Microphysiometry in the evaluation of cytotoxic drugs with special emphasis on the novel cyanoguanidine CHS 828

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_____________________________ _____________________________

Microphysiometry in the Evaluation of Cytotoxic Drugs

with Special Emphasis on the Novel Cyanoguanidine

CHS 828

BY

SARA EKELUND

ACTA UNIVERSITATIS UPSALIENSIS UPPSALA 2001

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ABSTRACT

Ekelund, S. 2001. Microphysiometry in the evaluation of cytotoxic drugs with special emphasis 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, in the in vitro evaluation of cytotoxic drugs, using the lymphoma cell line U-937 GTB and primary cultures of tumour cells from patients as main model systems. The method was specifically 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/II clinical trials.

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

In U-937 cells, CHS 828 induced a cytotoxic effect characterised by a steep concentration-response relationship followed by a plateau. After 24 h of incubation the DNA and 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 the structurally related mitochondrial inhibitor m-iodobenzylguanidine (MIBG). The CHS 828 induced acidification was observed in cell lines as well as in cells from various tumour types from patients and probably originates from increased glycolytic flux. The effects may be secondary to block of oxidative phosphorylation in the mitochondria, but the relevance of the early acidification is not clear. CHS 828 seemed to induce a late, at approximately 15 h, inhibition of the glycolysis followed by loss of ATP and subsequent cell death. After exposure to MIBG the loss of ATP and cell death occurred earlier and in parallel. The effects of CHS 828 were not found to resemble those of the structurally related polyamine biosynthesis inhibitor methylglyoxal-bis(guanylhydrazone) (MGBG). Thus, CHS 828 may represent a new and interesting mode of cytotoxic action worthwhile for further development.

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

It is concluded that measurement in the Cytosensor^®microphysiometer of early cellular 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 metabolism, CHS 828, guanidino-containing compound.

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

Printed in Sweden by Tryck & Medier, Uppsala 2001

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To myself...

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Papers discussed

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

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

II S Ekelund, G Liminga, F Björkling, E Ottosen, C Schou, L Binderup and R Larsson; Early stimulation of acidification rate by novel cytotoxic pyridyl cyanoguanidines in human tumor cells: comparison with 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 containing drug 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 cytotoxic 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 between the new cytotoxic drug CHS 828 and amiloride and mitomycin C in a human tumour cell line and in tumour cells from patients.

Manuscript, 2001.

Reprints were made with the permission of the publishers.

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ALL Acute lymphocytic leukaemia

AML Acute myelocytic leukaemia

AraC Cytarabine

ATP Adenosine triphosphate

BCECF 2´,7´-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein

CHS 828 N-(6-(4-chlorophenoxy)hexyl)-N´-cyano-N´´-4-pyridylguanidine

CI Combination index

CisP Cisplatinum

CLL Chronic lymphocytic leukaemia

CNCn α-Cyano-4-hydroxy cinnamic acid

CO² Carbon dioxide

2DG 2-deoxy-D-glucose

DHEA Dehydroisoandrosterone

DIDS Diisothiocyanatostilbene-2,2´-disulfonic acid

DMSO Dimethyl sulphoxide

Dox Doxorubicin

FAD Flavin adenine nucleotide

FDA Fluorescein diacetate

FMCA Fluorometric microculture cytotoxicity assay G6PD Glucose 6-phosphate dehydrogenase

GTP Guanosine 5´-triphosphate

h Hour(s)

Melph Melphalan

MGBG Methylglyoxal-bis(guanylhydrazone)

MIBG m-Iodobenzylguanidine

min Minute(s)

MMC Mitomycin C

MTT 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide NAD+/NADH Nicotine adenine dinucleotide

NADP Nicotine adenine dinucleotide phosphate

NCI National Cancer Institute

NTP Nucleoside 5´-triphosphate

O² Molecular oxygen

ODC Ornithine decarboxylase

Pac Paclitaxel

PARP Poly(ADP-ribose) polymerase

PBMC Peripheral blood mononuclear cells

PBS Phosphate buffered saline

PET Positron emission tomography

pHi Intracellular pH

pH^e Extracellular pH

PPP Pentose phosphate pathway

s Second(s)

SAMDC S-adenosyl methionine decarboxylase

SEM Standard error of the mean

SI Survival index

TCA Tricarboxylic acid cycle

Topo Topotecan

Vcr Vincristine

VP16 Etoposide

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Introduction

Chemotherapy for cancer

The development of effective and specific medical treatment for malignant diseases has been difficult since no general and major differences between tumour and normal cells that could be exploited for targeting therapy have been found. Thus, a major problem is to direct the anticancer therapy to tumour cells only, without causing any, or only limited, toxicity to normal cells. 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 from a fairly limited number of chemical structures. In Sweden, there are currently 45 registered cytotoxic drugs that can be classified according to their chemical structure and/or putative mechanism of action (Table 1) [2].

Classically, most of these drugs have been described to interact with cell replication, e.g., DNA synthesis, transcription, or mitosis.

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

Alkylating agents Antimetabolites Cytotoxic antibiotics and related substances

Topoisomerase interactive agents

Platinum agents

Antimicrotubule agents

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

Miscellaneous agents

Amsacrine Asparaginase Altretamine

Hydroxicarbamide Miltefosin

Estramustin

Table 1.

Cytotoxic drugs approved for marketing in Sweden according to the Swedish

FASS, 2001

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Chemotherapy for cancer often uses combinations of agents with different mechanisms of action rather than single agent therapy. This is because of the therapeutic advantage (better effect, fewer side-effects) that combinations usually provide over single agents [1]. When a new anticancer agent is being developed, it is important to investigate potential candidates for combination therapy. The presently used clinical protocols for cancer combination therapy were mainly obtained empirically from clinical trials [4]. A better strategy for selection of combinations of chemotherapeutic agents is needed.

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

Mechanistic classes of cytotoxic drugs

The main groups of cytotoxic anticancer agents currently registered and used in Sweden, and their putative mechanisms of action, are briefly described below:

The alkylating agents belong to the oldest class of anticancer drugs and the first clinical studies of nitrogen mustards were initiated in 1942 [5]. The alkylating agents are chemically diverse drugs that may undergo transformation to produce reactive intermediates that can bind covalently to electron-rich moieties on biological molecules, e.g., DNA. Alkylation of bases 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 acid analogues are all antimetabolites. These are drugs that have been synthesised to inhibit crucial biochemical pathways, usually leading to inhibition of DNA or RNA synthesis and these drugs tend to be cell-cycle dependent [6].

Microtubules are components of the mitotic spindle with an important function during cell division. Microtubules are composed of two subu- nits of tubulin, which form a heterodimer. Tubulin heterodimers and

microtubules are in a state of dynamic equilibrium with continuous formation and degradation and this process is an important cellular target for cancer chemotherapy. Both the vinca alkaloids and the taxanes belong to the class of antimicrotubule agents. The former acts by binding to the protein tubulin, thus inhibiting its polymerization to form microtubules. The taxanes bind to and stabilise the microtubules and, thereby, the depolyme- risation to tubulin is prevented and essential mitotic functions are inhibited [1].

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The chemical structure of the anthracycline antibiotics consists of a pigmented tetracyclic ring structure, an unusual sugar and a lateral chain.

The first anthracyclines were produced by the Streptomyces species, but the second-generation anthracyclines are synthetic. This class of drugs is one of the most clinically used anticancer agents, but their clinical use is limited by irreversible cardiac toxicity [1]. These compounds act by intercalating with DNA and many functions of the DNA are affected, including DNA and RNA synthesis, leading to single and double-strand breaks. The anthracyclines are also known to interact with the function of the nuclear enzymes topoisomerases (type I and II), forming stabile complexes between enzyme and DNA, thus leading to permanent breaks of the DNA strands [5].

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

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

Cell death and apoptosis

Above, the classical mechanisms of action for cytotoxic drugs are described. With increased understanding of tumour cell biology and with new techniques, the picture has become more complicated, with additional aspects to consider. One important finding is that most of the cytotoxic drugs kill tumour cells by inducing programmed cell death, i.e., apoptosis [7]. Probably, the drugs induce damage to vital cellular functions that in turn initiate apoptosis. However, the connection between apoptosis and the mechanisms of cytotoxic drug action is not clear. The alternative way for cells to die is by necrosis.

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

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membrane rupture, which triggers an inflammatory process [8].

The complete apoptotic programme has been shown to involve energy-requiring steps [11, 12] and the balance between death by apoptosis and necrosis appears to depend upon the intensity of the injury [12, 13]. Thus, ATP levels have been shown to serve as a switch between apoptosis and necrosis [12-14] and in endothelial cells glutamine could partially restore ATP levels after H²O²exposure, resulting in a significant increase in apoptosis instead of necrosis [15]. In cultured neurones it was demonstrated that intracellular energy levels were rapidly dissipated in necrosis, but not in apoptosis [16]. In fact, it is necessary to deplete tumour ATP levels by as much 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 be important to study the effects of cytotoxic drugs on cellular energy metabolism. The principal cellular pathways for energy metabolism are, therefore, summarised below.

Cellular energy metabolic pathways

An important characteristic of fast-growing cancer cells is their elevated rate of glycolysis and the formation of lactate from glycolytic pyruvate [18, 19]. It has been demonstrated that as much as 90% of glycolytic pyruvate in cancer cells can be reduced to lactate [20].

Mitochondrial respiration is the major source of ATP from glucose under most conditions in vivo, but glycolysis usually predominates in vitro [21]. The ATP yield from glucose through lactate formation is much lower than 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. The glycolytic pathway contains a different isozymic composition from the normal cell and the activity of key enzymes of regulation is often increased [23]. Other factors discussed are, e.g., facilitated glucose transport and high glutaminase activity [23, 24].

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

Glycolysis

Glucose enters the cells by passive transport through carrier proteins [25].

The first step in glucose metabolism is the glycolysis, which takes place in the cytoplasm. Glycolysis is composed of two phases divided in nine steps with the net gain of 2 molecules of ATP per glucose molecule. The first phase concerns glucose phosphorylation and its conversion into glyceraldehyde-3-phosphate. The second is the conversion of glyceraldehyde-3-phosphate 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

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cycle take place in the mitochondria. Pyruvate enters the mitochondria and is converted to the acetyl groups of the chemically reactive acetyl CoA by decarboxylation. In the TCA the acetyl groups are oxidized to produce CO², NADH, FAD and GTP. The CO²produced diffuses from the mitochondrion and leaves the cell [25].

Respiratory chain - oxidative phosphorylation

It 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 molecular oxygen (O² from water) to produce ATP and H²O [25].

Acetyl COA

Tricarboxylic acid cycle

Mitochondria

Oxidative phosphorylation NADH + H⁺

Complex II Complex Complex III

I

2H⁺ 2H⁺ 2H⁺

H⁺ F0

F1

ATP ADP+ Pi H⁺

NADH + H⁺

Glucose

Pyruvate Lactate

Glycolysis

Cytosol

Cell membrane

Glucose

6-phosphate Ribose 5-phosphate

4 carbon 5 carbon 7 carbon

Pentose phosphate

pathway ADP⁺

ATP

NADPH

e^- transport chain

(mitochondria inner membrane) CO2

CO2

Figure 1.

The principal energy-yielding pathways in cells are schematically described

and the different waste-products indicated.

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

Pentose phosphate pathway

The first step in the glycolysis results in glucose-6-phosphate. This is an intermediate 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, whereas ribose-5-phosphate is used in the synthesis of RNA, DNA and nucleotide enzymes. There is an interplay between the PPP and glycolytic pathway that enables the levels of NADPH, ATP and building blocks such as ribose- 5-phosphate and pyruvate to be continuously adjusted to meet the cellular needs [26].

Energy yield and acid production of the metabolic pathways

In a comparison between the principle energy-yielding metabolic pathways and with glucose as the carbon source, respiration through oxidative phosphorylation is the pathway that yields most ATP (36), followed by the combination of PPP + glycolysis + oxidative phosphorylation (27).

Furthermore, glycolysis produces most protons per ATP as compared with respiration through oxidative phosphorylation or through the combination of PPP + glycolysis + oxidative phosphorylation [21].

Regulation of intra- and extracellular pH

The new drug investigated in this thesis, N-(6-(4-chlorophenoxy)hexyl)-N´- cyano-N´´-4-pyridylguanidine (CHS 828), was found to stimulate cellular metabolism and increase the production of acidic waste-products that could subsequently decrease extracellular pH (pH^e). Such effect on pHe might also have implications for intracellular pH (pHⁱ). Therefore, regulation of pH^eand pHⁱare briefly discussed below.

Due to the poor vascularisation in most solid tumours, there are hypoxic and anoxic areas heterogeneously distributed within the tumour mass [27]. The metabolic rate of tumour cells is usually high, leading to a deficiency of both nutrients and oxygen, and the production of increased amounts of waste-products like protons, lactate and CO². These circum- stances will together result in an acidic pH^e [27]. Thus, tumour pH is

known to be approximately 0.5 units lower than in normal tissues [28, 29].

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The activities of a large number of intracellular enzymes that take part in the cellular metabolism are pH-sensitive. Protein, DNA and RNA synthesis are affected by pH and increase with increasing intracellular pH within the physiological range [30]. Thus, in order to survive and prolifera- te under acidic conditions, tumour cells become dependent on mechanisms to maintain pHⁱwithin the physiological range. There are three major membrane-based transport systems that are involved in the regulation of pHⁱ, 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 all mammalian cells. This exchanger acts to increase pHⁱand catalyses an electroneutral exchange of Na+ and H+ across the cell membrane [31]. The driving force for the H+ extrusion is provided by the Na+ gradient across the membrane [32]. The exchanger is inhibited by amiloride, which acts on the 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 reactions: H+ + HCO³- ↔ H²CO³ ↔ H²O + CO²[31]. This anion exchanger is present in U-937 cells [33], a cell line frequently used in this thesis. The exchanger 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 must be transported out of cells. Lactic acid is dissociated into a lactic anion and a proton at physiological pH and therefore diffusion across the cell membrane is probably minimal. Instead, lactate may be transported by the Na+

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

Nucleus

HCO₃^-

Cl^- Na⁺

Na⁺

H⁺

Lactate 3- -

Na

⁺

-dependent

HCO /Cl exchanger Na

^{+ +}

/H antiport

Cytoplasm

Cell membrane

H

⁺

/ lactate symport

Figure 2.

The main mechanisms considered to regulate acid excretion in cells.

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

Effects of cytotoxic drugs on cellular metabolism

The effect of cytotoxic drugs on cellular energy metabolism has so far not been a major field of research. However, some knowledge has accumulated.

The alkylating agents have been described to cause a decrease of tumour NAD+ levels. In addition, direct inhibition of purified glycolytic enzymes has also been demonstrated by some alkylating agents, but no causative relationship between pharmacological effects and the degree of glycolytic inhibition was observed [35].

Bleomycin and cytarabine (AraC) were shown to be inactive on both glycolysis and respiration, while doxorubicin (Dox) had a stimulating effect on respiration [36]. However, Dox has also been described to inhibit cellular respiration [37].

Actinomycin-D has been shown to induce an early and transient increase in ATP and GTP contents, while anthracyclines induced a corres- ponding increase in the total nucleoside triphosphate pool (NTP). This effect was suggested to be characteristic for these drugs, since cisplatinum (CisP) and AraC did not alter the NTP content [38]. In another study the ATP 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 in decreased ATP levels [39, 40]. Other cytotoxic drugs, e.g., paclitaxel (Pac) and etpopside (VP16), elicited similar effects, indicating the possibility of a general effect of drugs inducing apoptosis [40].

Pac has been reported to alter rat hepatocyte metabolism by inhibiting the respiratory chain, causing an immediate reduction of oxygen consumption and increased glycolytic flux [41]. The ATP levels were not enough to compensate for the inhibition of oxidative phosphorylation and cell viability was subsequently impaired. This ATP reduction might be a consequence of decreased activity of regulatory glycolytic enzymes [42].

A build-up of glycolytic intermediates and decrease in cellular energy content was reported for cells exposed to Dox, explained by the inhibition 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 the regulation of metabolism, Pac and vinblastine were both found to decrease the glycolytic rate [44]. This was surprising since these drugs are known to induce opposite effects on microtubule assembly. Based on these findings it was suggested that both drugs produce their effect on glycolytic rate by competing with the glycolytic enzymes for binding sites on the tubulin molecule [44].

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

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48]. These cell lines exhibited an increased rate of glycolysis, oxidative phosphorylation and PPP with a subsequent increased O²consumption as well as an increased production of CO², lactate and ATP.

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

MIBG

MIBG has a molecular mass of 324.1 g/mol and was synthesised over 20 years ago [57]. The structure of MIBG is shown in figure 3. It is the guanidino-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-seeking radiopharmaceutical agent for the diagnosis and treatment of neuro-endo- crine tumours [60].

A concentration of 10 µg/ml (equals 31 µM) MIBG inhibited the mitochondrial respiration of leukaemia L1210 cells [49, 61]. In studies using the human neuroblastoma cell line SK-N-BE(2c), optimal arrest of proliferation was seen at MIBG concentrations of 25 µM, but the mitochondrial 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 on mitochondrial oxidative phosphorylation [50, 51].

MIBG also has also demonstrated antitumour effects in animal models. MIBG alone causes hyperglycaemia and induces a lowering of tumour extracellular pH [62]. In addition, a stimulatory effect on tumour cell glycolysis in vivo was indicated by a downshift of tumour pH and an increase in plasma lactate levels in tumour-bearing animals with the decrease in pH limited to malignant tissue [63].

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

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MGBG

The molecular mass of MGBG is 257.1 g/mol (Figure 3) and the molecule consists of two polar aminoguanidino-groups separated by an aliphatic skeleton. There are similarities between MGBG and the natural polyamines, especially with spermidine. The polyamines spermidine, spermine and their precursor putrescine are present in all mammalian cells, but their physiological functions are not well understood [53, 66]. They are involved in many biologic processes [67], and an increased level of polyamines has been reported in several malignant diseases [68]. Clinical trials with MGBG were initiated in the early 1960s [69]. The drug is now known to be an antileukaemic agent and has also been reported to have activity against some solid tumours [66, 70].

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

In several human and murine cell types, MGBG has been reported to produce ultrastructural damage to the mitochondria [74, 75]. On the other hand, another study showed that MGBG was without effect on mitochondrial respiration, as there was no effect on oxygen consumption, ATP content or lactate production in a study with human and mouse neuroblastoma and lymphosarcoma cell lines [54]. In yet another study, MGBG was found to exert two effects on mitochondria; protection of mitochondria at low concentrations and aggregation at higher concentrations [76]. Most of the effects of MGBG on cultured cells can be prevented by the concurrent administration of spermidine at equimolar or higher concentrations [77].

NH N

N NH

N

O

Cl H2N NH

NH N

CH3

N NH NH2

NH H2NN NH

N H

I

MIBG MGBG

CHS 828

N

Figure 3.

Chemical structures for the guanidino-containing compounds MIBG, MGBG

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

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CHS 828

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

In the 1970s there was an interest in the hypotensive effect of a structural prototype, the potassium channel opener, pinacidil and its analogues [59]. Later it was discovered that a number of related pyridyl cyanoguanidines showed antitumour activity in a routine screening programme in a rat model [78]. This finding inspired further investigation of the guanidines in order to find structure-activity relationship for this class of substances. Crucial for the antiproliferative effect was the cyanoguanidine moiety but also the length of the carbon chain [78]. Optimisation and testing 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 no hypotensive effect. The chemical structure of CHS 828 is shown in figure 3.

CHS 828 has demonstrated interesting properties as a potential anti- cancer agent and has been found active against many tumour cell lines in vitro [78, 79]. A differential pattern of antitumour activity was revealed when investigating the cytotoxicity of CHS 828 in a panel of 10 human tumour cell lines representing defined mechanisms of resistance, including those associated with expression of P-glycoprotein, altered topoisomerase II, increased GSH levels and tubulin defects [79]. Activity was observed in the nM to µM range and the typical shape of the dose-response curves was a concentration dependent decrease in cell survival followed by a plateau.

This plateau was cell proliferation independent, since the phenomenon was also observed in non-proliferative cell systems, e.g., peripheral blood

mononuclear 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 tumour cells 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 the haematological malignancies. Solid tumour cells appeared less responsive.

Similar results were also demonstrated in an in vitro hollow fibre assay, in which high activity of CHS 828 was shown in CLL, with less activity towards solid ovarian carcinoma [82].

After oral administration in several animal models, CHS 828 demonstrated a broad spectrum of activity while causing only little or no toxicity to the animals. The xenograft models of MCF-7 breast cancer and NYH small cell lung cancer in nude mice, were both found sensitive to CHS 828 [79]. These data are interesting since the NYH xenograft model was not very sensitive to standard drugs.

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Cytotoxic drug development and evaluation

Principles for cytotoxic drug development

There are many different ways to proceed in the design and development of anticancer drugs. New compounds can be discovered by screening of

various chemicals. This approach has long been adopted by the National Cancer Institute (NCI). Today, NCI has an established primary screen in which compounds are tested for the ability to inhibit growth of 60 different 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 selected for further testing based on several different criteria such as disease-type specificity, unique structure, potency and demonstration of unique pattern of cytotoxicity in the cell line panel, indicating a new mechanism of action [84].

Another strategy in drug discovery is the modification of already existing compounds [85]. These agents can be altered to enhance their activities and to decrease their unwanted toxic effects. A third approach is rational drug design based on knowledge on specific biochemical or molecular targets in tumour cells. This approach might become fruitful as indicated by the recently developed tyrosine kinase inhibitor STI571 for treatment 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 treatment of cancer can only derive from the clinical trials programme.

However, the capacity to test new treatments in patients is limited in comparison with the great number of promising new drugs. For safety reasons and for optimising the chance of finding active new drugs, a lot of information needs to be collected during the pre-clinical phase of the development programme for a new cytotoxic drug [87].

Briefly, the pre-clinical programme includes studies in vitro, so far mostly on tumour cells lines and subcellular model systems, to characterise the activity profile of the new drug and mechanisms of action and resistance. Furthermore, some information should be obtained on suitable exposure 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, therapeutic index and schedule dependency of the new drug. In addition, extensive studies 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 available from the pre-clinical programme prior to clinical trials, the experience indicates that very few of the promising drug candidates will progress to

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become clinically useful drugs after extensive and resource consuming testing in patients [88]. A major deficiency in the pre-clinical development programmes is that the information provided on the antitumour activity is mostly not sufficiently predictive for the final clinical efficacy. Pre-clinical in 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 types and settings in which the new drug could be expected to be active.

The use of in vitro assays based on cells prepared from patient tumours, might be an approach to provide clinically more predictive information on the anticancer activity. Several assays based on the concept of total cell kill during short-term culture are available for this [89, 90].

Furthermore, the testing of possible new drug combinations can, as a first step, preferably be done in vitro [91]. Assays, developed to examine the cytotoxicity or biochemical effects of drugs on cultured cells, could indicate the therapeutic effect of combined agents as well as provide information on possible mechanisms of drug interactions. Actually, in vitro studies are uni- que in their ability to quantitatively evaluate synergistic or antagonistic interactions [4].

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

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Aims of the thesis

In general, the studies comprising this thesis were focused on the evaluation of cytotoxic drugs using the Cytosensor^®microphysiometer, with special emphasis on the novel cytotoxic cyanoguanidine CHS 828. Additional methods were used to further characterise the effect of CHS 828 and related drugs on, e.g., metabolic pathways, cytotoxicity, pHⁱand DNA/protein synthesis.

The more detailed aims of this thesis were to:

• Investigate the feasibility of the Cytosensor^® for evaluation of drugs used in the treatment of cancer, with a special emphasis on the novel cytotoxic drug CHS 828.

• Compare the metabolic and cytotoxic effects induced by CHS 828 with those of the established guanidino-containing compounds MIBG and MGBG.

• Characterise over time the metabolic events induced by CHS 828 and thereby increase the understanding of the mechanism of action of the drug.

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

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Materials and methods

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

Cells and cell culture

The most frequently used cell line was the human histiocytic lymphoma cell line, U-937 GTB [93]. Other cell lines used were the CHS 828 resistant subline U-937 CHS, the myeloma cell line RPMI 8226/S, the small cell lung cancer cell line NCI-H69 and the leukemic cell line CCRF-CEM. In some experiments tumour cells from patients with haematological or solid tumours were used. The samples were obtained from bone marrow/peripheral blood sampling, routine surgery or diagnostic biopsy. The sampling was approved by the local ethical committee at Uppsala University

Hospital. Cell preparations from these tumours have been described in detail earlier [89, 94].

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

Experimental drugs

CHS 828 and seven structurally related pyridylcyanoguanidines, supplied from Leo Pharmaceutical Products in 10 mM stock solutions, were dissolved in dimethyl sulphoxide (DMSO) and stored frozen at –20°C. The drugs were diluted ten times with 33% DMSO and sterile water and further dilu- tions were made using sterile phosphate buffered saline (PBS). MIBG and MGBG were from Sigma. MIBG was dissolved in 10% DMSO and sterile water and MGBG in sterile water to stock solutions of 1 mg/ml. They were stored 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 and topotecan (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 diluted as prescribed.

Measurement of extracellular acidification

A silicon microphysiometer, the Cytosensor^® (Molecular Devices

Corporation, Sunnyvale, CA, USA), was used to measure the excretion rate of metabolic waste-products.

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

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peristaltic pump and passes through a debubbler/degasser, a selection valve and then through the flow sensor chamber (Figure 4). In the measurement chamber the non-adherent cells were immobilised in 25% agarose

(Molecular Devices) to ensure that the cells are not washed away to one side of the chamber by the flow of medium. The optimal number of cells ranges from 10⁴to 10⁶cells [95] and in the present experiments there were 1.5 x 10⁵cells 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, in aqueous 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 to detect small changes of extracellular acidification rate [21, 97, 98]. Once each s the LAPS makes a voltage measurement of the acidification rate that is linearly related to pH and calculated by the Cytosoft program as -µV/s.

Medium is constantly pumped through two parallel channels at a flow rate of 100 µl/min. When the fluid flows (90 s), the sensor output is stable and reflects a pH near that of medium entering the flow chamber.

During flow-off periods (30 s), the acid released from the cells accumulates in the measuring chamber and the rate of release is quantified by fitting the sensor data to a straight line with the least-squares method. The slope of this line represents the acidification rate. No significant perturbations of cell physiology is caused during a cycle [95]. The change in acidification is

Pump

Debubbler Valve

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.

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

Cells were allowed to adapt in the Cytosensor^®during one h before the acidification rate for each measurement chamber was set to 100%

(baseline acidification). Thereafter, drugs were added. The acidification rates induced by drugs are presented as per cent of baseline acidification.

This compensated for small baseline differences and permitted comparison within 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 buffering capacity since it lacks both bicarbonate and Hepes. Thus, changes in acidification rate could readily be observed and were due only to the metabolic response of cells to the drugs. The osmotic balance was preserved by the addition of 6 ml/l 4 M NaCl. The medium also contained 10 ml/l 200 mM L-glutamine, 60 mg/l penicillin and 50 mg/l streptomycin.

Measurement of cytotoxicity

Viability of cells was determined by the fluorometric microculture cytotoxicity assay (FMCA). The assay is based on measurement of fluorescence generated from hydrolysis of fluorescein diacetate (FDA) to fluorescein by cells 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 final experimental concentration into the plates. The plates are then kept frozen at –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 10⁴and 1-3 x 10⁴cells/well for haematological and solid tumours, respectively), before incubation during 72 h at 37°C and 5% CO². Blank wells received culture medium only and control wells 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 incubated for 40 min. The fluorescence generated in each well was then measured at 538 nm in a Fluoroscan II (Labsystems Oy, Helsinki, Finland).

Measurement of DNA and protein synthesis

Protein and DNA synthesis were measured with a Cytostar-T plate, available in Amersham´s “In Situ mRNA Cytostar-T assay” kit, (Amersham International, Buckinghamshire, UK), a pre-made scintillating 96-well microtiter plate, with scintillation fluid moulded into the bottom of the wells [99, 100]. Cells were suspended in fresh media containing 111 nCi/ml [¹⁴C]Thymidine (Amersham CFA.532 56 mCi/mmol, 50 µCi/ml) for DNA synthesis 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

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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 methionine decarboxylase (SAMDC) were investigated in cells exposed to CHS 828 and MGBG as measurements of the polyamine biosynthesis. Cells were incubated with drugs during 1 or 20 h in cell culture flasks. After centrifugation, cells were resuspended in 1 ml cold PBS, centrifuged once more and sonicated 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/ml Trasylol, 10 mM benzamidine and 0.1 mg/ml albumin. The homogenates were centrifuged for 10 min at 4°C before an equal volume of supernatant and incubation buffer was mixed. For ODC measurements, the buffer contained 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 mM

NaPO4, 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. After 60 min incubation at 37 °C, reactions were terminated by addition of 100 µl 5 M H²SO⁴ and 250 µl Hyamine was used to trap liberated ¹⁴CO². After addition of 0.4 M Na²HPO⁴to liberate CO², Unisolve scintillation fluid was added and the radioactivity quantified by scintillation counting [101, 102].

Measurement of glucose consumption

Medium glucose concentration was used as an indirect measurement of glucose consumption of cells exposed to drug during different time periods ranging from 30 min to 50 h. 2.5 x 10⁵ cells/ml were incubated in culture flasks at 37°C. Samples were withdrawn from drug-exposed cells and from control cells. After centrifugation, 2 ml of the supernatants were transfer- red to a tube and stored at 8°C until analysis. The samples were analysed at the University Hospital laboratory using the routine analysis of glucose levels in plasma and other body fluids. Briefly, the level of glucose was indirectly determined by measuring the absorbance at 340 nm, which reflects the amount of NADH produced when glucose is enzymatically oxidized.

Measurement of lactate production

The production of lactate by cells exposed to CHS 828 and MIBG was measured using commercially available lactate reagents and a standard curve prepared from a lactate standard solution (40 mg/dl) (Sigma).

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

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SPECTRAmax^®PLUS (Molecular Devices). Blank values were subtracted and 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 based on bioluminescent measurements of ATP utilising the luciferase enzyme for formation of light from ATP and luciferin. Cell suspension was plated in a flat-bottomed, white 96-well microplate, which was incubated under standard cell culture conditions. At different time points, ranging from 5 min to 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 releasing reagent to each well before measurements were performed at 540 nm.

Measurement of G6PD

Measurment of G6PD was made using enzymatic analysis. CHS 828 was added to the cell suspension (5 x 10⁵cells/ml) after 0, 2, 8 and 24 h incubation 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 more and the supernatant collected and frozen until the enzyme activity analysis was performed at the University Hospital routine laboratory. Briefly, the rate of NADPH formation was measured spectrophotometrically and related to the enzyme activity [103]. The activity was presented using the unit nkatal/100µl cell suspension.

Measurement of changes in intracellular pH

Measurement of pHⁱchanges was performed using a fluorescence spectrophotometer 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 10⁶cells were loaded with 2 µM BCECF AM during 30 min at 37°C. Thereafter, cells were centrifuged and washed once with medium before dilution in a 10 ml standard Hepes buffer (Sigma-Aldrich) supple- mented with 10 mM glucose and 0.75 mM calcium chloride and with pH adjusted 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 spectrophotometer. 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. The concepts of these methods are described below, and in greater detail in Paper V.

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The isobole method is a commonly used method. Isobole means iso- effect curve and the construction of isoboles requires experimental data for the agents used alone and in various dose combinations at equi-effect levels [104]. This is a generally valid procedure for analysing interactions between agents irrespective of their mechanism of action or the nature of their dose-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 product of the individual survival fractions is employed [106]. Zero interaction of the combination is expected to be equal to the product of the survival indices of its constituents [105].

Presentation of results

The main results are summarised below with key findings illustrated in figures. Generally, figure data are presented as mean values ± SEM. For additional details on experimental performance and data presentation, see the individual papers.

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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 were investigated. Each drug induced a reproducible and characteristic response pattern, from which key features, e.g., stimulation and inhibition of acidification, the time point when the response curves of the drugs fell below the control curve, and the maximum inhibition of acidification at 20 h, could be quantified (Figure 5).

Vcr and VP16 induced distinct peaks of acidification during the first h, after which the acidification rate eventually declined. The shape of the response curves for AraC and Topo were similar to those of Vcr, but with smaller peaks. Melph and Dox followed the control curve several h before the acidification rate declined. For CisP and Pac, on the other hand, the acidification remained above or very close to the control curve for the complete duration of the 20 h experiments. For the drugs producing a reduced metabolic rate at 20 h, a concentration-response relationship was observed.

The qualitative nature of the response pattern induced by Vcr was found to be similar in the T-cell leukaemia cell line CCRF-CEM (not shown). To what extent the response patterns obtained are independent of cell type remains to be established.

The drug concentrations used are known to be highly cytotoxic after 72 h of continuous drug exposure. In Cytosensor^® experiments a decrease in acidification rate might be due either to a decreased number of viable cells and/or to a decrease in metabolic activity per cell and the proportions of these two alternatives are likely to be time dependent. Judging from the drug concentrations used in this study, long exposures inevitably lead to an increased number of dead cells.

To mimic the time frame of the Cytosensor^® experiments, the cytotoxic effects of the drugs were measured at 24 h in the FMCA. At 20-24 h of drug exposure the two assays correlated reasonably well (Figure 6).

There were two exceptions in Melph and Vcr, for which cell survival was high despite inhibition observed in the Cytosensor^®. This discrepancy may be related to differences in the overall conditions of tests. In the

Cytosensor^® experiments, the temperature of the medium is 37°C and continuously pumped through the sensor chambers so that metabolites and waste products are removed and new drug continuously added. In the microtiter plate-based assay, the plates are incubated at 37°C for 72 h with no new addition of drug and no change of medium. Melph and Vcr

decompose rapidly at 37°C [107], which may result in lower exposure in the FMCA than in the Cytosensor^®. This problem may arise also for other unstable drugs. These features of the Cytosensor^® have to be remembered when 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

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messengers and morphological or proliferative changes [108]. In a recent comparison with the established tetrazolium salt assay for assessment of cytotoxicity, the Cytosensor^® method was found more sensitive to the effects 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

5 10 15 20

0 25 50 75 100 125

Control AraC 0.5µg/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 2.5µg/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

^®

.

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reduction in acidification rate after 24 h of drug exposure and the IC⁵⁰ values calculated compared well with IC⁵⁰values obtained from [³H]thymidine uptake and Calcein AM fluorescence [110]. Most of the drugs tested inhibited cell metabolism and acidification rate, but ethanol enhanced the activity during the initial 4 h of exposure, but then irreversibly inhibited the metabolism after 24 h [110]. Similarly, in our study several of the cytotoxic drugs investigated initially stimulated the acidification rate. This phenomenon might possibly reflect an energy-requiring defence or response of cells as a first reaction to the drug.

The feasibility of using the Cytosensor^® for measurement of cytotoxic drug activity profile in primary cultures of patient ovarian carcinoma cells was also demonstrated in the present study. The response patterns of Dox was qualitatively similar to those obtained in the U-937 cell line, but CisP showed an initial stimulation of acidification. This type of tumour is known to be CisP sensitive but it remains to be clarified if this response reflects the presence of cell type specific response elements mediating apoptosis.

Early classification of new agents according to pharmacological similarity or dissimilarity to standard drugs with known mechanism of action is desirable in new drug discovery and development. The drug-specific 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 and characterisation could be the detailing of schedule dependency and drug interactions. The feasibility of easy change of exposure protocols in eight parallel channels makes the Cytosensor^® especially well suited for such applications.

Vcr Pac CisP Melph Dox VP16 AraC Topo 0

25 50 75 100

125 Acidification rate

Cell viability

Figure 6.

Microphysiometry in the evaluation of cytotoxic drugs with special emphasis on the novel cyanoguanidine CHS 828