05%

M1

58.80%

M1

5.91%

M1

24.72%

M1

3.82%

M1

3.32%

M1

72.27%

M1

3.79%

M1

3.61%

M1

51.61%

Fluorescence Intensity

45

A. B.

Figure 10. zVAD-fmk inhibits late drug-induced apoptosis. (A) Jurkat cells were cultured innormal growth medium and then either left untreated (medium) or treated with 4-hydoxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml) and methotrexate (MTX 30µg/ml) in the presence or absence of zVAD-fmk (100µM) respectively at 37 0C in a CO2 incubatorfor the indicated time points. Dead cells, DNA fragmentation, annexin V and PI positive cells wereassessed by flow cytometry. Data were the mean of triplicates with standard deviation. Simlarresults were obtained in 3 separated experiments. (B) Representative profiles of DNAfragmentation induced by anticancer drugs alone or in the presence of zVAD-fmk (100µM) for theindicated time points.

0

20

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12 24 48Time (hours)

0

20

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12 24 48Time (hours)

0

20

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12 24 48Time (hours)

0

20

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12 24 48Time (hours)

12h 24h 48h

MTX

Medium

4-HCP

4-HCP +zVAD

Doxo

Doxo +zVAD

MTX +zVAD

M15.09%

M110.49%

M14.08%

M1

3.62%

M16.03%

M13.99%

M126.07%

M14.26%

M119.46%

M15.66%

M174.33%

M117.55%

M148.77%

M112.78%

M1

4.95%

M1

12.91%

M1

76.32%

M15.76%

M13.56%

M14.55%

M118.41%

Fluorescence Intensity

46

fragmentation, but also cell morphologic changes and loss of membrane integrity. The

different apoptosis assays presented the same patterns. The ability of zVAD-fmk to

effectively inhibit caspase function suggested that anticancer drug-induced apoptosis is a

caspase-mediated event. This means that in all drugs investigated induction of apoptosis

largely depends on activation of caspases. However partial inhibition at late time points

indicated an involvement of additional caspase independent death pathways.

3.4. Disturbance of mitochondrial function induced

by anticancer drugs

3.4.1. Alterations of mitochondrial transmembrane potential

(∆Ψm)

3.4.1.1. Time course of drugs-induced ∆Ψm loss

Mitochondrial alterations are a central coordinating event in apoptosis signaling. In order

to investigate difference in apoptosis induction by different drugs, we analyzed induction

of mitochondrial membrane changes. One of these membrane changes involves disruption

of the inner membrane transmembrane potential (∆Ψm) through the opening of

permeability transition pores, known as the mitochondrial permeability transition pore

(PT). After pore opening, the normally impermeable inner membrane becomes permeable

to ions and solutes, and the negatively charged environment of the matrix is lost.

Intermembrane permeabilization can be assessed indirectly, by determining a reduction in

the ∆Ψm . For this, cells are incubated with lipophilic cationic fluorochromes such as

DiOC6(3) (3,3'dihexyloxacarbocyanine iodide), a lipophilic cation, which accumulates in

the mitochondria matrix, driven by the electrochemical gradient. According to the Nernst

equation, every 61.5-mV increase in ∆Ψm (usually 120-170 mV, negative inside) leads to a

10-fold increase in cation concentration in the mitochondrial matrix. Therefore, the

concentration of such cation is two to three logs higher in the matrix than in the cytosol. A

reduction in fluorescence intensity in the emission spectrum, as measured by flow

cytometery, is then interpreted as an indication of ∆Ψm dissipation.

47

A. B.

Figure 11. Anticancer drugs and anti-CD95 antibody cause ∆Ψm disruption. (A)Jurkat cellswere cultured in normal growth medium and then either left untreated (medium, as control) ortreated with etoposide (Eto 30µg/ml), cytarabine (Ara-c 30µg/ml), 4-hydroxy-cyclophosphamide(4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml), methotrexate (MTX 30µg/ml) and anti-CD95antibody (20ng/ml) plus protein A (2.5ng/ml) respectively at 370C in a CO2 incubator for theindicated time points. ∆Ψm loss was determined using 40 nM DiOC6(3) on flow cytrometry. Datawere the mean of triplicates with standard deviations. Similar results were obtained in 3 separatedexperiments. (B) Representative profiles of ∆Ψm loss induced by the anticancer drugs.

Medium

Eto

Ara-c

Medium

4-HCP

Doxo

MTX

6h 12h 24h

M1 M23.64%

M1M2

10.36%

M2M1

43.58%

M1M2

3.59%

M1 M2

67.53%

M2M1

24.80%

M1 M275.32%

M1M2

7.21%

M1M2

2.27%

12h 24h 48h

M1 M2

4.20%

M1 M2

7.62%

M1M2

6.60%

M1 M2

10.12%

M1 M2

3.77%

M1 M2

32.30%

M1 M23.79%

M1 M222.93%

M1 M2

13.41%

M1 M222.93%

M1 M2

22.40%

M1

M2

15.93%

Fluorescence Intensity

0

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0 6 12 18 24

Time (hours)

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Time (hours)

Medium

Apo20ng

48

A. B.

Figure 12. Effects of zVAD-fmk on ∆Ψm alteration induced by anticancer drugs and anti-CD95 antibody. (A) Jurkat cells were cultured in normal growth medium and then either leftuntreated (medium, as control) or treated with etoposide (Eto 30µg/ml), cytarabine (Ara-c30µg/ml), 4-hydroxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml),methotrexate (MTX 30µg/ml) and anti-CD95 antibody (20ng/ml) plus protein A (2.5ng/ml) in thepresence or absence of zVAD-fmk respectively at 370C in a CO2 incubator for the indicated timepoints. ∆Ψm alteration was determined using 40 nM DiOC6(3) on flow cytrometry as described inMaterials and Methods. Data were the mean of triplicates with standard deviation. Similar resultswere obtained in 3 separated experiments. (B) Representative profiles of ∆Ψm alteration inducedby etoposide and 4-hydroxy-cyclophosphamide in presence or absence of zVAD-fmk.

0

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6 12 24

Time (hours)

MediumEtoEto+zVADAra-cAra-c+zVAD

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12 24 48

Time (hours)

Medium4-HCP4-HCP+zVADDoxoDoxo+zVADMTXMTX+zVAD

0

20

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3 6 9

Time (hours)

MediumAnti-CD95Anti-CD95+zVAD

Fluorescence Intensity

4-HCP 4-HCP+zVAD

6h

12h

24h

12h

24h

48h

Eto Eto+zVAD

M2M1

7.62%

M1

M2

10.33%

M2M1

32.30%

M1M2

46.85%

M1 M2

50.64%

M1M2

57.14%

M2M

1

43.58% M1 M2

27.40%

M2M1

67.53%

M2M1

75.32%

M1 M272.30%

M2

68.14%

M1

49

Cells were treated with etoposide, cytarabine, 4-hydoxy-cyclophosphamide, doxorubicin,

methotrexate and anti-CD95 in indicated doses. At the appropriate time point, the cells

were stained with DiOC6 (3). DiOC6 (3) targets to the negative environment of the matrix in

metabolically active mitochondria, where it emits intensely in the FL1 channel during

apoptosis, dissipation of ∆Ψm leads to leakage of DiOC6 (3) from the matrix, which can be

measured by flow cytometry a decrease in the fluorescence intensity of DiOC6 (3). This is

visualized as ∆Ψm low cell subpopulation, which shift to the left in the FACS histogram.

In the absence of treatment (medium alone), ∆Ψm fluorescence was apparent (∆Ψm high

cells), indicating retention of the dye in mitochondria and an intact ∆Ψm as shown in

Figure 11. Jurkat cells treated with stimuli exhibited a reduction in the retention of DiOC6

(3) seen as a shift in the population from DiOC6 (3) high cells to DiOC6 (3) low cells. This

shift indicated a compromise in ∆Ψm integrity. Generally, anti-CD95 antibody-induced

∆Ψm loss was more rapid and efficient. Anti-CD95 antibody-induced ∆Ψm low cells nearly

equal to annexin V positive cells in live cells in 3 hours, approximate 45%. In contrast,

etoposide-induced ∆Ψm low cells was much higher (47%) than annexin V positive cells

(23%) in live cells in 6 hours. These results suggest that maybe etoposide induces

breakdown of ∆Ψm in a different mechanism from anti-CD95. On the other hand, it means

etoposide-induced ∆Ψm loss is the same early event of apoptosis as externalization of PS.

3.4.1.2. Caspase inhibitor abrogated anti-CD95-induced ∆Ψm loss but not drug-

mediated ∆Ψm loss

To investigate whether the mitochondria transmembrane potential disruption was caspase-

dependent, in a parallel experiment, the cells were pretreated with zVAD-fmk. And then

the cells were treated with various stimuli: etoposide, cytarabine, 4-hydroxy-

cyclophosphamide, doxorubicin, methotrexate and anti-CD95 antibody in the indicated

dose and time points as shown in Figure 12.

The data showed that zVAD-fmk completely abrogated ∆Ψm collapse induced by anti-

CD95 from 3 hours to 9 hours. The pan caspase inhibitor zVAD-fmk partially blocked

∆Ψm loss induced by early drugs (etoposide, cytarabine) in 6 hours and 12 hours, but not in

24 hours; whereas zVAD-fmk enhanced ∆Ψm disruption induced by late drugs (4-hydroxy-

cyclophosphamide, doxorubicin, methotrexate). Thus, in all drugs analyzed, ∆Ψm reduction

50

Figure 13. Cytochrome c release from mitochondria into cytosol induced by anticancer drugsand anti-CD95 antibody. (A) Jurkat cells were cultured in normal growth medium and then eitherleft untreated (medium, as control) or treated with etoposide (Eto 30µg/ml), cytarabine (Ara-c30µg/ml), 4-hydroxy-cyclophosphamide (4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml),methotrexate (MTX 30µg/ml) or anti-CD95 antibody (20ng/ml) plus protein A (2.5ng/ml) at 370Cin a CO2 incubator for the indicated time points. Total cellular protein 40µg per lane was separatedby 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred tonitrocellulose, and probed with antibodies that recognize mouse anti-cytochrome c monoclonalantibody. Arrows indicate the position of cytochrome c or β-actin. Equal protein loading wasconfirmed by reprobing the stripped blots with mouse anti-β-actin monoclonal antibody. Theresults shown are representative of at least three independent experiments.

Time (hours) 0 6 12 24 0 6 12 24

cytosol mitochondria

Cytochrome c

Cytochrome c

β-actin

Eto

Ara-c

4-HCPCytochrome c

Cytochrome c

Cytochrome c

β-actin

Doxo

MTX

Time (hours) 0 12 24 48 0 12 24 48

cytosol mitochondria

Cytochrome c

β-actin

Ant-CD95

Time (hours) 0 1 3 6 9 0 1 3 6 9

cytosol mitochondria

51

A. B.

Figure 14. Cytochrome c release quantified by flow cytometry. (A) Jurkat cells were cultured innormal growth medium and then either left untreated (medium, as control) or treated withetoposide (Eto 30µg/ml), cytarabine (Ara-c 30µg/ml), 4-hydroxy-cyclophosphamide (4-HCP3µg/ml), doxorubicin (Doxo 0.15µg/ml), methotrexate (MTX 30µg/ml) and anti-CD95 antibody(20ng/ml) plus protein A (2.5ng/ml) respectively at 370C in a CO2 incubator for the indicated timepoints. The cells were harvested and stained with anti-cytochrome c. Control cells were stainedwith isotype-matched antibody or secondary antibody alone. The cytochrome c low cells werequantified by flow cytometry using CellQuest software as described in Materials and Methods. Thedata were the mean of triplicates with standard deviation. (B) Representative Profiles ofcytochrome c release induced by etoposide during the incubation times.

Cytochrome c-FITC

Eto 6h

Eto 12h

Eto 24h

Medium

Eto

Isotype AbSecond Ab

0

20

40

60

80

100

0 6 12 18 24

Time (hours)

MediumEtoAra-c

0

20

40

60

80

100

0 12 24 36 48

Time (hours)

Medium4-HCPDoxoMTX

0

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0 3 6 9

Time (hours)

Medium

Anti-CD95

52

was not caspase dependent, while in anti-CD95 antibody-induced apoptosis, ∆Ψm loss was

caspase dependent.

3.4.2. Cytochrome c release induced by anticancer drugs

Recent evidence has demonstrated that mitochondria participate in the execution of

apoptosis by release of cytochrome c. Binding of cytochrome c to Apaf-1 results in the

cleavage of procaspase-9 or other caspase, which in turn activate caspase-3. To directly

examine mitochondrial involvement, release of cytochrome c into the cytoplasm was

determined.

After Jurkat cells were treated with stimuli, mitochondria fraction was isolated from

cytosol. The cytochrome c was detected in mitochondria fraction and cytosol fraction by

western blot. Similar to apoptosis pattern, anticancer drugs- and anti-CD95 induced

cytochrome c release was time-dependent. Figure 13 Showed cytochrome c increase in

cytosol over the course of incubation time; in the same time, cytochrome c decrease

gradually in mitochondria. These results suggest ∆Ψm loss induced by anticancer drugs and

anti-CD95 lead to cytochrome c translocation from mitochondria into cytosol. In the same

time, cytochrome c release was quantified by flow cytrometry as shown in Figure 14. The

pattern was similar to the results of Western Blot.

3.4.3. Anticancer drugs cleave Bid and Bcl-2

Bid, a member of the Bcl-2 family, was recently identified as a physiological substrate of

caspase-8 that is responsible for mitochondrial damage. Similar to anti-CD95, anticancer

drugs induced proteolysis of Bid, 24kDa, into the mature p15 Bid fragment as shown in

figure 16. Although all cancer drugs induced Bid cleavage, etoposide and anti-CD95 were

much more efficient as indicated.

The anti-apoptosis protein Bcl-2 reported to convert to proapoptosis regulator following

cleavage by caspase-3. Figure 16 showed the Bcl-2 protein, 26kDa, was cleaved into p21

fragmentation by anti-CD95 and anticancer drugs. The pattern was like Bid cleavage,

53

Figure 15. Bid and Bcl-2 cleaved by anticancer drugs and anti-CD95 antibody. Jurkat cellswere cultured in normal growth medium and then either left untreated (medium, as control) ortreated with etoposide (Eto 30µg/ml), cytarabine (Ara-c 30µg/ml), 4-hydroxy-cyclophosphamide(4-HCP 3µg/ml), doxorubicin (Doxo 0.15µg/ml), methotrexate (MTX 30µg/ml) and anti-CD95antibody (20ng/ml) plus protein A (2.5ng/ml) respectively at 370C in a CO2 incubator for theindicated time points. Total cellular protein 40µg per lane was separated by 12% sodium dodecylsulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to nitrocellulose, and probedwith rabbit anti-bid polyclonal antibody and mouse anti-Bcl-2 monoclonal antibody. Arrowsindicated the position of full length bid, bcl-2 and their cleavage fragments. Equal protein loadingwas confirmed by reprobing the stripped blots with mouse anti-β-actin monoclonal antibody.

Medium Eto Ara-c Anti-CD95

Time (hours) 0 6 12 24 6 12 24 3 6 9

Medium 4-HCP Doxo MTX

Time (hours) 0 12 24 48 12 24 48 12 24 48

β-actin

Bid

p15

Bcl-2

p21

β-actin

Bid

p15

β-actin

Bcl-2

p21

β-actin

54

etoposide and anti-CD95 were much more efficient; Bcl-2 was less strongly cleaved by

Cytarabine, cyclophosphamide, doxorubicin and methotrexate.

55

4. Discussion

Etoposide, cytarabine, cyclophosphamide, doxorubicin, and methotrexate are commonly

used antileukemic drugs. Despite their generalized use for more than 30 years, their

mechanisms of cytotoxicity have been a long-time matter of debate. During the past 20

years, several hypotheses have been formulated, including DNA intercalation/binding,

inhibition of topoisomerase II, free-radical generation, and damage to cell membranes. In

recent years, it has been found that anticancer drugs induce the intrinsic program of cell

death known as apoptosis (Arends et al., 1991; Mesner et al., 1997).

The molecular signaling pathways which are initiated in response chemotherapy-induced

cellular damage, and lead to the eventual apoptotic death of the cell, are largely undefined.

Although the ability to induce apoptosis has been examined in a variety of human tumor

cell types using one or several anticancer drugs, the molecular steps of anticancer drug-

induced apoptosis and its determinants have not been comprehensively evaluated in the

human acute leukemia cells. The elucidation of these pathways and different activation of

apoptotsis signaling molecules may reveal promising new targets for anticancer therapies,

and should contribute to design a drug's schedule of administration and new combination

chemotherapy protocol. The parameters of caspase activation and mitochondial apoptosis

signaling induced by anticancer drugs may be used for monitoring drug efficacy in primary

leukemia cells.

4.1. Different drugs induce apoptosis in a different

time and dose fashion and represent different

feature of apoptosis

Different drugs are used for the treatment of different cancer cells and different drugs

produce different side effects in clinic. Our data show that anticancer drugs induced

apoptosis in a different time and dose fashion. As a whole, anticancer drugs induced time-

56

dependent apoptosis, but for each drug, the different pattern was broad. Etoposide- and

cytarabine-induced apoptosis appear earlier. At 12 hours, the observed maximal apoptosis

for etoposide at the maximal concentration was about 50%, for cytarabine at the

concentration of 1-10 µg/ml was more than 35%. In contrast, at same time point, the

observed maximal apoptosis for 4-hydroxy-cyclophosphamide, doxorubicin, and

methotrexate at their maximal concentration was less than 15%. This data suggested that

taking effect of etoposide and cytarabine was earlier than 4-hydroxy-cyclophosphamide,

doxorubicin and methotrexate.

Interestingly, higher dose of cytarabine induced less apoptosis, whereas lower dose of

cytarabine induced more apoptosis. This just is a phenomenon observed in this experiment

system. It is need to observe in clinic. It is also need to investigate the mechanism of

producing this phenomenon. If this phenomenon is common, it means the dose escalation

approach is no use for cytarabine.

The duration of apoptosis is relatively short and variable depending on cell type, inducer of

apoptosis. Each of the methods used in present experiment has its advantages and suffers

limitations. So combination of several apoptosis assay can provide a more definite

identification of apoptotic cells.

Recent studies have shown that a critical event during apoptosis appears to be the

acquisition of plasma membrane changes that allows phagocytes to recognize and engulf

these cells before the rupture (Martin et al., 1995; Bino et al., 1999). One of these plasma

membrane changes is the translocation of phosphatidylserine to the outer layer, where it

becomes exposed to the external surface of the cell. Once on the cell surface, PS can be

detected by staining with FITC conjugate of annexin V, a protein with high affinity for PS.

Regardless of the apoptotic stimulus, externalization of PS occurs earlier than the nuclear

changes. Thus, the FITC conjugated annexin V binding assay of PS detects early phase of

apoptosis before the loss of cell membrane integrity (chan et al., 1998; Trauth et al., 2000).

In the present study, the four assays detecting apoptosis demonstrated roughly similar

pattern, in which the number of apoptotic cells increased with the increasing cocurlture

time period in presence of anticancer drugs. But in these four assays also differences could

be detected. Similar to anti-CD95 antibody, early drugs, especially etoposide, induced

57

apoptosis more strongly and rapidly in annexin V binding assay than in other assays. In

contrast, similar to γ-radiation, late drugs induced apoptosis with a similar pattern in all

four assays. After further analysis of the annexin V single positive cells, we can find that

similar to anti-CD95 antibody, the number of induced by early acting drugs was much

greater than annexin V/PI double positive cells. In contrast, late drugs-induced annexin V

single positive cells were much less than annexin V/PI double positive cells. Collectively,

early drugs and late drugs induced the translocation of phosphatidylserine from the inner

membrane to the out layer in a different pattern. In other words, the assay's sensitivity was

relevant to the apoptotic stimulus. For early drugs, the annexin V binding assay could

detect early phase of apoptosis before the loss of cell membrane integrity; for late drugs,

externalization of PS was detected at approximately the same time as the loss of cell

membrane integrity.

With these different features of anticancer drugs-induced apoptosis, we can more precisely

evaluate the different anticancer drugs-induced apoptosis when using different assays.

4.2. CD95-associated signaling molecules and

anticancer drugs

In 1989 two groups independently isolated mouse-derived antibodies that were cytolytic

for various human cell lines (Trauth et al., 1989; Yonehara et al., 1989). The cell surface

proteins recognized by the antibodies were designated Apo-1and Fas, respectively. The

antibody to Apo-1 was an immunoglobulin G (IgG3) antibody, whereas the antibody to Fas

(anti-Fas) was classified as IgM. Sequencing and cloning of the Apo-1/Fas proteins and

cDNAs, respectively, showed that receptor of Apo-1 and Fas were identical. The 5th

Workshop on Leukocyte Typing suggested the name CD95 for the receptor. CD95 (Apo-

1/Fas) belongs to the subfamily of death receptor, which is part of the TNF-receptor

(TNF-R) superfamily. Members of this family are characterized by two to five copies of

cysteine-rich extracellular death domain (DD). The DD is essential for transduction of the

apoptotic signal.

CD95 is a widely expressed glycosylated cell surface molecule of approximately 45 to 52

kDa (335 amino acids). It is a type I transmembrane receptor and can also occur in several

58

soluble forms (Itoh et al., 1991; Oehm et al., 1992; Cheng et al., 1994). The human gene

for CD95, APT, was localized to chromosome 10q23 and the mouse gene to chromosome

19 (Lichter et al., 1992; Watanabe-Fukunaga et al., 1992). Expression of the CD95 gene

and cell surface protein are enhanced by IFN-r and TNF and by activation of lymphocytes

(Klas et al., 1993; Leithäuser et al., 1993). Apoptosis can be triggered by agonistic

antibodies and by the natural ligand of the receptor, CD95L, expressed in a more restricted

way than CD95. CD95L was cloned from the cDNA of a killer cell (PC60-d10S) and show

to be a TNF-related type II transmembrane molecule. The mouse and human CD95L genes

were mapped to chromosome 1 (Takahashi et al., 1994a,b). Killer cells expressing CD95L

were shown to kill target cells in a Ca2+ -independent fashion via CD95-CD95L interaction

(Anel et al., 1995). In addition, human CD95L overexpressed in COS cells was found in

the supernatant and induced apoptosis in a soluble form. Soluble CD95L is found as a

trimer and is generated from the transmembrane form by the activity of a metalloprotease

(Krammer et al., 1995; Nagata et al., 1995; Peter et al., 1999).

The main death pathway initiated from Fas activation involves a series of death-associated

molecules, including FADD (Fas-associated-death domain-containing protein), which is an

adaptor protein that is recruited to Fas receptor upon its engagement. FADD then binds to

and activates procaspase-8, which is believed to be the first step of a proteolytic cascade

that triggers activation of other caspases. RIP (receptor interacting protein) was another

containing death domain protein to bind to CD95, which contains an N-terminal region

with homology to protein kinases and a C-terminal region containing a cytoplasmic death

domain present in both Fas and TNF-R1. These molecules were all cloned in the yeast two

hybrid systems with the cytoplasmic part of CD95 used as a bait. The two death domain-

proteins FADD and RIP bind to the CD95-death domain directly. Overexpression of

FADD and RIP causes apoptosis (Chinnaiyan et al., 1995; Juo et al., 1999; Boldin et al.,

1995; Stanger et al., 1995).

Based on the concept of activation-induced death in T cells, the cytotoxicity of anticancer

treatment using cytotoxic drugs or γ-radiation has been studied with respect to involvement

of CD95 receptor/ligand interaction mostly in cell lines derived from patients with T-cell

acute lymphoblastic leukemia (Friesen et al., 1996). Doxorubicin and other cytotoxic drugs

used in the chemotherapy of the leukemias were found to induce CD95L expression in cell

lines and patient derived leukemia cells. The cell lines used constitutively express CD95

59

and are sensitive to CD95-mediated apoptosis triggered via anti-Apo-1 antibody or the

natural ligand. Drug-induced apoptosis was strongly diminished by blocking CD95

receptor/ligand interaction with an anti Apo-1 F (ab)2 fragment or using cell lines in which

CD95 was downregulated by prolonged exposure to an ant-CD95 antibody, suggesting that

activation of CD95L/receptor interaction critically contributes to drug-induced cell death.

The contribution of doxorubicin-CD95L receptor interaction to cytotoxicity was most

pronounced at the lower concentrations of doxorubicin (up to 50-100ng/ml) which may be

achieved during therapy in vivo. In doxorubicin resistant cell lines derived from parental

sensitive cells no induction of CD95L was found. Induction of CD95L and activation of

CD95L/CD95 interaction has also been found with bleomycin in hepatoblastoma cells, 5-

fluorouracil (5-FU) in colon carcinoma cells and various chemotherapeutic drugs in

medulloblastoma and neuroblastoma cells (Fulda et al., 1997a,b; Houghton et al., 1997;

Müller et al., 1997). In addition to CD95L, increased expression of CD95 is induced in

cells with a low level of CD95 expression. These findings indicated that activation of

death-inducing ligands, such as CD95L and CD95, are part of the cellular responds to

cytotoxic treatments that damage DNA, disturb metabolism or affect the mitotic apparatus.

Our results show that, similar to anti-CD95 antibody, anticancer drugs upregulated Fas

Ligand and FADD expression and cleave RIP over the incubation time.

The role of CD95-associated signaling pathway in anticancer drugs-induced apoptosis will

depend on a more detailed understanding of the mechanisms of Fas-mediated apoptosis. It

seems clear that CD95/CD95L and a number of chemotherapeutic agents utilize the

caspase cascade as an effector of apoptosis. However, contrasting findings regarding the

role of the CD95/CD95L pathway in the response to chemotherapy suggests a

heterogeneity in the regulation of this pathway, which needs to be further explored

(Micheau et al., 1997; Wesselbor et al., 1999; Findley et al., 1999).

4.3. The central role of caspase in drug-induced

apoptosis

60

Caspases are a family of mammalian cysteine aspartic proteases that play a central role in

the death process. The 14 caspases so far identified in mammalian cells, of which 12

human enzymes are know, are synthesized as inactive proenzymes that must be cleaved at

key aspartate residues to be activated (Nicholson et al., 1999). X-ray analyses have shown

that activated enzymes form a tetramer containing two large and two small subunits

(Walker et al., 1994; Rotonda et al., 1996). Based on their structural and functional

homologies, mammalian caspases have been classified in two sub-families. Members of

the caspase-1 sub-family (caspase-1, -4, -5, -11, -12 and -14) are mainly involved in

cytokine maturation and inflammation, though they could contribute to some apoptotic

pathways. Members of the caspase-3 sub-family (caspase-2, -3, -6, -7, -8, -9, -10) play a

central role in apoptosis. Further subdivision can be made in this latter subfamily,

depending on the size of their prodomain.

Because caspases exist as latent zymogens, the question remains as to how the zymogens

are activated. Current evidence suggests that activation may proceed by autoactivation,

transactivation, or proteolysis by other proteinases (Wolf BB et al., 1999). Adapter

molecules link apotpotic sensors such as death receptors and mitochondria to procaspases.

To accomplish this, adapters generally contain one domain that couples the adapter to the

sensor and another that binds to long prodomain procaspases. These domains include death

domains (DDs), death effector domains (DEDs), and caspase recruitment domains

(CARDs) (Huang et al., 1996; Eberstadt et al., 1998; Chou et al., 1998).

The central role of caspases in drug-induced apoptosis is suggested by the observation that

several procaspases are cleaved in their active fragments during the cell death process. In

addition, extracts from drug-treated cells cleave peptide substrates that mimic the sequence

specifically recognized by several enzymes of this family. In U937 cells, casase-3 and

caspase-6 appear to play a central role in apoptosis triggered by topoisomerase inhibitors

while the various isoforms of caspase-2 modulate their activity (Dubrez et al., 1996;

Dubrez et al., 1998; Droin et al., 1998; Sakahira et al., 1999). In addition, data from

caspase-9 and Apaf-1 knockout mice have suggested that the generation of a caspase-9-

containing apoptosome complex was crucial for drug-induced apoptosis. Caspase-3 is

required for some typical hallmarks of apoptosis such as DNA fragmentation and

membrane blebbing (Sakahira et al., 1998; Sahara et al., 1999). The other caspases

61

involved in the cell death process could vary, depending on the cell type and the apoptotic

stimulus.

Our results here showed that similar to anti-CD95 antibody, anticancer drugs induce

activation of caspases-8, -3. The cleavage pattern of the individual caspases and PARP did

not differ between the anticancer drugs and anti-CD95 antibody stimulation. However,

corresponding to the different kinetics of apoptosis, Anti-CD95 antibody-induced caspase

activation was more rapid and efficient than etoposide and cytarabine; 4-hydroxy-

cyclophosphamide-, doxorubicin- and methotrexate-induced caspase activation occurs

latest. Collectively, these data show that caspases are involved in drug-mediated apoptosis.

Besides from directly observed drug-induced caspase activation, we also investigated the

effect of the broad-spectrum tripeptide caspase inhibitor zVAD-fmk on anticancer drug-

induced apoptosis. The results show that zVAD-fmk blocked not only externalization of

phosphatidylserine and DNA fragmentation, but also cell morphologic changes and

membrane integrity loss. The patterns was similar to zVAD-fmk inhibited the anti-CD95-

induced changes of apoptosis in Jurkat cells. These observations provide additional

evidence that anticancer drug-induced these changes of apoptosis depend on the function

of caspases.

Comparable results have been found in other cell types or with anticancer drugs. Treatment

of promyelocytic HL-60 cells with clinically achievable doses of etoposide caused

caspase-9 activation and apoptosis. Apoptosis was blocked by cotreatment with zVAD-

fmk. Complete inhibition of apoptotic phenotype and cell death by zVAD-fmk was also

observed in promonocytic THP.1 cells treated with etoposide, and activation of caspases-2,

-3, -6, and -7. Etoposide-induced activation of caspases-3, -7, -8, and -9 in IMR90E1A

cells expressing a dominant-negative mutant of caspase-9 was blocked, but cells still died

(Perkins et al., 1998; Fearhead et al., 1998).

In line with previous studies, our results demonstrate that induction of apoptosis by

anticancer drugs was entirely dependent on the intracellular activation of caspases, because

(a) cell death was completely prevented by zVAD -fmk, a broad caspases inhibitor, and (b)

proteolytic cleavage of the multiple procaspases to their active enzymes, as well as

cleavage of the caspase-specific substrates, could be observed.

62

4.4. Disturbance of mitochondrial function induced

by anticancer drugs

Mitochondria are now recognized as being important in the control of cell survival and

death (green et al., 1989; gottlieb, 2000; Kroemer et al., 2000; Costantini et al., 2000). The

apoptotic changes of mitochondria consist in ∆Ψm loss, transient swelling of the

mitochondrial matrix, mechanical rupture of the outer membrane and/or its nonspecific

permeabilization by giant protein-permeation pores, and release of soluble intermembrane

proteins through the outer membrane.

In addition to a role of the CD95 system in anticancer drug-induced apoptosis, alteration of

mitochondrial functions such as PT (permeability transition) have been found to play a

major role in the chemotherapeutic agent-induced apoptosis. Mitochondria undergoing PT

release apoptogenic proteins such as cytochrome c or AIF from the mitochondrial

intermembrane space into the cytosol, where they can activate caspases and endonucleases

(Liu et al., 1996; Vier et al., 1999). The connection of mitochondrial PT to activation of the

caspase cascade appears to be complex. On one hand, recombinant caspases can induce

PT, probably via a direct effect on the PT pore complex. On the other hand, caspases are

activated by mitochondrial intermembrane protein, suggesting that caspase can act either

upstream or downstream of mitochondria.

Mitochondrial function during apoptosis seems to be controlled by the Bcl-2 family of

proteins. Bcl-2 and several of its homologues have been localized to intracellular

membranes, including the mitochondrial membrane. Overexpression of the antiapoptotic

molecules Bcl-2 and Bcl-XL has been found to confer resistance to anticancer treatment

(Yang et al., 1997; Kluck et al., 1997). Bcl-2 and Bcl-XL may inhibit apoptosis through the

capacity to prevent PT and/or to stabilize the barrier function of the outer mitochondrial

membrane. Both Bcl-2 and Bcl-XL have been shown to prevent opening of the purified PT

pore complex reconstituted in liposomes. In addition, it has been suggested that Bcl-2 and

Bcl-XL can bind cytosolic caspases via Apaf-1 to the mitochondrial membrane, thereby

preventing their activation. Bcl-XL has recently been reported to prevent Apaf-1-dependent

caspase-9 activation via interaction with Apaf-1.

63

Betulinic acid, a pentacyclic triterpene, is a novel experimental anticancer drug. It

possesses an antitumoral activity in vitro and in vivo in melanoma, neuroectodermal

tumors, and glioma cell lines. Fulda et al (Fulda et al., 1998) have shown that betulinic acid

induces apoptosis via direct mitochondrial alterations. All of these effects have been

observed in intact cells and in cell-free systems. When added to isolated mitochondria,

betulinic acid directly induces loss of ∆Ψm in a way that is not affected by the caspase

inhibitor zVAD-fmk and yet is inhibited by Bongkrekic acid.

A currently open question is whether the breakdown of ∆Ψm is an 'assassin or accomplice'

(Green et al., 1989) since under some circumstances the breakdown of ∆Ψm occur

downstream of caspase activation. Similarly, time course analysis of ∆Ψm breakdown and

cytochrome c release revealed contradictory results (Heibei et al., 1999). Whereas in some

models the breakdown of ∆Ψm occurred considerably later than cytochrome c release and

caspase activation, other showed that ∆Ψm loss and the release of cytochrome c occurred

rather simultaneously. Our data reveal that anticancer drugs induce a breakdown of ∆Ψm

independently of caspase activation. The time course of caspase activation and the

breakdown of ∆Ψm are very similar.

Previous studies suggested that a decline of ∆Ψm may be an early event in apoptosis,

including Fas-dependent signaling. In present results, anticancer drug-induced ∆Ψm loss,

similar to anti-CD95-induced ∆Ψm loss, occurs at the same time points as PS

externalization and DNA fragmentation. Anti-CD95 antibody-induced ∆Ψm loss can be

blocked by zVAD-fmk. In contrast, anticancer drug-induced-∆Ψm loss cannot be blocked

by zVAD-fmk. These results suggested that anti-CD95 antibody-induced ∆Ψm loss was via

caspase activation; whereas anticancer drug-induced ∆Ψm loss was not via caspase

activation. In other words, the direct effect of anticancer drugs on mitochondria leads to

mitochondrial transmembrane potential dissipation. These results are fundamentally

different from those reported by Gamen et al (Gamen et al., 2000), who used doxorubicin

as stimulus in Jurkat cells. Their results showed that doxorubicin induced ∆Ψm loss was

prevented by coincubation with zVAD-fmk.

Numerous reports have described the ability of various specific and nonspecific DNA-

damaging agents to stimulate the release of mitochondrial cytochrome c. In contrast to

death receptor-mediated apoptosis, during which caspase-8 activity is often responsible for

64

the cleavage of a cytosolic substrate, e.g. Bid, which targets mitochondria triggering the

release of cytochrome c, this event is traditionally accepted as caspase-independent in

chemical- and /or DNA damage-induced apoptosis.

In the present study, anti-CD95 antibody-induced cytochrome c release occurs at 3 hours;

etoposide and cytarabine at 6 hours; while 4-hydroxy-cyclophosphamide, doxorubicin and

methatrexate nearly at 12 hours, at the same time of the appearance of cells displaying

apoptotic morphologic changes, PS translocation, DNA fragmentation and loss of cell

membrane integrity.

Cytochrome c, once released from mitochondria, is believed to form a complex with Apaf-

1 and caspase-9. This 'apoptosome' then mediates activation of caspase-3 in an ATP-

dependent fashion. We do not have any direct evidence that cytochrome c release during

anticancer drug-mediated apoptosis form a complex with Apaf-1 and caspase-9.

Nevertheless, such an interaction has been described for Jurkats in response to the CD95

pathway, and so it is not unreasonable to assume that such a process may occur in these

cells in response to anticancer drugs. Since anticancer drugs are capable of directly

activating caspase-3 in intact cells, one possible role for the Apaf-1/cytochrome c/caspase-

9 complex is to amplify the cascade.

Proteins of the Bcl-2 family are major regulators of the mitochondria-initiated caspase

activation pathway. The anti-apoptotic members of this family, including Bcl-2 and Bcl-

XL, preserve mitochondrial integrity and prevent the release of cytochrome c in the

presence of apoptotic stimuli (Kluck et al., 1997; Yang et al., 1997). Conversely, the

proapoptotic members of this family such as Bad, Bax, Bid, and Bim move from other

cellular compartments to mitochondria in response to apoptotic stimuli and promote

cytochrome c release (Reed, 1998; Antonsson et al., 2000; Gross A et al., 1999).

Cleavage of Bid was observed in anticancer drug- as well as anti-CD95-mediated

apoptosis. Bid is most likely cleaved by low concentrations caspase-8 but can be cleaved

by higher concentrations of caspase-3 (Sun et al., 1999). Thus in anticancer drug-induced

apoptosis, inhibition of Bid cleavage by zVAD-fmk may be due to inhibition of

processing/activity of either caspase-8 at a later stage of the apoptotic process, resulting in

cleavage of Bid. The cleaved bid moves to mitochondrial form cytosol, insert in

65

mitochondrial membrane and promotes release of mitochondrial cytochrome c ant other

pro-apototic proteins such as AIF, SMAC, DIABLO (Du C et al., 2000; Verhagen AM et

al., 2000), thereby further amplifying the apoptotic program.

The bcl-2 gene was originally isolated from the t(14;18) chromosomal breakpoint in

follicular B-lymphoma cells ( Tsujimoto et al., 1985). This gene has been shown to prevent

apoptosis induced by growth-factor deprivation in certain haematopoietic cell lines. 26 kDa

Bcl-2 protein, the product of bcl-2 gene, is an integral intracellular membrane protein that

inhibits programmed cell death induced by multiple insults in a wide variety of cell types.

Both biochemical and genetic evidence indicates that Bcl-2 family member can regulate

cell death induced by caspases . A number of substrates for the caspase proteases have now

been identified, including protein kinases, the retinoblastoma protein, cytoskeletal proteins,

and several autoantigens. Cleavage of these proteins by caspase may either activate or

inactivate essential functions or produce cleavage products with altered activities. Previous

research showed that when bcl-2 was cleaved by capases, the conversion of Bcl-2 became

a Bax-like death effector. Other investigators also reported the cleaved Bcl-2 fragment

increased the sensitivity to VP-16 (Etoposide). In current studies, Bcl-2 was cleaved in

anti-CD95 antibody- and anticancer drug-induced apoptosis, in agreement with previous

observations. The anti-apoptosis protein Bcl-2 was converted to proapoptosis regulators

following cleavage by caspase-3 (Cheng et al., 1997; Wang et al., 2001).

Besides of death receptor signaling pathway, caspase activation and mitochodrial signaling

pathway, there are several other mechanisms involved in drug-induced apoptosis, such as

p53 tumor suppressor gene, lipid-dependent signaling pathway and generation of oxygen

radicals (Solary E et al., 2000).

Overall, understanding the mechanisms of anticancer drug-induced apoptosis is of

principal importance for developing effective strategies in tumor therapy. Our results show

that etoposide- and cytarybine-induced apoptosis was early; 4-hydroxy-cyclophosphamide-

, doxorubicin-, and methotrexate-induced apoptosis was late. Higher doses of cytarabine

induce lower rate of apoptosis than lower doses of cytarabine. In all drugs tested, apoptosis

could be inhibited by caspases, suggesting induction of caspase dependent cell death by all

drugs. Also mitochondrial transmembrane potential loss and cytochrome c release were

induced by all drugs. The different characteristics of anticancer drugs e.g. time of apoptosis

66

induction are probably due to events upstream of the common mitochondrial and caspase

signaling pathways. Since all tested drugs induced caspase activation and mitochondrial

apoptosis signaling, measurement of these parameters can be used for assessment of drug

efficacy in primary leukemia cells.

67

5. Summary

It is known that in empirical medicine, different anticancer drugs have different

characteristic concerning specific anti-tumor or anti-leukemic efficacy and side effects on

normal tissue. While solid tumors are often treated with cisplatin, treatment of leukemia is

based on the use of anthracyclines and antimetabolites. Many drugs also have unique

toxicities affecting normal tissues, such as the cardiotoxicity associated with the

anthracyclines, the hemorrhagic cystitis associated with the cyclophosphamide and

ifosfamide. The peripheral neuropathy from vincristine, and the coagulopathy from L-

asparaginase. We therefore hypothesized that the different clinical used anticancer drugs

might induce apoptosis in a drug specific manner. Thus the clinical observed difference

could be reflected by different activation of apoptosis signaling pathways. The molecular

signaling pathways which are initiated in response to anticancer drug-induced cellular

damage, and lead to the eventual apoptosis death of the cell, are largely undefined.

Previous studies show the caspase family are the critical executioners of apoptosis and

anticancer drug-induced apoptosis by activating two major cell-intrinsic pathways, one that

begins with ligation of cell surface death receptors, such as CD95, and another that

involves mitochondrial release of cytochrome c. We therefore investigated induction of

apoptosis, activation of caspases and involvement of mitochondrial signaling in the well

defined Jurkat human leukemic T cells by five conventional used anticancer drugs:

etoposide, cytarabine, 4-hydroxy-cyclophosphamide, doxorubicin, and methotrexate, in

order to identify drug specific activation of distinct apoptosis pathways.

In present studies, we found some difference of apoptosis induced by these five anticancer

drugs: (1) the anticancer drug-induced apoptosis appeared in different time kinetics,

etoposide and cytarabine were early acting drugs, while 4-hydroxy-cyclophosphamide,

doxorubicin, and methotrexate were late acting drugs. (2) Interestingly, higher doses of

cytarabine induced less apoptosis, whereas lower doses of cytarabine induced more

apoptosis. (3) Similar to CD95, early drug-induced PS externalization was earlier than

drug-induced membrane integrity loss, whereas late drug-induced PS externalization and

membrane integrity loss occurred in the same time, similar to γ-radiation-induced

68

apoptosis. (4) Etoposide strongly induced caspases activation, compared to cytarabine, 4-

hydroxy-cyclophosphamide, doxorubicin and methotrexate. Besides these differences, we

found that all drugs induced apoptosis in a similar manner by all anticancer drugs. All

tested drugs can activate caspase-8, caspase-3 and cleave PARP. Furthermore, these five

drug-induced apoptosis could be inhibited by the pan-caspase inhibitor zVAD-fmk,

suggesting that drugs induce apoptosis in a caspase dependent way. All drugs induce

mitochondrial transmembrane potential reduction and all drug-induced ∆Ψm loss can not be

blocked by caspase inhibitor zVAD-fmk, indicating that caspases are not required for early

mitochondrial changes. All drugs induced cytochrome c translocation from mitochondria

into cytosol. This suggests that mitochondrial signaling and caspase activation are

commonly activated in programmed cell death induced by cytostatic drugs used for anti-

cancer treatment. Since all tested drugs induced caspase activation and mitochondrial

apoptosis signaling, measurement of these parameters can be used for assessment of drug

efficacy in primary leukemia cells. The different characteristics of anticancer drugs e.g.

time of apoptosis induction are probably due to events upstream of the common

mitochondrial and caspase signaling pathways.

69

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81

7. Acknowledgements

I would like to express my gratitude to Prof. Dr. Debatin for accepting me as a M.D.

student under his supervision and providing scientific support during my thesis. I am

benefit a lot from the ''work in process'' and ''journal club'' on every Monday and Friday

during these two years, which were organized and promoted by Prof. Dr. Debatin. These

seminars helped me to understand science and will affect my whole scientific career.

I am greatly indebted to Dr. Stahnke, who provided me the ideas and project, supervised

whole research process, and reviews the manuscript. Without his help, I wouldn't have

completed my work.

My thanks also go to Ms. Dravits for her teaching me laboratory techniques, to all of my

colleagues of the research laboratory of the children's hospital for the help during these two

years ' work.

I thank Prof. Dr. Dr. Dr. A. Grünert, who provide me many useful supports before and

during my study in Ulm University.

Finally, I am deeply grateful to my wife, Chen Longgui and my daughter, Liu Chang. With

their support and love, I could overcome every difficulty in the past two years.

82

Curriculum Vitae

Name Jiahao Liu

Date of Birth 10. 06. 1961

Place of Birth Hubei, P. R. China

Gender Male

Marital Status married with one child

Nationality Chinese

Education2000 - 2001 University of Ulm, Ulm, Germany

Doctor Degree of Medicine (Dr. med)

1986 - 1989 Southeast University, Nanjing, China

Master Degree of Medicine

1978 - 1983 Tongji Medical University, Wuhan, China

Bachelor Degree of Medicine

Professional Experience

2000 - present University of Ulm, Ulm, Germany

Research laboratory of children's hospital

1989 - 1999 Southeast University, Nanjing, China

Associate professor

1983 - 1986 Southeast University, Nanjing, China

Resident, Dept. of pediatrics, Zhong-Da hospital