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
10
20
30
40
50
60
70
80
0 6 12 18 24 30 36Time (hours)
Medium
4-HCP0.1µg
4-HCP0.5µg
4-HCP1µg
4-HCP3µg
4-HCP5µg
0
10
20
30
40
50
60
70
80
0 6 12 18 24 30 36Time (hours)
Medium
Doxo0.01µg
Doxo0.1µg
Doxo0.2µg
Doxo0.5µg
Doxo1µg
0
10
20
30
40
50
60
70
80
0 6 12 18 24 30 36
Time (hours)
Medium
MTX1µg
MTX5µg
MTX10µg
MTX100µg
MTX500µg
0
10
20
30
40
50
60
70
80
0 6 12Time (hours)
Medium
Ara-c1µg
Ara-c5µg
Ara-c10µg
Ara-c100µg
Ara-c500µg
0
10
20
30
40
50
60
70
80
0 6 12
Time (hours)
Medium
Eto1µg
Eto5µg
Eto10µg
Eto100µg
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.
0
20
40
60
80
100
0 6 12 18 24
Time (hours)
Medium
Eto
Ara-c
0
20
40
60
80
100
0 6 12 18 24
Time (hours)
Medium
Eto
Ara-c
0
20
40
60
80
100
0 6 12 18 24
Time (hours)
Medium
Eto
Ara-c
0
20
40
60
80
100
0 6 12 18 24
Time (hours)
MediumEto
Ara-c
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.
0
20
40
60
80
100
0 12 24 36 48
Time (hours)
Medium
4-HCP
Doxo
MTX
0
20
40
60
80
100
0 12 24 36 48
Time (hours)
Medium
4-HCP
Doxo0.
MTX
0
20
40
60
80
100
0 12 24 36 48
Time (hours)
Medium
4-HCP
Doxo
MTX
0
20
40
60
80
100
0 12 24 36 48
Time (hours)
Medium
4-HCP
Doxo
MTX
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.
0
20
40
60
80
100
0 3 6 9Time (hours)
MediumAnti-CD95
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)
Medium
4-HCP
Doxo
MTX
Anti-CD95
R1
R290.18%
9.76%
R1
R255.08%
44.88%
32.70%
3.22%
39.51%
58.09%
Anti-CD95
3h
6h
R1
R289.58%
10.42%
R1R2
62.52%
37.46%
7.92%
2.04%
52.82%
8.67%
12h
48h
4-HCP 4-HCP
2.35%
26.36%
39.89%
37.74%
R1
R2
54.94%
45.03%
R1
R287.79%
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
60
80
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
60
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
20
40
60
80
100
6 12 24Time (hours)
0
20
40
60
80
100
6 12 24Time (hours)
0
20
40
60
80
100
6 12 24Time (hours)
0
20
40
60
80
100
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
40
60
80
100
12 24 48Time (hours)
0
20
40
60
80
100
12 24 48Time (hours)
0
20
40
60
80
100
12 24 48Time (hours)
0
20
40
60
80
100
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
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
20
40
60
80
100
0 3 6 9
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
20
40
60
80
100
6 12 24
Time (hours)
MediumEtoEto+zVADAra-cAra-c+zVAD
0
20
40
60
80
100
12 24 48
Time (hours)
Medium4-HCP4-HCP+zVADDoxoDoxo+zVADMTXMTX+zVAD
0
20
40
60
80
100
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
20
40
60
80
100
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
6. References
Alnemri ES: Mammlian cell death proteases: a family of highly conserved aspartate
specific cysteine proteases. J Cell Biochem 64: 33-42 (1997)
Anel A, Simon AK, Auphan N, Buferne M, Boyer C, Golstein P, Schmitt-Verhulst AM:
Two signaling pathways can lead to Fas ligand expression in CD8+ cytotoxic T
lymphocyte clones. Eur J Immunol 25: 3381-3387 (1995)
Antonsson B, Martinou JC: The Bcl-2 Protein family. Experimental Cell Research 256: 50-
57 (2000)
Arends MJ, Wyllie AH: Apoptosis: Mechanisms and roles in pathology. Int Rev ExpPathol
32: 223-254 (1991)
Bacso Z, Eversion RB, Eliasion JF: The DNA of annexin V-binding apoptotic cells is
highly fragmented. Cancer Res 60: 4623-4628 (2000)
Bino GD, Darzynkiewicz Z, Degraef C, Mosselmans R, Fokan D, Galand P: comparison of
methods based on annexin-V binding, DNA content or TUNEL for evaluating cell death in
HL-60 and adherent MCF-7 cells. Cell Prolif 32: 25-37 (1999)
Boldin MP, Varfolomeev EE, Pancer Z, Mett IL, Camonis, JH, Wallach D: A novel protein
that interacts with the death domain of Fas/APO1 contins a sequence motif related to the
death domain. J Biol Chem 270: 7795-7798 (1995)
Budihardjo I, Oliver H, Lutter M, Luo X Wang X: Biochemical pathways of caspase
activation during apoptosis. Annu Rev Cell Dev Biol 98: 47-58 (1998)
70
Chan A, Reiter R, Wiese S, Fertig G, Gold R: Plasma membrane phospholipid asymmetry
precedes DNA fragmentation in different apoptotic cell models. Hitochem Cell Biol 110:
553-558 (1998)
Cheng EH, Kirsch DG, Clem RJ, Ravi R, Kasten MB, Bedi A, Ueno K, Hardwick JM:
Conversio of Bcl-2 t o a Bax-like death effector by caspases. Science 278:1966-1968
(1997)
Cheng J, Ahou T, Liu C, Shapiro JP, Brauer MJ, Kiefer MC, Barr PJ Mountz JD:
Protection from Fas-mediated apoptosis by a soluble form of the Fas molecule. Science
263: 1749-1762 (1994)
Chinnaiyan AM, O'Rourke K, Tewari M, Dixit VM: FADD , a novel death domain-
containing protein, interacts with the death domain of Fas and initiates apoptosis. Cell 81:
505-512 (1995)
Chinnaiyan AM, O'RourkeK, Lane BR, Dixit VM: Ineraction of Ced-4 with Ced-3 and
CED-9: a molecular framework for cell death. Science 275: 112-1126 (1997)
Chou JJ, Matsuo H, Duan H, Wagner G: Solution structure of the RAIDD CARD and
model for CARD/CARD interaction in caspase-2 and caspase-9 recruitment. Cell 94: 171-
180 (1998)
Costantini P, Jacotot E Decaudin D Kroemer G: Mitochondria as novel target of anticancer
chemotherapy. J Natl Cancer Inst 92: 1042-1053 (2000)
Costantini P, Jacotot E Decaudin D Kroemer G: Mitochondria as novel target of anticancer
chemotherapy. J Natl Cancer Inst 92: 1042-1053 (2000)
Debatin KM: Activation of apoptosis pathways by anticancer drugs. Adv Exp Med Biol
547: 237-244 (1999)
71
Depraetere V, Golstein P: Dismantling in cell death: molecular mechanisms and
relationship to caspase activation. Scand J Immunol 47: 523-531 (1998)
Droin N, Bubrez L, Eymin B, Renvoize C, Breard J, Dimanche-boitrel MT, Solary E:
Upregulation of CASP genes in human tumor cells undergoing etoposide-induced
apoptosis. Oncogene 16: 2885-2894 (1998)
Dubrez L, Savoy I, Hamman A, Solary E: Pivotal role of a DEVD-sensitive step in
etoposide-induced and Fas medated apoptotic pathways. EMBO J 15: 5504-5512 (1996)
Du C, Fang M, Li Y, Li L, Wang X: Smac, a mitochondrial protein that promotes
cytocchorme c-dependent caspase activation by eliminating IAP inhibition. Cell 102: 33-
42 (2000)
Dubrez L, Eymin B, Sordet O, Droin N, Turhan AG, Solary E: BCR-ABL delays apoptosis
upstream of procaspase-3 activation. Blood 91: 2415-2422 (1998)
Earnshaw WC, Martins LM, Kaufmann SH: Mammalian caspasese: structure, activation,
substrates, and functions during apoptosis. Annu Rev Biochem 68: 383-424 (1999)
Eberstadt M, Huang B, Chen Z, Meadows RP, Ng SC, Zheng L, Lenardo MJ, Fesik SW:
NMR structure and mutagenesis of the FADD (Mortl) death -effector domain. Nature 392:
941-945 (1998)
Ellis RE, Jacobson DM, Horvitz HR: Genes required for the engulfment of cell corpses
during programmed cell death in Caenorhabditis elegans. Genetics. 129:79-94 (1991)
Evans VG: Multiple pathways to apoptosis. Cell Bio Int 17: 461-476 (1993)
Fadok VA, Henson PM: Apoptosis: getting rid of the bodies. Curr Biol 8: R693-R695
(1998)
72
Fearhead HO, Rodriguez J, Govek EE, Guo W, Kobayashi R, Hannon G, Lazebnik YA:
Oncogene-dependent apoptosis is mediated by caspase-9. Poc Natl Acad Sci USA 95:
13664-13669 (1998)
Findley HW, Zhou M: The clinical significance of fas expression in leukemia: questions
and controversies. Leukemia 13: 147-149 (1999)
Franc NC, Dimarcq JL, Lagueux M, Hoffmann J, Ezekowitz RA: Croquemort, a novel
Drosophila hemocyte/macrophage receptor that recognizes apoptotic cells. Immunity 4:
431-443 (1996)
Friesen C, Herr I, Krammer PH, Debatin KM: Involvement of the CD95 (APO-1/FAS)
receptor/ligand system in drug-induced apoptosis in leukemia cells. Nature Med 2: 574-
577 (1996)
Friesen C, Herr I, Krammer PH, Debatin KM: Involvement of the CD95 (APO-1/FAS)
receptor/ligand system in drug-induced apoptosis in leukemia cells. Nature Med 2: 574-
577 (1996)
Fulda S, Friesen c, Los M, Scaffidi, CA, Mier W, Benedict M, Nunez G, Krammer PH,
Peter ME, Debatin KM: Betuliic acid triggers CD95 (APO-1/Fas)- and p53 independent
apoptosis via activation of caspases in neuroectodermal tumors. Cancer Res 57: 4956-4964
(1997)
Fulda S, Friesen c, Los M, Scaffidi, CA, Mier W, Benedict M, Nunez G, Krammer PH,
Peter ME, Debatin KM: Betuliic acid triggers CD95 (APO-1/Fas)- and p53 independent
apoptosis via activation of caspases in neuroectodermal tumors. Cancer Res 57: 4956-4964
(1997a)
Fulda S, Sieverts H, Friesen C, Herr I, Debatin KM: The CD95 (APO-1/FAS) system
mediates drug-induced apoptosis in neuroblastoma cells. Cancer Res 57: 3823-3829
(1997b)
73
Fulda S, Susin SA, Kroemer G, Debatin KM: Molecular ordering of apoptosis induced by
anticancer drugs in neuroblastma cells. Cancer Res 1998, 58: 4453-60 (1998)
Gamen S, Anel A, Perez-Galan P, Lasierra P, Johnson D, Pineiro A, Naval J: Doxorubicin
treatment activates a z-VAD-sensitive caspase, which causes DYm loss, caspase-9
activity, and apoptosis in Jurkat cells. Exp Cell Res 258: 223-235 (2000)
Garcia-Calvo M, Peterson EP, Rasper DM, Vaillancourt JP, Zamboni R, Nicholson DW
Thornberry NA: Purification and catalytic properties of human caspase family members.
Cell Death Differ 6: 362-369 (1999)
Gavrieli Y, Sherman Y, Ben-Sasson SA: Identification of programmed cell death in situ
via specific labeling of nuclear DNA fragmentation. J Cell Biol 119: 493-501 (1992)
Gottlieb RA: Mitochondria: execution central. FEBS L 482: 6-12 (2000)
Green DG, Reed JC: Mitochondria and apoptosis. Science 281: 1309-1312 (1989)
Häcker G: The morphology of apoptosis. Cell Tissue Res 301:5-17 (2000)
Heibei JA, Barry M, Motyka B, Bleackley RC: Granzyme B-induced loss of mitochondrial
inner membrane potential (DYm) and cytochrome c release are caspase independent. J
Immunol 163: 4683-4693 (1999)
Houghton JA, Harwood FG, Tillman DM: Thymineless death in colon carcinoma cells is
mediated via FAS signaling. Proc Natl Acad Sci USA 94: 8144-8149 (1997)
Huang B, Eberstadt M, Olejniczak ET, Meadows, RP, Fesik SW: NMR structure and
mutagenesis of the Fas (APO-1/CD95) death domain. Nature 384: 638-641 (1996)
Itoh N, Yonehara S, Ishii A, Yonehara M, Mizushima S, Sameshima M, Hase A, Seto Y,
Nagata S: The polypettide encoded by teh cDNA for human cell surface antigen Fas can
mediate apoptosis. Cell 66: 233-243 (1991)
74
Juo P, Kuo CJ, Yuan J, Blenis J: Essential requirement for caspase-8/FLICE in the
initiation of the Fas-induced apoptotic cascade. Curr Biol 8: 1001-1008 (1998)
Juo P, Woo MSA, Kuo CJ, Signorelli P, Biemann HP, Hannun YA, Blenis J: FADD is
required for multiple signaling events downstream of the receptor fas. Cell Growth Differ
10:797-804 (1999)
Kaufmann SH, Earnshaw WC: Induction of apoptosis by cancer chemotherapy. Exp Cell
Res 256: 42-49 (2000)
Kawabata Y, Hirokawa M, Kitabayashi A, Horiuchi T, Kuroki J, Miura AB: Defective
apoptositic signal transduction pathway downstream of caspase-3 in human B-lymphoma
cells: a novel mechanism of nuclear apoptosis resistance. Blood 94: 3523-3530 (1999)
Kerr JF, Wyllie AH, Currie AR: Apoptosis: a basis biological phenomenon with wide-
ranging implications in tissue kinetics. Br J Cancer 26: 239-257 (1972)
Klas C, Debatin KM, Jonker RR Krammer PH: Activation interferes with the APO-1
pathway in mature human T cells. Int. Immunol 5: 625-630 (1993)
Kluck RM, Bossy-Wetzel E, Green DR, Newmeyer DD: The release of cytochrome c
frome mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science 275: 1132-
1136 (1997)
Kluck RM, Bossy-Wetzel E, Green DR, Newmeyer DD: The proapoptotic function of
Drosophila Hid is conserved in mammalian cells. Pro. Natl. Acad. Sci. USA 96: 4936-4941
(1997)
Krammer PH, Dhein J, Walczak H, Behrmann I, Mariani S, Matiba B, Fath, M, Daniel PT,
Knipping E Westendorp MO, et al: The role of APO-1-mediated apoptosis in the immune
system. Immunol Rev 142: 175-191 (1994)
75
Krammer PH: The CD95(APO-1/Fas)-mediated apoptosis: live and let die. Adv Immunol
71:163-210 (1999)
Kroemer G, Dallaporta B, Resche-Rigon M: The mitochondrial death/life regulator in
apoptosis and necrosis. Annu Rev Physiol. 60: 619-642 (1998)
Kroemer G, Reed JC: Mitochondrial control of cell death. Nature Med 6: 513-519 (2000)
Leithäuser F, Dhein J, Mechtersheimer G, Koretz K, brüderlein S, Henne C, Schmidt A,
Debatin KM, Krammer PH, Möller P: Constitutive and induced expression of APO-1, a
new member of the nerve growth factor/tumor necrosis factor receptor superfamily, in
normal and neoplastic cells. Lab Invest 69: 415-429 (1993)
Li P, Nijhawan D, Budihardjo I, Srinivasula SM, Ahmad M, Alnemri ES, Wang X:
Cytochrome c and dAT-dependent formation of Apaf-1/caspase-9 complex initiates an
apoptotic protease cascade. Cell 91: 479-489 (1997)
Lichter P, Walczak H, Weitz s, Behrmann I, Krammer PH: The human APO-1 (APT)
antigen maps to 10q23, a region that is syntenic with mouse chromosome 19. Gnomics
14:179-180 (1992)
Liu X, Caryn Kim CN, Yang J, Jemmerson R, Wang X: Induction of apoptotic program in
cell-free extracts: requirement for dATP and cytochrome c. Cell 86: 147-157 (1996)
Los M, Wesselborg S, Schulze OK: The role of caspases in development, immunity, and
apoptotic signal transdcution: lessons from knockout mice. Immunity 10: 629-639 (1999)
Majno G, Joris I: Apoptosis, oncosis, and necrosis. An overview of cell death. Am J Pathol
146: 3-15 (1995)
Makin G, Kickman JA: Apoptosis and cancer chemotherapy. Cell Tissue Res 301: 143-152
(2000)
76
Martin SJ, Reutelingsperger CP, McGahon AJ, Rader JA, van Schie RC, LaFace DM,
Green DR: Early redistributio of plasma membrne phosphatidyl serine is a general feature
of apoptosis regardless of the initiating stimulus: inhibition by over expression of Bcl-2
and Abl. J Exp Med 182: 1545-1556 (1995)
Mesner P, Budihardjo I Kaufmann SH: Chemotherapy-induced apoptosis. Adv Pharmacol
41:461-499 (1997)
Micheau O, Solary E, Hammann A, Dimanche-Boitrel MT: Fas ligand-independent
FADD-mediated activation of the Fas Death pathway by anticancer drugs. J Bio Chem
274: 7987-7992 (1999)
Mills JC, Stone NL, Erhardt J, Pittman RN: Apoptotic membrane blebbin is regulated by
myosin light chain phosphorylation. J Cell Biol 140: 627-636 (1998b)
Müller M, Stand S, Hug H, Heinemann EM, Walczak H, Hofmann WJ, Stremmel W,
Krammer PH, Galle P: Drug-induced apotosis in hepatoma cells is mediated by the CD95
(APO-1/Fas) receptor/ligand system and involves activation of wild-type p53. J Clin Invest
99: 403-413 (1997)
Nagata S, Glstein P: The Fas death factor. Science 267: 1449-1456 (1995)
Nagata S: Apoptosis by death factor. Cell 88:355-365 (1997)
Nicholson DW: Caspase structure, proteolytic substrates, and function during apoptotic
cell death. Cell Death Differ 6: 1028-1042 (1999)
Oehm A, Behrmann I, Falk w, Pawlita M, Maier g, Klas c, Li-Weber M, Richards S, Dhein
J, Trauth BC: Purification and molecular cloning of the APO-1 cell surface antigen, a
member of the tumor necrosis facor/nerve growth factor receptor superfmily. Sequenc
identity with the Fas antigen. J Biol Chem 267: 10709-10715 (1992)
77
Perkins C, Kim CN, Bhalla KN: Overexpression of Apaf-1 promotes apoptosi of untreated
and paclitaxelor etoposide-treated HL-60 cells. Cancer Res 58: 4561-4566 (1998)
Peter ME, Scaffidi C, Medema JP, Kischkel F, Drammer PH: The death receptors. Results
Probl Cell Differ 23: 25-63 (1999)
Reed JC: Bcl-2 family proteins. Oncogene 17: 3225-3236 (1998)
Rotello Rj, Fernandez PA, Yuan J: Anti-apogens and anti-engulfens: monoclonal
antibodies reveal specific antigens on apoptotic and engulfment cells during chicken
embryonic development. Development 120: 1421-1431 (1994)
Rotonda J, Nicholson DW, Fazil KM, Gallant M, Gareau Y, Labelle M, Peterson EP,
Rasper DM, Ruel R, Vaillancourt JP, Thornberry NA, Becker JW: The three-dimensional
structure of apopain/CPP32, a key mediator of apoptosis. Nat Struct Biol 3: 619-625
(1996)
Sahara S, Aoto M, Eguchi Y, Imamoto N, Yoneda Y, Tsujimoto Y: Acinus is a caspase-3-
activated protein required for apoptotic chromatin condensation. Nature 401: 168-173
(1999)
Sakahira H, Enari M, Nagata S: Cleavage of CAD inhibitor in CAD activation and DNA
degradation during apoptosis. Nature 391: 96-99 (1998)
Sakahira H, Enari M, Nagata S: Cleavage of CAD inhibitor in CAD activation andDNA
degradation during apoptosis. Nature 391: 96-99 (1998)
Salvesen GS Dixit VM: Caspasee: intracellular signaling by proteolysis. Cell 91: 443-446
(1997)
Savill J: Phagocte recognition of apoptotic cells. Biochem Soc Trans 24: 1065-1069 (1996)
Schulze-Osthoff K, Ferrari D, Low M, Wesselborg S, Peter ME: Apoptosissignaling by
death receptors. Eur J Biochem 254: 439-459 (1998)
78
Sordet O, Bettaieb A, Bruey JM, Eymin B, Droin N, Ivarsson M, Garrido C, Solary E:
Selective inhibition of apoptosis by TPA-induced differentiation of U937 leukemic cells.
Cell Death Differ 6: 351-361 (1999)
Solary E, Droin N, Bettaieb A, Corcos L, Dimanche-Boitrel MT, Garrido C: Positive and
negative regulation of apoptotic pathways by cytotoxic agents in hematological
malignancies. Leukemia 14: 1833-1849 (2000)
Stanger BZ, Leder P, Lee TH, Kim E, Seed B: RIP: A novel protien containing a death
domain that interacts with Fas/APO-1 (CD95) in yeast and causes cell death. Cell 81: 513-
523 (1995)
Stennicke HR, Deveraux QL, Humke EW, Reed JC, Dixit VM Savesen GS: Caspase-9 can
be activated without proteolytic processing. J Biol Chem 274: 8359-8362 (1999)
Sun XM, MacFarlane M, Zhuang J, Wolf BB, Green DR, Cohen GM: Distinct casapase
cascades are initiated in receptor-mediated and chemical-induced apoptosis. J Biol Chem
274: 5053-5060 (1999)
Takahashi T, Tanaka M, Brannan CI, Jenkins NA, Copeland NG, Suda T, Nagata S:
Generalized lymphoproliferative disease in mice, caused by a point mutation in the Fas
ligand. Cell 76: 969-976 (1994a)
Takahashi T, Tanaka M, Inazawa J, Abe T, Suda T, Nagata S: Human Fas ligand: Gene
structure, chromosomal location and species specificity. Int Immunol 6: 1567-1574
(1994b)
Tewari M, Quan LT, O'Rourke K, Desnoyers, Zeng Z, Beidler DR, Poirier GG, Salvesen
GS, Kixit VM: Yama/CPP32 beta, a mammalian homolog of CED-3, is a CrmA-
inhibitable protease that cleaves the death substrate poly(ADP-ribose) polymerase. Cell 81:
801-809 (1995)
Thornberry NA Lazebnik Y: Caspases: enemies within. Science 281: 1312-1316 (1998)
79
Trauth BC, Klas C, Peters AM, Matzku S, Möller P, Falk W, Debatin KM, Krammer PH:
Monoclonal antibody-mediated tumor regression by induction of apoptosis. Science
245:301-305 (1989)
Tsujimoto Y, Cossman J, Jaffe E, Croce CM: Involvement of the bcl-2 gene in human
folicular lymphoma. Science 228: 1440-1443 (1985)
Ucker DS, Obermiller PS, Eckhart W, Apgar JR, Berger NA Meyer J: Genome degestion
is a dispensable consequence of physiological cell death mediated by cytotoxic T
lymphocytes. Mol Cell Biol 12: 3060-3069 (1992)
Vergen AM, Ekert PG, Pakusch M, Silke J, Connolly LM, Reid GE, Moritz RL, Simpson
RJ, Vaux DL. Cell 102: 43-53 (2000)
Vier J, Linsinger G, Häcker G: Cytochrome c is despensable for Fas-induced caspase
activation and apoptosis. Biochem Biophys Res Commun 261:71-78 (1999)
Walker NP, Talanian RV, Brady KD, Dang LC, Bump NJ, Ferenz CR, Franklin S, Ghayur
T, Hckett MC, Hammill LD, Herzog L, Hugunin M, Houy W, Mankovich JA, McGuiness
L, Orlewicz E, Paskind M, Pratt CA, Reis P, Summani A, Terranova M, Welch JP, Xiong
L, Möller A. Tracey DE, Kamen R, Wong WW: Crystal structure of teh cysteine protease
interleukin-1 beta-converting enzyme: a (p20/p10)2 homodimer. Cell 78: 343-352 (1994)
Wang GQ, Gastman BR, Wieckowski E, Goldstein LA, Rabinovitz A, Yin XM,
Rabinowich H: Apoptosis-resistant mitochondria in T cells selected for resistance to Fas
signaling. The Journal of Biological Chemistry 276: 3610-3619 (2001)
Watanabe-Fukunaga R, Brannan CI, Itoh N, Yonehara S, Copeland NG, Jenkins NA,
Nagata S: The cDNA structure, expression, and chromosomal assignment of the mouse Fas
antigen. J Immunol 148: 1274-1279 (1992)
80
Wesselborg S, Engels ICH, Rossmann E, Los M, Schulze-Osthoff K: Anticancer drugs
induce caspase-8/FLICE activation and apoptosis in the absence of CD95 receptor/ligand
interaction. Blood 93: 3053-3063 (1999)
Wolf BB, Green DR: Suicidal tendencies: apoptotic cell death by caspase family
proteinases. J. Biol. Chem. 274: 20049-20052 (1999)
Wyllie AH: Glucocorticoid-induced thymocyte apotosis is associated with endogenous
endonuclease activation. Nature 284: 555-556 (1980)
Yang J, Liu X, Bhalla K, Naekyung K, Ibrado AM, Cai J, Peng TI, Jones DP, Wang X:
Prevention of apoptosis by Bcl-2: Release of cytochrome c from mitochondria blocked.
Science 275:1129-1132 (1997)
Yonehara S, Ishii A, Uonehar M: A cell-killing nomoclonal antibody (anti-Fas) to a cell
surface antigen codownregulated with the receptor of tumor necrosis factor. J Exp Med
169: 1747-1756 (1989)
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