Microphysiometry in the Evaluation of Cytotoxic Drugs with …160922/FULLTEXT01.pdf ·...

Comprehensive Summaries of Uppsala Dissertationsfrom the Faculty of Medicine 1068

_____________________________ _____________________________

Microphysiometry in theEvaluation of Cytotoxic Drugswith Special Emphasis on the

Novel CyanoguanidineCHS 828

BY

SARA EKELUND

ACTA UNIVERSITATIS UPSALIENSISUPPSALA 2001

Dissertation for the Degree of Doctor of Philosophy (Faculty of Medicine) in ClinicalPharmacology presented at Uppsala University in 2001.

ABSTRACT

Ekelund, S. 2001. Microphysiometry in the evaluation of cytotoxic drugs with specialemphasis on the novel cyanoguanidine CHS 828. Acta Universitatis Upsaliensis.Comprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine 1068,59 pp. Uppsala. ISBN 91-554-5105-5.

This thesis describes the use of a new technology, the Cytosensor® microphysiometer, inthe in vitro evaluation of cytotoxic drugs, using the lymphoma cell line U-937 GTB andprimary cultures of tumour cells from patients as main model systems. The method wasspecifically applied to study the metabolic effects of the novel cyanoguanidine N-(6-(4-chlorophenoxy)hexyl)-N´-cyano-N´´-4-pyridylguanidine, CHS 828, currently in phase I/IIclinical trials.

The Cytosensor® measures metabolic effects as changes in the rate of extracellularacidification of cells exposed to a drug by perfusion. A number of standard cytotoxicdrugs were found to produce typical and reproducible acidification response patternsduring observation times up to 20 h. There seemed to be a relationship between a decreasein acidification and cytotoxicity, measured in the fluorometric microculture cytotoxicityassay (FMCA), after 20-24 h of continuous drug exposure.

In U-937 cells, CHS 828 induced a cytotoxic effect characterised by a steep con-centration-response relationship followed by a plateau. After 24 h of incubation the DNAand protein synthesis were turned off. CHS 828 was found to produce a rapid and pro-longed increase in extracellular acidification and lactate production similar to that of thestructurally related mitochondrial inhibitor m-iodobenzylguanidine (MIBG). The CHS 828induced acidification was observed in cell lines as well as in cells from various tumourtypes from patients and probably originates from increased glycolytic flux. The effectsmay be secondary to block of oxidative phosphorylation in the mitochondria, but the rele-vance of the early acidification is not clear. CHS 828 seemed to induce a late, at approxi-mately 15 h, inhibition of the glycolysis followed by loss of ATP and subsequent celldeath. After exposure to MIBG the loss of ATP and cell death occurred earlier and inparallel. The effects of CHS 828 were not found to resemble those of the structurally rela-ted polyamine biosynthesis inhibitor methylglyoxal-bis(guanylhydrazone) (MGBG). Thus,CHS 828 may represent a new and interesting mode of cytotoxic action worthwhile forfurther development.

In combinatory studies, a synergistic interaction was demonstrated between CHS828 and the non-toxic drug amiloride. Additive-to-synergistic effects were also seen bet-ween CHS 828 and the bioreductive cytotoxic drug mitomycin C. In U-937 cells as well asin tumour cells from patients, CHS 828 demonstrated synergistic interactions in combina-tion with melphalan and etoposide.

It is concluded that measurement in the Cytosensor® microphysiometer of earlycellular metabolic changes is a feasible and potentially valuable complement to more con-ventional methods used in the evaluation of anticancer agents.

Key words: Cytotoxic drug development, Cytosensor® microphysiometer, cellular metabo-lism, CHS 828, guanidino-containing compound.

Sara Ekelund, Clinical Pharmacology, Department of Medical Sciences, UniversityHospital, SE-752 85 Uppsala, Sweden

© Sara Ekelund 2001

ISSN 0282-7476ISBN 91-554-5105-5

Printed in Sweden by Tryck & Medier, Uppsala 2001

“Why are things as they are and not otherwise?”Johannes Kepler (1571-1630), German atronomer

To myself...

Table of contents

Abstract 1

Papers discussed 6

List of abbreviations 7

Introduction 8Chemotherapy for cancer 8

Mechanistic classes of cytotoxic drugs 9Cell death and apoptosis 10

Cellular energy metabolic pathways 11Glycolysis 11Tricarboxylic acid cycle (TCA) 11Respiratory chain - oxidative phosphorylation 12Pentose phosphate pathway 13Energy yield and acid production of the metabolic pathways 13

Regulation of intra- and extracellular pH 13Effects of cytotoxic drugs on cellular metabolism 15

MIBG 16MGBG 17CHS 828 18

Cytotoxic drug development and evaluation 19Principles for cytotoxic drug development 19Role of pre-clinical models in development of cytotoxic drugs 19

Aims of the thesis 21

Materials and methods 22Cells and cell culture 22Experimental drugs 22Measurement of extracellular acidification 22Measurement of cytotoxicity 24Measurement of DNA and protein synthesis 24Measurement of polyamine biosynthesis 25Measurement of glucose consumption 25Measurement of lactate production 25Measurement of ATP 26Measurement of G6PD 26Measurement of changes in intracellular pH 26Interaction analyses 26Presentation of results 27

4

Sara Ekelund: Microphysiometry in the Evaluation of CHS 828

Results and discussion 28Cellular metabolic responses to cytotoxic drugs as measuredby the Cytosensor® (Paper I) 28Metabolic effects of CHS 828 - Comparison with MIBG and MGBG (Papers II & III) 31Further aspects of the metabolic effects induced by CHS 828 (Paper IV) 35Investigation of CHS 828-combinations (Paper V) 38

Combinations between standard drugs and CHS 828 intumour samples from patients (unpublished data) 41

Overall discussion and concluding remarks 42The Cytosensor® 42CHS 828 43

Conclusions 47

Acknowledgements 48

References 50

5

Papers discussed

This thesis is based on the following original papers, which will be referredto in the text by their Roman numerals:

I S Ekelund, P Nygren and R Larsson; Microphysiometry: new tech-nology for evaluation of anticancer drug activity in human tumourcells in vitro. Anti-Cancer Drugs 1998; 9: 531-538.

II S Ekelund, G Liminga, F Björkling, E Ottosen, C Schou, L Binderupand R Larsson; Early stimulation of acidification rate by novel cyto-toxic pyridyl cyanoguanidines in human tumor cells: comparisonwith m-iodobenzylguanidine. Biochemical Pharmacology 2000; 60:839-849.

III S Ekelund, Å Sjöholm, P Nygren, L Binderup and R Larsson;Cellular pharmacodynamics of the cytotoxic guanidino containingdrug CHS 828. Comparison with methylglyoxal-bis(guanylhydrazo-ne). Eur J Pharmacol 2001; 418: 39-45.

IV S Ekelund, R Larsson and P Nygren; Metabolic effects of the cyto-toxic guanidino-containing drug CHS 828 in human U-937 lympho-ma cells. Manuscript, 2001.

V S Ekelund, I Persson, R Larsson and P Nygren; Interactions betweenthe new cytotoxic drug CHS 828 and amiloride and mitomycin C ina human tumour cell line and in tumour cells from patients.Manuscript, 2001.

Reprints were made with the permission of the publishers.

6


List of abbreviationsALL Acute lymphocytic leukaemia AML Acute myelocytic leukaemia AraC CytarabineATP Adenosine triphosphateBCECF 2´,7´-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein CHS 828 N-(6-(4-chlorophenoxy)hexyl)-N´-cyano-N´´-4-pyridylguanidineCI Combination indexCisP CisplatinumCLL Chronic lymphocytic leukaemiaCNCn α-Cyano-4-hydroxy cinnamic acidCO2 Carbon dioxide2DG 2-deoxy-D-glucose DHEA Dehydroisoandrosterone DIDS Diisothiocyanatostilbene-2,2´-disulfonic acid DMSO Dimethyl sulphoxideDox DoxorubicinFAD Flavin adenine nucleotide FDA Fluorescein diacetate FMCA Fluorometric microculture cytotoxicity assayG6PD Glucose 6-phosphate dehydrogenaseGTP Guanosine 5´-triphosphateh Hour(s)Melph MelphalanMGBG Methylglyoxal-bis(guanylhydrazone) MIBG m-Iodobenzylguanidine min Minute(s)MMC Mitomycin CMTT 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromideNAD+/NADH Nicotine adenine dinucleotide NADP Nicotine adenine dinucleotide phosphateNCI National Cancer InstituteNTP Nucleoside 5´-triphosphate O2 Molecular oxygenODC Ornithine decarboxylase Pac PaclitaxelPARP Poly(ADP-ribose) polymerasePBMC Peripheral blood mononuclear cells PBS Phosphate buffered salinePET Positron emission tomographypHi Intracellular pHpHe Extracellular pHPPP Pentose phosphate pathways Second(s)SAMDC S-adenosyl methionine decarboxylaseSEM Standard error of the meanSI Survival indexTCA Tricarboxylic acid cycle Topo TopotecanVcr VincristineVP16 Etoposide

7

Introduction

Chemotherapy for cancerThe development of effective and specific medical treatment for malignantdiseases has been difficult since no general and major differences betweentumour and normal cells that could be exploited for targeting therapy havebeen found. Thus, a major problem is to direct the anticancer therapy totumour cells only, without causing any, or only limited, toxicity to normalcells. The therapeutic index of most chemotherapeutic agents is narrow,leading to a number of unpleasant and potentially life-threatening side-effects [1].

The currently available drugs for treatment of cancer originate froma fairly limited number of chemical structures. In Sweden, there are cur-rently 45 registered cytotoxic drugs that can be classified according to theirchemical structure and/or putative mechanism of action (Table 1) [2].Classically, most of these drugs have been described to interact with cellreplication, e.g., DNA synthesis, transcription, or mitosis.

Despite some progress, chemotherapy for cancer is mostly far fromsuccessful. Thus, for the major tumour types in the advanced setting, che-motherapy mainly provides palliation, i.e., symptom relief and modest pro-longation of life [3]. There is, thus, a great need for development of drugswith improved efficacy and a broader spectrum of activity as well findingof drugs with new mechanisms of action. A better understanding of themechanisms underlying the transformation process to malignant cells aswell as of the action of cytotoxic drugs is required if new drugs are to bemore effectively developed.

8


Alkylating agents Antimetabolites Cytotoxic antibioticsand related substances

Topoisomeraseinteractive agents

Platinumagents

Antimicrotubuleagents

Cyclophosphamide

Chlorambucile

Melphalan

Iphosphamide

Busulfan

Thiotepa

Lomustine

Temozolomide

Dacarbazin

Methotrexate

Mercaptopurine

Thioguanine

Cladribine

Fludarabine

Cytarabine

Fluorouracil

Gemcitabine

Capecitabine

Daktinomycin

Doxorubicin

Daunorubicin

Epirubicin

Idarubicin

Mitoxantrone

Bleomycin

Mitomycin C

Etoposide

Teniposide

Topotecan

Irinotecan

Vinblastine

Vincristine

Vindesin

Vinorelbin

Paclitaxel

Docetaxel

Cisplatinum

Carboplatinum

Oxaliplatinum

Miscellaneousagents

Amsacrine

Asparaginase

Altretamine

Hydroxicarbamide

Miltefosin

Estramustin

Table 1. Cytotoxic drugs approved for marketing in Sweden according to the Swedish

FASS, 2001

Chemotherapy for cancer often uses combinations of agents withdifferent mechanisms of action rather than single agent therapy. This isbecause of the therapeutic advantage (better effect, fewer side-effects) thatcombinations usually provide over single agents [1]. When a new antican-cer agent is being developed, it is important to investigate potential candi-dates for combination therapy. The presently used clinical protocols forcancer combination therapy were mainly obtained empirically from clinicaltrials [4]. A better strategy for selection of combinations of chemotherapeu-tic agents is needed.

Currently there is great optimism for the future with respect to deve-lopment of new concepts for the treatment of cancer. The cumulated newknowledge on, e.g., the genetic basis for cancer development, properties ofthe immune system, mechanisms for cellular signal-transduction and newvessel formation in tumours (angiogenesis) has opened up for new and per-haps fruitful approaches in cancer treatment. However, the path from basicresearch to established treatment is long and it seems reasonable to believethat the old concept of cytotoxic drugs will be part of cancer treatment forquite some time [3]. Thus, new methods and concepts for more efficientdevelopment of these drugs seem worthwhile.

Mechanistic classes of cytotoxic drugsThe main groups of cytotoxic anticancer agents currently registered andused in Sweden, and their putative mechanisms of action, are briefly descri-bed below:

The alkylating agents belong to the oldest class of anticancer drugsand the first clinical studies of nitrogen mustards were initiated in 1942[5]. The alkylating agents are chemically diverse drugs that may undergotransformation to produce reactive intermediates that can bind covalentlyto electron-rich moieties on biological molecules, e.g., DNA. Alkylation ofbases in DNA appears to be the major cause of lethal toxicity [6].

The antimetabolites are among the best characterised and most ver-satile of all chemotherapeutic drugs [1]. Pyrimidine-, purine- and folic acidanalogues are all antimetabolites. These are drugs that have been synthesi-sed to inhibit crucial biochemical pathways, usually leading to inhibition ofDNA or RNA synthesis and these drugs tend to be cell-cycle dependent [6].

Microtubules are components of the mitotic spindle with an impor-tant function during cell division. Microtubules are composed of two subu-nits of tubulin, which form a heterodimer. Tubulin heterodimers andmicrotubules are in a state of dynamic equilibrium with continuous forma-tion and degradation and this process is an important cellular target forcancer chemotherapy. Both the vinca alkaloids and the taxanes belong tothe class of antimicrotubule agents. The former acts by binding to the pro-tein tubulin, thus inhibiting its polymerization to form microtubules. Thetaxanes bind to and stabilise the microtubules and, thereby, the depolyme-risation to tubulin is prevented and essential mitotic functions are inhibited[1].

9

Introduction

The chemical structure of the anthracycline antibiotics consists of apigmented tetracyclic ring structure, an unusual sugar and a lateral chain.The first anthracyclines were produced by the Streptomyces species, but thesecond-generation anthracyclines are synthetic. This class of drugs is one ofthe most clinically used anticancer agents, but their clinical use is limitedby irreversible cardiac toxicity [1]. These compounds act by intercalatingwith DNA and many functions of the DNA are affected, including DNAand RNA synthesis, leading to single and double-strand breaks. Theanthracyclines are also known to interact with the function of the nuclearenzymes topoisomerases (type I and II), forming stabile complexes betweenenzyme and DNA, thus leading to permanent breaks of the DNA strands[5].

The topoisomerase interactive agents take part in vital cellular func-tions by interacting with topoisomerases. The mechanisms of these enzy-mes involve DNA cleavage and strand passage through the break, followedby religation of the cleaved DNA [1]. Etoposide and teniposide are semi-synthetic podophyllotoxin derivatives similar in action forming a complexwith toposiomerase II and DNA, which results in double-stranded DNAbreaks as religation of DNA. The camptothecin analogues, topotecan andirinotecan, interact with the complex between topoisomeras I and DNA [1,5].

The platinum-containing agents are unusual since they are inorganiccompounds, containing a metallic element. Cisplatinum was the first plati-num-containing agent and it was discovered by accident. It is one of themost important drugs for treatment of solid tumours and acts by a mecha-nism similar to the alkylating agents [1].

Cell death and apoptosis Above, the classical mechanisms of action for cytotoxic drugs are descri-bed. With increased understanding of tumour cell biology and with newtechniques, the picture has become more complicated, with additionalaspects to consider. One important finding is that most of the cytotoxicdrugs kill tumour cells by inducing programmed cell death, i.e., apoptosis[7]. Probably, the drugs induce damage to vital cellular functions that inturn initiate apoptosis. However, the connection between apoptosis and themechanisms of cytotoxic drug action is not clear. The alternative way forcells to die is by necrosis.

The processes of apoptosis and necrosis are regulated by many ofthe same biochemical intermediates [8] and under pathological conditionsin vivo apoptosis and necrosis may often coexist [9]. Apoptotic cell death isan active process, characterised by chromosomal condensation, cellularshrinkage, cytoplasmic blebbing, internucleosomal DNA fragmentation andintact membranes [10]. It occurs in a variety of cellular systems and inresponse to many different stimuli including those of antineoplastic drugs[7, 8]. The morphological changes in cells undergoing necrosis are quitedifferent in that necrosis is a passive process involving cell swelling and

10


membrane rupture, which triggers an inflammatory process [8]. The complete apoptotic programme has been shown to involve ener-

gy-requiring steps [11, 12] and the balance between death by apoptosis andnecrosis appears to depend upon the intensity of the injury [12, 13]. Thus,ATP levels have been shown to serve as a switch between apoptosis andnecrosis [12-14] and in endothelial cells glutamine could partially restoreATP levels after H2O2 exposure, resulting in a significant increase in apop-tosis instead of necrosis [15]. In cultured neurones it was demonstratedthat intracellular energy levels were rapidly dissipated in necrosis, but notin apoptosis [16]. In fact, it is necessary to deplete tumour ATP levels by asmuch as 85% of normal levels or otherwise cell viability may be maintai-ned [17].

Thus, for understanding the action of cytotoxic drugs it seems to beimportant to study the effects of cytotoxic drugs on cellular energy meta-bolism. The principal cellular pathways for energy metabolism are, therefo-re, summarised below.

Cellular energy metabolic pathwaysAn important characteristic of fast-growing cancer cells is their elevatedrate of glycolysis and the formation of lactate from glycolytic pyruvate [18,19]. It has been demonstrated that as much as 90% of glycolytic pyruvatein cancer cells can be reduced to lactate [20].

Mitochondrial respiration is the major source of ATP from glucoseunder most conditions in vivo, but glycolysis usually predominates in vitro[21]. The ATP yield from glucose through lactate formation is much lowerthan from the complete oxidation of glucose through the respiratory chain.Still, this inefficient way of obtaining energy is favoured in tumour cells[22]. This metabolic behaviour probably depends on several factors. Theglycolytic pathway contains a different isozymic composition from the nor-mal cell and the activity of key enzymes of regulation is often increased[23]. Other factors discussed are, e.g., facilitated glucose transport andhigh glutaminase activity [23, 24].

The principal cellular energy-yielding pathways are summarisedbelow (Figure 1).

GlycolysisGlucose enters the cells by passive transport through carrier proteins [25].The first step in glucose metabolism is the glycolysis, which takes place inthe cytoplasm. Glycolysis is composed of two phases divided in nine stepswith the net gain of 2 molecules of ATP per glucose molecule. The firstphase concerns glucose phosphorylation and its conversion into glyceralde-hyde-3-phosphate. The second is the conversion of glyceraldehyde-3-phos-phate into pyruvate or lactate coupled to ATP formation [20].

Tricarboxylic acid cycle (TCA)The next step in glucose degradation is the TCA. All of the reactions in the

11

Introduction

cycle take place in the mitochondria. Pyruvate enters the mitochondria andis converted to the acetyl groups of the chemically reactive acetyl CoA bydecarboxylation. In the TCA the acetyl groups are oxidized to produceCO2, NADH, FAD and GTP. The CO2 produced diffuses from the mito-chondrion and leaves the cell [25].

Respiratory chain - oxidative phosphorylationIt is in the respiratory chain, the final stage in the degradation of glucose,that most of the ATP is generated. In a complicated series of steps that reli-es on electron transport, NADH produced in the TCA reacts with molecu-lar oxygen (O2 from water) to produce ATP and H2O [25].

12


Acetyl COA

Tricarboxylicacid cycle

Mitochondria

Oxidativephosphorylation

NADH + H+

Complex

II

Complex

III

Complex

I

2H+ 2H+ 2H+

H+

F0

F1

ATP

ADP

+ Pi

H+

NADH + H+

Glucose

Pyruvate Lactate

Glycolysis

Cytosol

CellmembraneGlucose

Glucose

6-phosphateRibose 5-phosphate

4 carbon

5 carbon

7 carbon

PentosephosphatepathwayADP+

ATP

NADPH

e- transport chain

(mitochondria inner membrane)

CO2

CO2

CO2

Figure 1. The principal energy-yielding pathways in cells are schematically described

and the different waste-products indicated.

NADH acts as a source of readily transferable electrons in cells. Theoxidation of NADH involves the transfer of a pair of electrons, issued fromtwo hydrogen atoms, down through the electron transport chain in theinner membrane of the mitochondria to reduce two molecules of oxygen toform H2O. During the transport energy is released at different couplingsites, called complex I, II and III, generating an “electrochemical protongradient”. This gradient in turn drives a flux of protons to re-enter thematrix through a special enzyme complex, F1F0-ATP synthase. Thereby ATPis generated inside the mitochondrion along with H2O and the newly madeATP is then transported from the mitochondrion to the rest of the cell [20,25].

Pentose phosphate pathwayThe first step in the glycolysis results in glucose-6-phosphate. This is anintermediate that can also enter the pentose phosphate pathway (PPP) [23].The two major products of the PPP are NADPH and ribose-5-phosphate.NADPH is used in reductive biosynthesis as an electron acceptor, whereasribose-5-phosphate is used in the synthesis of RNA, DNA and nucleotideenzymes. There is an interplay between the PPP and glycolytic pathwaythat enables the levels of NADPH, ATP and building blocks such as ribose-5-phosphate and pyruvate to be continuously adjusted to meet the cellularneeds [26].

Energy yield and acid production of the metabolic pathwaysIn a comparison between the principle energy-yielding metabolic pathwaysand with glucose as the carbon source, respiration through oxidative phos-phorylation is the pathway that yields most ATP (36), followed by thecombination of PPP + glycolysis + oxidative phosphorylation (27).Furthermore, glycolysis produces most protons per ATP as compared withrespiration through oxidative phosphorylation or through the combinationof PPP + glycolysis + oxidative phosphorylation [21].

Regulation of intra- and extracellular pHThe new drug investigated in this thesis, N-(6-(4-chlorophenoxy)hexyl)-N´-cyano-N´´-4-pyridylguanidine (CHS 828), was found to stimulate cellularmetabolism and increase the production of acidic waste-products thatcould subsequently decrease extracellular pH (pHe). Such effect on pHemight also have implications for intracellular pH (pHi). Therefore, regula-tion of pHe and pHi are briefly discussed below.

Due to the poor vascularisation in most solid tumours, there arehypoxic and anoxic areas heterogeneously distributed within the tumourmass [27]. The metabolic rate of tumour cells is usually high, leading to adeficiency of both nutrients and oxygen, and the production of increasedamounts of waste-products like protons, lactate and CO2. These circum-stances will together result in an acidic pHe [27]. Thus, tumour pH isknown to be approximately 0.5 units lower than in normal tissues [28, 29].

13

Introduction

The activities of a large number of intracellular enzymes that takepart in the cellular metabolism are pH-sensitive. Protein, DNA and RNAsynthesis are affected by pH and increase with increasing intracellular pHwithin the physiological range [30]. Thus, in order to survive and prolifera-te under acidic conditions, tumour cells become dependent on mechanismsto maintain pHi within the physiological range. There are three majormembrane-based transport systems that are involved in the regulation ofpHi, by excreting acid from the inside to the outside of the cells [31](Figure 2).

The Na+/H+ exchanger or antiport is a glycoprotein present in allmammalian cells. This exchanger acts to increase pHi and catalyses an elec-troneutral exchange of Na+ and H+ across the cell membrane [31]. Thedriving force for the H+ extrusion is provided by the Na+ gradient acrossthe membrane [32]. The exchanger is inhibited by amiloride, which acts onthe extracellular side of the membrane [31].

The Na+ dependent Cl-/HCO3- exchanger acts to increase pHi.

Entry of HCO3- into the cell allows buffering of H+ according to the reac-

tions: H+ + HCO3- ↔ H2CO3 ↔ H2O + CO2 [31]. This anion exchanger is

present in U-937 cells [33], a cell line frequently used in this thesis. Theexchanger is inhibited by the stilbene derivative, diisothiocyanatostilbene-2,2´-disulfonic acid (DIDS), but is insensitive to amiloride [31, 32].

The end product of the glycolytic pathway is lactic acid that mustbe transported out of cells. Lactic acid is dissociated into a lactic anion anda proton at physiological pH and therefore diffusion across the cell mem-brane is probably minimal. Instead, lactate may be transported by the Na+

dependent anion exchanger, but also via the lactate/proton symport thatspecifically transports lactate and other monocarboxylic acids such as

14


Nucleus

HCO3

-

Cl-

Na+

Na+

H+

H+

Lactate

3

- -Na

+-dependent

HCO /Cl exchanger/HNa+ +

antiport

Cytoplasm

Cell membraneH+/ lactate

symport

Figure 2. The main mechanisms considered to regulate acid excretion in cells.

pyruvate [34]. The transport is electroneutral and probably involves sym-port with H+ and thereby contributes to the removal of protons [31]. Thissymport is inhibited by α-cyano-4-hydroxy cinnamic acid (CNCn) [34].

Effects of cytotoxic drugs on cellular metabolismThe effect of cytotoxic drugs on cellular energy metabolism has so far notbeen a major field of research. However, some knowledge has accumulated.

The alkylating agents have been described to cause a decrease oftumour NAD+ levels. In addition, direct inhibition of purified glycolyticenzymes has also been demonstrated by some alkylating agents, but nocausative relationship between pharmacological effects and the degree ofglycolytic inhibition was observed [35].

Bleomycin and cytarabine (AraC) were shown to be inactive on bothglycolysis and respiration, while doxorubicin (Dox) had a stimulating effecton respiration [36]. However, Dox has also been described to inhibit cellu-lar respiration [37].

Actinomycin-D has been shown to induce an early and transientincrease in ATP and GTP contents, while anthracyclines induced a corres-ponding increase in the total nucleoside triphosphate pool (NTP). Thiseffect was suggested to be characteristic for these drugs, since cisplatinum(CisP) and AraC did not alter the NTP content [38]. In another study theATP levels in cells exposed to CisP decreased by one-third within 20 min[39]. The mitochondrias were suggested to be the primary target for CisP,with inhibition of complexes I to IV of the respiratory chain, resulting indecreased ATP levels [39, 40]. Other cytotoxic drugs, e.g., paclitaxel (Pac)and etpopside (VP16), elicited similar effects, indicating the possibility of ageneral effect of drugs inducing apoptosis [40].

Pac has been reported to alter rat hepatocyte metabolism by inhibi-ting the respiratory chain, causing an immediate reduction of oxygen con-sumption and increased glycolytic flux [41]. The ATP levels were notenough to compensate for the inhibition of oxidative phosphorylation andcell viability was subsequently impaired. This ATP reduction might be aconsequence of decreased activity of regulatory glycolytic enzymes [42].

A build-up of glycolytic intermediates and decrease in cellular ener-gy content was reported for cells exposed to Dox, explained by the inhibi-tion of glycolysis at the level of glyceraldehyde-3-phosphate dehydrogenase,probably by NAD+ depletion following poly(ADP-ribose) polymerase(PARP) activation [43]. When investigating the role of microtubules in theregulation of metabolism, Pac and vinblastine were both found to decreasethe glycolytic rate [44]. This was surprising since these drugs are known toinduce opposite effects on microtubule assembly. Based on these findings itwas suggested that both drugs produce their effect on glycolytic rate bycompeting with the glycolytic enzymes for binding sites on the tubulinmolecule [44].

Basic alterations in glucose metabolism have been found to be asso-ciated with drug resistance in several tumour cell lines resistant to Dox [45-

15

Introduction

48]. These cell lines exhibited an increased rate of glycolysis, oxidativephosphorylation and PPP with a subsequent increased O2 consumption aswell as an increased production of CO2, lactate and ATP.

The class of compounds known as guanidines, notably m-iodoben-zylguanidine (MIBG) and methylglyoxal-bis(guanylhydrazone) (MGBG),are described to affect cellular metabolism. MIBG is known to inhibit cel-lular respiration at complex I and III [49-51], while MGBG is believed toinhibit the biosynthesis of polyamines [52, 53]. These guanidino-containingdrugs were shown to have antitumour properties several decades ago andover the years have been subjected to both preclinical and clinical evalua-tion [52, 54-56]. When exploring the metabolic and cytotoxic effects of thenewly discovered cytotoxic pyridyl cyanoguanidine CHS 828, MIBG andMGBG were used as comparisons. These substances are therefore dealtwith in greater detail in the following sections.

MIBGMIBG has a molecular mass of 324.1 g/mol and was synthesised over 20years ago [57]. The structure of MIBG is shown in figure 3. It is the gua-nidino-group of MIBG that is essential for the cytotoxic effect of the drug[58] and it is also this part that resembles CHS 828 [59].

MIBG is a structural and functional analogue of the natural neuro-transmitter norepinephrine, but it does not act like a false hormone [58].In its radio-iodinated form, MIBG is used clinically as a tumour-seekingradiopharmaceutical agent for the diagnosis and treatment of neuro-endo-crine tumours [60].

A concentration of 10 µg/ml (equals 31 µM) MIBG inhibited themitochondrial respiration of leukaemia L1210 cells [49, 61]. In studiesusing the human neuroblastoma cell line SK-N-BE(2c), optimal arrest ofproliferation was seen at MIBG concentrations of 25 µM, but the mito-chondrial respiratory chain was almost completely inhibited at 10 µM [50].Similar results were also described in a leukaemic cell line, Molt-4 [51].This suggests that MIBG induces proliferation arrest not only by effects onmitochondrial oxidative phosphorylation [50, 51].

MIBG also has also demonstrated antitumour effects in animalmodels. MIBG alone causes hyperglycaemia and induces a lowering oftumour extracellular pH [62]. In addition, a stimulatory effect on tumourcell glycolysis in vivo was indicated by a downshift of tumour pH and anincrease in plasma lactate levels in tumour-bearing animals with the decrea-se in pH limited to malignant tissue [63].

The exact mechanism by which MIBG induces cytotoxicity and celldeath is not yet fully known, but there are several proposals. Progressiveacidification of the culture medium, by lactate and other metabolic waste-products, has suggested that MIBG primarily affects mitochondrial respira-tion with subsequent compensation in glycolytic flux [49, 50, 54, 64]. Thestimulation of glycolysis occurs when the mitochondria can not produceATP through the oxidative phosphorylation [65].

16


MGBGThe molecular mass of MGBG is 257.1 g/mol (Figure 3) and the moleculeconsists of two polar aminoguanidino-groups separated by an aliphaticskeleton. There are similarities between MGBG and the natural polyami-nes, especially with spermidine. The polyamines spermidine, spermine andtheir precursor putrescine are present in all mammalian cells, but their phy-siological functions are not well understood [53, 66]. They are involved inmany biologic processes [67], and an increased level of polyamines hasbeen reported in several malignant diseases [68]. Clinical trials with MGBGwere initiated in the early 1960s [69]. The drug is now known to be anantileukaemic agent and has also been reported to have activity againstsome solid tumours [66, 70].

In cultured cells MGBG induces a reduction in the concentrations ofspermidine and spermine [52, 71-73]. In parallel, DNA synthesis and cellproliferation are inhibited [53]. MGBG also exerts other effects not associ-ated with inhibition of biosynthesis of polyamines, e.g., an antimitochon-drial action, inhibition of carnitine dependent oxidation of long chain fattyacids and blocking of intestinal diamine oxidase [52].

In several human and murine cell types, MGBG has been reportedto produce ultrastructural damage to the mitochondria [74, 75]. On theother hand, another study showed that MGBG was without effect on mito-chondrial respiration, as there was no effect on oxygen consumption, ATPcontent or lactate production in a study with human and mouse neuroblas-toma and lymphosarcoma cell lines [54]. In yet another study, MGBG wasfound to exert two effects on mitochondria; protection of mitochondria atlow concentrations and aggregation at higher concentrations [76]. Most ofthe effects of MGBG on cultured cells can be prevented by the concurrentadministration of spermidine at equimolar or higher concentrations [77].

17

NH

N

N

NH

N

O

Cl

NH2

NH

NH

N

CH3

NNH

NH2

NH

NNH2

NH

N H

I

MIBG MGBG

CHS 828

N

N

Figure 3. Chemical structures for the guanidino-containing compounds MIBG, MGBG

and CHS 828.The guanidino-groups are indicated with circles.

Introduction

CHS 828The focus of this thesis is CHS 828, an interesting and promising com-pound with the potential to become a new anticancer agent. The first clini-cal trial was initiated in 1998 and the first phase II study is ongoing inpatients with chronic lymphocytic leukaemia (CLL).

In the 1970s there was an interest in the hypotensive effect of astructural prototype, the potassium channel opener, pinacidil and its analo-gues [59]. Later it was discovered that a number of related pyridyl cyano-guanidines showed antitumour activity in a routine screening programmein a rat model [78]. This finding inspired further investigation of the guani-dines in order to find structure-activity relationship for this class of sub-stances. Crucial for the antiproliferative effect was the cyanoguanidinemoiety but also the length of the carbon chain [78]. Optimisation and tes-ting of the antitumour activity resulted in the selection of a drug candidate,CHS 828. CHS 828 has a molecular weight of 371.87 g/mol and has nohypotensive effect. The chemical structure of CHS 828 is shown in figure3.

CHS 828 has demonstrated interesting properties as a potential anti-cancer agent and has been found active against many tumour cell lines invitro [78, 79]. A differential pattern of antitumour activity was revealedwhen investigating the cytotoxicity of CHS 828 in a panel of 10 humantumour cell lines representing defined mechanisms of resistance, includingthose associated with expression of P-glycoprotein, altered topoisomeraseII, increased GSH levels and tubulin defects [79]. Activity was observed inthe nM to µM range and the typical shape of the dose-response curves wasa concentration dependent decrease in cell survival followed by a plateau.This plateau was cell proliferation independent, since the phenomenon wasalso observed in non-proliferative cell systems, e.g., peripheral bloodmononuclear cells (PBMC) [80, 81].

In an in vitro study of 156 primary cell cultures from haematologi-cal and solid tumours, CHS 828 showed a high activity against tumourcells from CLL as well as from acute leukaemia and high-grade lymphoma[80]. In addition, CHS 828 was less active in PBMC as compared with thehaematological malignancies. Solid tumour cells appeared less responsive.Similar results were also demonstrated in an in vitro hollow fibre assay, inwhich high activity of CHS 828 was shown in CLL, with less activitytowards solid ovarian carcinoma [82].

After oral administration in several animal models, CHS 828demonstrated a broad spectrum of activity while causing only little or notoxicity to the animals. The xenograft models of MCF-7 breast cancer andNYH small cell lung cancer in nude mice, were both found sensitive toCHS 828 [79]. These data are interesting since the NYH xenograft modelwas not very sensitive to standard drugs.

18


Cytotoxic drug development and evaluation

Principles for cytotoxic drug development There are many different ways to proceed in the design and development ofanticancer drugs. New compounds can be discovered by screening ofvarious chemicals. This approach has long been adopted by the NationalCancer Institute (NCI). Today, NCI has an established primary screen inwhich compounds are tested for the ability to inhibit growth of 60 diffe-rent human tumour cell lines [83]. Included in the panel are cell lines origi-nating from leukaemias and cancers of the breast, prostate, lung, colon,ovary, kidney and central nervous system. Active compounds are selectedfor further testing based on several different criteria such as disease-typespecificity, unique structure, potency and demonstration of unique patternof cytotoxicity in the cell line panel, indicating a new mechanism of action[84].

Another strategy in drug discovery is the modification of alreadyexisting compounds [85]. These agents can be altered to enhance their acti-vities and to decrease their unwanted toxic effects. A third approach isrational drug design based on knowledge on specific biochemical or mole-cular targets in tumour cells. This approach might become fruitful as indi-cated by the recently developed tyrosine kinase inhibitor STI571 for treat-ment of chronic myeloid leukaemia [86]. Finally, serendipitous observa-tions might lead to the discovery of new drugs [85].

Role of pre-clinical models in development of cytotoxic drugs The final information on the true performance of new drugs for the treat-ment of cancer can only derive from the clinical trials programme.However, the capacity to test new treatments in patients is limited in com-parison with the great number of promising new drugs. For safety reasonsand for optimising the chance of finding active new drugs, a lot of infor-mation needs to be collected during the pre-clinical phase of the develop-ment programme for a new cytotoxic drug [87].

Briefly, the pre-clinical programme includes studies in vitro, so farmostly on tumour cells lines and subcellular model systems, to characterisethe activity profile of the new drug and mechanisms of action and resistan-ce. Furthermore, some information should be obtained on suitable exposu-re times and cell-cycle dependency that could be of guidance for the treat-ment schedule in vivo.

In vivo studies, mainly in rodent tumour-bearing experimental ani-mals, provide additional information on the antitumour activity, therapeu-tic index and schedule dependency of the new drug. In addition, extensivestudies in non-tumour bearing animals provide necessary information on,e.g., toxicology, pharmacokinetics and local tolerance.

Although extensive knowledge on properties of a new drug is availa-ble from the pre-clinical programme prior to clinical trials, the experienceindicates that very few of the promising drug candidates will progress to

19

Introduction

become clinically useful drugs after extensive and resource consuming tes-ting in patients [88]. A major deficiency in the pre-clinical developmentprogrammes is that the information provided on the antitumour activity ismostly not sufficiently predictive for the final clinical efficacy. Pre-clinicalin vitro and in vivo models with improved clinical relevance are desired,e.g., to be able to focus the clinical trials on patients with tumour typesand settings in which the new drug could be expected to be active.

The use of in vitro assays based on cells prepared from patienttumours, might be an approach to provide clinically more predictive infor-mation on the anticancer activity. Several assays based on the concept oftotal cell kill during short-term culture are available for this [89, 90].Furthermore, the testing of possible new drug combinations can, as a firststep, preferably be done in vitro [91]. Assays, developed to examine thecytotoxicity or biochemical effects of drugs on cultured cells, could indicatethe therapeutic effect of combined agents as well as provide information onpossible mechanisms of drug interactions. Actually, in vitro studies are uni-que in their ability to quantitatively evaluate synergistic or antagonisticinteractions [4].

Unfortunately, these possibilities to obtain pre-clinical data morerelevant for the clinical situation have so far been very little utilised in thedevelopment programmes for new cytotoxic drugs [92].

20


Aims of the thesis

In general, the studies comprising this thesis were focused on the evalua-tion of cytotoxic drugs using the Cytosensor® microphysiometer, with spe-cial emphasis on the novel cytotoxic cyanoguanidine CHS 828. Additionalmethods were used to further characterise the effect of CHS 828 and rela-ted drugs on, e.g., metabolic pathways, cytotoxicity, pHi and DNA/proteinsynthesis.

The more detailed aims of this thesis were to:

• Investigate the feasibility of the Cytosensor® for evaluation of drugsused in the treatment of cancer, with a special emphasis on the novelcytotoxic drug CHS 828.

• Compare the metabolic and cytotoxic effects induced by CHS 828 withthose of the established guanidino-containing compounds MIBG andMGBG.

• Characterise over time the metabolic events induced by CHS 828 andthereby increase the understanding of the mechanism of action of thedrug.

• Investigate drugs suitable for combination with CHS 828 to enhancethe cytotoxic effect of the drug.

21

Aims

Materials and methods

The methods used in this thesis are briefly described in this section. Themain method, i.e., measurement of acidification rate (the Cytosensor®), isdescribed in greater detail.

Cells and cell cultureThe most frequently used cell line was the human histiocytic lymphomacell line, U-937 GTB [93]. Other cell lines used were the CHS 828 resistantsubline U-937 CHS, the myeloma cell line RPMI 8226/S, the small cell lungcancer cell line NCI-H69 and the leukemic cell line CCRF-CEM. In someexperiments tumour cells from patients with haematological or solidtumours were used. The samples were obtained from bone marrow/perip-heral blood sampling, routine surgery or diagnostic biopsy. The samplingwas approved by the local ethical committee at Uppsala UniversityHospital. Cell preparations from these tumours have been described indetail earlier [89, 94].

Patient cells and the cell lines were grown under standard cell-cultu-re conditions (5% CO2, 37°C). Cell culture medium was RPMI 1640 sup-plemented with 10% fetal calf serum, 2 mM glutamine, 50 µg/ml streptom-ycin and 60 µg/ml penicillin (Sigma-Aldrich Co. Ltd., Irvine, UK). Growthand morphology of the cell lines were monitored two or three times aweek.

Experimental drugsCHS 828 and seven structurally related pyridylcyanoguanidines, suppliedfrom Leo Pharmaceutical Products in 10 mM stock solutions, were dissol-ved in dimethyl sulphoxide (DMSO) and stored frozen at –20°C. The drugswere diluted ten times with 33% DMSO and sterile water and further dilu-tions were made using sterile phosphate buffered saline (PBS). MIBG andMGBG were from Sigma. MIBG was dissolved in 10% DMSO and sterilewater and MGBG in sterile water to stock solutions of 1 mg/ml. They werestored at -70°C until use and further dilution was in PBS.

Standard cytotoxic drugs investigated were mitomycin C (MMC),vincristine (Vcr), Pac, CisP, melphalan (Melph), Dox, VP16, AraC andtopotecan (Topo). Other chemicals used were 2-deoxy-D-glucose (2DG),spermidine, dehydroisoandrosterone (DHEA), amiloride, DIDS and CNCn.All these compounds were obtained from commercial sources and were dil-uted as prescribed.

Measurement of extracellular acidificationA silicon microphysiometer, the Cytosensor® (Molecular DevicesCorporation, Sunnyvale, CA, USA), was used to measure the excretion rateof metabolic waste-products.

The Cytosensor® consists of two units with altogether eight parallelmeasurement channels. Culture medium is pumped from a reservoir by a

22


peristaltic pump and passes through a debubbler/degasser, a selection valveand then through the flow sensor chamber (Figure 4). In the measurementchamber the non-adherent cells were immobilised in 25% agarose(Molecular Devices) to ensure that the cells are not washed away to oneside of the chamber by the flow of medium. The optimal number of cellsranges from 104 to 106 cells [95] and in the present experiments there were1.5 x 105 cells in each capsule.

Cells are retained in a micro-volume (2.8 µl) flow chamber in a disc-shaped region between two microporous polycarbonate membranes, inaqueous diffusive contact with the surface of a pH sensitive silicon chip[96]. The chip, together with a reference electrode and other components,forms a light-addressable potentiometric sensor (LAPS), that is used todetect small changes of extracellular acidification rate [21, 97, 98]. Onceeach s the LAPS makes a voltage measurement of the acidification rate thatis linearly related to pH and calculated by the Cytosoft program as -µV/s.

Medium is constantly pumped through two parallel channels at aflow rate of 100 µl/min. When the fluid flows (90 s), the sensor output isstable and reflects a pH near that of medium entering the flow chamber.During flow-off periods (30 s), the acid released from the cells accumulatesin the measuring chamber and the rate of release is quantified by fitting thesensor data to a straight line with the least-squares method. The slope ofthis line represents the acidification rate. No significant perturbations ofcell physiology is caused during a cycle [95]. The change in acidification is

23

Pump

DebubblerValve

Printer and computer

Drug/medium

supply

LED

Sensor

chamber

Reference

electrode

Waste

Waste

Figure 4. A schematic diagram of the Cytosensor‚ system. Each sensor chamber is con-

nected to two fluid reservoirs. A pump delivers fluid (medium with or without drug)

through a debubbler to the selection valve, which determines which of the fluids is direc-

ted to the sensor chamber and which is directed to the waste. After passing the cells in

the sensor chamber, the fluid passes the reference electrode and leaves the system.


indicated as a change in applied voltage (mV) (a decrease of 61 mV isapproximately equal to an acidification of 1 pH unit at 37°C) [95].

Cells were allowed to adapt in the Cytosensor®during one h beforethe acidification rate for each measurement chamber was set to 100%(baseline acidification). Thereafter, drugs were added. The acidificationrates induced by drugs are presented as per cent of baseline acidification.This compensated for small baseline differences and permitted comparisonwithin and between experiments.

The pH of the experimental medium (National Veterinary Institute,Uppsala, Sweden) was set to 7.35-7.39. The medium has no bufferingcapacity since it lacks both bicarbonate and Hepes. Thus, changes in acidi-fication rate could readily be observed and were due only to the metabolicresponse of cells to the drugs. The osmotic balance was preserved by theaddition of 6 ml/l 4 M NaCl. The medium also contained 10 ml/l 200 mML-glutamine, 60 mg/l penicillin and 50 mg/l streptomycin.

Measurement of cytotoxicity Viability of cells was determined by the fluorometric microculture cytotoxi-city assay (FMCA). The assay is based on measurement of fluorescencegenerated from hydrolysis of fluorescein diacetate (FDA) to fluorescein bycells with intact plasma membranes.

This method has been described in detail previously [94]. Briefly,experimental V-shaped, 96-well microtiter plates were prepared in advance,using a pipetting robot (Pro/Pette; Perkin Elmer, Norwalk, CT, USA).Triplets of 20 µl of drug solution were dispensed at ten times the finalexperimental concentration into the plates. The plates are then kept frozenat –70°C until further use.

The cell suspension was added to the plates, 180 µl/well (2 x 104

cell/well for cell lines and 5-10 x 104 and 1-3 x 104 cells/well for haemato-logical and solid tumours, respectively), before incubation during 72 h at37°C and 5% CO2. Blank wells received culture medium only and controlwells contained cell suspension and PBS but no drug. After incubation,drugs and medium were removed and the cells were washed once with PBS.100 µl of 10 µg/ml FDA, was added to each well and the plates were incu-bated for 40 min. The fluorescence generated in each well was then measu-red at 538 nm in a Fluoroscan II (Labsystems Oy, Helsinki, Finland).

Measurement of DNA and protein synthesisProtein and DNA synthesis were measured with a Cytostar-T plate, availa-ble in Amersham´s “In Situ mRNA Cytostar-T assay” kit, (AmershamInternational, Buckinghamshire, UK), a pre-made scintillating 96-wellmicrotiter plate, with scintillation fluid moulded into the bottom of thewells [99, 100]. Cells were suspended in fresh media containing 111 nCi/ml[14C]Thymidine (Amersham CFA.532 56 mCi/mmol, 50 µCi/ml) for DNAsynthesis or 222 nCi/ml [14C]Leucine (Amersham CFB.183, 56 mCi/mmol,50 µCi/ml) for protein synthesis. Cell suspension was added to each well

24


and radioactivity was measured with a Wallac 1450 MicroBeta trilux liqu-id scintillation counter (Wallac OY, Turku, Finland).

Measurement of polyamine biosynthesis The activities of ornithine decarboxylase (ODC) and S-adenosyl methioninedecarboxylase (SAMDC) were investigated in cells exposed to CHS 828and MGBG as measurements of the polyamine biosynthesis. Cells wereincubated with drugs during 1 or 20 h in cell culture flasks. After centrifu-gation, cells were resuspended in 1 ml cold PBS, centrifuged once more andsonicated in 50 µl of a lysis buffer containing 50 mM Tris-HCl (pH 7.4),0.1% Triton-X 100, 4 mM EDTA, 5 mM dithiothreitol, 200 000 U/mlTrasylol, 10 mM benzamidine and 0.1 mg/ml albumin. The homogenateswere centrifuged for 10 min at 4°C before an equal volume of supernatantand incubation buffer was mixed. For ODC measurements, the buffer con-tained 2 mM pyridoxal phosphate, 0.4 mM L-ornithine and 10 µCi/ml L-[1-14C]ornithine and for measurement of SAMDC, the buffer 5 mMNaPO4, 0.4 mM S-adenosyl-L-methionine and 4 µCi/ml S-adenosyl-L-[car-boxyl-14C]methionine. The assays were performed in small glass vials. After60 min incubation at 37 °C, reactions were terminated by addition of 100µl 5 M H2SO4 and 250 µl Hyamine was used to trap liberated 14CO2. Afteraddition of 0.4 M Na2HPO4 to liberate CO2, Unisolve scintillation fluidwas added and the radioactivity quantified by scintillation counting [101,102].

Measurement of glucose consumptionMedium glucose concentration was used as an indirect measurement ofglucose consumption of cells exposed to drug during different time periodsranging from 30 min to 50 h. 2.5 x 105 cells/ml were incubated in cultureflasks at 37°C. Samples were withdrawn from drug-exposed cells and fromcontrol cells. After centrifugation, 2 ml of the supernatants were transfer-red to a tube and stored at 8°C until analysis. The samples were analysedat the University Hospital laboratory using the routine analysis of glucoselevels in plasma and other body fluids. Briefly, the level of glucose wasindirectly determined by measuring the absorbance at 340 nm, whichreflects the amount of NADH produced when glucose is enzymatically oxi-dized.

Measurement of lactate productionThe production of lactate by cells exposed to CHS 828 and MIBG wasmeasured using commercially available lactate reagents and a standardcurve prepared from a lactate standard solution (40 mg/dl) (Sigma).

Cells were incubated at a density of 1 x 106 cells/ml in culture flasksat 37°C. After different exposure times samples, were taken out and centri-fuged at 200g for 5 min. Thereafter, 10 µl of supernatant was added to acuvette together with 990 µl lactate reagents. After 5 min incubation theabsorbance was measured at 540 nm in a spectrophotometer,

25


SPECTRAmax®PLUS (Molecular Devices). Blank values were subtractedand values transformed to µg/ml using the standard curve equation.

Measurement of ATP The commercially available ApoGlow™ (LumiTech Ltd., Nottingham, UK)kit was used to determine the levels of intracellular ATP. The kit is basedon bioluminescent measurements of ATP utilising the luciferase enzyme forformation of light from ATP and luciferin. Cell suspension was plated in aflat-bottomed, white 96-well microplate, which was incubated under stan-dard cell culture conditions. At different time points, ranging from 5 minto 72 h, drugs were added to the plate. Thereafter, the automatically dis-pensing luminometer, Mediators PhL (Mediators Diagnostica, Vienna,Austria) added 100 µl of a mixture of nucleotide monitoring and releasingreagent to each well before measurements were performed at 540 nm.

Measurement of G6PDMeasurment of G6PD was made using enzymatic analysis. CHS 828 wasadded to the cell suspension (5 x 105 cells/ml) after 0, 2, 8 and 24 h incu-bation at 37°C. The cells were harvested by centrifugation and the cell-pel-lets were resuspended in 500 µl sterile water and allowed to freeze at–70°C for 10 min. After thawing, the samples were centrifuged once moreand the supernatant collected and frozen until the enzyme activity analysiswas performed at the University Hospital routine laboratory. Briefly, therate of NADPH formation was measured spectrophotometrically and rela-ted to the enzyme activity [103]. The activity was presented using the unitnkatal/100µl cell suspension.

Measurement of changes in intracellular pHMeasurement of pHi changes was performed using a fluorescence spectrop-hotometer F 2000 (Hitachi Ltd., Tokyo, Japan) and the tetraacetoxymethyl(AM) ester of 2´,7´-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein(BCECF, Sigma).

10 x 106 cells were loaded with 2 µM BCECF AM during 30 min at37°C. Thereafter, cells were centrifuged and washed once with mediumbefore dilution in a 10 ml standard Hepes buffer (Sigma-Aldrich) supple-mented with 10 mM glucose and 0.75 mM calcium chloride and with pHadjusted to 7.4. Portions of 2 ml cell suspension were incubated with con-stant stirring at 37°C in a 1 cm cuvette in the fluorescence spectrophotome-ter. The fluorescence after each addition of drug was monitored until stabi-lity was attained.

Interaction analyses There are many different models for in vitro analysis of drug interactions.In this thesis the isobole method and the additive model were used. Theconcepts of these methods are described below, and in greater detail inPaper V.

26


The isobole method is a commonly used method. Isobole means iso-effect curve and the construction of isoboles requires experimental data forthe agents used alone and in various dose combinations at equi-effect levels[104]. This is a generally valid procedure for analysing interactions bet-ween agents irrespective of their mechanism of action or the nature of theirdose-response relations [104, 105].

In the additive model it is assumed that each agent acts independent-ly of the other and it is a consequence of probability theory that the pro-duct of the individual survival fractions is employed [106]. Zero interactionof the combination is expected to be equal to the product of the survivalindices of its constituents [105].

Presentation of resultsThe main results are summarised below with key findings illustrated infigures. Generally, figure data are presented as mean values ± SEM. Foradditional details on experimental performance and data presentation, seethe individual papers.

27


Results and discussion

Cellular metabolic responses to cytotoxic drugs as measu-red by the Cytosensor® (Paper I) Eight standard cytotoxic drugs with different mechanisms of action wereinvestigated. Each drug induced a reproducible and characteristic responsepattern, from which key features, e.g., stimulation and inhibition of acidifi-cation, the time point when the response curves of the drugs fell below thecontrol curve, and the maximum inhibition of acidification at 20 h, couldbe quantified (Figure 5).

Vcr and VP16 induced distinct peaks of acidification during the firsth, after which the acidification rate eventually declined. The shape of theresponse curves for AraC and Topo were similar to those of Vcr, but withsmaller peaks. Melph and Dox followed the control curve several h beforethe acidification rate declined. For CisP and Pac, on the other hand, theacidification remained above or very close to the control curve for the com-plete duration of the 20 h experiments. For the drugs producing a reducedmetabolic rate at 20 h, a concentration-response relationship was observed.The qualitative nature of the response pattern induced by Vcr was found tobe similar in the T-cell leukaemia cell line CCRF-CEM (not shown). Towhat extent the response patterns obtained are independent of cell typeremains to be established.

The drug concentrations used are known to be highly cytotoxic after72 h of continuous drug exposure. In Cytosensor® experiments a decreasein acidification rate might be due either to a decreased number of viablecells and/or to a decrease in metabolic activity per cell and the proportionsof these two alternatives are likely to be time dependent. Judging from thedrug concentrations used in this study, long exposures inevitably lead to anincreased number of dead cells.

To mimic the time frame of the Cytosensor® experiments, the cyto-toxic effects of the drugs were measured at 24 h in the FMCA. At 20-24 hof drug exposure the two assays correlated reasonably well (Figure 6).There were two exceptions in Melph and Vcr, for which cell survival washigh despite inhibition observed in the Cytosensor®. This discrepancy maybe related to differences in the overall conditions of tests. In theCytosensor® experiments, the temperature of the medium is 37°C and cont-inuously pumped through the sensor chambers so that metabolites andwaste products are removed and new drug continuously added. In themicrotiter plate-based assay, the plates are incubated at 37°C for 72 h withno new addition of drug and no change of medium. Melph and Vcrdecompose rapidly at 37°C [107], which may result in lower exposure inthe FMCA than in the Cytosensor®. This problem may arise also for otherunstable drugs. These features of the Cytosensor® have to be rememberedwhen comparing results obtained with cytotoxicity assays.

Data collected with the Cytosensor® have been compared with stan-dard in vitro assays and found to compare well with assays for second

28


messengers and morphological or proliferative changes [108]. In a recentcomparison with the established tetrazolium salt assay for assessment ofcytotoxicity, the Cytosensor® method was found more sensitive to theeffects on cellular metabolism [109]. In addition, in a human liver cell line,the toxic effects of 10 drugs were shown to produce a time-dependent

29

5 10 15 20

0

25

50

75

100

125

Control

AraC 0.5 µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

CisP 2.5µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

Dox 2.5µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

Melph 2.5µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

Pac 0.5µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

Topo 2.5 µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

Vcr 0.5 µg/ml

Time (h)

5 10 15 20

0

25

50

75

100

125

Control

VP16 5µg/ml

Time (h)

Addition of drug

Figure 5. The effects in U-937 cells of eight standard cytotoxic drugs, with different post-

ulated mechanism of actions, on the extracellular acidification rate as measured using

the Cytosensor®.


reduction in acidification rate after 24 h of drug exposure and the IC50

values calculated compared well with IC50 values obtained from [3H]thymi-dine uptake and Calcein AM fluorescence [110]. Most of the drugs testedinhibited cell metabolism and acidification rate, but ethanol enhanced theactivity during the initial 4 h of exposure, but then irreversibly inhibitedthe metabolism after 24 h [110]. Similarly, in our study several of the cyto-toxic drugs investigated initially stimulated the acidification rate. This phe-nomenon might possibly reflect an energy-requiring defence or response ofcells as a first reaction to the drug.

The feasibility of using the Cytosensor® for measurement of cytotox-ic drug activity profile in primary cultures of patient ovarian carcinomacells was also demonstrated in the present study. The response patterns ofDox was qualitatively similar to those obtained in the U-937 cell line, butCisP showed an initial stimulation of acidification. This type of tumour isknown to be CisP sensitive but it remains to be clarified if this responsereflects the presence of cell type specific response elements mediating apop-tosis.

Early classification of new agents according to pharmacologicalsimilarity or dissimilarity to standard drugs with known mechanism ofaction is desirable in new drug discovery and development. The drug-speci-fic patterns generated by the Cytosensor® as demonstrated by this study,may provide unique and important additional information in this respect.Other potentially valuable applications for anticancer drug evaluation andcharacterisation could be the detailing of schedule dependency and druginteractions. The feasibility of easy change of exposure protocols in eightparallel channels makes the Cytosensor® especially well suited for suchapplications.

30


Vcr Pac CisP Melph Dox VP16 AraC Topo0

25

50

75

100

125Acidification rate

Cell viability

Figure 6. Comparison of the extracellular acidification rate and cell survival in U-937

GTB cells after 20 h and 24 h of drug exposure, respectively.

Metabolic effects of CHS 828 - Comparison with MIBG and MGBG(Papers II & III)CHS 828 induced a rapid and sustained increase in acidification rate. Themetabolic pattern produced by CHS 828 resembled the pattern induced byMIBG, but not that of MGBG (Figure 7). MIBG is known to inhibit mito-chondrial respiration with a compensatory increase in the glycolytic flux,subsequent increase in lactate production, decrease in O2 consumption [49,50, 54] and lowering of tumour extracellular pH [62]. In correspondencewith these changes, the rapid change in acidification rate of CHS 828might be interpreted as a reaction to a mitochondrial block and the pro-longed increase in acidification rate as an enhanced glycolytic activity. Theincrease in acidification rate for CHS 828 was dose-dependent, as a higherconcentration both increased the magnitude of the acidification rate andshortened the time period of the stimulated metabolic activity.

Unpublished data demonstrated that another mitochondrial inhibi-tor, Oligomycin B, an inhibitor of F0F1 ATPase, induced a similar patternwith an instant and enhanced metabolic rate as seen for CHS 828 (Figure7). This further supports the theory of mitochondrial inhibition as anexplanation of the CHS 828 stimulated acidification rate.

An interesting finding was that the acidification induced by CHS828 could be abolished by substituting glucose for pyruvate (10 mM), fur-ther supporting the theory that stimulated glycolysis was the source of theincreased metabolic activity (Figure 8). After 5 h in pyruvate-containingmedium, switching back to glucose medium rendered an overshoot of theextracellular acidification rate. In contrast, after 5 h in pyruvate medium

31

5 101

50

75

100

125

150

17 20

Control

CHS 828 10 µM

MIBG 10 µg/ml

MGBG 10 µg/ml

Oligomycin B 1 µM

Time (h)

Figure 7. Extracellular acidification of the guanidines CHS 828, MIBG, MGBG and

Oligomycin B in U-937 cells using the Cytosensor microphysiometer during continuous

drug exposure for the indicated time period.


and exposure to 10 µM CHS 828, the decreased acidification rate of theexposed cells could not be reversed with glucose medium. Probably, with ablock in both glycolysis (pyruvate) and mitochondria (CHS 828) for thisperiod of time, damage to the energy producing machinery was too severeto allow restored metabolism. When investigating the cytotoxic effect,removal of glucose paradoxically reduced CHS 828-induced cytotoxicitywhile potentiating that of MIBG (Figure 9). With a block in both mito-chondria and glycolysis, an increased cell death would be the expected sce-nario. Also for MGBG the cytotoxic effect was inhibited, but not to thesame extent as for CHS 828.

Without pyruvate, glycolysis is blocked and the cells are dependenton other metabolic pathways for their energy production. In glucose-freemedium substituted with pyruvate, the acidification rate rapidly decreasedto 15-20% of baseline acidification (Figure 8). Thus, glycolysis contributesat least 80% of the acid production in U-937 GTB cells. A similar resultwas seen in murine hepatoma cells using glucose-free medium where theextracellular acidification rate rapidly decreased to approximately 60% ofbaseline activity [111].

The relationship between an increased acidification rate and CHS828-induced toxicity has not yet been fully elucidated, but there are severalindications that these effects are not causally related. Comparable increasesin the acidification rate were evident also in a subline with >1000-foldresistance to the cytotoxic actions of CHS 828. Furthermore, a series ofclosely related CHS 828 analogues were tested and compared regardingboth influence on the rate of acidification and cytotoxicity. However, no

32


5 101

0

50

100

150

200

20

Addition of drug

↓

Glucose control

Pyruvate control

CHS 828 10 µM

CHS 828 10 µM

Glucose medium Glucose medium

Glucose mediumPyruvate medium

Time (h)

Figure 8. Effect of CHS 828 on the acidification rate in U-937 GTB cells during exposure

to 10 mM glucose medium or medium without glucose but supplemented with 10 mM

pyruvate.

apparent relationship between cytotoxicity and acidification could be dis-cerned (R=0.47). These data, together with the fact that concentrationswell below those inducing acidification are cytotoxic, indicate that acidifi-cation is not necessary for the cytotoxic effect of CHS 828. In addition, inmedium containing pyruvate, where no stimulation of the extracellular aci-dification was seen, CHS 828 still induced cytotoxicity, although to a lesserextent than in glucose medium.

However, it can not be excluded that CHS 828-induced stimulationof acidification rate may contribute to the cytotoxic response. The pro-nounced increase of the acidification rate was also observed in a variety ofdifferent human tumour cell lines (RPMI 8226/S, CCRF-CEM, NCI-H69,H69AR) as well as in tumour cells from patients (ovarian carcinoma, CLLand PBMC), of which all were sensitive to the cytotoxic effect of CHS 828.Apparently, acidification appears to be a general effect of the drug. Withthis reasoning we speculate that mitochondrial inhibition is one of severalbiochemical mechanisms for the cytotoxic action of CHS 828. Indeed,maintained ATP level is an important factor that may determine the modeof cell death in U-937 cells (see “Further aspects of the metabolic effectsinduced by CHS 828 (Paper IV)”).

33

10-4 10-3 10-2 10-1 100 101

0

25

50

75

100

glucose

pyruvate

A

Concentration (µM)

10-3 10-2 10-1 100 101 102

0

25

50

75

100

glucose

pyruvate

Concentration ( µg/ml)

B

10-3 10-2 10-1 100 101 102

0

25

50

75

100

glucose

pyruvate

C

Concentration ( µg/ml)

Figure 9. The effect of CHS 828 (A), MIBG (B) and MGBG (C) on U-937 GTB cell survi-

val in medium containing either 10 mM glucose (�) or 10 mM pyruvate (❏).


The effect of CHS 828 on extracellular acidification and cytotoxicitywas investigated using samples from patients with CLL (n=5-7) and PBMC(n=7-9) (unpublished data). The mean exposure of CLL cells to CHS 828(10 µM) resulted in a rapid increase in the extracellular acidification rate toapproximately 280% of the baseline value, ranging from 200 to 600%(Figure 10). No other cell system has been seen to give such high increases.The corresponding value for PBMC was less than 150%. For CHS 828 at1 µM, the stimulation was 150% and 120% for CLL and PBMC, respecti-vely (not shown). CLL cells were also more sensitive than PBMC to the

34


5 10 15

0

50

100

150

200

250

300

CHS 828 10 µM - CLL

Control

CHS 828 10 µM - PBMC

Time (h)

Figure 10. Effect of 10 µM CHS 828 on the acidification rate in patient CLL cells (n=7)

and PBMC (n=9).

Figure 11. The cytotoxic effect of 10 µM CHS 828 in patient CLL cells (n=5) and PBMC

(n=7).

10-3 10-2 10-1 100 101

0

25

50

75

100 PBMC

CLL

Concentration of CHS 828 (µM)

cytotoxic action of CHS 828 (Figure 11). This is in line with previous fin-dings, demonstrating that CLL is particularly sensitive to CHS 828 [80,82].

MGBG did not stimulate acidification, but rather followed the con-trol curve (Figure 7). The mechanism of action described for MGBG pri-marily involves inhibition of the biosynthesis of the polyamines. It has alsobeen suggested that the cytotoxic effect of MGBG is exerted by the preven-tion of mitochondrial spermine flux [112]. Yet, other reports suggest thatMGBG is also involved in inhibition of mitochondrial respiration and ATPproduction [74, 113, 114]. However, according to Cytosensor® data,MGBG does not seem to involve mitochondrial inhibition since glycolysiswas unaffected. This is in agreement with a previous study demonstratingthat MGBG did not affect glycolysis during the first 24 h of drug exposurein Ehrlich ascites cells [75].

The concentration-response relationship for cytotoxicity of CHS 828resembled that of MGBG, but not that of MIBG (Figure 9). For CHS 828and MGBG, the longer the time of drug exposure the higher the degree ofcell death was observed in a dose-dependent manner, reaching a plateau ofmaximum effect at each exposure time. Also with respect to inhibition ofDNA and protein synthesis, CHS 828 and MGBG demonstrated similarkinetics. The synthesis began to decline after 20 h of drug exposure. Thecourse of events for MIBG was earlier.

MGBG significantly enhanced the activity of the enzymes (SAMDCand ODC) involved in the biosynthesis of the polyamines while CHS 828did not. Addition of spermidine fully reversed the cytotoxic effects ofMGBG, but there was no effect on survival of cells exposed to CHS 828.In conclusion, CHS 828 does not appear to share the MGBG effects onpolyamine metabolism.

Further aspects of the metabolic effects induced by CHS828 (Paper IV)To investigate the cause of the CHS 828-induced acidification, glucose con-sumption and lactate production were measured. For cells exposed to CHS828 there was no increase in glucose consumption measured as glucoselevel in the cell culture medium. However, with an increase in the glycolyticactivity/induced acidification, the consumption of glucose is expected toincrease. Probably, the method used for detection of glucose consumptionwas too insensitive. The excess of glucose (10 mM) in the cell culture medi-um and an already high uptake of glucose might be another explanation ofthe absence of increased glucose consumption. Acid production has beendemonstrated to reach 75% of maximum effect already at a glucose con-centration of 1.25 mM [115]. There was also no enhancement of the cyto-toxic action of CHS 828 when the glucose concentration was doubled(unpublished data). On the other hand, there was an increase in lactateproduction for both MIBG and CHS 828 exposed cells (Figure 12). MIBGhas previously been demonstrated to inhibit mitochondrial respiration and,

35


as a secondary phenomenon, stimulate glycolysis and the production of lac-tic acid [49, 54]. In the light of these results a similar mechanism is plausi-ble also for CHS 828.

An alternative source of extracellular acidification is the PPP, butthis pathway is normally not a major source of ATP [21]. The activity ofthe first and rate-limiting enzyme, G6PD, which connects this pathwaywith glycolysis, was measured in cells exposed to CHS 828. No increase inactivity was found. Thus, the PPP is probably not of crucial importance forthe cytotoxic effect of CHS 828.

Three inhibitors of the major proton translocation mechanisms werecombined with CHS 828 and MIBG. For CHS as well as MIBG, combina-tion with DIDS paradoxically increased the extracellular acidification,while combination with amiloride produced no change in acidification rate.Only CNCn seemed to decrease the CHS 828- and MIBG-induced stimula-tions in acidification rate, compatible with the lactate symport being themajor mediator of outward proton transport [34].

When investigating the temporal effects of CHS 828 and MIBG oncellular ATP levels and cytotoxic effect, there was a delay in both effectsfor CHS 828 compared with MIBG (Figure 13). ATP levels in CHS 828-exposed cells were somewhat reduced at 24 h, but then dropped dramati-cally, which coincided with inhibition of glucose consumption and increa-sed cytotoxic effect. In comparison, the ATP levels in MIBG-exposed cellsdropped earlier and in parallel with the onset of the cytotoxic effect. Thisis in accordance with previous studies where ATP levels in MIBG-exposedcells were reported to decrease early. A more than 50% decrease after only4 h [49] and a ATP to ADP ratio reduced by more than half after 12 h ofMIBG exposure have been observed [50].

Most of the standard anticancer drugs in use today induce apoptosis

36


0 5

90

110

130

20 30 40 50

MIBG 10 µg/ml

CHS 828 10 µM

Time (h)

Figure 12. The effect over time on lactate production in U-937 GTB cells exposed to 10

µM CHS 828 and 10 µg/ml MIBG.

[7]. The complete apoptotic programme is dependent on a certain level ofATP in order to proceed [8]. If ATP levels are too low, necrosis dominates[16]. It has recently been demonstrated that ATP levels can serve as aswitch between apoptosis and necrosis [11, 116]. In other words, thebalance between apoptosis and necrosis depends on the intensity of theinjury and the level of intracellular ATP [13].

The importance of ATP levels for cell death induced by CHS 828was recently described [117]. When combining CHS 828 with 3-aminoben-zamide, which is reported to preserve ATP in several cell systems [118],cells became 100-fold resistant to the cytotoxic effect of CHS 828. Theincrease in glycolytic activity was prolonged and ATP levels maintained foran extended time period. Under these conditions, the cell death pattern tur-ned to more apoptotic features, thus probably facilitated by the preservedenergy levels.

CHS 828 and MIBG seem similar in their initial action, both cau-sing an early effect on mitochondrial respiration with subsequent increase

37

0 10 20 30 40 50 60 70

0

200

400

600

800

1000

1200

1400

ATP

Cytotoxic effect

0

25

50

75

100A

Time (h)

0 10 20 30 40 50 60 70

0

200

400

600

800

1000

1200

1400

Cytotoxic effect

ATP

0

25

50

75

100

B

Time (h)

Figure 13. The levels of ATP and the cytotoxic effects in U-937 GTB cells exposed to 10

µM CHS 828 (A) or 10 µg/ml MIBG (B).


in glycolytic activity and lactate production. However, CHS 828 appears tocause a late inhibition of glycolysis leading to a subsequent sharp drop inATP levels. This ATP depletion is accompanied by inhibited glucose con-sumption, and the previously reported inhibitory effects on DNA and pro-tein synthesis appear with subsequent cell death [64, 119]. This metabolicinsult appears much earlier for MIBG, where ATP levels decline simultane-ously with growth inhibition and cytotoxicity.

Investigation of CHS 828-combinations (Paper V)Amiloride is an inhibitor of the Na+/H+ antiport, one of several mecha-nisms in cells used to regulate pHi [120]. The antiporter responds to a fallin pHe by extruding protons in exchange with extracellular Na+ [30].Exposure to amiloride does not affect cell survival at neutral pHe, but redu-ces cell viability at lower pHe [121]. Since cells exposed to CHS 828 excre-te acid, this might lead to a decrease in pHe. Thereby, the blocking of a pHi

regulating mechanism might possibly enhance the cytotoxic effect of CHS828.

Amiloride alone affected neither cell survival nor acidification rateat the concentrations tested (0.1-10 µM; Figure 14). Synergy was evidentfor all concentrations of amiloride together with CHS 828 0.5 µM accor-ding to the additive interaction model. There was a 0.5 µM decrease in theEC50 value, but the shape of the dose-response curve was similar to that ofCHS 828 alone. The maximum effect of the drug was unaltered and inCytosensor® experiments the response for the combined drugs was similarto that of CHS 828 alone.

A theoretical explanation of the synergistic interaction between CHS

38


Figure 14. The cytotoxic effect in U-937 GTB cells of increasing concentrations of CHS

828 alone or in combination with 0,1 (�), 1 (�) or 10 (�) µM amiloride.

10-2 10-1 100 101 102

0

25

50

75

100

125

CHS 828 alone

+ amiloride 10 µM

+ amiloride 1 µM

+ amiloride 0.1µM

amiloride alone

CHS 828 concentration (µM)

828 and amiloride might be the blocking of the Na+/H+ channels [122].The block could possibly lead to a decrease in pHi with a subsequent decre-ase in cell viability. However, this scenario is not likely since no effect ofthe combination on pHi was observed. In other cell systems, amiloride hasbeen reported to affect acidification rate, but higher concentrations (0.5-1mM) were used [111, 115]. A more probable explanation of the synergyobserved is that amiloride potentiates the effect of CHS 828 by increasingthe cellular uptake of the drug. This effect of amiloride has been shown forother drugs, either secondary to changes in pHi or by a pHi independentmechanism [123, 124].

To study the interaction between CHS 828 and MMC, the isobolemethod was used. Five combinations for effects in the range of 30-70%were chosen to cover the steep part of the dose-response curves where aninteraction is most likely to be detected. MMC is a well-known bioreducti-ve drug that requires enzymatic activation, a process favoured by low pH,generating an enhanced cytotoxic activity of MMC [125]. The interactionsfor the CHS 828/MMC combinations were mostly additive but synergisticat the highest effect level (Figure 15). In addition, the additive model wasused on the same set of data and a fairly good agreement between the twomethods was shown in a previous study [126] as well as in this report.

An enhanced antitumour effect of MMC in vivo was demonstratedwhen the drug was administered under hyperglycaemic conditions thatwould lower pHe and create a favourable environment for bioreduction ofMMC by reduction of pHi [125]. CHS 828 is known to dose-dependentlystimulate extracellular acidification (paper II), which hypothetically wouldincrease MMC bioreduction via changes in pHi. This scenario was compa-tible with the concentration-dependent tendency towards synergy betweenCHS 828 and MMC. However, the increase in extracellular acidificationinduced by CHS 828 was not enough to affect pH inside the cells (notshown). Furthermore, with only a minor increase in acidification, thiseffect might be abolished by the buffering capacity of the medium. Thus,the pattern of interaction observed between CHS 828 and MMC might bepH independent. Also for other cytotoxic drugs, higher effect levels seem toproduce synergy [127].

Tumour cells from patients (n=78) were tested for the individual andcombined effects of 1 µM CHS 828 and 2.5 µg/ml MMC. The additivemodel was used since there were no concentration-response curves for thedrugs. The haematological tumour samples (n=48) were generally moresensitive than the solid tumour samples (n=30), with 10% of the samplesshowing synergistic interactions and 6% antagonistic, while the figures forsolid samples were 7% synergistic and 27% antagonistic interactions.

The combinations of CHS 828 and seven standard anticancer agentswere investigated using both the isobole method and the additive model inU-937 GTB cells (unpublished data) (Figure 15). The methods correlatedreasonably well, except in the case of Pac, where the additive model indica-ted that the response of the cells to the combination was higher than that

39


of the isobole model. The combinations of CHS 828 with VP16 or Melphclearly demonstrated synergistic interactions at all effect levels tested, whe-reas, for the other drugs, the interaction patterns were more inhomogene-ous but generally closer to additive effects.

40


CHS 828 + MMC

10 20 30 40 50 60 70 80 90

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

CHS 828 + AraC

40 50 60 70 80 90

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

CHS 828 + Dox

10 20 30 40 50 60 70 80 90

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

CHS 828 + VP16

75 76 77 78 79 80

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

CHS 828 + Pac

75 80 85 90 95

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

CHS 828 + Topo

30 40 50 60 70 80 90

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

2.0

CI

SI ratio

Effect level (%)

CHS 828 + Melph

60 70 80

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

CHS 828 + CisP

0 10 20 30 40 50 60 70 80 90

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

CI

SI ratio

Effect level (%)

Figure 15. Types of interactions between CHS 828 and eight standard cytotoxic drugs,

at five different levels of effect in U-937 GTB cells according to the isobole method and

the additive model.

Combinations between standard drugs and CHS 828 in tumour sam-ples from patients (unpublished data)In a preliminary study, single drugs and combinations of CHS 828 andseven standard cytotoxic agents were tested in a total of 129 patient sam-ples; 76 from haematological and 53 from solid malignancies. The domina-ting cell types among the haematological tumour samples were AML(n=37) and ALL (n=23), whereas for the solid tumour samples ovarian car-cinoma (n=25) was the most common tumour type.

Compared with solid tumour samples, the interactions classified assynergistic according to the additive model were essentially similar butfewer were antagonistic in haematological tumour samples (Table 2). Thecombinations most frequently producing synergy were with Melph (56%),VP16 (37%) and Dox (34%). The combination producing fewest synergis-tic interactions was with Topo (5%), which also had the highest frequencyof antagonistic interactions (10%). The combinations of CHS 828 withMelph or CisP did not produce any antagonistic interactions at all. In gene-ral, the haematological tumour samples were more sensitive to the combi-nations than the solid tumour samples. Although there were synergisticinteractions for all combinations investigated in the solid tumour samples,they were counterbalanced by the high frequency of antagonistic interac-tions. All combinations in the solid samples produced antagonistic interac-tions with the lowest frequencies for Melph (13%) and Dox (9%). Thesecombinations also produced most synergistic interactions with 56% forMelph and 45% for Dox.

More patient samples are being continuously analysed and a defini-tive analysis will be performed in the near future. Whether this is a validapproach for further clinical development of CHS 828 can only be determi-ned by testing of the combinations in vivo.

41

Drug combinations

Synergistic%

Antagonistic%

34

56

5

25

21

21

37

1

-

10

6

-

6

2

HAEMATOLOGICAL

Synergistic%

Antagonistic%

SOLID

45

56

18

20

21

17

22

9

13

16

20

17

15

12

CHS 828 1 µM+ Dox 0.5 µg/ml

+ Melph 2.5 µg/ml

+ Topo 2.5 µg/ml

+ MMC 2.5 µg/ml

+ CisP 2.5 µg/ml

+ AraC 0.5 µg/ml

+ VP16 5.0 µg/ml

Table 2. Types of interactions for the combination of CHS 828 and eight standard cyto-

toxic agents in a variety of 76 haematological and 53 solid tumour samples as analyzed

by the additive model


Overall discussion and concluding remarks

The Cytosensor®

A wide variety of receptor stimulations trigger metabolic changes and theCytosensor® microphysiometer has mostly been used for studying receptorpharmacology [95, 96]. Microphysiometry has also been used to detectbacterial antibiotic sensitivity and to discriminate between bacteriostaticand bacteriocidal concentrations [128]. Recently, tumour specimens derivedfrom surgery were placed on a micro-sensor chip and reading of metabolicactivity was performed [129]. Other conceivable areas for the use of theCytosensor® technique are, e.g., in screening of novel chemotherapeuticagents, testing of drug combinations and when investigating the mecha-nisms of action for current and new drugs.

A unique feature of the Cytosensor® is the possibility of real-timemonitoring of cellular metabolism, which enables quantitative kinetic ana-lysis and reversibility studies. A disadvantage is that the Cytosensor® is arelatively new technique and there are still fairly few reports in the literatu-re on the use of the Cytosensor® in research of cytotoxic drugs. In otherwords, a lot of work remains to be done, where, e.g., Cytosensor® experi-ments are combined with parallel investigations on more established end-points such as induction of cell death and inhibition of tumour growth.

In clinical oncology, the positron emission tomography (PET) tech-nology for measurement of the uptake of 18F-fluoro-2-deoxy-D-glucose, hasrecently been proposed to provide an early prediction of the clinical effectof chemotherapy [130]. It might be speculated that the Cytosensor®, orsimilar technology for measurement of early metabolic effects of cytotoxicdrugs, could become a parallel to PET at the in vitro level. Thus, it mightprovide early and relevant data on the effect of various cytotoxic drugs oreven antibodies and biological response modifiers, that would simplify andoptimise the development process in this field.

In the papers included in this thesis, most experiments were perfor-med using the U-937 GTB lymphoma cell line. This cell line was establis-hed and characterised in Uppsala in 1976 [93]. The U-937 cell line waschosen because of its sensitivity against most standard anticancer agentsand its widespread use in cancer research. Another advantage with U-937is the easy handling of the cells as well as the convenient 24h regrowth-cycle.

Retrospectively, it could be argued that the use of suspension cellswas perhaps not the optimal choice. In Cytosensor® experiments, adherentcells can be grown directly on the membrane of the capsule cup. In the caseof non-adherent cells, like U-937, cells are immobilised in the cup usingagarose. However, the use of agarose does not seem to be optimal formaintaining cell viability, since the control curves tended to decrease to60% of initial baseline values over 20 h experiments.

42


CHS 828Most of the current anticancer drugs kill cancer cells by apoptosis [7]. CHS828 is partly an exception since the features of cell death induced by thisdrug include both apoptosis and necrosis [119, 131, 132]. Cells exposed toCHS 828 showed a decline in glycolysis at approximately 15 h but wereunaffected in growth and viability after 24 h or more, but with subsequentdecrease in ATP levels, glucose consumption and decreased lactate produc-tion (paper IV). These events were associated with a sharp inhibition ofDNA and protein synthesis and subsequent cell death (paper II) [119]. Inelectron microscopy after 24 h the ultrastructure was indistinguishablefrom that of untreated control, but it was affected after 48–64 h of exposu-re [119]. A significant hyperpolarisation of the mitochondrial membranepotential was evident after 24 h, and there were no major increases in cas-pase-8 or –9 activities. Only caspase-3 exhibited a modest increase at 48 h,which could not be blocked by caspase inhibition, indicating atypical featu-res of apoptosis [132]. Similarly, when investigating the cytotoxic effects inthe breast cancer cell line MCF-7, DNA fragmentation was induced, butother markers of typical classic apoptosis were lacking [131]. Derangedenergy metabolism was suggested as one possible explanation of the absen-ce of late morphological signs of apoptosis of the cells [131].

A similar course of events as seen for CHS 828 has also beendemonstrated in murine neocortical cell cultures exposed to a-monochloro-hydrin [133]. There was a delayed cell death with decreased ATP levelsfirst after 36 h of continuous exposure, followed by progressive cell death.This was described to be a consequence of inhibition of the key upstreamglycolytic enzymes glyceraldehyde-3-phosphate dehydrogenase and triosep-hosphate isomerase. These enzymes have not been measured in this thesis,but the scenario described could possibly serve as an explanation also ofthe effects induced by CHS 828. This has to be further explored.

Not much is known about the effects of other cytotoxic drugs oncellular metabolism, and data are sometimes contradictory and inconclusi-ve. When investigating the effect of CHS 828 on cellular ATP levels overtime, the results were only compared with those of MIBG. However, theeffects of seven standard cytotoxic drugs on DNA and protein synthesishave recently been investigated in U-937 cells [134]. These are energy requ-iring processes and thus mirror the energy state of drug-exposed cells [25].For Dox, VP16, Topo and AraC, the DNA synthesis was shut off alreadyafter 4 h, while for CisP, Vcr and Pac, the curves diverged from that of thecontrol after 10 h of drug exposure [134]. This was also the case withrespect to protein synthesis. In Cytosensor® experiments, CisP and Pac donot decrease acidification below baseline within the first 20 h of drugexposure (paper II). In addition, cell survival for these drugs after 24 hdrug of exposure, was high. These findings indicate that the onset of thecytotoxic effect for most investigated standard drugs is earlier than obser-ved for CHS 828, which might indicate the possibility of a new mechanismof action for this drug.

43


The effect of CHS 828 on the mitochondria was indirectly demon-strated by comparison with the more investigated and structurally relatedmitochondrial inhibitor, MIBG. In analogy with MIBG, CHS 828 producedan increased extracellular acidification rate. This increase was shown tooriginate from an increased glycolytic activity since it could be completelyabolished by the use of pyruvate to block the glycolysis (paper II). The ideaof a CHS 828-induced mitochondrial block is further supported by theCHS 828 induced lower acid production compared to control in glucose-free pyruvate-supplemented medium (Figure 8). Furthermore, the F1F0

ATPase inhibitor Oligomycin B also induced an increased acidification rate,similar to those of CHS 828 and MIBG (unpublished data). Also, wheninvestigating CHS 828-exposed cells with electron microscopy, the mito-chondria appeared swollen and affected by the drug [119]. These data sup-port the idea of the mitochondria as a target for CHS 828 cytotoxicity.

Despite similarities between CHS 828 and MIBG in acidificationrates and other properties, the cytotoxic onset of the drugs differed asMIBG affected cell viability early and CHS 828 later. The effects over timeof CHS 828 based on observations in this thesis are speculatively outlinedin figure 16. Unfortunately, MIBG were not included in all CHS 828 expe-riments, e.g., on glucose consumption and morphology, making it difficultto perform a complete comparison of the two drugs.

The pre-clinical data on antitumour effect of CHS 828 are indeedpromising but there still remains a huge step to the cancer treatment ofhumans. The present studies of the metabolic events following exposure toCHS 828 shed some light on what precedes the cytotoxic actions of thedrug and thereby provide clues to its mechanism of action. However, meta-bolic studies have to be connected to, and strengthened by, results fromstandard methods before the importance of the information obtained fromobservations of early metabolic changes can be assessed.

44


45

CHS 828Cells

Mitochondrial block

Increased glycolytic activity

&

Increased lactate production

Instant increase in extracellular

acidification rate as measured

in the Cytosensor

Increased glycolysis tocompensate for impairedenergy production

Decreased glycolytic activityThe acidification rate for CHS

828 falls below the curve of

untreated control

Late block in glycolytic activity

Decrease in DNA synthesis

&

protein synthesis

Decrease in [14C]Thymidine &

[14C]Leucine incorporation

No energy left to proceed withthese energy-requiringprocesses

Decreased cell viability Measurement of cytotoxicity in the

FMCA

0 h

0-15 h

15 h

24-30 h

30- h

Exposure

24-30 h

Decrease in glucose

concentration in culture medium

& decrease in cellular ATP

Glycolysis is turned offNo ATP production due to blockin both glycolysis andmitochondria

Observation - interpretation

Decreased ATP levels

Glucose consumption turned off

&

Secondary to increased glycolysis

Increase in acidification rate over

control & increase in lactate

production

Figure 16. Outline of observed and putative effects of CHS 828 in U-937 GTB cells over

time.


46


Conclusions

The main conclusions of this thesis are, in brief:

• The new technology, the Cytosensor® microphysiometer, which detectsextracellular acidification as a measure of cellular metabolic activity, wasfound to be a feasible and potentially valuable complement to conventio-nal methods used in the development of anticancer agents. Continuous“on-line” measurements of cellular metabolism may add valuable know-ledge for optimal development and use of anticancer drugs.

• A number of standard cytotoxic drugs produced typical and reproducibleacidification response patterns in the Cytosensor®. The relevance of thesefindings remains to be determined.

• The novel cyanoguanidine CHS 828 was found to exert a pronouncedand prolonged increase in extracellular acidification similar to that of thestructurally related mitochondrial inhibitor MIBG. The results indicatethat the acid produced originated from increased glycolytic flux andincreased lactate production, effects that may be secondary to the block-ing of oxidative phosphorylation in the mitochondria.

• The CHS 828-induced acidification was observed in several cell types,including cells from various tumour types from patients.

• The cellular effects induced by CHS 828 were not found to resemblethose of the structurally related polyamine synthesis inhibitor MGBG.CHS 828 did not affect the key enzymes involved in the synthesis of poly-amines.

• The changes in energy metabolism after exposure to CHS 828 seem to berelated to its cytotoxic effect. The relevance of the early acidification isnot clear, but the delayed cell death is associated with inhibition of gluco-se utilisation and a drop in ATP levels.

• In combinatory studies, a synergistic interaction was demonstrated bet-ween CHS 828 and the non-toxic drug amiloride, known to inhibit onemechanism for the regulation of pHi. Furthermore, additive-synergisticeffects were seen between CHS 828 and the bioreductive drug MMC.The mechanisms for these interactions are not clear, but seem indepen-dent of the extracellular acidification induced by CHS 828.

• In the U-937 cell line, as well as in tumour samples from patients, CHS828 demonstrated synergistic interactions in combination with bothMelph and VP16. Thus, Melph and VP16 might be interesting for combi-nation with CHS 828 in future clinical trials.

47

Conclusions

Acknowledgements

The present studies were performed at Uppsala University, at the Instituteof Medical Sciences, Department of Clinical Pharmacology, during theyears 1997-2001. In writing this thesis I have had help from others in different ways, bothscientifically and not so scientifically. Therefore, I would like to express mysincere gratitude to many people…

My tutors, Peter Nygren and Rolf Larsson, for giving me the opportunityto work together with you and for sharing your great scientific knowledge.Peter, for your reliability, analytical skills and ability to ask the right ques-tions. Rolf, for all your splendid thoughts, ideas and enthusiasm.

Co-authors, for help with manuscripts and discussions about results andscience in general. Åke Sjöholm, for helping me out in the lab and for theinterest you have shown in my work. Leo Pharmaceutical Products inBallerup, Denmark, for their support and collaboration concerning CHS828.

All the PhDs and post-graduate students at the lab, Gunnar and Jocke forsharing your knowledge and thoughts with me. Petra for co-writing andElin, Henrik, Sumeer, Sadia, Anna and Britt-Marie for making my time atthe department challenging and exciting.

The lab-girls for their skilful work with patient samples, Kicki and Maria,Carina my sunny room-mate and Lena for great help with the Cytosensor.Annika, for valuable administrative help at the department.

My degree project students Pia, Ingrid and Emica, for helping me with datacollection, inspiration and good company at the lab. Marie Sandström, forsharing two degree project students with me and for introducing me to thedifficulties of combinatory studies.

The Swedish Cancer Foundation and the Lions Cancer Foundation fortheir financial support in general. The Swedish Academy of PharmaceuticalSciences, the IF Foundation for Pharmaceutical Research, the RektorFoundation and the Bristol-Myers Squibb travel scholarship for their speci-fic financial support, which made it possible for me to attend scientificmeetings in Europe and the United States.

All my friends (including those not mentioned below)! Lena, for always being there for discussions, listening, encouraging, gossi-ping and spinning. Pia, Markus and Victor for friendship and trips to themountains. Annika, my third-generation great friend. The Vikingstad gang,Nicke, Ulle, Danne and Limpan, from Mulle/kindergarten and onward.

48


Lise-Lotte, Jocke and Amanda for tenta studying, gossiping and good coo-king. Mia, Malin and Lena for all our girl dinners. The wine-tasting gangfor wine sensations.

Mormor and my late morfar for always showing sincere interest in me andfollowing me on my path through life. And all the rest of the family, whomean a lot to me.

Mamma, pappa and Micke – the best you can get! Words can not express what you mean to me… you are always there forme.

Fredrik - my darling sambo and best friend! For all your love, support and encouragement.

Ph.inally D.one

49

Acknowledgements

References

1.DeVita JDT, Hellman S and Rosenberg SA, Pharmacology of cancerchemotherapy. In: Cancer - principles and practice of oncology, Vol. 1(Eds. DeVita JDT, Hellman S and Rosenberg SA), pp. 375-514.Lippincott-Raven, Philadelphia, 1997.

2.FASS - The Swedish Drug Compendium, Stockholm, 2001.3.Nygren P, What is cancer chemotherapy? Acta Oncol 40: 166-74, 2001.4.Fan W, Johnson KR and Miller III MC, In vitro evaluation of combi-

nation chemotherapy against human tumor cells. Oncol Rep 5: 1035-1042, 1998.

5.Chabner BA, Allegra CJ, Curt GA and Calabresi P, Antineoplasticagents. In: The pharmacological basis of therapeutics (Eds. HardmanJG, Limbird LE, Molinoff PB, Ruddon RW and Goodman Gilman A),pp. 1233-1287. McGraw-Hill, New York, 1996.

6.Erlichman C, Pharmacology of anticancer drugs. In: The basic scienceof oncology (Eds. Tannock IF and Hill RP), pp. 317-337. McGraw-Hill, New York, 1992.

7.Hickman JA, Apoptosis induced by anticancer drugs. Cancer andMetastasis Reviews 11: 121-139, 1992.

8.McConkey DJ, Biochemical determinants of apoptosis and necrosis.Toxicol Lett 99: 157-168, 1998.

9.Leist M, Gantner F, Bohlinger I, Tiegs G, Germann P and Wendel A,Tumor necrosis factor-induced hepatocyte apoptosis precedes liver fai-lure in experimental murine shock models. Am J Pathol 146: 1220-34,1995.

10.Kerr JFR, Wyllie AH and Currie AR, Apoptosis: a basic biologicalphenomenon with wide-ranging implications in tissue kinetics. Br JCancer 26: 239-257, 1972.

11.Leist M and Nicotera P, The shape of cell death. Biochem Biophys ResCommun 236: 1-9, 1997.

12.Nicotera P, Leist M and Ferrando-May E, Intracellular ATP, a switchin the decision between apoptosis and necrosis. Toxicol Lett 102-103:139-142, 1998.

13.Renvoizé C, Biola A, Pallardy M and Bréard J, Apoptosis: identifica-tion of dying cells. Cell Biol Toxicol 14: 111-120, 1998.

14.Leist M, Single B, Naumann H, Fava E, Simon B, Kuhnle S andNicotera P, Inhibition of mitochondrial ATP generation by nitric oxideswitches apoptosis to necrosis. Exp Cell Res 249: 396-403, 1999.

15.Lelli JL, Becks LL, Dabrowska MI and Hinshaw DB, ATP convertsnecrosis to apoptosis in oxidant-injured endothelial cells. Free RadicalBiology & Medicine 25: 694-702, 1998.

16.Ankarcrona M, Dypbukt JM, Bonfoco E, Zhivotovsky B, Orrenius S,Lipton SA and Nicotera P, Glutamate-induced neuronal death: a suc-cession of necrosis or apoptosis depending on mitochondrial function.Neuron 15: 961-973, 1995.

50


17.Martin DS, Spriggs D and Koutcher AJ, A concomittant ATP-deple-ting strategy markedly enhances anticancer agent activity. Apoptosis 6:125-131, 2001.

18.Warburg O, On the origin of cancer cells. Science 123: 309-314,1956.

19.Aisenberg AC, Part I: Anaerobic and aerobic glycolysis of normal andtumor tissues. In: The glycolysis and respiration of tumors (Ed.Aisenberg AC), pp. 1-54. Academic press, New York, 1961.

20.Baggetto LG, Deviant energetic metabolism of glycolytic cancer cells.Biochimie 74: 959-974, 1992.

21.Owicki JC and Parce WJ, Biosensors based on the energy metabolismof living cells: The physical chemistry and cell biology of extracellularacidification. Biosensors & Bioelectronics 7: 255-272, 1992.

22.Macbeth RLA and Bekesi JG, Oxygen consumption and anaerobicglycolysis of human malignant and normal tissue. Cancer Res 22: 244-248, 1962.

23.Board M, Humm S and Newsholme EA, Maximum activities of keyenzymes of glycolysis, glutaminolysis, pentose phosphate pathway andtricarboxylic acid cycle in normal, neoplastic and suppressed cells.Biochem J 265: 503-509, 1990.

24.Argilés JM and López-Soriano FJ, Why do cancer cells have such highglycolytic rate? Med Hypotheses 32: 151-155, 1990.

25.Alberts B, Bray D, Lewis J, Raff M, Roberts K and Watson JD,Molecular biology of the cell. Garland publishing, New York, 1994.

26.Stryer L, Pentose phosphate pathway and gluconeogenesis. In:Biochemistry, pp. 427-448. W H Freeman and company, New York,1988.

27.Vaupel P, Kallinowski F and Okunieff P, Blood flow, oxygen andnutrient supply, and metabolic microenvironment of human tumors: Areview. Cancer Res 49: 6449-6465, 1989.

28.Ashby BS, pH studies in human malignant tumours. Lancet 2: 312-315, 1966.

29.Wike-Hooley JL, Haveman J and Reinhold HS, The relevance oftumour pH to the treatment of malignant disease. Radiother Oncol 2:343-366, 1984.

30.Madshus IH, Regulation of intracellular pH in eukaryotic cells.Biochem J 250: 1-8, 1988.

31.Tannock IF and Rotin D, Acid pH in tumors and its potential for the-rapeutic exploitation. Cancer Res 49: 4373-4384, 1989.

32.Frelin C, Vigne P, Ladoux A and Lazdunski M, The regulation of theintracellular pH in cells from vertebrates. Eur J Biochem 174: 3-14,1988.

33.Ladoux A, Miglierina R, Krawice I, Cragoe EJ, Abita JP and Frelin C,Single-cell analysis of the intracellular pH and its regulation during themonocytic differentiation of U937 human leukemic cells. Eur JBiochem 175 (3): 455-460, 1988.

51

References

34.Poole RC and Halestrap AP, Transport of lactate and other monocar-boxylates across mammalian plasma membranes. Am J Physiol 264:C761-C782, 1993.

35.Arese P and Bosia A, Drugs affecting glycolysis. In: Fundamentals ofcell pharmacology (Ed. Dikstein S), pp. 102-. Charles C Thomas,Springfield, 1973.

36.Bossa R, Galatulas I, Leotta L and Tofanetti O, Effects on variousanti-tumor drugs on energetic processes. Pharmacol Res Commun 8:369-77, 1976.

37.Gosalvez M, Blanco M, Hunter J, Miko M and Chance B, Effects ofanticancer agents on the respiration of isolated mitochondria andtumor cells. Eur J Cancer 10: 567-74, 1974.

38.Neeman M, Eldar H, Rushkin E and Degani H, Chemotherapy-indu-ced changes in the energetics of human breast cancer cells; 31P- and13C-NMR studies. Biochim Biophys Acta 1052: 255-263, 1990.

39.Kruidering M, van de Water B, de Heer E, Mulder GJ and NagelkerkeJF, Cisplatin-induced nephrotoxicity in porcine proximal tubular cells:mitochondrial dysfunction by inhibition of complexes I to IV of therespiratory chain. JPET 280: 638-649, 1997.

40.Melendez-Zajgla J, Cruz E, Maldonado V and Espinoza AM,Mitochondrial changes during the apoptotic process of HeLa cellsexposed to cisplatin. Biochem Mol Biol Int 47: 765-71, 1999.

41. Manzano A, Roig T, Bermundez J and Bartrons R, Effects of Taxol onisolated rat hepatocyte metabolism. Am J Physiol 271: C1957-62, 1996.

42.Glass-Marmor L and Beitner R, Taxol (paclitaxcel) induces a detach-ment of phosphofructokinase from cytoskeleton of melanoma cellsand decreases the levels of glucose 1,6-bisphosphate, fructose 1,6-bis-phosphate and ATP. Eur J Pharmacol 370: 195-199, 1999.

43. Ronen SM, DiStefano F, McCoy CL, Robertson D, Smith TAD, Al-Saffar NM, Titley J, Cunningham DC, Griffiths JR, Leach MO andClarke PA, Magnetic resonance detects metabolic changes associatedwith chemotherapy-induced apoptosis. Br J Cancer 80: 1035-41, 1999.

44.Lloyd PG and Hardin CD, Role of microtubules in the regulation ofmetabolism in isolated cerebral microvessels. Am J Physiol 277:C1250-62, 1999.

45.Fanciulli M, Bruno T, Castiglione S, Del Carlo C, Paggi MG andFloridi A, Glucose metabolism in adriamycin-sensitive and -resistantLoVo human colon carcinoma cells. Oncol Res 5: 357-62, 1993.

46.Fanciulli M, Bruno T, Giovannelli A, Gentile FP, Padova M, Rubiu Oand Floridi A, Energy metabolism of human LoVo colon carcinomacells: correlation to drug resistance and influence of lonidamine. ClinCancer Res 6: 1590-97, 2000.

47.Kaplan O, Navon G, Lyon RC, Faustino PJ, Starka EJ and Cohen JS,Effects of 2-deoxyglucose on drug-sensitive and drug-resistant humanbreast cancer cells: toxicity and magnetic resonance spectroscopy stu-dies of metabolism. Cancer Res 50: 544-51, 1990.

52


48.Miccadei S, Fanciulli M, Bruno T, Paggi MG and Floridi A, Energymetabolism of adriamycin-sensitive and -resistant Ehrlich ascitestumor cells. Oncol Res 8: 27-35, 1996.

49.Loesberg C, Van Rooij H, Nooijen WJ, Meijer AJ and Smets LA,Impaired mitochondrial respiration and stimulated glycolysis by m-iodobenzylguanidine (MIBG). Int J Cancer 46: 276-281, 1990.

50.Cornelissen J, Wanders RJ, Van den Bogert C, Van Kuilenburg AB,Elzinga L, Voute PA and Van Gennip AH, Meta-iodobenzylguanidine(MIBG) inhibits malate and succinate driven mitochondrial ATP syn-thesis in the human neuroblastoma cell line SK-N-BE(2c). Eur JCancer 31A(4): 582-586, 1995.

51.Cornelissen J, Wanders RJA, van Gennip AH, van den Bogert C,Voute PA and van Kuilenburg ABP, Meta-iodobenzylguanidine inhibitscomplex I and III of the respiratory chain in the human cell line Molt-4. Biochem Pharmacol 49: 471-477, 1995.

52. Jänne J, Alhonen L and Leinonen P, Polyamines: from molecular bio-logy to clinical applications. Ann Med 23: 241-259, 1991.

53.Pegg AE and McCann PP, Polyamine metabolism and function. Am JPhysiol 243: C212-C221, 1982.

54.Loesberg C, Van Rooij H, Romijn JC and Smets LA, Mitochondrialeffects of the guanidino group-containing cytostatic drugs, m-iodoben-zylguanidine and methylglyoxal bis (guanylhydrazone). BiochemPharmacol 42: 793-798, 1991.

55.Troncone L and Rufini V, 131I-MIBG therapy of neural crest tumors.Anticancer Res 17: 1823-1832, 1997.

56.Von Hoff DD, MGBG: teaching an old drug new tricks. Ann Oncol 5:487-493, 1994.

57.Wieland DM, Wu J-I, Brown LE, Mangner TJ, Swanson DP andBeierwaltes WH, Radiolabeled adrenergic neuron-blocking agents:adrenomedullary imaging with [131I]iodobenzylguanidine. J Nucl Med21: 349-353, 1980.

58.Smets LA, Bout B and Wisse J, Cytotoxic and antitumor effects of thenorepinephrine analogue meta-iodo-benzylguanidine (MIBG). CancerChem Pharmacol 21 (1): 9-13, 1988.

59.Petersen HJ, Kaegaard Nielsen C and Arrigoni-Martelli E, Synthesisand hypotensive activity of N-alkyl-N´´-cyano-N´pyridylguanidines. JMed Chem 21: 773-781, 1978.

60.Hoefnagel CA, Voute PA, de Kraker J and Marcuse HR, Radionuclidediagnoses and therapy of neural crest tumors using iodine-131 metaio-dobenzylguanidine. J Nucl Med 28: 308-314, 1987.

61.Kuin A, Aalders M, Lamfers M, van Zuidam DJ, Essers M, H BJ andSmets LA, Potentiation of anti-cancer drug activity at low intratumo-ral pH induced by the mitochondrial inhibitor m-iodobenzylguanidine(MIBG) and its analogue benzylguanidine (BG). Br J Cancer 79: 793-801, 1999.

62.Kuin A, Smets L, Volk T, Paans A, Adams G, Atema A, Jähde E, Maas

53

References

A, Rajewsky MF and Visser G, Reduction of intratumoral pH by themitochondrial inhibitor m-iodobenzylguanidine and moderate hyper-glycemia. Cancer Res 54: 3785-3792, 1994.

63. Jähde E, Volk T, Atema A, Smets L, Glüsenkamp K-H and RajewskyMF, pH in human tumor xenografts and transplanted rat tumors:effect of insulin, inorganic phosphate and m-iodobenzylguanidine.Cancer Res 52: 6209-6215, 1992.

64.Ekelund S, Nygren P and Larsson R, Guanidino-containing drugs incancer chemotherapy: biochemical and clinical pharmacology.Biochem Pharmacol 61: 1183-1193, 2001.

65.Hatefi Y, The mitochondrial electron transport and oxidative phos-phorylation system. Ann Rev Biochem 54: 1015-1069, 1985.

66.Pegg AE, Polyamine metabolism and its importance in neoplasticgrowth and as a target for chemotherapy. Cancer Res 48: 759-774,1988.

67.Warrell JRP and Burchenal JH, Methylglyoxal-bis(guanylhydrazone)(methyl-GAG): current status and future prospects. J Clin Oncol 1:52-65, 1983.

68. Jänne J, Pösö H and Raina A, Polyamines in rapid growth and cancer.Biochem Biophys Acta 473: 241-293, 1978.

69.Freedlander BL and French FA, Carcinostatic action of polycarbonylcompounds and their derivatives. II Glyoxal bis(guanylhydrazone) andderivatives. Cancer Res 18: 360-363, 1958.

70.Marsoni S, Hoth D, Simon R, Leyland-Jones B, De Rosa M andWittes RE, Clinical drug development: An analysis of phase II trials,1970-1985. Cancer Treatment Reports 71: 71-80, 1987.

71.Svensson F, Kockum I and Persson L, Diethylglyoxal bis(guanylhydra-zone), a potent inhibitor of mammalian S-adenosylmethionine decar-boxylase. Effects on cell proliferation and polyamine metabolism inL1210 leukemia cells. Mol Cell Biochem 124: 141-147, 1993.

72.Snyder RD and Bhatt S, Alterations in repair of alkylating agent-indu-ced DNA damage in polyamine-depleted human cells. Cancer Lett 72:83-90, 1993.

73.Higaki I, Matsui-Yuasa I, Terakura M, Kinoshita H and Otani S,Increased spermidine levels is essential for hepatocyte growth factor-induced DNA synthesis in cultured rat hepatocytes. Gastroenterology106: 1024-1031, 1994.

74.Mikles-Robertson F, Feuerstein B, Dave C and Porter CW, The genera-lity of methylglyoxal bis(guanylhydrazone)-induced mitochondrialdamage and the dependence of this effect on cell proliferation. CancerRes 39: 1919-1926, 1979.

75.Alhonen-Hongisto L, Pösö H and Jänne J, Inhibition by derivatives ofdiguanidines of cell proliferation in Ehrlich ascites cells grown in cul-ture. Biochem J 188: 491-501, 1980.

76.Diwan JJ, Yune HH, Bawa R, Haley T and Mannella CA, Enhanceduptake of spermidine and methylglyoxal-bis(guanylhydrazone) by rat

54


liver mitochondria following outer membrane lysis. BiochemPharmacol 37: 957-961, 1988.

77.Porter CW, Dave C and Mihich E, Inhibition of S-adenosylmethioninedecarboxylase as an approach in cancer therapeutics. In: Polyaminesin biology and medicine (Eds. Morris DR and Marton LJ), pp. 407-436. Marcel Dekker, New York, 1981.

78.Schou C, Ottosen ER, Petersen HJ, Björkling F, Latini S, Hjarnaa PV,Bramm E and Binderup L, Novel cyanoguanidines with potent oralantitumour activity. Bioorganic & Medical Chemistry Letters 7: 3095-3100, 1997.

79.Vig Hjarnaa P-J, Dhar S, Latini S, Jonsson E, Larsson R, Bramm E,Skov T and Binderup L, CHS 828, novel pyridyl cyanoguanidine withpotent antitumor activity in vitro and in vivo. Cancer Res 59: 5751-5757, 1999.

80.Åleskog A, Bashir-Hassan S, Hovstadius P, Kristensen J, Höglund M,Tholander B, Binderup L, Larsson R and Jonsson E, Activity of CHS828 in primary cultures of human hematological and solid tumors invitro. In press Anti-Cancer Drugs:, 2001.

81.Hassan SB, Jonsson E, Larsson R and Karlsson MO, Model for time-dependency of the cytotoxic effect of CHS 828 in vitro suggests twodifferent mechanisms of action. Manuscript, 2001.

82.Hassan SB, de la Torre M, Nygren P, Karlsson MO, Larsson R andJonsson E, A hollow fiber model for in vitro studies of cytotoxic com-pounds: activity of the cyanoguanidine CHS 828. Anticancer Drugs12: 33-42, 2001.

83.Weinstein JN, Myers TG, O´Connor PM, Friend SH, Fornace Jr AJ,Kohn KW, Fojo T, Bates SE, Rubenstein LV, Anderson NL,Buolamwini JK, van Osdol WW, Monks AP, Scudiero DA, SausvilleEA, Zaharevitz DW, Bunow B, Viswanadhan VN, Johnson GS, WittesRE and Paull KD, An information-intensive approach to the molecularpharmacology of cancer. Science 275: 343-349, 1997.

84.Boyd MR, The NCI anticancer drug discovery screen. In: Anticancerdrug development guide (Ed. Teicher BA), pp. 23-42. Humana Press,Totowa, 1997.

85.Schacter LP, Anderson C, Canetta RM, Kelly S, Nicaise C, Onetto N,Rozencweig M, Smaldone L and Winograd B, Drug discovery anddevelopment in the pharmaceutical industry. Seminars in Oncology19: 613-621, 1992.

86.Druker BJ, Talpaz M, Resta DJ, Peng B, Buchdunger E, Ford JM,Lydon NB, Kantarjian H, Capdeville D, Ohno-Jones S and SawyersCL, Efficacy and safety of a specific inhibitor of the bdr-abl tyrosinekinase in chronic myeloid leukemia. N Eng J Med 344: 1031-37,2001.

87.Committee for proprietary medicinal products (CPMP), Note for gui-dance on the pre-clinical evaluation of anticancer medicinal products.CPMP/SWP/997/96: 1-5, 1998.

55

References

88.Venkatesh S and Lipper RA, Role of the development scientist in com-pound lead selection and optimization. J Pharm Sci 89: 145-54, 2000.

89.Nygren P, Fridborg H, Csoka K, Sundström C, De La Torre M,Kristensen J, Bergh J, Hagberg H, Glimelius B, Rastad J, Tholander Band Larsson R, Detection of tumor-specific cytotoxic drug activity invitro using the fluorometric microculture cytotoxicity assay and pri-mary cultures of tumor cells from patients. Int J Cancer 56: 715-720,1994.

90.Fruehauf JP and Bosanquet AG, In vitro determination of drugresponse: A discussion of clinical applications. Principles & Practice ofOncology, updates 7: 1-16, 1993.

91.Larsson R, Fridborg H, Kristensen J, Sundström C and Nygren P, Invitro testing of chemotherapeutic drug combinations in acute myelocy-tic leukaemia using the fluorometric microculture cytotoxicity assay(FMCA). Br J Cancer 67: 969-974, 1993.

92.Weisenthal LM, Antineoplastic drug screening belongs in the laborato-ry, not in the clinic. J Natl Cancer Inst 84: 466-69, 1992.

93.Sundström C and Nilsson K, Establishment and characterization of ahuman histiocytic lymphoma cell line (U-937). Int J Cancer 17: 565-577, 1976.

94.Larsson R, Kristensen J, Sandberg C and Nygren P, Laboratory deter-mination of chemotherapeutic drug resistance in tumor cells frompatients with leukemia, using a fluorometric microculture cytotoxicityassay (FMCA). Int J Cancer 50: 177-185, 1992.

95.McConnell HM, Owicki JC, Parce JW, Miller DL, Baxter GT, WadaHG and Pitchford S, The Cytosensor microphysiometer: Biologicalapplications of silicon technology. Science 257: 1906-1912, 1992.

96.Hafner F, Cytosensor microphysiometer: technology and recent appli-cations. Biosens Bioelectron 15: 149-58, 2000.

97.Owicki JC, Bousse LJ, Hafeman DG, Kirk GL, Olson JD, Wada HGand Parce JW, The light-addressable potentiometric sensor: Principlesand biological applications. Annu Rev Biophys Biomol Struct 23: 87-113, 1994.

98.Hafeman DG, Parce JW and McConnell HM, Light-addressablepotentiometric sensor for biochemical systems. Science 240: 1182-1185, 1988.

99.Harris DW, Kenrick MK, Pither RJ, Anson JG and Jones DA,Development of a high-volume in situ mRNA hybridization assay forthe quantification of gene expression utilizing scintillating microplates.Anal Biochem 243: 249-256, 1996.

100. Graves R, Davies R, Brophy G, O´Beirne G and Cook N,Noninvasive, real-time method for the examination of thymidine upta-ke events: application of the method to V-79 cell synchrony studies.Anal Biochem 248: 251-257, 1997.

101. Sjöholm Å, Bucht E, Theodorsson E, Larsson R and Nygren P,Polyamines regulate human medullary thyroid carcinoma TT-cell pro-

56


liferation and secretion of calcitonin and calcitonin gene-related pepti-de. Mol Cell Endocrinol 103: 89-94, 1994.

102. Sjöholm Å, Larsson R and Nygren P, Difluoromethylornithine andethylglyoxal bis(guanylhydrazone) as inhibitors of human renal carci-noma cell proliferation and polyamine metabolism. Anticancer Res 13:979-984, 1993.

103. Löhr GW and Waller HD, Glucose-6-phosphate dehydrogenase. In:Methods of enzymatic analysis, Vol. 2 (Ed. Bermeyer HU), pp. 636-543. Academic Press Inc, Weinheim, 1974.

104. Suhnel J, Evaluation of synergism or antagonism for the combinedaction of antiviral agents. Antiviral Res 13: 23-40, 1990.

105. Berenbaum MC, What is synergy? Pharmacological reviews 41: 93-132, 1989.

106. Valeriote F and Lin H, Synergistic interaction of anticancer agents: acellular perspective. Cancer Chemother Rep 59: 895-900, 1975.

107. Fridborg H, Feasiability of the fluorometric microculture cytotoxicityassay (FMCA) in the preclinical evaluation of anticancer drugs. In:Thises in Clinical Pharmacology, pp. 66. Uppsala University, Uppsala,1996.

108. Hirst MA and Pitchford S, Use of a single assay system to assess func-tional coupling of a variety of receptors. J NIH Res 5: 69, 1993.

109. Cook D and O´Kennedy R, Comparison of the tetrazolium salt assayfor succinate dehydrogenase with the Cytosensor microphysiometer inthe assessment of compound toxicities. Analytical Biochemistry 274:188-194, 1999.

110. Cao CJ, Mioduszewski RJ, Menking DE, Valdes JJ, Cortes VI,Eldefrawi ME and Eldefrawi AT, Validation of the Cytosensor for invitro cytotoxicity studies. Toxicology in vitro 11: 285-293, 1997.

111. Twiner MJ, Hirst M, Valenciano A, Zacharewski TR and Dixon SJ,N,N-Dimethylformamide modulates acid extrusion from murine hepa-toma cells. Toxicol Appl Pharmacol 153: 143-151, 1998.

112. Toninello A, Dalla Via L, Di Noto V and Mancon M, The effects ofmethylglyoxal-bis(guanylhydrazone) on spermine binding and trans-port in liver mitochondria. Biochem Pharmacol 58: 1899-1906, 1999.

113. Toninello A, Siliprandi D, Castagnini P, Novello MC and Siliprandi N,Protective action of methylglyoxal bis (guanylhydrazone) on the mito-chondrial membrane. Biochem Pharmacol 37: 3395-3399, 1988.

114. Chaffee RRJ, Arine RM and Rochelle RH, The possible role of intra-cellular polyamines in mitochondrial metabolic regulation. BiochemBiophys Res Commun 86: 293-299, 1979.

115. Pelster B and Niederstätter H, pH-dependent proton secretion in cul-tured swim bladder gas gland cells. Am J Physiol 273: R1719-25,1997.

116. Eguchi Y, Shimizu S and Tsujimoto Y, Intracellular ATP levels determi-ne cell death fate by apoptosis or necrosis. Cancer Res 57: 1835-1840,1997.

57

References

117. Lövborg H, Gullbo J, Ekelund S, Nygren P and Larsson R,Modulation of cell death and metabolic effects by 3-aminobenzamidein U-937 GTB cells exposed to CHS 828. Manuscript, 2001.

118. Colussi C, Albertini MC, Coppola S, Rovidati S, Galli F and GhibelliL, H2O2-induced block of glycolysis as an active ADP-ribosylationreaction protecting cells from apoptosis. FASEB 14: 2266-2276, 2000.

119. Martinsson P, Liminga G, Dhar S, de la Torre M, Lukinius A, JonssonE, Bashir-Hassan S, Binderup L, Kristensen J and Larsson R,Temporal effects of the novel antitumor pyridyl cyanoguanidine (CHS828) on human lymphoma cells. Eur J Cancer 37: 260-267, 2001.

120. Sariban-Sohraby S and Dale JB, The amiloride-sensitive sodium chan-nel. Am J Physiol 250: C175-C190, 1986.

121. Rotin D, Wan P, Grinstein S and Tannock I, Cytotoxicity of com-pounds that interfere with the regulation of intracellular pH: A poten-tially new class of anticancer drugs. Cancer Res 47: 1497-1504, 1987.

122. Boyer MJ and Tannock IF, Regulation of intracellular pH in tumorcell lines: Influence of microenvironmental conditions. Cancer Res 52:4441-4447, 1992.

123. Parkins CS, Chadwick JA and Chaplin DJ, Inhibition of intracellularpH control and relationship to cytotoxicity of chloambucil and vin-blastine. Br J Cancer 74: S75-S77, 1996.

124. Wunsch S, Sanchez CP, Gekle M, Grosse-Wortmann L, Wiesner J andLanzer M, Differential stimulation of the Na+/H+ exchanger determi-nes chloroquine uptake in plasmodium falciparum. J Cell Biol 140:335-345, 1998.

125. Sakamoto N, Toge T and Nishiyama M, Tumor-specific synergistictherapy of mitomycin C: modulation of bioreductive activation.Hiroshima J Med Sci 46: 67-73, 1997.

126. Kristensen J, Nygren P, Liliemark J, Fridborg H, Killander A,Simonsson B, Öberg G and Larsson R, Interactions between cladribine(2-chlorodeoxyadenosine) and standard antileukemic drugs in primarycultures of human tumor cells from patients with acute myelocyticlymphoma. Leukemia 8: 1712-1717, 1994.

127. Kanzawa F, Nishio K, Fukuoka K, Fukuda M, Kunimoto T and SaijoN, Evaluation of synergism by a novel three-dimensional model forthe combined action of cisplatin and etoposide on the growth of ahuman small-cell lung-cancer cell line, SBC-3. Int J Cancer 71: 311-319, 1997.

128. Baxter GT, Bousse LJ, Dawes TD, Libby JM, Modlin DN and OwickiJC, Microfabrication in silicon microphysiometry. Clin Chem 40:1800-4, 1994.

129. Henning T, Brischwein M, Baumann W, Ehret R, Freund I, KammererR, Lehmann M, Schwinde A and Wolf B, Approach to a multiparame-tric sensor-chip-based tumor chemosensitivity assay. Anticancer Drugs12: 21-32, 2001.

130. Bomanji JB, Costa DC and Ell PJ, Clinical role of positron emission

58


tomography in oncology. Lancet Oncology 2: 157-64, 2001.131. Mork Hansen C, Hansen D, Kaae Holm P, Larsson R and Binderup L,

The cyanoguanidine CHS 828 induces programmed cell death withapoptotic features in human breast cancer cells in vitro. AnticancerRes 6: 4211-4220, 2000.

132. Martinsson P, de la Torre M, Binderup L, Nygren P and Larsson R,Cell death with atypical features induced by the novel antitumoraldrug CHS 828, in human U-937 GTB cells. Eur J Pharmacol 417:181-187, 2001.

133. Sheline CT and Choi DW, Neuronal death in cultured murine cortialcells is induced by inhibition of GAPDH and triosephosphate isomera-se. Neurobiology of Diseases 5: 47-54, 1998.

134. Liminga G, Martinsson P, Jonsson B, Nygren P and Larsson R,Apoptosis induced by calcein acetoxymethyl ester in the human histio-cytoc lymphoma cell line U-937 GTB. Biochem Pharmacol 15: 1751-9, 2000.

59

References

Microphysiometry in the Evaluation of Cytotoxic Drugs with …160922/FULLTEXT01.pdf ·...

Documents

Transcript of Microphysiometry in the Evaluation of Cytotoxic Drugs with …160922/FULLTEXT01.pdf ·...