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EVALUATION OF THE EFFECT OF P53 IN

CELLULAR RESPONSE FROM ELECTRON

MICROSCOPY IMAGES

ANA CATARINA FREITAS DA SILVA DE JESUS

JUNHO 2010

EVALUATION OF THE EFFECT OF P53 IN CELLULAR

RESPONSE FROM ELECTRON MICROSCOPY IMAGES

Report of the Course Practical Works, Master Course in Biomedical

Engineering Program, Faculty of Engineering of University of Porto

Ana Catarina Freitas da Silva de Jesus

Graduated in Biochemistry (2000)

Faculty of Science of University of Porto

Graduated in Nuclear Medicine (2006)

Superior School of Allied Health Sciences

Polytechnic Institute of Porto

Supervisor:

João Manuel R. S. Tavares

Assistant Professor of the Mechanical Engineering Department

Faculty of Engineering of University of Porto

ACKNOWLEDGEMENTS

To Professor João Manuel R. S. Tavares for the support provided throughout

this work, particularly for guidance, support and availability, essential for the proper

and constructive development of the same.

To all of those who make possible the development of this work.

SUMMARY

The theme of this practical work falls in the computational vision domain,

particularly in the area of the processing and analysis of biomedical images.

The objective of the correspondent MSc thesis is to perform the computational

analysis of cells represented in microscopic images. For that, the preprocessing of the

input images assumes particularly relevance. This first image preprocessing step is the

main concern of this practical work. In a later stage, that is, during the thesis project,

will be considered the segmentation of the input images and motion tracking and

analysis of cells submitted to irradiation.

In this practical work, it is emphasized the importance of the cell cycle

regulation, namely the cell death mechanisms. The associate checkpoints are

particularly important when the cells are irradiated. For this reason, in this work is

made a description of the harmful effect of radiation on cells and tissues. In addition,

the cell cultures and the adequate means to obtain reasonable laboratory culture of

cells, without contamination, for subsequent use to study the effect of radiation on

cells, are discussed.

The experimental results obtained through image processing and analysis

highlight the changes in intracellular content due to irradiation of cells and emphasize

the effects of the lack of cell regulation, specifically detecting the location of p53 and

changes in its content.

CONTENTS

CONTENTS

EVALUATION OF THE EFFECT OF P53 IN CELLULAR RESPONSE FROM ELECTRON MICROSCOPY IMAGES i

CHAPTER I – INTRODUCTION TO THE THEME AND REPORT ORGANIZATION 1

1.1 – Introduction 3

1.2 - Main Objectives 4

1.3 - Report Organization 6

1.4 - Major Contributions 8

CHAPTER II – CELL CYCLE REGULATION AND APOPTOSIS 9


2.2 - Cell Life Cycle 12

2.2.1 – Interphase 12

2.2.2 - DNA Replication 13

2.2.3 - Cell Division 14

2.2.3.1 – Mitosis 14

2.2.3.2 – Cytokinesis 16

2.2.4 – Meiosis 16

2.3 - Progression of the cell cycle 19

2.4 - Growth characteristics of malignant cells 26

2.4.1 - Phenotypic Alterations in Cancer Cells 27

2.4.2 - Immortality of Transformed Cells in Culture 28

2.4.3 - Decreased Requirement for Growth Factors 29

2.4.4 - Loss of Anchorage Dependence 29

2.4.5 - Loss of Cell Cycle Control and Resistance to Apoptosis 30

2.5 - Cell Cycle Regulation 31

CONTENTS

EVALUATION OF THE EFFECT OF P53 IN CELLULAR RESPONSE FROM ELECTRON MICROSCOPY IMAGES ii

2.5.1 - CDK Inhibitors 32

2.5.2 – Cyclins 33

2.5.3 - Cell Cycle Checkpoints 34

2.5.4 - Cell Cycle Regulatory Factors as Targets for Anticancer Agents 37

2.6 – Apoptosis 39

2.6.1 - Biochemical Mechanism of Apoptosis 41

2.6.2 – Caspases 44

2.6.3 - Bcl-2 Family 45

2.6.4 – Anoikis 45

2.7 - Resistance to Apoptosis in Cancer and Potential Targets for Therapy 47

2.8 – Summary 49

CHAPTER III – RADIATION EFFECT ON NORMAL AND NEOPLASTIC TISSUES 51


3.2 - Irradiation Carcinogenesis 54

3.2.1 - Ionizing Radiation 54

3.2.2 - Ultraviolet Radiation 55

3.3 - Cell Death in Mammalian Tissues 56

3.4 - Nature of Cell Populations in Tissue 59

3.5 - Cell Population Kinetics and Radiation Damage 60

3.5.1 - Growth Fraction and its significance 61

3.6 - Cell Kinetics in Normal Tissues and Tumors 62

3.7 - Models for Radiobiological Sensitivity of Neoplastic Tissues 63

CONTENTS

EVALUATION OF THE EFFECT OF P53 IN CELLULAR RESPONSE FROM ELECTRON MICROSCOPY IMAGES iii

3.7.1 - Hewitt Dilution Assay 64

3.7.2 - Lung Colony Assay System 67

3.8 - Tumor Growth and Tumor “Cure” Models 68

3.8.1 - Tumor Volume Versus Time 68

3.8.2 - TCD50, Tumor Cure 70

3.9 - Radiobiological Responses of Tumors 70

3.10 - Hypoxia and Radiosensitivity in Tumor Cells 71

3.11 – Summary 74

CHAPTER IV – CELL CULTURE AND FLOW CYTOMETRY 75


4.2 - Cell-Culture Laboratory 77

4.3 - Maintaining Cultures 78

4.3.1 – Medium 79

4.3.2 - The use of medium in analysis and alternatives 83

4.4 - Cytogenetic Analysis of Cell Lines 84

4.4.1 - The Utility of Cytogenetic Characterization 84

4.5 - Methods to Induce Cell Cycle Checkpoints 85

4.6 - Methods for Synchronizing Mammalian Cells 86

4.7 - Analysis of the Mammalian Cell Cycle by Flow Cytometry 88

4.8 – Conclusion 89

CONTENTS

EVALUATION OF THE EFFECT OF P53 IN CELLULAR RESPONSE FROM ELECTRON MICROSCOPY IMAGES iv

CHAPTER V – MATERIALS AND METHODS 92


5.2 - Materials and Methods of the paper “Lack of p53 function promotes

radiation-induced mitotic catastrophe in mouse embryonic fibroblast cells” 94

5.2.1 - Cell Culture 94

5.2.2 - Light microscopy 94

5.2.3 - Bivariate BrdUrd-PI (bromodeoxiuridine-propidium iodide)

flow cytometry 95

5.2.4 - Bivariate cyclin B1-PI flow cytometry 95

5.2.5 - Western blotting 95

5.3 - Materials of the paper “Nuclear accumulation and activation of p53

in embryonic stem cells after DNA damage” 96

5.3.1 - Cell lines and their treatments 96

5.3.2 - Immunofluorescence staining 97

5.3.3 - RT-PCR 97

5.3.4 - Beta-Galactosidase staining 97

5.3.5 - MTT-assay 98

5.3.6 - Colony assay 98

5.3.7 - Apoptosis Assay by Annexin V staining 98

CHAPTER VI – IMAGE PROCESSING AND ANALYSIS 99


6.2 - Image processing and Analysis 102

CONTENTS

EVALUATION OF THE EFFECT OF P53 IN CELLULAR RESPONSE FROM ELECTRON MICROSCOPY IMAGES v

6.3 - Comparison between original and processed images 112

6.4 - Summary 129

CHAPTER VII – CONCLUSIONS AND FUTURE WORKS 131

7.1 - Final Conclusions 133

7.2 - Future Works 134

REFERENCES 135

CHAPTER I

INTRODUCTION TO THE THEME AND REPORT ORGANIZATION

CHAPTER I – INTRODUCTION TO THE THEME AND REPORT ORGANIZATION

EVALUATION OF THE EFFECT OF P53 IN CELLULAR RESPONSE FROM ELECTRON MICROSCOPY IMAGES 3

1.1 – INTRODUCTION

One of the most widely used steps in the process of obtaining information from

images is image segmentation: dividing the input image into regions that hopefully

correspond to structural units in the scene or distinguish objects of interest (Russ,

1998).

Segmentation is often described by analogy to visual processes as a

foreground/background separation, implying that the selection procedure

concentrates on a single kind of feature and discards the rest. This is not quite true for

computer systems, which can generally deal much better than humans with scenes

containing more than one type of feature of interest (Russ, 1998).

In computational vision, segmentation refers to the process of partitioning a

digital image into multiple segments (sets of pixels, also known as superpixels). The

goal of segmentation is to simplify and/or change the representation of an image into

something that is more meaningful and easier to analyze. Image segmentation is

typically used to locate objects and boundaries (lines, curves, etc.) in images. More

precisely, image segmentation is the process of assigning a label to every pixel in an

image such that pixels with the same label share certain visual characteristics. The

result of image segmentation is a set of segments that collectively cover the entire

image, or a set of contours extracted from the image. Each of the pixels in a region is

similar with respect to some characteristic or computed property, such as color,

intensity, or texture. Adjacent regions are significantly different with respect to the

same characteristic(s) (Shapiro, 2001).

In this work, the goal is to study the morphology changes of irradiated cells,

more precisely the cellular changes in p53 content upon irradiation. Therefore, the

cells images are studied and processed using the image processing toolbox of the

MATLAB program, with the intent of highlight the cellular differences between the

control and irradiated cells in terms of the p53 quantity inside the cell.



1.2 – MAIN OBJECTIVES

The study report in the paper: “Nuclear accumulation and activation of p53 in

embryonic stem cells after DNA damage”, of Valeriya Solozobova, ALexandre

Rolletschek and Christine Blattner, BMC Cell Biology 2009 10:46, is based on the fact

that:

Cells are continuously subjected to DNA lesions arising both from

environmental conditions and from the intrinsic metabolism of a cell.

Such lesions can lead to mutations and large-scale genome alterations

that may be deleterious for cellular function. To maintain genomic

stability cell cycle checkpoints exist that can detect errors during DNA

replication. If errors are encountered, cell division is paused and repair

mechanisms and/or cell death ensues.

The p53 tumor suppressor protein plays an important role in this

process. By being part of a signal transduction process, p53 relays

information leading to cellular responses such as cell cycle arrest and

apoptosis, resulting from DNA lesions. P53 activity is regulated mainly at

the protein level. In response to DNA lesions, p53 is rescued from

targeted degradation, which leads to a strong increase in the amount of

the otherwise short lived tumor suppressor protein, and the protein is

intensively modified. Cells deficient in p53 fail to undergo apoptosis or

cell cycle arrest in response to DNA damage which increases the rates of

tumorigenicity and genomic instability in these animals.

Whereas the study report in the paper: “Lack of p53 function promotes

radiation-induced mitotic catastrophe in mouse embryonic fibroblast cells”, of

Fiorenza Ianzini, Alessandro Bertoldo, Elizabeth A Kosmacek, Stacia L Philips and

Michael A Mackey, Cancer Cell International 2006 6:11, is based on the fact that:

Mitotic catastrophe (MC) has been observed following alterations in

specific cellular proteins, or by treatment of cells with chemicals, heat,

and/or ionizing radiation. MC is characterized by an aberrant nuclear

morphology observed following premature entry into mitosis and often

results in the generation of aneuploid and polyploidy cell progeny. The



initiating event in this process involves the premature entry of cells into

mitosis; cells undergo a spontaneous chromosome condensation that

produces chromosome morphologies very similar to those observed

when metaphase cells are fused with cells located late in the cell cycle.

Thus, one consider these abnormal mitotic figures as indicative of cells

undergoing spontaneous premature chromosome condensation (SPCC).

These cells entering into mitosis prematurely often either fail to achieve

cytokinesis or divide and fuse shortly thereafter, and later exhibit the

features of MC.

These cells almost always die; however, some studies have suggested

that a small fraction of cells might survive long enough to establish a

growing population of cells, and one study demonstrated a high

frequency of surviving clones containing an elevated incidence of MC.

These results may indicate that a small fraction of cells can survive MC.

Stress-induced SPCC and subsequent MC is observed under conditions

where cyclin B1/cdc2 kinase is activated while cells are delayed in S or

G2 phases, indicating that stress-induced MC is the result of abrogation

of cell cycle regulatory pathways, in particular G2 checkpoint pathways.

There are many proteins that play a role in the regulation of checkpoint

functions in G2, both inhibitory and stimulatory abrogation of the G2/M

checkpoint, due to over accumulation of cyclin B1 protein and

premature activation of cyclin B1/cdc2 kinase, plays a critical role in the

induction of SPCC and subsequent MC. Cyclin B1 biosynthesis

contributes to the regulation of mitotic entry, as cyclin B1 levels are cell

cycle regulated, with the gene being expressed only in S and G2 phases

in human and rodent cells.

At the later stages of mitosis, proteosome-mediated degradation of

cyclin B1 begins, and new cyclin B1 synthesis is required for entry into

the next mitosis. Thus, the cyclic rise and fall of cyclin B1 levels provides

for one level of regulation of this promitotic protein. Cells arrested late

in the cell cycle are located at that point in the cycle when cyclin B1

gene expression is at its peak value. Under these conditions it has been



shown that the p53 tumor suppressor gene product is a negative

regulator of cyclin B1 transcription, perhaps providing for negative

feedback regulation of cyclin B1 levels under abnormal conditions. If the

induction of MC in cells post-irradiation is due to cyclin B1 over

accumulation, a role for p53 in this response might be expected.

In this study one present data which describe such a role for p53 in the

induction of MC mediated by over accumulation of cyclin B1 occurring

during delay of cells late in the cell cycle.

In both studies, the role of the p53 is enhanced and, based on the results of

these studies I will withdraw information from the resulting images supported on the

image processing of them.

In the end of this practical work, the main goal is to identify, study and compare

techniques of image processing and analysis to performing the extraction of relevant

information from the images contained in the above mentioned papers, in order to

validate their results in an automate and robust manner.

1.3 – REPORT ORGANIZATION

It was intended to organize this document in a self-directed and self-regulating

approach to improve the access to various topics structured in seven chapters. So, it

will be described very succinctly what is treated in each remaining chapter:

Chapter II – Cell cycle regulation and apoptosis

In this chapter takes place a description of key concepts related to the cell cycle

checkpoints, to the behavior of the malignant cells and to the cellular death

mechanisms among other information related to the normal and malignant cells.



Chapter III – Radiation effect on normal and neoplastic tissues

In this chapter it is presented a description of the irradiated carcinogenesis as

well as the cell death mechanisms. It is also described important issues regarding the

cellular behavior upon irradiation.

Chapter IV – Cell culture and flow cytometry

In this fourth chapter it is performed an approach of some important issues

regarding the safety manipulation and maintenance of cells when performing cell

culture techniques. It is also described the methods to induce cell cycle checkpoints

and the flow cytometry technique.

Chapter V – Materials and Methods

In this fifth chapter a description on the materials and methods used in the

articles in which this work is based on.

Chapter VI – Image Processing and Analysis

In this chapter it is performed the analysis and the segmentation of the images,

using the MATLAB image processing toolbox, obtained from the papers used as based

for the execution of this practical work.

Chapter VII – Final Conclusions and Future Works

In the last chapter it is presented the final conclusions of the work performed,

as well as the future perspectives regarding the execution of the correspondent thesis.



1.4 – MAJOR CONTRIBUTIONS

This work has helped to highlight the contribution of techniques image

processing and analysis to obtain additional and complementary information to the

two papers used as basis. Additionally, knowledge about cell cycle regulation and

checkpoints that help to understand the behavior of cells when they are irradiated was

gained. This information will be helpful to study the electron microscopy images of

breast cancer cells submitted to brachytherapy for the thesis work.

CHAPTER II

CELL CYCLE REGULATION AND APOPTOSIS

CHAPTER II – CELL CYCLE REGULATION AND APOPTOSIS


2.1 - INTRODUCTION

The development of knowledge about the biochemistry and cell biology of

cancer comes from a number of disciplines. Some of this knowledge has come from

research initiated a century or more ago. There has been a flow of information about

genetics into a knowledge base about cancer, starting with Gregor Mendel and the

discovery of the principle of inherited traits and leading through Theodor Boveri’s work

on the chromosomal mode of heredity and chromosomal damage in malignant cells to

Avery’s discovery of DNA as the hereditary principle, Watson and Crick’s determination

of the structure of DNA, the human genome project, DNA microarrays, and

proteomics. Not only has this information provided a clearer understanding of the

carcinogenic process, it has also provided better diagnostic approaches and new

therapeutic targets for anticancer therapies (Ruddon, 2007).

Cancer cells contain many alterations, which accumulate as tumors develop.

Over the last 25 years, considerable information has been gathered on the regulation

of cell growth and proliferation leading to the identification of the proto-oncogenes

and the tumor suppressor genes. The proto-oncogenes encode proteins, which are

important in the control of cell proliferation, differentiation, cell cycle control and

apoptosis. Mutations in these genes act dominantly and lead to a gain in function. In

contrast the tumor suppressor genes inhibit cell proliferation by arresting progression

through the cell cycle and block differentiation. They are recessive at the level of the

cell although they show a dominant mode of inheritance. In addition, other genes are

also important in the development of tumors. Mutations leading to increase genomic

instability suggest defects in mismatch and excision repair pathways. Genes involved in

DNA repair, when mutated, also predispose the patient to developing cancer

(Macdonald, 2005).

A crucial decision in every proliferating cell is the decision to continue with a

further round of cell division or to exit the cell cycle and return to the stationary phase.

Similarly quiescent cells must make the decision, whether to remain in the stationary

phase (G0) or to enter into the cell cycle. Entry into the cycle occurs in response to

mitogenic signals and exit in response to withdrawal of these signals. To ensure that



DNA replication is complete and that any damaged DNA is repaired, cells must pass

through specific checkpoints. Tumor cells undergo uncontrolled proliferation either

due to mutations in the signal transduction pathways or because of mutations in the

regulatory mechanism of the cell cycle (Macdonald, 2005).

In this chapter, it is provided a detailed description of the cell cycle, its

progression and the cellular events involved in transforming normal cells into

malignant cells. For this purpose, the chapter starts with the explanation of the cell

cycle followed by the description of the progression of the cell cycle, the growth

characteristics of the malignant cells and the cell cycle regulation. After this, the

chapter focuses the importance of the apoptosis phenomena and ends referring the

resistance to apoptosis in cancer cells and potential targets for therapy.

2.2 – CELL LIFE CYCLE

The cell life cycle includes the changes a cell undergoes from the time it is

formed until it divides to produce two new cells. The life cycle of a cell has two stages,

an interphase and a cell division stage, Figure 2.1 (Seelev, 2004).

Figure 2.1 – Cell cycle (from (Seeley, 2004))

2.2.1 – Interphase

Interphase is the phase between cell divisions. Ninety percent or more of the

life cycle of a typical cell is spent in interphase and, during this time the cell carries out



the metabolic activities necessary for life and performs its specialized functions such as

secreting digestive enzymes. In addition, the cell prepares to divide which includes an

increase in cell size; because many cell components double in quantity, and a

replication of the cell’s DNA. Consequently, the centrioles within the centrosome are

also duplicated, when the cell divides, each new cell receives the organelles and DNA

necessary for continued functioning. Interphase can be divided into three subphases,

called G1, S, and G2. During G1 (the first gap phase) and G2 (the second gap phase), the

cell carries out routine metabolic activities. During the S phase (the synthesis phase),

the DNA is replicated (new DNA is synthesized) (Seelev, 2004).

Many cells in the human body do not divide for days, months, or even years.

These “resting” cells exit and enter the cell cycle that is called the G0 phase, in which

they remain, unless, stimulated to divide (Seelev, 2004).

2.2.2 - DNA Replication

DNA replication is the process by which two new strands of DNA are made,

using the two existing strands as templates. During interphase, DNA and its associated

proteins appear as dispersed chromatin threads within the nucleus. When DNA

replication begins, the two strands of each DNA molecule separate from each other for

some distance, Figure 2.2. Then, each strand functions as a template, or pattern, for

the production of a new strand of DNA, which is formed as new nucleotides pair with

the existing nucleotides of each strand of the separated DNA molecule. The production

of the new nucleotide strands is catalyzed by DNA polymerase, which adds new

nucleotides at the 3` end of the growing strands. One strand, called the leading strand,

is formed as a continuous strand, whereas the other strand, called the lagging strand,

is formed in short segments going in the opposite direction. The short segments are

then spliced by DNA ligase. As a result of DNA replication, two identical DNA molecules

are produced, each of them having one strand of nucleotides derived from the original

DNA molecule and one newly synthesized strand (Seelev, 2004).



Figure 2.2 – Replication of DNA (from (Seelev, 2004))

2.2.3 - Cell Division

New cells necessary for growth and tissue repair are produced by cell division.

A parent cell divides to form two daughter cells, each of which has the same amount

and type of DNA as the parent cell. Because DNA determines cell structure and

function, the daughter cells have identical structure and perform the same functions as

the parent cell. Cell division involves two major events: the division of the nucleus to

form two new nuclei, and the division of the cytoplasm to form two new cells. Each of

the new cells contains one of the newly formed nuclei. The division of the nucleus

occurs by mitosis, and the division of the cytoplasm is called cytokinesis (Seelev, 2004).

2.2.3.1 - Mitosis

Mitosis is the division of the nucleus into two nuclei, each of which has the

same amount and type of DNA as the original nucleus. The DNA, which was dispersed

as chromatin in interphase, condenses in mitosis to form chromosomes. All human

somatic cells, which include all cells except the sex cells, contain 46 chromosomes,

which are referred to as a diploid number of chromosomes. Sex cells have half the

number of chromosomes as somatic cells (Seelev, 2004).

The 46 chromosomes in somatic cells are organized into 23 pairs of

chromosomes. Twenty-two of these pairs are called autosomes. Each member of an



autosomal pair of chromosomes looks structurally alike, and together they are called a

homologous pair of chromosomes. One member of each autosomal pair is derived

from the person’s father, and the other is derived from the mother. The remaining pair

of chromosomes is the sex chromosomes. In females, the sex chromosomes look alike,

and each is called an X chromosome. In males, the sex chromosomes do not look

similar. One chromosome is an X chromosome, and the other is smaller and is called a

Y chromosome. One X chromosome of a female is derived from her mother and the

other is derived from her father. The X chromosome of a male is derived from his

mother and the Y chromosome is derived from his father (Seelev, 2004).

Mitosis is divided into four phases: prophase, metaphase, anaphase, and

telophase. Although each phase represents major events, mitosis is a continuous

process, and no discrete jumps occur from one phase to another. Learning the

characteristics associated with each phase is helpful, but a more important concept is

how each daughter cell obtains the same number and type of chromosomes as the

parent cell. The major events of mitosis are summarized in Figure 2.3 (Seelev, 2004).

Figure 2.3 – Mitosis. (1) Interphase; (2) Prophase; (3) Metaphase; (4) Anaphase; (5) Telophase; (6) Interphase,

Cytokinesis (from (Seelev, 2004))



2.2.3.2 - Cytokinesis

Cytokinesis is the division of the cytoplasm of the cell to produce two new cells

(Figure 2.3). Cytokinesis begins in anaphase continues through telophase and ends in

the following interphase. The first sign of cytokinesis is the formation of a cleavage

furrow, or puckering of the plasma membrane, which forms midway between the

centrioles. A contractile ring composed primarily of actin filaments pulls the plasma

membrane inward, dividing the cell into two halves. Cytokinesis is complete when the

membranes of the two halves separate at the cleavage furrow to form two separate

cells (Seelev, 2004).

2.2.4 – Meiosis

All cells of the body are formed by mitosis, except sex cells that are formed by

meiosis. In meiosis the nucleus undergoes two divisions resulting in four nuclei, each

containing half as many chromosomes as the parent cell. The daughter cells that are

produced by cytokinesis differentiate into gametes, or sex cells.

The gametes are reproductive cells—sperm cells in males and oocytes (egg

cells) in females. Each gamete not only has half the number of chromosomes found in

a somatic cell but also has one chromosome from each of the homologous pairs

verified in the parent cell. The complement of chromosomes in a gamete is referred to

as a haploid number. Oocytes contain one autosomal chromosome from each of the

22 homologous pairs and an X chromosome. Sperm cells have 22 autosomal

chromosomes and either an X or Y chromosome. During fertilization, when a sperm

cell fuses with an oocyte, the normal number of 46 chromosomes in 23 pairs is

reestablished. The sex of the baby is determined by the sperm cell that fertilizes the

oocyte. The sex is male if a Y chromosome is carried by the sperm cell that fertilizes the

oocyte and female if the sperm cell carries an X chromosome (Seelev, 2004).

The first division during meiosis is divided into four phases: prophase I,

metaphase I, anaphase I, and telophase I, Figure 2.4. As in prophase of mitosis, the

nuclear envelope degenerates, spindle fibers form, and the already duplicated

chromosomes become visible. Each chromosome consists of two chromatids joined by

a centromere. In prophase I, however, the four chromatids of a homologous pair of



chromosomes join together, or synapse, to form a tetrad. In metaphase I the tetrads

align at the equatorial plane and in anaphase I each pair of homologous chromosomes

separate and move toward opposite poles of the cell (Seelev, 2004).

For each pair of homologous chromosomes, one daughter cell receives one

member of the pair, and the other daughter cell receives the other member. Thus each

daughter cell has 23 chromosomes, each of which is composed of two chromatids.

Telophase I with cytokinesis is similar to telophase of mitosis and two daughter cells

are produced. Interkinesis is the phase between the formation of the daughter cells

and the second meiotic division. No duplication of DNA occurs during this phase. The

second division of meiosis also has four phases: prophase II, metaphase II, anaphase II,

and telophase II. These stages occur much as they do in mitosis, except that 23

chromosomes are present instead of 46 (Seelev, 2004).

The chromosomes align at the equatorial plane in metaphase II, and their

chromatids split apart in anaphase II. The chromatids then are called chromosomes,

and each new cell receives 23 chromosomes. In addition to reducing the number of

chromosomes in a cell from 46 to 23, meiosis is also responsible for genetic diversity

for two reasons:

A random distribution of the chromosomes is received from each

parent. One member of each homologous pair of chromosomes was

derived from the person’s father and the other member from the

person’s mother. The homologous chromosomes align randomly during

metaphase I when they split apart, each daughter cell receives some of

the father’s and some of the mother’s chromosomes. The number of

chromosomes each daughter cell receives from each parent is

determined by chance;

However, when tetrads are formed, some of the chromatids may break

apart, and part of one chromatid from one homologous pair may be

exchanged for part of another chromatid from the other homologous

pair, Figure 2.5. This exchange is called crossing-over; as a result,

chromatids with different DNA content are formed, Figure 2.5.



With random assortment of homologous chromosomes and crossing-over, the

possible number of gametes with different genetic makeup is practically unlimited.

When the distinct gametes of two individuals unite, it is virtually certain that the

resulting genetic makeup never has occurred before and never will occur again. The

genetic makeup of each new human being is unique (Seelev, 2004).

Figure 2.4 – Meiosis (from (Seelev, 2004))



Figure 2.5 – Crossing-over (from (Seelev, 2004))

2.3 - PROGRESSION OF THE CELL CYCLE

The cell cycle is controlled by a complex pattern of synthesis and degradation of

regulators together with careful control of their spatial organization in specific

subcellular compartments. In addition, checkpoint controls can modulate the

progression of the cycle in response to adverse conditions such as DNA damage.

Cells either enter G1 from G0 in response to mitogenic stimulation or follow on

from cytokinesis if actively proliferating (i.e. from M to G1). Removal of mitogens

allows them to return to G0. The critical point between mitogen dependence and

independence is the restriction point or R which occurs during G1. It is here that cells

reach the ‘point of no return’ and are committed to a round of replication (Macdonald,

2005), Figure 2.6.

Figure 2.6 – Restriction point, R (from (Griffiths, 1999))



Synthesis of the D-type cyclins begins at the G0/G1 transition and continues so

long as growth factor stimulation persists. This mitogen stimulation of cyclin D is in

part dependent on RAS activation, a role which is highlighted by the ability of anti-RAS

antibodies to block the progression of the cell cycle if added to cells prior to mitogen

stimulation. The availability of cyclin D activates CDK4 and 6 and these complexes then

drive the cell from early G1 through R to late G1; largely by regulation of RB which

exists in a phosphorylated state at the start of G1 complexed to a large number of

proteins. Cyclin D-CDK4/6 activation begins phosphorylation of Rb during early G1. This

initial phosphorylation leads to release of histone deacetylase activity from the

complex alleviating transcriptional repression. The E2F transcription factor remains

bound to Rb at this stage but can still transcribe some genes including cyclin E.

Therefore, levels of cyclin E increase and lead to activation of CDK2, which can then

complete phosphorylation of Rb. Consequently, complete phosphorylation of Rb

results in the release of E2F to activate genes required to drive cells through the G1/S

transition (Macdonald, 2005), Figure 2.7.

Figure 2.7 – Regulation of the G1 to S transition (from (Griffiths, 1999))

The CKIs also play a role in control of cell cycle progression at this stage and in

response to antimitogenic signals, oppose the activity of the CDKs and cause cell cycle

arrest. INK4 inhibitors bind to CDK4/6 to prevent cyclin D binding and CIP/KIP

inhibitors similarly inhibit the kinase activity of cyclin ECDK2, Figure 2.8. CIP/KIP

inhibitors also interact with cyclin D-CDK4/6 complexes during G1, but rather than

blocking cell cycle progression, this interaction is required for the complete function of



the complex and allows G1 progression. This interaction sequesters CIP/KIP, preventing

its inhibition of cyclin E-CDK2 and thereby facilitating its full activation to contribute to

G1 progression. In the presence of an antimitogenic signal, levels of cyclin D-CDK4/6

are reduced, CIP/KIP is released, which can then interact with and inhibit CDK2 to

cause cell cycle arrest (Macdonald, 2005).

Cells which have suffered DNA damage are prevented from entering S phase

and are blocked at G1. This process is dependent on the tumor suppressor gene p53

and p21. Activation of p53 by DNA damage results in increased p21 levels which can

then inactivate cyclin E-CDK2 to prevent phosphorylation of Rb and inhibit the release

of E2F to promote transcription of genes involved in DNA synthesis, Figure 2.8. This

causes the cell cycle to arrest in G1. Clearly, loss or mutation of p53 will lead to loss of

this checkpoint control and cells will be able to enter S phase with damaged DNA. After

cells have entered S phase, cyclin E is rapidly degraded and CDK2 is released. In S

phase, a further set of cyclins and CDKs, cyclin A-CDK2, are required for continued DNA

replication. Two A-type cyclins have been identified to date: cyclin A1 is expressed

during meiosis and in early cleavage embryos whereas cyclin A2 is present in all

proliferating cells. Cyclin A2 is also induced by E2F and is expressed from S phase

through G2 and M until prometaphase when it is degraded by ubiquitin-dependent

proteolysis (Macdonald, 2005).

Cyclin A2 binds to two different CDKs. Initially, during S phase, it is found

complexed to CDK2 following its release from cyclin E and subsequently in G2 and M it

is found complexed to CDC2 (also known as CDK1). Cyclin A2 has a role in both

transcriptional regulation and DNA replication and its nuclear localization is crucial to

its function. Cyclin A regulates the E2F transcription factor and in S phase, when E2F

directed transcription is no longer required, cyclin A directs its phosphorylation by

CDK2 leading to its degradation. This down-regulation by cyclin A2 is required for

orderly S phase progression and in its absence apoptosis occurs. Recently, cyclin A as

well as cyclin E have been shown to be regulators of centrosome replication and are

able to do so because of their ability to shuttle between nucleus and cytoplasm, Figure

2.9 (Macdonald, 2005).



Figure 2.8 – Cell cycle arrest at G1/S, mediated by cdk inhibitors (from (Shapiro, 1999))

Figure 2.9 – Dynamics of the DNA synthesome (from (Frouin, 2003))



The final phase of the cycle is M phase, that comprises mitosis and cytokinesis.

The purpose of mitosis is to segregate sister chromatids into two daughter cells so that

each cell receives a complete set of chromosomes, a process that requires the

assembly of the mitotic spindle. Mitosis is split into a number of stages that includes

prophase, prometaphase, metaphase, anaphase and telophase (Macdonald, 2005).

Cytokinesis, the process of cytoplasmic cleavage, follows the end of mitosis and

its regulation is closely linked to mitotic progression. Mitosis involves the last of

cyclin/CDKs, cyclin B1 and CDC2 as well as additional mitotic kinases. These include

members of the Polo family (PLK1), the aurora family (aurora A, B and C) and the NIMA

family (NEK2) plus kinases implicated at the mitotic checkpoints (BUB1), mitotic exit

and cytokinesis (Macdonald, 2005).

Entry into the final phase of the cell cycle, mitosis, is signaled by the activation

of the cyclin B1-CDC2 complex also known as the M phase promoting factor or MPF.

This complex accumulates during S and G2, but is kept in the inactive state by

phosphorylation of tyrosine 15 and threonine 14 residues on CDC2 by two kinases,

WEE1 and MYT1. WEE1 is nuclear and phosphorylates tyrosine 15, whereas MYT1 is

cytoplasmic and phosphorylates threonine 14. At the end of G2, the CDC25

phosphatase is stimulated to dephosphorylate these residues thereby activating CDC2.

These enzymes are all controlled by DNA structure checkpoints which delay the onset

of mitosis if DNA is damaged. Regulation of cyclin B1-CDC2 is also regulated by

localization of specific subcellular compartments. It is initially localized to the

cytoplasm during G2, but is translocated to the nucleus at the beginning of mitosis. A

second cyclin B, cyclin B2, also exists in mammalian cells and is localized to the Golgi

and endoplasmic reticulum where it may play a role in disassembly of the Golgi

apparatus at mitosis (Macdonald, 2005).

A further checkpoint exists at the end of G2 which checks that DNA is not

damaged before entry into M. Once more p21 activation by p53 can arrest the cell

cycle as at the end of G1. In addition, the CHK1 kinase can phosphorylate CDC25 to

create a binding site for the 14–3–3 protein, a process which inactivates CDC25,

thereby preventing dephosphorylation of CDC2 and halting the cell cycle, Figure 2.10

(Macdonald, 2005).



Tumor cells can enter mitosis with damaged DNA, suggesting a defect in the

G2/M checkpoint. Tumor cell lines have been shown to activate the cyclin B-CDC2

complex irrespective of the state of the DNA. Activation of cyclin B1-CDC2 leads to

phosphorylation of numerous substrates including the nuclear lamins, microtubule-

binding proteins, condensins and Golgi matrix components that are all needed for

nuclear envelope breakdown, centrosome separation, spindle assembly, chromosome

condensation and Golgi fragmentation respectively. During prophase, the

centrosomes—structures which organize the microtubules and which were duplicated

during G2—separate to define the poles of the future spindle apparatus, a process

regulated by several kinases including the NIMA family member NEK2, as well as

aurora A. At the same time centrosomes begin nucleating the microtubules which

make up the mitotic spindle (Macdonald, 2005).

Chromatin condensation also occurs accompanied by extensive histone

phosphorylation to produce well defined chromosomes. Nuclear envelope breakdown

occurs shortly after centrosome separation. The nuclear envelope is normally

stabilized by a structure known as the nuclear lamin which is composed of lamin

intermediate filament proteins. This envelope is broken down as a result of

hyperphosphorylation of lamins by cyclin B-CDC2 (Macdonald, 2005).

During prometaphase, the microtubules are captured by kinetochores, the

structure which binds to the centromere of the chromosome. Paired sister chromatids

interact with the microtubules emanating from opposite poles resulting in a stable

bipolar attachment. Chromosomes then sit on the metaphase plate where they

oscillate during metaphase. Once all bipolar attachments are complete anaphase is

triggered. This is characterized by simultaneous separation of all sister chromatids.

Each chromosome must be aligned in the center of the bipolar spindle such that its

two sister chromatids are attached to opposite poles. If this is correct, the anaphase-

promoting complex (APC) together with CDC20 is activated to control degradation of

proteins such as securin. This in turn activates the separin protease which cleaves the

cohesion molecules between the sister chromatids allowing them to separate. At this

stage, there is one final checkpoint, the spindle assembly checkpoint, at the

metaphase to anaphase transition, which checks the correct assembly of the mitotic



apparatus and the alignment of chromosomes on the metaphase plate. The

gatekeeper at this checkpoint is the APC complex. Unaligned kinetochores are

recognized and associate with the MAD2 and BUB proteins which can prevent

activation of APC and cell arrest at metaphase preventing exit from mitosis. In tumor

cell abnormalities of spindle formation are found, suggesting that checkpoint control is

lost (Macdonald, 2005).

Mitotic exit requires that sister chromatids have separated to opposite poles.

During telophase, nuclear envelopes can begin to form around the daughter

chromosomes and chromatin decondensation occurs. The spindle is also disassembled

and cytokinesis is completed. The control of these processes requires destruction of

both the cyclins and other kinases such as NIMA and aurora family members by

ubiquitin dependent proteolysis mediated by APC. Daughter cells can now re-enter the

cell cycle (Macdonald, 2005).

Figure 2.10 – Cell cycle regulation of cyclin dependent kinase (Cdk1) Cyclin-B (CycB) complex (from (Novák, 2010))



2.4 - GROWTH CHARACTERISTICS OF MALIGNANT CELLS

Cancer can be characterized as a disease of genetic instability, altered cellular

behavior and altered cell–extracellular matrix interactions. These alterations lead to

dysregulated cell proliferation, and ultimately to invasion and metastasis. There are

interactions between the genes involved in these steps. For example, the genes

associated with loss of control of cell proliferation may also be involved in genetic

instability (rapidly proliferating cells have less time to repair DNA damage) and tumor

vascularization that leads to dysregulated proliferation of cells, which in turn eats up

more oxygen, creates hypoxia, and turns on HIF-1 and additional angiogenesis.

Similarly, genes involved in tumor cell invasion may also be involved in loss of growth

control (invasive cells have acquired the skills to survive in ‘‘hostile’’ new

environments) and evasion of apoptosis (less cell death even in the face of a normal

rate of cell proliferation produces more cells). The molecular genetic alterations of

cancer cells lead to cells that can generate their own growth-promoting signals are less

sensitive to cell cycle checkpoint controls, evade apoptosis, and thus have almost

limitless replication potential. This redundancy makes design of effective signal

transduction-targeted chemotherapeutic drugs that target a single pathway very

difficult indeed (Ruddon, 2007).

Cancer cells can also subvert the environment in which they proliferate.

Alterations in both cell–cell and cell–extracellular matrix interactions also occur,

leading to creation of a cancer-facilitating environment. For example, a common

alteration in epithelial carcinomas is alteration of E-cadherin expression, which is a

cell–cell adhesion molecule found on all epithelial cells. Cancer cells exhibit remarkable

plasticity and have the ability to mimic some of the characteristics of other cell types

as they progress and became less well differentiated. For example, cancer cells may

assume some of the structure and function of vascular cells. As cancer cells

metastasize, they may eventually take on a new phenotype such that the tissues of

origin may become unclear—so-called cancers of unknown primary site (Ruddon,

2007).



2.4.1 - Phenotypic Alterations in Cancer Cells

Treatment of animals or cells in culture with carcinogenic agents is a means of

studying discrete biochemical events that lead to malignant transformation, Figure

2.11. However, studies of cell transformation in vitro have many pitfalls. These ‘‘tissue

culture artifacts’’ include overgrowth of cells not characteristic of the original

population of cultured cells (e.g., overgrowth of fibroblasts in cultures that were

originally primarily epithelial cells), selection for a small population of variant cells with

continued passage in vitro, or appearance of cells with an abnormal chromosomal

number or structure (karyotype). Such changes in the characteristics of cultured cell

populations can lead to ‘‘spontaneous’’ transformation that mimics some of the

changes seen in populations of cultured cells treated with oncogenic agents. Thus, it is

often difficult to sort out the critical malignant events from the noncritical ones

(Ruddon, 2007).

Figure 2.11 – Cellular response (from (Gil, 2006))

Although closer to the carcinogenic process in humans, malignant

transformation induced in vivo by treatment of susceptible experimental animals with

carcinogenic chemicals or oncogenic viruses or by irradiation, is even more difficult

because it is hard to discriminate toxic from malignant events and to determine what

role a myriad of factors, such as the nutritional state of the animal, hormone levels, or

endogenous infections with microorganisms or parasites, might have on the in vivo

carcinogenic events. Moreover, tissues in vivo are a mixture of cell types, and it is



difficult to determine in which cells the critical transformation events are occurring

and what role the microenvironment of the tissue plays. Thus, most studies designed

to identify discrete biochemical events occurring in cells during malignant

transformation have been done with cultured cells, since clones of relatively

homogeneous cell populations can be studied and the cellular environment defined

and manipulated. The ultimate criterion that establishes whether cells have been

transformed, however, is their ability to form a tumor in an appropriate host animal.

The generation of immortalized ‘‘normal’’ cell lines of a given differentiated phenotype

from human embryonic stem cells, has enhanced the ability to study cells of a normal

genotype from a single source. Such cell lines may also be generated by transfection of

the telomerase gene into cells to maintain chromosomal length (Ruddon, 2007).

Over the past 60 years, much scientific effort has gone into research aimed at

identifying the phenotypic characteristics of in vitro transformed cells that correlate

with the growth of a cancer in vivo. This research has tremendously increased our

knowledge of the biochemistry of cancer cells. However, many of the biochemical

characteristics initially thought to be closely associated with the malignant phenotype

of cells in culture has subsequently been found to be dissociable from the ability of

those cells to produce tumors in animals. Furthermore, individual cells of malignant

tumors growing in animals or in humans exhibit marked biochemical heterogeneity, as

reflected in their cell surface composition, enzyme levels, immunogenicity, response to

anticancer drugs, and so on. This has made it extremely difficult to identify the

essential changes that produce the malignant phenotype (Ruddon, 2007).

2.4.2 - Immortality of Transformed Cells in Culture

Most normal diploid mammalian cells have a limited life expectancy in culture.

For example, normal human fibroblast lines may live for 50 to 60 population doublings

(the ‘‘Hayflick index’’), but then viability begins to decrease rapidly, unless they

transform spontaneously or are transformed by oncogenic agents. However, malignant

cells, once they become established in culture, will generally live for an indefinite

number of population doublings, provided the right nutrients and growth factors

(Ruddon, 2007).



It is not clear what limits the life expectancy of normal diploid cells in culture,

but it may be related to the continual shortening of chromosomal telomeres each time

cells divide. Transformed cells are known to have elevated levels of telomerase that

maintain telomere length. Transformed cells that become established in culture also

frequently undergo karyotypic changes, usually marked by an increase in

chromosomes (polyploidy), with continual passage. This suggests that cells with

increased amounts of certain growth-promoting genes are generated and/or selected

during continual passage in culture. The more undifferentiated cells from cancers of

animals or patients also often have an atypical karyology, thus the same selection

process may be going on in vivo with progression over time of malignancy from a lower

to a higher grade (Ruddon, 2007).

2.4.3 - Decreased Requirement for Growth Factors

Other properties that distinguish transformed cells from their non transformed

counterparts are decreased density-dependent inhibition of proliferation and the

requirement for growth factors for replication in culture. Cells transformed by

oncogenic viruses have lower serum growth requirements than do normal cells. Cancer

cells may also produce their own growth factors that may be secreted and activate

proliferation in neighboring cells (paracrine effect) or, if the same malignant cell type

has both the receptor for a growth factor and the means to produce the factor, self-

stimulation of cell proliferation (autocrine effect) may occur. One example of such an

autocrine loop is the production of tumor necrosis factor-alpha (TNF-α) and its

receptor TNFR1 by diffuse large cell lymphoma. Co-expression of TNF-α and its

receptor are negative prognostic indicators of survival, suggesting that autocrine loops

can be powerful stimuli for tumor aggressiveness and thus potentially important

diagnostic and therapeutic targets.

2.4.4 - Loss of Anchorage Dependence

Most freshly isolated normal animal cells and cells from cultures of normal

diploid cells do not grow well when they are suspended in fluid or a semisolid agar gel.



However, if these cells contact with a suitable surface they attach, spread, and

proliferate. This type of growth is called anchorage-dependent growth. Many cell lines

derived from tumors and cells transformed by oncogenic agents are able to proliferate

in suspension cultures or in a semi solid medium (methylcellulose or agarose) without

attachment to a surface. This is called anchorage-independent growth and this

property of transformed cells has been used to develop clones of malignant cells. This

technique has been widely used to compare the growth properties of normal and

malignant cells. Another advantage that has been derived from the ability of malignant

cells to grow in soft agar (agarose), is the ability to grow cancer cells derived from

human tumors to test their sensitivity to chemotherapeutic agents and to screen for

potential new anticancer drugs (Ruddon, 2007).

2.4.5 - Loss of Cell Cycle Control and Resistance to Apoptosis

Normal cells respond to a variety of suboptimal growth conditions by entering a

quiescent phase in the cell division cycle, the G0 state. There appears to be a decision

point in the G1 phase of the cell cycle, at which time the cell must make a commitment

to continue into the S phase, the DNA synthesis step, or to stop in G1 and wait until

conditions are more optimal for cell replication to occur. If this waiting period is

prolonged, the cells are said to be in a G0 phase. Once cells make a commitment to

divide, they must continue through S, G2, and M to return to G1. If the cells are blocked

in S, G2, or M for any length of time, they die. The events that regulate the cell cycle

are called cell cycle checkpoints (Ruddon, 2007).

The loss of cell cycle check point control by cancer cells may contribute to their

increased susceptibility to anticancer drugs. Normal cells have mechanisms to protect

themselves from exposure to growth-limiting conditions or toxic agents by calling on

these check point control mechanisms. Cancer cells, by contrast, can continue through

these checkpoints into cell cycle phases that make them more susceptible to the

cytotoxic effects of drugs or irradiation. For example, if normal cells accrue DNA

damage due to ultraviolet (UV) or X-irradiation, they arrest in G1 so that the damaged

DNA can be repaired prior to DNA replication. Another check point in the G2 phase

allows repair of chromosome breaks before chromosomes are segregated at mitosis,



Figure 2.12. Cancer cells, which exhibit poor or absent check point controls, proceed to

replicate the damaged DNA, thus accounting for persisting and accumulating

mutations (Ruddon, 2007).

2.5 - CELL CYCLE REGULATION

Cyclin-dependent protein kinases (CDKs), of which CDC2 is only one, are crucial

regulators of the timing and coordination of eukaryotic cell cycle events. Transient

activation of members of this family of serine/threonine kinases occurs at specific cell

cycle phases (Ruddon, 2007).

Figure 2.12 - Major pathways where Plks may play a role in intra-S-phase checkpoint in mammalian systems (from

(Suqing, 2005))

In budding yeast G1 cyclins encoded by the CLN genes, interact with and are

necessary for the activation of, the CDC2 kinase (also called p34cdc2), driving the cell

cycle through a regulatory point called START (because it is regulated by the cdc2 or

start gene) and committing cells to enter S phase. START is analogous to the G1

restriction point in mammalian cells. The CDKs work by forming active heterodimeric

complexes following binding to cyclins, their regulatory subunits. CDK2, 4, and 6, and



possibly CDK3 cooperate to push cells through G1 into S phase. CDK4 and CDK6 form

complexes with cyclins D1, D2, and D3, and these complexes are involved in

completion of G1. Cyclin D–dependent kinases accumulate in response to mitogenic

signals and this leads to phosphorylation of the Rb protein. This process is completed

by the cyclin E1- and E2-CDK2 complexes. Once cells enter S phase, cyclin E is degraded

and A1 and A2 cyclins get involved by forming a complex with CDK2. There are a

number of regulators of CDK activities; where they act in the cell cycle is depicted in

Figure 2.13 (Ruddon, 2007).

Figure 2.13 - Restriction point control and the G1-S transition (from (Ruddon, 2007))

2.5.1 - CDK Inhibitors

The inhibitors of CDKs include the Cip/Kip and INK4 family of polypeptides. The

Cip/Kip family includes p21cip1, p27kip1, and p57kip2. The actions of these proteins

are complex. Although the Cip/Kip proteins can inhibit CDK2, they are also involved in

the sequestration of cyclin D-dependent kinases that facilitates cyclin E-CDK2

activation necessary for G1/S transition (Ruddon, 2007).

The INK4 proteins target the CDK4 and CDK6 kinases, sequester them into

binary CDKINK4 complexes, and liberate bound Cip/Kip proteins. This indirectly inhibits

cyclin E–CDK and promotes cell cycle arrest. The INK4-directed arrest of the cell cycle

in G1 keeps Rb in a hypophosphorylated state and represses the expression of S-phase

genes. Four INK4 proteins have been identified: p16INK4a, p15INK4b, p18INK4c, and



p19INK4d. INKA4a loss of function occurs in a variety of cancers including pancreatic

and small cell lung carcinomas and glioblastomas. INK4a fulfills the criteria of a tumor

suppressor and appears to be the INK4 family member with the most active role in this

regard. The INK4a gene encodes another tumor suppressor protein called ARF

(p14ARF). Mice with a disrupted ARF gene have a high propensity to develop tumors,

including sarcomas, lymphomas, carcinomas, and CNS tumors. These animals

frequently die at less than 15 months of age. ARF and p53 act in the same pathway to

insure growth arrest and apoptosis in response to abnormal mitogenic signals such as

myc-induced carcinogenesis, Figure 2.14 (Ruddon, 2007).

Figure 2.14 - Regulation of the Rho pathway and the cytoskeleton by cyclin-dependent kinase (CDK) inhibitors (from

(Besson, 2004))

2.5.2 - Cyclins

The originally discovered cyclins, cyclin A and B, identified in sea urchins, act at

different phases of the cell cycle. Cyclin A is first detected near the G1/S transition and



cyclin B is first synthesized during S phase and accumulates in complexes with p34cdc2

as cells approach the G2-to-M transition. Cyclin B is then abruptly degraded during

mitosis. Thus, cyclins A and B regulate S and M phase, but do not appear to play a role

in G1 control points such as the restriction point (R point), which is the point where key

factors have accumulated to commit cells to enter S phase (Ruddon, 2007).

Three more recently discovered mammalian cyclins, C, D1, and E, are the

cyclins that regulate the key G1 and G1/S transition points. Unlike cyclins A and B,

cyclins C, D1, and E are synthesized during the G1 phase in mammalian cells. Cyclin C

levels change only slightly during the cell cycle but peak in early G1. Cyclin E peaks at

the G1–S transition, suggesting that it controls entry into S. Three distinct cyclin D

forms, D1, 2, and 3, have been discovered and are differentially expressed in different

mouse cell lineages. These D cyclins all have human counterparts and cyclin D levels

are growth factor dependent in mammalian cells: when resting cells are stimulated by

growth factors, D-type cyclin levels rise earlier than cyclin E levels, implying that they

act earlier in G1 than E cyclins. Cyclin D levels drop rapidly when growth factors are

removed from the medium of cultured cells. All of these cyclins (C, D, and E) form

complexes with, and regulate the activity of various CDKs and these complexes control

the various G1, G1–S, and G2–M transition points, Figure 2.15 (Ruddon, 2007).

Interestingly, negative growth regulators also interact with the cyclin-CDK system. For

example, TGF-b1, which inhibits proliferation of epithelial cells by interfering with G1-S

transition, reduced the stable assembly of cyclin E-CDK2 complexes in mink lung

epithelial cells, and prevented the activation of CDK2 kinase activity and the

phosphorylation of Rb. This was one of the first pieces of data suggesting that the

mammalian G1 cyclin-dependent kinases are targets for negative regulators of the cell

cycle (Ruddon, 2007).

2.5.3 - Cell Cycle Checkpoints

The role of various CDKs, cyclins, and other gene products in regulating

checkpoints at G1 to S, G2 to M, and mitotic spindle segregation have been described in

detail previously. Alterations of one or more of these checkpoint controls occur in

most, if not all, human cancers at some stage in their progression to invasive cancer. A



key player in the G1–S checkpoint system is the retinoblastoma gene Rb (Ruddon,

2007).

Figure 2.15 - Cell-cycle regulation (from (Charles, 2004))

Phosphorylation of the Rb protein by cyclin D–dependent kinase releases Rb

from the transcriptional regulator E2F and activates E2F function. Inactivation of Rb by

genetic alterations occurs in retinoblastoma and is also observed in other human

cancers, for example, small cell lung carcinomas and osteogenic sarcomas (Ruddon,

2007).

The p53 gene product is an important cell cycle checkpoint regulator at both

the G1–S and G2–M checkpoints but does not appear to be important at the mitotic

spindle checkpoint because gene knockout of p53 does not alter mitosis. The p53

tumor suppressor gene is the most frequently mutated gene in human cancer,

indicating its important role in conservation of normal cell cycle progression. One of

p53’s essential roles is to arrest cells in G1 after genotoxic damage, to allow for DNA

repair prior to DNA replication and cell division. In response to massive DNA damage,

p53 triggers the apoptotic cell death pathway. Data from short-term cell-killing assays,

using normal and minimally transformed cells, have led to the conclusion that mutated

p53 protein confers resistance to genotoxic agents (Ruddon, 2007).

The spindle assembly checkpoint machinery involves genes called bub (budding

uninhibited by benomyl) and mad (mitotic arrest deficient). There are three bub genes

and three mad genes involved in the formation of this checkpoint complex. A protein



kinase called Mps1 also functions in this checkpoint function. The chromosomal

instability, leading to aneuploidy in many human cancers, appears to be due to

defective control of the spindle assembly checkpoint. Mutant alleles of the human

bub1 gene have been observed in colorectal tumors displaying aneuploidy. Mutations

in these spindle checkpoint genes may also result in increased sensitivity to drugs that

affect microtubule function because drug-treated cancer cells do not undergo mitotic

arrest and go on to die (Ruddon, 2007).

Maintaining the integrity of the genome is a crucial task of the cell cycle

checkpoints. Two checkpoint kinases, called Chk1 and Chk2 (also called Cds1), are

involved in checkpoint controls that affect a number of genes involved in maintenance

of genome integrity. Chk1 and Chk2 are activated by DNA damage and initiate a

number of cellular defense mechanisms that modulate DNA repair pathways and slow

down the cell division cycle to allow time for repair. If DNA is not successfully mended,

the damaged cells usually undergo cell death via apoptosis. This process prevents the

defective genome from extending its paternity into daughter cells (Ruddon, 2007).

Upstream elements activating the checkpoint signaling pathways such as those

turned on by irradiation or agents causing DNA double strand breaks include the ATM

kinase, a member of the phosphatidylinositol 3-kinase (PI3K) family, which activates

Chk2 and its relative ATR kinase that activates Chk1. There is also cross talk between

ATM and ATR that mediates these responses. Chk1 and Chk2 phosphorylate CDC25A

and C, which inactivate them. In its dephosporylated state CDC25A activates the CDK2-

cyclin E complex that promotes progression through S phase. It should be noted that

this is an example of dephosphorylation rather than phosphorylation activating a key

biological function. This is in contrast to most signal transduction pathways, where the

phosphorylated state of a protein (often a kinase) is the active state and the

dephosphorylated state is the inactive one. In addition, Chk1 renders CDC25A

unstable, which also diminishes its activity. CDC25A also binds to and activates CDK1-

cyclin B, which facilitates entry into mitosis. G2 arrest induced by DNA damage induces

CDC25A degradation and, in contrast, G2 arrest is lost when CDC25A is overexpressed.

A number of proteins are now known to act as mediators of checkpoint responses by

impinging on the Chk1 and 2 pathways. These include the BRCT domain–containing



proteins 53BP1, BRCA1, and MDC1.These proteins are involved in activation of Chk1

and Chk2 by acting through protein–protein interactions that modulate the activity of

these checkpoint kinases. In general, these modulators are thought to be tumor

suppressors (Ruddon, 2007).

Chk1 and 2 have overlapping roles in cell cycle regulation, but different roles during

development. Chk1 but not Chk2 is essential for mammalian development, as

evidenced by the early embryonic lethality of Chk1 knockout mice. Chk2-deficient mice

are viable and fertile and do not have a tumor-prone phenotype unless exposed to

carcinogens, and this effect is more evident later in life. As illustrated in Figure 2.16,

there are interactions between the Chk kinases and the p53 pathway. Chk2

phosphorylates threonine-18 or serine-20 on p53, which attenuates p53’s interaction

with its inhibitor MDM2, thus contributing to p53 stabilization and activation.

However, Chk2 and p53 only have partially overlapping roles in checkpoint regulation

because not all DNA-damaging events activate both pathways, Figure 2.16 (Ruddon,

2007).

2.5.4 - Cell Cycle Regulatory Factors as Targets for Anticancer Agents

The commonly observed defects in cell cycle regulatory pathways in cancer

cells distinguish them from normal cells and provide potential targets for therapeutic

agents. One approach is to inhibit cell cycle checkpoints in combination with DNA-

damaging drugs or irradiation. The rationale for this is that normal cells have a full

complement of checkpoint controls, whereas tumor cells are defective in one or more

of these and thus are more subject to undergoing apoptosis in response to excessive

DNA damage. This has been accomplished by combining ATM/ATR inhibitors such as

caffeine or Chk1 inhibitors in combination with DNA-damaging drugs. So far this

approach has not been demonstrated clinically, and indeed is somewhat counter

intuitive, since p53 mutant tumor cells are more resistant to many chemotherapeutic

drugs. p53 is a key player in causing cell death in drug treated, DNA-damaged cells

(one exception to that is the microtubule inhibitor paclitaxel), and active, unmutated

p53 is needed for this response (Ruddon, 2007).



Figure 2.16 - Simplified scheme of cell-cycle checkpoint pathways induced in response to DNA damage (here

DSBs), with highlighted tumor suppressors shown in red and proto-oncogenes shown in green (from (Kastan,

2004))

Another approach is to target the cyclin dependent kinases directly. Alteration

of the G1–S checkpoint occurs in many human cancers. Cyclin D1 gene amplification

occurs in a subset of breast, esophageal, bladder, lung, and squamous cell carcinomas.

Cyclins D2 and D3 are overexpressed in some colorectal carcinomas. In addition, the

cyclin D–associated kinases CDK4 and CDK6 are over expressed or mutated in some



cancers. Mutations or deletions in the CDK4 and CDK6 inhibitor INK4 have been

observed in familial melanomas, and in biliary tract, esophageal, pancreatic, head and

neck, non small cell lung, and ovarian carcinomas. Inactivating mutations of CDK4

inhibitory modulators p15, p16, and p18 have been observed in a wide variety of

human cancers. Cyclin E is also amplified and overexpressed in some breast and colon

carcinomas and leukemias (Ruddon, 2007).

Human cancers have a variety of mutations in cell cycle regulatory genes. This

includes overexpression of D1 and E1 cyclins and CDKs (mainly CDK4 and CDK6) as

noted above. Loss of CDK inhibitory functions (mainly INK4a and 4b and Kip1) also

occurs, as does loss of Rb, one of the first tumor suppressor genes identified. Loss of

Kip1 function and overexpression of cyclin E1 occur frequently and are associated with

poor prognosis in breast and ovarian cancers (Ruddon, 2007).

The mitogen-stimulated proliferation of cells is mediated via a retinoblastoma

(Rb) pathway that involves phosphorylation of Rb, its dissociation form and activation

of the E2F family of transcription factors, and subsequent turn-on of genes involved in

G1–S transition and DNA synthesis. Disruption of this pathway by overexpression of

cyclin D1, loss of the INK4 inhibitor p16, mutation of CDK4 to a p16-resistant form, or

loss or mutation of Rb is frequently seen in cancer cells. The activation of CDK

inhibitory factors such as p16INK4 or p27kip1 and inhibition of cyclin dependent

kinases are, therefore, potential ways to interdict the overactive cell proliferation

pathways in cancer cells. Thus, inhibition of cyclins D1 and E and CDKs, especially CDK4

and CDK6, could be targets for inhibiting growth of cancers. As more knowledge of the

complicated steps in cell cycle regulation is gained, more potential targets become

available (Ruddon, 2007).

2.6 - APOPTOSIS

Apoptosis (sometimes called programmed cell death) is a cell suicide

mechanism that enables multicellular organisms to regulate cell number in tissues and

to eliminate unneeded or aging cells as an organism develops. The biochemistry of

apoptosis has been well studied in recent years, and the mechanisms are now

reasonably well understood (Ruddon, 2007).



The apoptosis pathway involves a series of positive and negative regulators of

proteases called caspases, which cleave substrates, such as poly (ADP-ribose)

polymerase, actin and lamin. In addition, apoptosis is accompanied by the

intranucleosomal degradation of chromosomal DNA, producing the typical DNA ladder

seen for chromatin isolated from cells undergoing apoptosis. The endonuclease

responsible for this effect is called caspase-activated DNase, or CAD (Ruddon, 2007).

A number of ‘‘death receptors’’ have also been identified, they are cell surface

receptors that transmit apoptotic signals initiated by death ligands, Figure 2.17. The

death receptors sense signals that tell the cell that it is in an uncompromising

environment and needs to die. These receptors can activate the death caspases within

seconds of ligand binding and induce apoptosis within hours. Death receptors belong

to the tumor necrosis factor (TNF) receptor gene superfamily and have the typical

cystine rich extracellular domains and an additional cytoplasmic sequence termed the

death domain (Ruddon, 2007).

Figure 2.17 - Apoptosis signaling through death receptors (from (Frederik, 2002))



The best-characterized death receptors are CD95 (also called Fas or Apo1) and

TNF receptor TNFR1 (also called p55 or CD120a). The importance of the apoptotic

pathway in cancer progression is seen when there are mutations that alter the ability

of the cell to undergo apoptosis and allow transformed cells to keep proliferating

rather than die. Such genetic alterations include the translocation of the bcl-2 gene in

lymphomas that prevents apoptosis and promotes resistance to cytotoxic drugs. Other

genes involved as players on the apoptosis stage include c-myc, p53, c-fos, and the

gene for interleukin-1b-converting enzyme (ICE). Various oncogene products can

suppress apoptosis, like the adenovirus protein E1b, ras, and n-abl (Ruddon, 2007).

Mitochondria plays a pivotal role in the events of apoptosis by at least three

mechanisms:

1) Release of proteins, e.g., cytochrome c, that triggers activation of caspases;

2) Alteration of cellular redox potential;

3) Production and release of reactive oxygen species after mitochondrial

membrane damage.

Another mitochondrial link to apoptosis is implied by the fact that Bcl-2, the

anti-apoptotic factor, is a mitochondrial membrane protein that appears to regulate

mitochondrial ion channels and proton pumps, Figure 2.18 (Ruddon, 2007).

2.6.1 - Biochemical Mechanism of Apoptosis

Multicellular organisms, from the lowest to the highest species, must have a

way to get rid of excess cells or cells that are damaged in order for the organism to

survive. Apoptosis is the mechanism that they use to do this. It is the way that the

organism controls cell numbers and tissue size and protects itself from ‘‘rogue’’ cells.

A simplified version of the apoptotic pathways can be visualized in Figure 2.19

(Ruddon, 2007).



Figure 2.18 - Apoptosis signaling through mitochondria (from (Frederik, 2002))

The death receptor–mediated pathway is turned on by members of the death

receptor superfamily of receptors including Fas receptor (CD95) and TNF receptor 1,

which are activated by Fas ligand and TNF, respectively. Interaction of these ligands

with their receptors induces receptor clustering, binding of the receptor clusters to

Fas-associated death domain protein (FADD), and activation of caspase-8, Figure 2.20.

This activation step is regulated by c-FLIP. Caspase-8, in turn, activates caspase-3 and

other ‘‘executioner’’ caspases, which induce a number of apoptotic substrates. The

DNA damage–induced pathway invokes a mitochondrial-mediated cell death pathway

that involves pro-apoptotic factors like Bax (blocked by the anti-apoptotic protein Bcl-

2). This results in cytochrome c release from the mitochondria and triggering of

downstream effects facilitating caspase-3 activation, which is where the two pathways

intersect. There are both positive and negative regulators that also interact on these

pathways (Ruddon, 2007).



Figure 2.19 - The two main apoptotic signaling pathways (from (Frederik, 2002))

Figure 2.20 - Illustration of the main TNF receptor signaling pathways (from (Dash, 2003))



2.6.2 - Caspases

Caspases are a family of cysteine proteases that are activated specifically in

apoptotic cells. This family of proteases is highly conserved through evolution all the

way from hydra and nematodes up to humans. Over 12 caspases have been identified

and although most of them appear to function during apoptosis, the function of all of

them is not yet clear. The caspases are called cysteine-proteases because they have a

cysteine in the active site that cleaves substrates after asparagines in a sequence of

asp-X, with the four amino acids amino-terminal to the cleavage site determining a

caspase’s substrate specificity (Ruddon, 2007).

The importance of the caspases in apoptosis is demonstrated by the inhibitory

effects of mutation or drugs that inhibit their activity. Caspases can either inactivate a

protein substrate by cleaving it into an inactive form or activate a protein by cleaving a

pro-enzyme negative regulatory domain. In addition, caspases themselves are

synthesized as pro-enzymes and are activated by cleavage at asp-x sites. Thus, they can

be activated by other caspases, producing elements of the ‘‘caspase cascade’’ shown

in Figure 2.21.

Figure 2.21 – Caspase activation (from (Dash, 2003))



Also, as illustrated in Figure 2.21, caspases are activated in a number of steps

by proteolytic cleavage by an upstream caspase or by protein–protein interactions,

such as that seen for the activation of caspase-8 and the interaction of cytochrome c

and Apaf-1 in the activation of caspase-9. A number of important substrates of

caspases have been identified, including the caspase-activated DNase (CAD), noted

above, which is the nuclease responsible for the DNA ladder of cells undergoing

apoptosis. Activation of CAD is mediated by caspase-3 cleavage of the CAD-inhibitory

subunit. Caspase-mediated cleavage of other specific substrates has been shown to be

responsible for other typical changes seen in apoptotic cells, such as the cleavage of

nuclear lamins required for nuclear shrinkage and budding, loss of overall cell shape by

cleavage of cytoskeleton proteins, and cleavage of PAK2, a member of the p21-

activated kinase family, that mediates the blebbing seen in dying cells.

2.6.3 - Bcl-2 Family

Mammalian Bcl-2 was first identified as anti-apoptotic protein in lymphomas

cells. It turned out to be a homolog of an anti-apoptotic protein called Ced-9 described

in C. elegans and protects from cell death by binding to the pro-apoptotic factor Ced-4.

Similarly, in mammalian cells, Bcl-2 binds to a number of pro-apoptotic factors such as

Bax, Figure 2.22. One concept is that pro- and anti- apoptotic members of the Bcl-2

family of proteins form heterodimers, which can be looked on as reservoirs of plus and

minus apoptotic factors waiting for the appropriate signals to be released (Ruddon,

2007).

2.6.4 - Anoikis

Anoikis is a form of apoptosis that occurs in normal cells that lose their

adhesion to the substrate or extracellular matrix (ECM) on which they are growing.

Adherence to a matrix is crucial for the survival of epithelial, endothelial, and muscle

cells. Prevention of their adhesion usually results in rapid cell death, which occurs via

apoptosis. Thus, anoikis is a specialized form of apoptosis caused by prevention of cell

adhesion (Ruddon, 2007).



Figure 2.22 – Apoptotic pathways. Two major pathways lead to apoptosis: the intrinsic cell death pathway

controlled by Bcl-2 family members and the extrinsic cell death pathway controlled by death receptor signaling

(from (Zhang, 2005))

The term anoikis means ‘‘homelessness’’ in Greek and although the observation

of this phenomen occurs only with cultured cells, it is likely to occur also in vivo

because it is known that cell-cell and cell-ECM interactions are crucial to cell

proliferation, organ development, and maintenance of a differentiated state. This may

be a way that a multicellular organism protects itself from free-floating or wandering

cells (such as occurs in tumor metastasis). The basic rule for epithelial and endothelial

cells appears to be ‘‘attach or die’’. Interestedly, cells that normally circulate in the

body such as hematopoietic cells do not undergo anoikis (Ruddon, 2007).

Cell attachment is mediated by integrins, and ECM integrin interactions

transduce intracellular signaling pathways that activate genes involved in cell



proliferation and differentiation. Although the cell death pathways induced by

disruption of these cell attachment processes are not clearly worked out, cell

detachment–induced anoikis does result in activation of caspases-8 and -3 and is

inhibited by Bcl-2 and Bcl-XL, indicating some similarities to the typical apoptosis

mechanisms. In addition, integrin-ECM interaction activates focal adhesion kinase

(FAK) and attachment-mediated activation of PI3-kinase. Both of these steps protect

cells from anoikis, whereas inhibition of the PI3-kinase pathway induces anoikis

(Ruddon, 2007).

Disruption of cell-matrix interactions also turns on the JNK /p38 pathway, a

stress-activated protein kinase. The mitogen-activated kinase system may also be

involved, since caspase mediated cleavage of MEKK-1 occurs in cells undergoing

anoikis. As stated earlier, one of the hallmarks of malignantly transformed cells

growing in culture is their ability to grow in an anchorage independent manner,

whereas normal cells do not. Thus, cancer cells may develop resistance to anoikis. This

may be a way that metastatic cancer cells can survive in the bloodstream until they

seed out in a metastatic site (Ruddon, 2007).

2.7 - RESISTANCE TO APOPTOSIS IN CANCER AND POTENTIAL TARGETS FOR THERAPY

It would be a mistake to portray apoptosis as only a mechanism to kill cells

damaged by some exogenous insult such as DNA-damaging toxins, drugs, or

irradiation. Apoptosis is, in fact, a usual mechanism used by all multicellular organisms

to facilitate normal development, selection of differentiated cells that the organism

needs, and control of tissue size. For example, studies of nematodes (C. elegans), fruit

flies, and mice indicate that apoptotic-mediated mechanisms similar to those

described here are intrinsic and required for normal development. Dysfunction of

these pathway results in developmental abnormalities and disease states (Ruddon,

2007).

In the human, development of the immune system is perhaps the best example

of the role for apoptosis in normal development. In the immune system, apoptosis is a

fundamental process that regulates T- and B-cell proliferation and survival and is used

to eliminate immune cells that would potentially recognize and destroy host tissues



(‘‘anti-self ’’). Mechanisms involving Apo-1/FAS (CD95)-mediated signaling of the

caspase cascade are employed in lymphocytic cell selection. In the case of T

lymphocytes, pre-T cells are produced in the bone marrow and circulate to the thymus

where they differentiate and rearrange their T-cell receptors (TCRs). Those cells that

fail to rearrange appropriately their TCR genes, and thus cannot respond to self–major

histocompatibility complex (MHC)–peptide complexes, die by ‘‘neglect’’, Figure 2.23.

Those T cells that pass the TCR selection tests mature and leave the thymus to become

the adult peripheral T-cell pool. The mature T-cell pool thus passes through a number

of selection steps to ensure self-MHC restriction and self-tolerance. Apoptosis also is

used to delete mature peripheral T cells that are insufficiently stimulated by positive

growth signals, and this is a mechanism to downregulate, or terminate, an immune

response (Ruddon, 2007).

B lymphocytes undergo selection and maturation in the bone marrow and

germinal centers of the spleen and other secondary lymphoid organs. Those with low

antigen affinity or those autoreactive are eliminated by apoptosis. Those that pass this

test mature into memory B cells and long-lived plasma cells. The ability of lymphoid

progeny cells to avoid apoptosis may lead to lymphatic leukemias or lymphomas. In

addition, cancers develop multiple mechanisms to evade destruction by the immune

system such as a decreased expression of MHC molecules on cancer cell surfaces and

production of immunosuppressive cytokines. Several cell proliferations promoting

events take place in cancer cells as they evolve over time into growth dysregulated,

invasive, metastatic cell types. These events include activation of proliferation-

promoting oncogenes such as ras and myc, overexpression of cell cycle regulatory

factors such as cyclin D, increased telomerase to overcome cell senescence, and

increased angiogenesis to enhance blood supply to tumor tissue (Ruddon, 2007).

The cancer-related alterations in the apoptotic pathway provide a number of

cancer chemotherapeutic targets.



Figure 2.23 - The role of apoptosis in the development and function of T lymphocytes. Major pro-apoptotic and anti-

apoptotic signals/molecules (from (Zhang, 2005))

2.8 - SUMMARY

At the end of this chapter is possible to conclude that many of the controls that

govern the transition between quiescence and active cell cycling in mammals operate

in G1 phase. Loss of R point control appears to be a common, possibly even universal

step in tumor development, and a number of genetic lesions that can contribute to this

deregulation have been identified.

Loss of survival proteins can also contribute to apoptosis. The antiapoptotic

gene, BCL2, has been shown to be repressed by p53 and, therefore, contributes to

apoptosis by blocking survival signals mediated by BCL2. The choice as to whether a

cell undergoes apoptosis or cell cycle arrest and DNA repair depends on a number of

factors. Some may be independent of p53 such as extracellular survival factors, the

existence of oncogenic alterations and the availability of additional transcription

factors. However, the extent of DNA damage may also contribute to the choice by

affecting the level of activity of p53 induced. Activation of apoptosis has been

associated with higher levels of p53 than those required for cell cycle arrest which may

reflect a lower affinity of cell cycle arrest target gene promoters for p53. In addition,

the type of cell may affect the response to p53. Importantly, it is vital to identify why

transformed cells die in response to p53, whereas normal cells undergo cell cycle



arrest and DNA repair as this may be of great potential for the development of cancer

therapies (Macdonald, 2005).

This loss of cell cycle check point control by cancer cells may contribute to their

increased susceptibility to anticancer drugs. Normal cells have mechanisms to protect

themselves from exposure to growth-limiting conditions or toxic agents by calling on

these check point control mechanisms. Cancer cells, by contrast, can continue through

these checkpoints into cell cycle phases that make them more susceptible to the

cytotoxic effects of drugs or irradiation (Ruddon, 2007).

Apoptosis occurs in most, if not all, solid cancers. Ischemia, infiltration of

cytotoxic lymphocytes, and release of TNF may all play a role in this and it would be

therapeutically advantageous to tip the balance in favor of apoptosis over mitosis in

tumors, if that could be done.

Clearly, a number of anticancer drugs induce apoptosis in cancer cells but the

problem is that they usually do this in normal proliferating cells as well. Therefore, the

goal should be to manipulate selectively the genes involved in inducing apoptosis in

tumor cells, although understanding how those genes work may go a long way to

achieving this goal.

CHAPTER III

RADIATION EFFECT ON NORMAL AND NEOPLASTIC TISSUES

CHAPTER III – RADIATION EFFECT ON NORMAL AND NEOPLASTIC TISSUES



When cells are exposed to ionizing radiation the standard physical effects

between radiation and the atoms or molecules of the cells occur first and the possible

biological damage to cell functions follows later. The biological effects of radiation

result mainly from damage to the DNA, which is the most critical target within the cell;

however, there are also other sites in the cell that, when damaged, may lead to cell

death (Suntharalingam, 2002).

Many aspects of the response of tissue systems are strongly affected by the

state of the cell in its cycle, for example, the state of oxygenation of the cell. The

supply of metabolic substrates and the removal of metabolic products also play a role

in modifying the response of tissue systems. The most significant aspect of the

radiosensitivity of a tissue or organ system centers on the state of reproductive activity

and this proliferative state varies widely among the tissues of any mammalian species.

At one extreme are the tissues of the central nervous system, some of which rarely, if

ever, undergo division during the organism's adult life, and for which loss of clonogenic

ability is an irrelevant end point. At the other extreme are the blood forming organs,

which are proliferating at a rate approaching that of an exponentially growing, in vitro

culture (Alpen, 1998).

In this chapter, it is provided a detailed description of the effects of radiation

on normal and neoplastic tissues. For this purpose, the chapter starts with the

irradiation carcinogenesis which includes the description of ionizing and UV radiation

effect followed by the description of the types of cell death in mammalian cells. After

this item, it is performed a description of the nature of cell population in tissues and of

the cell population kinetics and radiation damage. Subsequently, the chapter focuses

on the cell kinetics in normal and tumor tissues, on the models for radiobiology

sensitivity of neoplastic tissues and the tumor growth and “cure” models. Finally, it

ends with a description of the radiobiological responses, hypoxia and radiosensitivity

of the tumor cells.



3.2 - IRRADIATION CARCINOGENESIS

X-rays and ultraviolet (UV) radiation produce damage to DNA inducing DNA

repair processes, some of which are error prone and may lead to mutations. The

development of malignant transformation in cultured cells after irradiation requires

cell proliferation to ‘‘fix’’ the initial damage into a heritable change and then to allow

clonal proliferation and expression of the typical transformed phenotype. Fixation

appears to be complete after the first post irradiation mitotic cycle, thus, a promotion

phase is required for full expression of the initiated malignant alteration. Moreover,

when low doses of chemical carcinogens and X-rays are used together, these two types

of agents act synergistically to produce malignant transformation (Raddon, 2007).

When cells are exposed to UV light in the 240 to 300 nm range, the bases

acquire excited energy states, producing photochemical reactions between DNA bases.

The principal products in DNA at biologically relevant doses of UV light are cyclobutane

dimers formed between two adjacent pyrimidine bases in the DNA chain. Both

thymine–thymine and thymine–cytosine dimmers are formed. That formation of these

dimers is linked to mutagenic events (Raddon, 2007).

3.2.1 - Ionizing Radiation

The history of radiation carcinogenesis goes back a long way. The harmful

effects of X-rays were observed soon after their discovery in 1895 by W. K. Roentgen.

The first observed effects were acute, such as reddening and blistering of the skin

within hours or days after exposure. By 1902, it became apparent that cancer was one

of the possible delayed effects of X-ray exposure. These cancers, which included

leukemia, skin cancers, lymphomas, and brain tumors, were usually seen in radiologists

only after long-term exposure before adequate safety measures were adopted, thus it

was thought that there was a safe threshold for radiation exposure. The hypothesis

that small doses of radiation might also cause cancer was not adopted until the 1950s,

when data from atomic bomb survivors in Japan and certain groups of patients treated

with X-rays for noncancerous conditions, such as enlarged thyroids, were analyzed.



These and other data led to the concept that the incidence of radiation-induced

cancers might increase as a linear, nonthreshold function of dose (Raddon, 2007).

In radiation carcinogenesis, the damage to DNA, and hence its mutagenic and

carcinogenic effect, is due to the generation of free radicals as the radiation passes

through tissues. The amount of radical formation and ensuing DNA damage depend on

the energy of the radiation. In general, X-rays and gamma rays have a low rate of linear

energy transfer, generate ions sparsely along their tracks, and penetrate deeply into

tissue. This profile contrasts with that of charged particles, such as protons and α

particles, which have a high linear energy transfer, generate many more radical ions

locally, and have low penetration through tissues. The damage to DNA can include

single- and double-strand breaks, point mutations due to misrepair deletions, and

chromosomal translocations, Figure 3.1 (Raddon, 2007).

The molecular genetics events that follow radiation damage to cells include:

1) Induction of early-response genes such as c-jun and Egr-1;

2) Induction of later-response genes such as tumor necrosis factor-α (TNF-α),

fibroblast growth factor (FGF), and platelet-derived growth factor-α (PDGF-α);

3) Activation of interleukin-1 (IL-1) PKC;

4) Activation of oncogenes such as c-myc and K-ras.

Induction of these genes may be involved in the cellular responses to

irradiation and in the longer-range effects that lead to carcinogenesis. At any rate, the

production of clinically detectable cancers in humans after known exposures generally

occurs after long latent periods. Estimates of these latent periods are 7 to 10 years for

leukemia, 10–15 years for bone, 27 years for brain, 20 years for thyroid, 22 years for

breast, 25 years for lung, 26 years for intestinal, and 24 years for skin cancers (Raddon,

2007).

3.2.2 - Ultraviolet Radiation

Ultraviolet radiation–induced lesions, generated by UV-B (280–320 nm

wavelength) or UV-A (320–400 nm wavelength), result from DNA damage, which is

converted to mutations during cellular repair processes. UB-B and UV-A generate

different types of DNA damage and DNA repair mechanisms (Raddon, 2007).



Figure 3.1 - Direct DNA damage radiation model (from (Price, 2009))

Irradiation with UV-B produces cyclobutane pyrimidine dimers that are

repaired by nucleotide excision repair. If left unrepaired, C T and CC TT base

transitions occur. UV-A induced DNA damage produces mostly oxidative lesions via

photosensitization mechanisms and is repaired by base excision repair (Raddon, 2007).

UV-B and UV-A also produce distinct effects on the immune system and elicit

different transcriptional and inflammatory responses. While the specific mechanisms

by which UV radiation induces basal cell or squamous cell carcinomas or melanoma are

not clear, a number of signal transduction pathways are affected that can either lead

to apoptosis or to increased cell proliferation (Raddon, 2007).

UV irradiation activates receptor tyrosine kinases and other cell surface

receptors. It also enhances phosphorylation by ligand-independent mechanisms via

inhibition of protein tyrosine phosphatase activity. Ligand dependent cell surface

receptor activation can occur as well by activation of autocrine or paracrine release of

growth factors from keratinocytes, melanocytes, or neighboring fibroblasts (Raddon,

2007).

3.3 – CELL DEATH IN MAMMALIAN TISSUES

The clonogenic potential is the essential element for the maintenance of a cell

line, either in vitro or in organized tissues, although there are other important issues in



the maintenance associated with complex tissue systems. Normal senescence of cells

is one of these important issues and the other is the removal of cells that are in the

wrong place at the wrong time. Examples of this would be the metastatic arrival of

tumor cells transported from a primary tumor elsewhere or the resolution of

inflammatory processes (Alpen, 1998).

It is possible to define at least two different types of cell death that go beyond

the end point of clonogenic potential and involve the actual disappearance of the cell:

necrosis and apoptosis (Alpen, 1998).

Necrosis is characterized by a tendency for cells to swell and ultimately to lyse,

which allows the cell's contents to flow into the extracellular space, this is usually

accompanied by an inflammatory response. In the case of neoplasms, necrosis is most

often seen in rapidly growing tumors, where the tumor mass outgrows its blood supply

and regions of the tumor become undernourished in oxygen and energy sources. In

this case inflammation is not a characteristic of the necrotic process (Alpen, 1998).

Apoptosis involves shrinkage of the nucleus and cytoplasm, followed by

fragmentation and phagocytosis of these fragments by neighboring cells or

macrophages. The contents of the cell do not usually leak into extracellular space, so

there is no inflammation. Since there is no inflammation accompanying apoptosis, the

process is histologically quite inconspicuous (Alpen, 1998).

Figure 3.2 - Structural changes of cells undergoing necrosis or apoptosis (from (Goodlett, 2001))



The concept of apoptosis as a mechanism for the control of cell population

numbers and cell senescence has been around for several decades, but the

mechanisms of apoptosis have received extensive research attention only in the

nineties. This interest in apoptosis was engendered by the discovery that tumor

suppressor genes and oncogenes were central control agents for the process. The

principal focus of these studies has been the role of the p53 tumor suppressor gene,

already described in chapter II. The p53 gene is a transcriptional activator that may

include activation of genes that regulate genomic stability, cell cycle progression, and

cellular response to DNA damage. The synthesis of the p53 product is known to be

responsible for the induction of apoptosis in many cell lines in which this gene is

present in unmutated form. The mutational absence of this gene is often accompanied

by the inability of a cell line to initiate apoptosis. For radiation pathology, the

important finding is that even small amounts of DNA damage in G1 cells cause

synthesis of the p53 product and ultimate apoptosis of the cells. It is pertinent for

radiation pathology that cells of the lymphoid system generate high concentrations of

p53 gene product after cell damage. This is particularly true for low doses of ionizing

radiation. Clearly, the generation of the p53 product is not sufficient for the onset of

apoptosis, but it is certainly necessary (Alpen, 1998).

Another significant gene involved in apoptosis is the bcl-2 gene (described in

chapter II). This gene encodes a protein that blocks physiological cell death (apoptosis)

in many mammalian cell types, including neurons, myeloid cells, and lymphocytes. This

gene is able to prevent cell death after the action of many noxious agents (Alpen,

1998).

The role of apoptosis as a mechanism for cell death following ionizing radiation

exposure remains unclear at this time, particularly the relative importance of the

agonistic role of p53 and the antagonistic role of bcl-2. However, it must be important,

as that the detection of small nicks and errors in the DNA of G1 cells is crucial to the

recovery of irradiated tissues and the reduction of genomic misinformation (Alpen,

1998).



3.4 – NATURE OF CELL POPULATIONS IN TISSUE

One of the earlier systematic overviews of the nature of cell population kinetics

in normal and malignant tissues was that of Gilbert, 1965. Their classification of the

various kinetic systems found in mammalian (and, incidentally, in other organisms)

organs and tissues is shown in Figure 3.3 (Alpen, 1998).

Figure 3.3 - Classification of cell kinetic types in the system of Gilbert, 1965 (from (Alpen, 1998)).

From Figure 3.3, the definitions of each of the systems are the following (the

double arrows in classifications D, E, and F, are meant to signify the mitotic division of

one of the cells of the compartment, giving rise to two daughter cells):

A. Simple transit population. Fully functional cells are added to the

compartment while a population of either aging or randomly destroyed cells disappear

from the pool. There are many examples of functional end cells that are in this

category. Examples are spermatozoa, which are constantly being replaced, as well as

red cells or other end cells of the blood.

B. Decaying population. The cell numbers decrease with time without

replacement. The population of oocytes in the mammalian female is often quoted as

an example, if not the only example. Populations of this classification are rare in

mammalian systems, but not in insects.

C. Closed, static population. There is neither decrease nor increase in cell

numbers during life. It is unlikely that such a population truly exists. The differentiated

neurons of the central nervous system are quoted as an example of a static

population, but there is probably a decline in cell numbers even in this population.



D. Dividing, transit population. In addition to the transiting cells, division of the

cells within the compartment occurs that leads to a larger number leaving than

entering. It is assumed in this model that the number of cells in the compartment

remains more or less static. The differentiating and proliferating blood cell types (for

example, the proerythroblast of the bone marrow) that follow the stem cell are

examples of this type of population.

E. Stem cell population. A self-sustaining population, that relies on self-

maintenance for its continued existence. All the progeny of this type of cell line

depend upon the continued existence of the stem cell pool. Every self-maintaining,

dividing cell population must have such a precursor pool. Examples are the stem cells

responsible for sustained spermatogenesis or hematopoiesis.

F. Closed, dividing population. Such a population is best represented by

neoplastic growth. No cells enter or leave the compartment in the early stages of

tumor growth. In the long run, neoplastic growth is probably best represented as a

stem cell population, since as the tumor enlarges, there is cell death, suppression of

growth by metabolic and other nutrient shortages, and a highly variable rate of

division. The epithelial cells responsible for cell renewal in the lens of the eye are

another example of this type of population (Alpen, 1998).

3.5 – CELL POPULATION KINETICS AND RADIATION DAMAGE

It should be almost self-evident that the kinetic types represented by D, E, and

F of Figure 3.3 will be most vulnerable to radiation damage. It has been established

that for clonogenic death of the cell the principal target of ionizing radiation is the

genome, and the genome is certainly at its most vulnerable to radiation damage during

G2 and mitosis (M), when replication has been completed. The principal outcome of

disturbances to the dynamic replicative activity of the genome is altered clonogenic

ability. That is indeed the case, and the most critically sensitive of these systems would

be the stem-cell-type tissue (E), which depends for its continuing function on its own

continued clonogenic potential, since there is no precursor compartment to replace

deficiencies (Alpen, 1998).



The ultimate functional viability of a tissue that is dependent on stem cell

activity will be determined by whether, after radiation exposure, there are adequate

numbers of surviving and still clonogenic stem cells to repopulate the compartment

and finally to produce functionally competent progeny. The most resistant tissues are

those that require neither input of cells from a prior compartment nor division within

the compartment. The closed static model is such a case, and in the case of the central

nervous system, its high degree of radioresistance can be attributed to its lack of need

for cell replication and replacement (Alpen, 1998).

3.5.1 – Growth Fraction and its significance

The concept of growth fraction as a descriptive parameter for the kinetics of

proliferating tissue appears to have been first proposed by Mendelsohn (1962) as the

result of his observations that all cells in a growing tumor are not in the active process

of proliferation as determined by the cellular incorporation of radioactive labels of

DNA synthesis. Lajtha (1963), based on his own studies as well as those of others,

proposed the concept of the G0 phase of the cell cycle, a state of the cell in which the

cell was not engaged in active proliferation, but in which the cell could reenter the

proliferative state. The G0 cell was visualized as a cell that has been removed from the

actively dividing population by regulatory activities rather than as a result of metabolic

deprivation. Subsequently, it became apparent that cells also could be removed from

active division in a reversible manner by deprivation of oxygen, glucose, or other

metabolites (Hlatky et al., 1988). Restoration of the lacking nutrient led to reentry of

the cell into active proliferation (Alpen, 1998).

Figure 3.4 – Cell cycle phases (from (Goldwein, 2006))



The growth fraction is defined as the fraction of the total cellular population

that is clonogenically competent and is actually in the active process of DNA replication

and cell division. The growth fraction may be estimated by any one of several

techniques, most of which depend on incorporation of a radioactively labeled DNA

precursor into those cells that are actively dividing. One of the simpler methods for

determination of the growth fraction is the exposure of a growing culture of cells, in

vitro or in vivo, to an appropriate radioactive label for the synthesis of DNA. A typical

and frequently used label is 3H-thymidine. The cells are exposed to the radioactive

label in the medium or by injection into the intact animal for at least the full length of a

cell cycle (and usually for half again as long). Under these conditions, all cells that

synthesize DNA, thus indicating their passage through the S period of the cell cycle, are

labeled and can be identified by autoradiography. The percentage of cells that is

labeled constitutes the growth fraction, since every cell in cycle will have passed

through the S period at least once during exposure to the radioactive label (Alpen,

1998).

The radiobiological significance of the growth fraction was unclear until the

appearance of new data in the late 1980s. In 1980, Dethlefsen indicated that the role

of quiescent cells in radiobiological response was not satisfactorily delineated. Recent

studies indicate that cells that are out of cycle are capable of a more significant

amount of repair of potentially lethal damage, simply because there is more time

before the cell is called on to replicate its DNA. It is possible, but by no means proved,

that the concentration of enzymes necessary for repair of DNA damage may be

depleted in the noncycling cell, but, in spite of this, the additional time allows effective

repair to proceed with the lower concentration of repair enzymes (Alpen, 1998).

3.6 – CELL KINETICS IN NORMAL TISSUES AND TUMORS

Both normal and neoplastic tissues have a cellular kinetic pattern that follows

the accepted model of a G1-S-G2-M cycle, and, indeed, the cell cycle parameters are

not very different for tumors as compared to other growing tissues. The total cycle

time and the time devoted to DNA synthesis in the S period are very much alike for

both tissue types. However, there are, significant differences in some of the



characteristics of the kinetic pattern as the tumor reaches a size where vascularization

is required for continued tumor growth. The orderly vascularization of normal tissues

that originates in embryonic life and that is maintained throughout the existence of

normal, nonpathological function assures that the supply of oxygen and nutrients is

adequate for survival of cells. Most, if not all, tumors, on the other hand, originate as

nonvascularized aggregations of cells and develop a vascular supply sometime after

the origination of tumor growth. The development of vascular supply in a tumor

depends on the activities of angiogenic factors that occur in normal tissues. The newly

developing vascular supply is, at best, chaotic and disorganized (Alpen, 1998).

Some parts of the tumor tissue will be so far from the source of oxygen and

nutrients that cell survival will be impossible, Figure 3.5. Other parts of the tumor will

have nutrient and oxygen supplies that are adequate only for survival of cells without

replication. The lack of oxygen and glucose can lead to a decrease in the growth

fraction, and probably to cell death and necrosis. Several nutrients and metabolic

products, including oxygen, glucose, and lactic acid, play an important role in the

determination of quiescent and proliferating cells in tumors (Alpen, 1998).

One important difference between normal tissues and tumor tissues is the

determinant of the fraction of quiescent cells in the organ or tumor. Because of the

orderly vascular architecture of normal tissue, the movement of cells from the

proliferating to the quiescent compartment is probably not the result of nutrient lack,

but, rather, the result of the activity of normal soluble growth factors and naturally

occurring inhibitors that regulate the growth and development of the tissue (Alpen,

1998).

3.7 – MODELS FOR RADIOBIOLOGICAL SENSITIVITY OF NEOPLASTIC TISSUES

The earliest attempts to assay the sensitivity of organized tissue systems were

directed at establishing the radiosensitivity of tumor tissues. This was partly because

these tissues offered opportunities for analysis that were not available for normal

tissues. The possibility for syngeneic transplantation of the cell lines from host to

recipient animal was the most important characteristic of these in vivo tissue systems.



Figure 3.5 - Role of hypoxia in tumour angiogenesis (from (Carmeliet, 2000))

After irradiation of the tumor in the host in which it was growing, it was

possible to transplant the tumor cells to an unirradiated recipient animal and to

observe the growth response of the irradiated tumor cells. There was also strong

interest in understanding tumor biology arising from the treatment of cancer by

radiotherapy. It was important to establish the role of oxygen in the sensitivity of

cancer cells, as well as the importance of the fraction of G0 cells and repair or

repopulation in these tissues. The overall goal was practical: to maximize the

effectiveness of radiotherapy for cancer control in patients, while reducing damage to

normal tissues in the radiation field (Alpen, 1998).

3.7.1 – Hewitt Dilution Assay

Probably the first in vivo assay for mammalian tissues was that developed by

Hewitt and Wilson (1959) with a syngeneic mouse tumor system. At that time a

number of tumor cell lines that were grown in the peritoneal cavity of mice had been



developed. The cells from these ascites tumors could be harvested or allowed to

continue to grow in the peritoneal cavity of the host, which would cause the death of

the animal. It occurred to Hewitt and Wilson that this end point - death of the host

animal could be used to measure the clonogenic potential of the tumor cells after

irradiation. Figure 3.6 shows the essentials of a Hewitt assay for a single dose point at

10 Gy (Alpen, 1998).

Figure 3.6 - Typical data set for a Hewitt dilution assay (from (Alpen, 1998))

Cells harvested from the mouse ascites tumor P388 and unirradiated cells were

collected from the donor and a series of dilutions was prepared from a stock

suspension of the tumor cells. A typical microbiological-type binary dilution was

carried out to produce cell suspensions with low concentrations of cells that will allow

the recipient animal to be injected with cell numbers that are correct for killing about

half of the animals. For the tumor line used, the usual cell dose required to kill half of

the animals is about two to three cells. A small number of animals (5-10) are injected

with the same cell dose and the survival is followed. The same procedure is used for

several additional cell doses. The resulting data on percent survival at each of the cell

doses are plotted as shown in Figure 3.6, and the LD50 (lethal dose for 50% of the

animals) is determined by graphical or analytical means. The procedure is repeated,

but with the cell suspension prepared from animals that were irradiated before cell

collection. Animals are irradiated at several doses and injections proceed as just

described for each dose. The LD50 values can be used to construct a survival curve.

Figure 3.6 shows an example for only one radiation dose on the right panel and for



unirradiated cells on the left panel, with the calculated surviving fraction. The surviving

fraction is estimated for each of the other doses, and a survival curve of surviving

fraction against dose is plotted in the usual way (Alpen, 1998).

The Hewitt assay has been the tool used for a number of significant studies of

tumor cell sensitivity to radiation. Figure 3.7 is a very good example of such studies.

Andrews and Berry (1962) developed survival curves for three mouse tumors, two

leukemias, and a sarcoma. Some of the data were Berry's own previously unpublished

observations and some were provided by Hewitt. The clonogenic survival curves were

developed for both anoxic and oxic conditions. All three cell lines could be plotted on

the same curve for oxic cells or for anoxic cells as appropriate, and the line produced

was a good fit for the appropriate condition of oxygenation. The oxygen enhancement

ratio (OER) for these cells was about 2.4, which is not far from the 2.8 or so for cell

lines that are irradiated in vitro and analyzed for clonogenic survival in vitro. The Do for

the cells irradiated under oxic conditions was about 150 cGy, and the extrapolation

number was about 3-4 for this set of data (Alpen, 1998).

A significant shortcoming of the dilution assay system is that donor cells that

are grown in ascites fluid are usually irradiated when the cell number in the peritoneal

cavity is very large. Under these conditions, it is not always clear that the cells are fully

oxygenated at the time of irradiation. If that is indeed the case, there is the possibility

of significant anoxic protection of the cells and, subsequently, there is an

overestimation of the resistance of the cells to the irradiation. The data reported in the

Berry study do not seem to be affected by such hypoxia. The Do (oxic) is about 150 cGy,

a number quite consistent with that found for many cell systems in vitro. The OER of

2.4 or so is, again, not very different from the 2.5-2.8 seen for in vitro systems. We

must conclude, at least for the cell lines reported in this study, that adequate

oxygenation probably existed at the time of irradiation (Alpen, 1998).

Another shortcoming of the Hewitt method is that the irradiated tumor cells

must be capable of expressing clonogenic potential while growing in the ascites

medium. For example, most leukemias grow readily in this environment, and usually

require an inoculum of only 1-3 cells to cause the death of 50% of the recipient

animals. For the Berry data just described, the sarcoma cells required an inoculum of

more than 80 cells to kill 50% of the recipients. In many cases no cell growth is seen



and no assay is possible. To avoid this shortcoming, other assays have been developed

(Alpen, 1998).

Figure 3.7 - The survival curve obtained by Berry (1964) via the Hewitt assay method for two mouse leukemias and a sarcoma (from (Alpen, 1998))

3.7.2– Lung Colony Assay System

A modification to the Hewitt assay was developed by Hill and Bush (1969) to

measure clonogenic survival of cells derived from solid tumors. In principle, the assay

measures the clonogenic survival of tumor cells by determining their ability to form

colonies in the lung of recipient syngeneic mice. The cells from a tumor, irradiated

either in vivo or, after dissection and cell dissociation, in vitro, are injected into a

recipient mouse, and after 18-20 days the animals are killed, the lungs are dissected,

and the number of tumor colonies in the lung is counted. Hill and Bush were able to

demonstrate a linear relationship between cell number injected and the number of

colonies formed in the lung. A very large enhancement of the number of colonies in

the lung was found if, along with the experimentally irradiated cells, a large number of

heavily irradiated, nonclonogenic cells were injected. Typically, such a procedure

produced a 10-50-fold increase in the number of colonies formed from the clonogenic

survivors. Hill and Bush were not able to establish the mechanism of this

enhancement, but it was not due to an immune response on the part of the recipient.



Very consistent survival curves were obtained, and, for the KHT transplantable

sarcoma, the Do was 134 cGy, with an extrapolation number of about 9.5. Again, these

data were found to be quite consistent with the values found for the same tumor with

the Hewitt assay. Such an agreement not only validates the lung colony assay, it also

demonstrates that there was little protection from radiation damage due to partial

hypoxia for the KHT cells irradiated as solid tumors and tested by the dilution assay

(Alpen, 1998).

A significant limitation of the lung colony assay is that cells must be injected

into syngeneic recipient mice, that is, inbred mouse lines of the same genotype as that

from which the tumor is derived (Alpen, 1998).

3.8 – TUMOR GROWTH AND TUMOR “CURE” MODELS

Since there is a very limited set of models for examining the clonogenic

potential of tumor cells, much of the radiation biology of tumors has been developed

using a set of tools that was developed for general use in tumor biology. Therefore,

some of these tools have been more valuable than others for radiation effect studies

because of the inherent inability to effect precise quantitation.

3.8.1 – Tumor Volume Versus Time

A widely used and relatively powerful tool in tumor radiobiology is the tumor

growth curve after implantation of an inoculum of cells, usually in the flank region of

recipient syngeneic mice or rats. The simplest application of the growth curve for

implanted tumors is the analysis on the increase rate of the tumor volume. For analysis

of the radiation effect we can measure the time for the tumor to reach a preselected

volume. The measurements of tumor volume are at best imprecise. The volume is

usually determined from a caliper measurement of two or more diameters of the

growing tumor and calculation of the volume from the average diameter (Alpen,

1998).

After the tumor has been irradiated, the time course of volume change is as

shown in Figure 3.8. There may be a slowing of growth for a brief time, followed by a



period of decreasing tumor volume. This decrease is due to lack of replacement of the

normal cell loss from tumors, associated with local necrosis, nutrient lack, or other

causes unrelated to the radiation exposure. It is not due to the interphase death of

cells as the result of irradiation. As the surviving clonogenic cells repopulate the tumor,

regrowth will be observed; the surviving clonogenic cells will ultimately produce

progeny exceeding the cell-loss factor (Alpen, 1998).

Figure 3.8 - Tumor volume versus time (from (Alpen, 1998))

The criterion for measurement of the radiation dependent response is the time

for the cell volume to again reach the value observed at the time of irradiation. This

time is shown in Figure 3.8, and it is measured, as shown, as the time from irradiation

until the tumor volume achieves the value existing at the time irradiation occurred.

This time value is called the growth delay. The important limitation of the growth delay

model for testing the radiobiological response of tumors is that a significant number of

transplantable tumors does not show any decrease in the volume of tumor after

irradiation (Alpen, 1998).

Presumably, this failure to decrease in volume is the result of a small cell-loss

fraction in the growing tumor. When irradiation takes place, clonogenic activity is

reduced until repopulation from competent clonogenic cells occurs. During the period

before regrowth commences as the result of repopulation, the normally small cell-loss

fraction of the tumor does not lead to reduction in tumor volume. In these cases it is

necessary to revert to the simpler measure of tumor volume versus time and the use



of the time to reach a preset volume. Alternatively, differences in this time for control

and irradiated tumors may be taken as the end point (Alpen, 1998).

3.8.2 – TCD50, Tumor Cure

Another end point that is widely used in tumor biology is the dose required to

"cure" an implanted tumor. For this model, a large number of implanted tumors are

irradiated with graded doses at the same time period after implantation of the tumor

inoculum. The end point is the fraction of animals that has received a given dose in

which the growth of the tumor is controlled. This local control index can be plotted for

each of the doses, and the dose required to control tumor growth in 50% of the

animals is estimated by a variety of statistical techniques. This value is usually called

the 50% tumor cure dose -TCD50 (Alpen, 1998).

3.9 – RADIOBIOLOGICAL RESPONSES OF TUMORS

Using a number of end points, including dilution assay, lung colony assay,

primary cell cultures, and tissue derived in vitro cultures, it has been possible to define

rather clearly the radiobiological responsiveness of various tumor lines, both animal

and human. With only a few important exceptions, the various tumor cell lines in wide

and long term experimental use have been found to have clonogenic survival

characteristics that are generally stable and for which the relevant survival parameters

are not very variable, considering the range of cell types and tissues from which these

transformed and immortal cell lines have been derived (Alpen, 1998).

Rather different findings have been reported for the survival curve parameters

of freshly derived culture systems grown from naturally occurring malignant tumors.

Extensive efforts have been devoted to characterization of the radiosensitivity of cell

lines from human tumors. The best fit to the data for a large number of human cell

lines, both nontransformed fibroblasts and tumors, is the linear-quadratic (LQ) model.

The radiosensitivity of the various cell lines can be divided into three groups with a

very good correlation with the known responsiveness of the tumors to radiotherapy:

lymphomata, known to be highly curable, were the most radiosensitive of the derived



cell lines, and melanomata revealed to be the most resistant for tumor curability and

the most radioresistant in the survival of the cell lines in culture (Alpen, 1998).

It is important to realize that the immediate responsiveness of a tumor to

radiation, as determined by reduction in the tumor volume, does not necessarily

predict the curability of the tumor with high efficiency. The degree of responsiveness

will be determined by many of the cell kinetic parameters of the tumor system. A high

cell-loss factor and a high growth factor associated with a small fraction of cells out of

cycle and associated with inherent cellular radiosensitivity, will assure a high degree of

responsiveness of the tumor, as measured by volume changes. Curability, on the other

hand, will depend in a complex way on the ability of the few remaining clonogenic cells

to repopulate the tumor after irradiation is over (Alpen, 1998).

3.10 – HYPOXIA AND RADIOSENSITIVITY IN TUMOR CELLS

Under circumstances where severe anoxia can occur in tissues or cellular

preparations, one should expect to see significant protection from the effects of

ionizing radiation. It is expected to find conditions of moderate to severe anoxia in

growing tumors in vivo. For cells grown in suspension, careful attention to culture

conditions usually will prevent the development of such anoxic conditions with

concomitant radioprotection. For the tissue assay systems, such as the Hewitt dilution

assay and others, there is clearly a protective effect of oxygen lack under the correct

conditions. Figure 3.7 shows such radioprotection for cells deliberately made anoxic by

killing the host animal or by allowing the cell number for cells growing in the peritoneal

cavity to reach very high levels. Figure 3.9 demonstrates methods by which the

fraction of hypoxic cells in a mixture with fully oxygenated cells can be detected and

measured quantitatively. The radioresistant "tail" for the dashed line survival curve

shown in Figure 3.9 (10% anoxic cells) is a common observation for cells from tumors

and indicates the presence of a mixed population of cells, part of which have a

radioresistance relative to the remainder of the population. This resistant fraction may

be due to hypoxia and the radioprotection that this state affords (Alpen, 1998).



Figure 3.9 - Survival curve for the irradiation of a cell suspension containing a fraction of hypoxic cells (from (Alpen, 1998))

The well known work of Thomlinson and Gray (1955) laid the foundations for

our understanding of hypoxia as well as reoxygenation in tumors during growth and

regrowth. Figure 3.10 (from Thomlinson, 1967) illustrates the processes proposed by

this author. The very young tumor is well oxygenated, since it is so small that no cells

are beyond the effective diffusion distance of oxygen from nearby capillaries. As the

tumor continues to grow, portions of the tumor volume may be beyond easy access to

diffusing oxygen. The tumor must depend for its supply of oxygen on the development

of newly formed vessels that arise from the adjacent normal tissue and penetrate the

tumor volume. This neovascularization of the tumor is not as well organized as the

blood supply in normal tissues, and the expanding volume of tumor will contain

regions in which oxygen is inadequate for the maintenance of metabolism, and some

fraction of the cells will be anoxic. Figure 3.10 illustrates that the fraction of anoxic

cells in the growing tumor may rise to several percent and in some tumor types, to as



much as 10%. According to the model of Thomlinson, when the tumor is irradiated

(position R 1 in the figure) the more radiosensitive, fully oxygenated cells are killed,

and the remaining hypoxic cells are in an environment of dead and dying cells with

lesser demand for metabolic oxygen (Alpen, 1998).

Figure 3.10 - Development of hypoxia and reoxygenation in an irradiated tumor (from (Alpen, 1998))

Shrinking of the tumor volume and lowered oxygen demand allow for

reoxygenation of the hypoxic cells, which is indicated by a rapid fall to near zero for the

anoxic fraction. After this period of reoxygenation, tumor regrowth commences and

the complete cycle is repeated. The significance of the reoxygenation phase in

fractionated radiotherapy of human tumors is undergoing careful reexamination,

partly because treatment modalities designed to optimize the kill of anoxic cells (high

linear energy transfer (LET) radiation, radiation under hyperbaric oxygen conditions,

and so on) have not been particularly successful. According to Figure 3.10, the

optimum time for a second irradiation of a fractionated scheme would be at point H in

the curve, when the population of hypoxic clonogenic cells is at a minimum. Recent

data suggest that the reoxygenation phenomenon actually occurs very soon after

irradiation, and indeed may take place while the irradiation is in progress.



3.11 – SUMMARY

Human tumors strongly differ in radiosensitivity and radiocurability and this is

thought to stem from differences in capacity for repair of sub-lethal damage.

Radiosensitivity varies along the cell cycle, S being the most resistant phase and G2 and

M the most sensitive. Therefore, cells surviving an exposure are preferentially in a

stage of low sensitivity (G1), i.e. synchronized in a resistant cell cycle phase. They

progress thereafter together into S and then to the more sensitive G2 and M phases. A

new irradiation exposure at this time will have a larger biological effect (more cell kill).

However, while this synchronization effect has explained some experimental results,

redistribution has never been shown to play a measurable role in the clinic of

radiotherapy (Mazeron, 2005).

Cells surviving an irradiation keep proliferating, increasing the number of

clonogenic cells, i.e. the number that must eventually be sterilized to eradicate cancer.

An inappropriate development of intratumoral vasculature leads to a large proportion

of poorly oxygenated cells and the proportion of hypoxic cells increases with the tumor

size (Mazeron, 2005).

Acutely hypoxic cells are far more radioresistant than well oxygenated cells.

Hypoxic cells usually survive irradiation, but they progressively (re)oxygenate due to

the better supply of oxygen available after well oxygenated cells have died. This

restores radiosensitivity in the tumor by several mechanisms, but re-oxygenation

occurring at long intervals is probably due to tumor shrinkage leading to a reduction of

the intercapillar distance (Mazeron, 2005).

CHAPTER IV

CELL CULTURE AND FLOW CYTOMETRY

CHAPTER IV – CELL CULTURE AND FLOW CYTOMETRY



Cell culture is an invaluable tool for researchers in numerous fields. It facilitates

the analysis of biological properties and processes that are not readily accessible at the

level of the intact organism. Successful maintenance of cells in culture, whether

primary or immortalized, requires knowledge and practice of a few essential

techniques (Helgason, 2005).

The use of cells in analytical chemistry, engineering, and biology requires a

dedicated space for cell culture and maintenance. The proper handling of cells and

tissues requires a level of diligence and constant education, to mitigate health and

safety risks. Cell culture requires a system of mutual separation of sample and scientist

to avoid contamination of either. Each time a culture flask and the dish is opened is, in

essence, an opportunity for a single bacterium or fungal cell to ruin an experiment.

Likewise, every time cell cultures or tissues are handled, there is a risk to the scientist.

It is therefore needed to understand the protective countermeasures required to

handle cells properly (Pappas, 2010).

This chapter presents the importance of the laboratory conditions in the

manipulation and maintenance of cell culture. Subsequently, it is explained the

cytogenetic analysis of cell line and I performed a description of the methods to induce

cell cycle checkpoints. In the end of the chapter, it is presented a description of the

methods for synchronizing mammalian cells and the analysis of the mammalian cell

cycle by flow cytometry.

4.2 - CELL-CULTURE LABORATORY

Setting up a laboratory (or space within an existing lab) for cell culture is not a

daunting task, but requires some planning and strict adherence to regulations. Most

universities, research institutes, and hospitals have a safety committee (some

committees specialize in biosafety) that is in place in part to help a research establish a

cell lab. While the government guidelines typically set the standard for safety rules, the

research institution may have additional guidelines to follow. Therefore, the safety

committee is therefore indispensable in the planning and setting up of a cell lab, as

well as in the subsequent (and often frequent) safety inspections. The main issues



when setting up and maintaining a culture lab are safety, sterility, and contamination.

All three of these issues are linked by the common safe practices and proper use of

equipment, and all three require that individuals working in the lab are properly

educated (Pappas, 2010).

Working in the lab requires universal precautions, assuming that all cell cultures

and related materials may contain hazardous pathogens. This assumption maintains a

more vigilant attitude, and reduces the risk of accidental exposure to a real pathogen.

Moreover, the possibility that cultures can be cross-contaminated requires additional –

albeit similar – precautions. In short, careful procedures will result in productive

research in a safe environment for cells and individuals. For those new to cells and cell

culture, this chapter will not only serve as an introduction to the tools required for a

cell lab, but will also detail some of the practical aspects to setting up a culture facility.

For those with cell culture experience, the discussion of analytical equipment should

prove useful (Pappas, 2010).

4.3 – MAINTAINING CULTURES

The proper maintenance of cells includes homeostasis during culture, cell

storage and the correct preparation of cells for analysis. The latter case is of the most

importance, as often analysis and homeostasis are incongruent. Buffers must be

changed, different media used, and the cells, at times, are exposed to drastically

diverdse conditions for analysis. In some cases, the change in conditions can affect the

outcome of the experiment negatively. In other instances, the conditions suitable for

cell analysis are fatal to the cell (e.g., electron microscopy). There are many works

available on the culture of almost every cell type imaginable (Pappas, 2010).

When culturing primary or immortal cells for analysis, sterility and cross-

contamination must also be monitored at all times. A few bacteria in a sample can

wreak havoc in a short time, rendering any analytical data useless. The cross-

contamination of cultures is at best a nightmare, as extensive genetic testing is

required to purify cell populations and yield accurate data. Considering the cost of

cells, reagents, instrumentation, and lab upkeep, at least as much thought should be

placed on the maintenance of cell cultures for appropriate analysis. The type of



environment the cell encounters can directly affect the outcome of an analytical

experiment: cell-growth conditions, analysis buffers and reagents can affect the cell

phenotype, cell signaling, and a host of other parameters. By careful maintenance of

primary and immortal cells, accurate and reproducible cell analyses can be conducted

(Pappas, 2010).

4.3.1 – Medium

More than any other reagent in a cell-analysis laboratory, a steady supply of

culture medium – and the choice of correct medium type – is essential for cell analysis.

There are, in general, two classes of medium one can consider for cell analysis. First,

medium that is used to maintain a culture in between experiments, and second,

medium used in the analysis itself. Often these two can be one in the same, although

in some cases a modified medium or supplemented buffer is needed during the

analysis or processing phase (Pappas, 2010).

There are many types of medium available and the supplements that can be

added to them expand the palette of options even further. Table 4.1 lists some

medium types that are common to cellular analysis, by cell type. The table is not

inclusive, but serves to highlight the differences in medium types, and that some

medium formulations are applicable to many cell lines. In most cases, the medium in

Table 4.1 is used during the culture (maintenance) phase, and a different buffer or

medium may be used during the analysis itself (Pappas, 2010).

Medium can be classified as basic or complete, depending on whether or not

serum is included, respectively. Basic medium has many of the components required

for cell metabolism. Basic media, such as DMEM and RPMI 1640 (see Table 4.1),

contain salts (partly from buffer action), amino acids, vitamins (such as biotin, folic

acid, B-12, etc.), and molecules involved in energy production (glucose, pyruvate).

Basic medium also often contains other buffers (such as HEPES) and a colorimetric

acid–base indicator, such as phenol red. The latter serves as a quick visual inspection

of the “age” of the medium in culture. As cells consume nutrients and produce waste,

the culture medium acidifies, resulting in a shift in color for the pH indicator. The

formulations of most culture media are available and should be examined for potential



interference in the analysis. For example, staining using Annexin-V-based apoptosis

probes requires relatively high Ca2+ concentrations and at the same time, the presence

of phenol red in the medium will interfere with fluorescence measurements of

fluorescein, green fluorescent protein (GFP), and other fluorophores with similar

emission spectra. Fluorescence from phenol red itself makes sensitive fluorescence

measurements nearly impossible (Pappas, 2010).

Table 1 – Medium types common to cell analysis (from (Pappas, 2010))

Medium Serum Additives Cell lines

RPMI 1640 10% FBS Antibacterial-Antifungal

Jurkat, HuT 78, RPMI

8226, CCRF-CEM, U937,

HL-60

Dulbecco`s modified

Eagle Medium (DMEM) 10% FBS

Antibacterial-Antifungal,

L-Glutamine

NIH 3T3, RBL-1, HT-29,

HeLa

Clavcomb`s Medium 10% FBS

Antibacterial-Antifungal,

Norepunephrine, L-

Glutamine

HL-1

Cell Mab 0-10% FBS Varies

Designed for

monoclonal antibody

production

Leibovitz`s L-15 Hemolymph Bag neuronal cells

Eagle`s Minimum

Essential Medium 0-10% FBS L-Glutamine

F-12 0-10% FBS L-Glutamine Designed for primary

cells

Iscove`s Modified

DMEM 0-10% FBS L-Glutamine HuT 78 T Cells

FBS = Fetal Bovine Serum

Medium is, in essence, a man-made attempt to mimic the life support found in

vivo. It is, therefore, lacking in many essential compounds for cell growth. Many cell

lines can function in basic medium without additional materials, but for the most

routine culture and analysis, serum must be added to form the complete medium

(Pappas, 2010).



Serum is typically derived from animal sources, the most common being fetal

bovine serum (FBS). FBS and other sera contain growth factors such as epidermal

growth factor (EGF), some interleukins, and transferrin. Furthermore, present are

adhesion-promoting proteins and peptides, for example, fibronectin and laminin and

other components including insulin and various minerals. FBS and other animal-based

sera are by far the most common supplements used for culture maintenance (Pappas,

2010).

Being derived from animal sources, serum is inherently difficult to use from a

quality-control perspective and since it is derived from different animal types this can

affect experiment outcome. For example, the use of FBS instead of native rat serum

was shown to affect the outcome of rat leukocyte immunological response. In addition

to species variability, serum varies from lot to lot, as well as by country of origin, so if

cell products are to be analyzed over long time periods (months of experimentation) it

is best to purchase a large quantity of serum from one particular lot. Given the high

cost of medium, this may not always be practical since serum cost increases as the

level of quality control improves. The more consistent and well characterized the

medium, the higher the cost (Pappas, 2010).

Another negative aspect of dealing with serum is that the serum, or animal of

origin, is subject to contamination, just like any other primary derived material. Certain

viruses, bacteria, and mycoplasma have been shown to be transmitted via serum.

There are several replacement sera that can be substituted for FBS. For example, the

FetalClone series and Bovine Growth Serum, both from HyClone, are non-fetal animal

sera supplemented with various growth factors, minerals, and other compounds. Since

they are not derived from fetal animals, there is less variability between lots (especially

for the added compounds). None of the alternative sera offers much relief as far as

cost is concerned, but the increase in quality control is a major improvement (Pappas,

2010).

Some cells readily grow in serum-free medium; most, however, must be

acclimated to a serum-free environment. This requirement is especially true if the cell

line in question is already being cultured in serum-enriched medium (typically 10%

v/v). It is possible to reduce serum content in medium; in some cases, it is advisable to

do so, because reducing the amount of serum added can reduce costs, as serum is the



most expensive component of the complete medium. Reducing serum also lowers the

total protein content of the medium, facilitating collection of cell products, and

minimizing sources of contamination. For cells growing in serum-enriched medium, a

method of systematically reducing medium can be implemented (Pappas, 2010).

One must first consider the growth of cells in culture, before discussion of how

to achieve serum reduction can initiate, Figure 4.1. Cell growth in culture – whether

the cells are adherent or suspended – is characterized by several stages. The lag phase,

during which minimal or no cell division occurs, is a brief period after inoculation. The

lag phase occurs as cells adjust to a new cell-culture environment, and as adherent

cells begin the process of reattaching to the culture substrate. The lag phase is

followed by the log or exponential phase. This is the major phase of cell division. The

doubling time, an indicator of cell growth, is determined during this period (Pappas,

2010).

Figure 4.1 - Cell growth in culture (from (Pappas, 2010))

The time for the cell population to double, Figure 4.1, can be determined at any

point during the log phase, although it is most accurate at the center of that phase.

After the log phase, the culture reaches the stationary phase (Pappas, 2010).

High cell density, contact inhibition, and consumption of nutrients signal a

slowing of the cell cycle, and the cell concentration remains constant. Cell crowding,

depletion of nutrients and accumulation of waste eventually causes a sharp drop in cell

concentration, called the death phase. This latter phase can be confirmed by



microscopy, where the presence of a large number of dead cells, cell debris, and

acidified medium (if an indicator is present) can be observed (Pappas, 2010).

The glucose content of basic medium varies and is sometimes supplemented

with additional glucose. The high glucose content of many medium types is intended

to stimulate growth of the culture. However, some cell lines change phenotypic

properties in high or low glucose. When culturing for conditions close to those

encountered in vivo, the glucose concentration should be adjusted to reflecting the

physiological value as much as possible. Like serum reduction, the impact of changes in

glucose concentration can be monitored using the culture doubling time (Pappas,

2010).

When formulating complete medium, care must be taken to preserve sterility

of the final mixture. If all components are sterile to begin with, then aseptic handling in

the biosafety cabinet will prevent contamination of the complete medium. If any of

the reagents are not sterile at the onset, then filtration can be employed to remove

contaminating organisms.

4.3.2 – The use of medium in analysis and alternatives

Medium is primarily used to maintain cultures and samples before analysis. The

medium can also be used during the analysis; in other instances, components of the

medium may produce artifacts or otherwise interfere. The presence of several

components of medium can interfere with fluorescence measurements. Phenol red,

one of the most common pH indicators added to medium, has a broad absorption

band that interferes with most green fluorescence. Phenol red is also weakly

fluorescent, creating an additional problem for green-emitting fluorophores. If the cell

homeostasis is not required, then any buffer devoid of phenol red will work for

fluorescence. On the other hand, if the cells are to be kept alive for long periods, then

phenol-red-free medium is available from most medium manufacturers. In addition to

the weakly fluorescent properties of phenol red, other compounds present at

relatively high concentrations can interfere with fluorescence detection. Riboflavin is

also weakly fluorescent, but the relatively large volume of the medium contributes to

an unacceptable background signal. Proteins such as albumin, one of the major



components of serum, also contribute strongly to autofluorescence of medium. The

exact medium used for culture depends on the cell type, the culture conditions, and

the desired end result. For analysis, a similar selection process must be undertaken.

The final medium or buffer used for analysis must be of low background, minimal

interference, and – when possible – capable of sustaining cell viability and function for

the experiment duration (Pappas, 2010).

4.4 – CYTOGENETIC ANALYSIS OF CELL LINES

4.4.1 - The Utility of Cytogenetic Characterization

Countless cell lines have been established—more than 1000 from human

hematopoietic tumors alone —and the novelty and utility of each new example should

be proven prior to publication. For several reasons, karyotypic analysis has become a

core element for characterizing cell lines, mainly because of the unique key

cytogenetics provides for classifying cancer cells. Recurrent chromosome changes

provide a portal to underlying mutations at the DNA level in cancer, and cell lines are

rich territory for mining them. Cancer changes might reflect developmentally

programmed patterns of gene expression and responsiveness within diverse cell

lineages. Dysregulation of certain genes facilitates evasion of existing antineoplastic

controls, including those mediated by cell cycle checkpoints or apoptosis. The

tendency of cells to produce neoplastic mutations via chromosomal mechanisms,

principally translocations, duplications, and deletions, renders these changes

microscopically visible, facilitating cancer diagnosis by chromosome analysis. Arguably,

of all neoplastic changes, those affecting chromosomal structures combine the

greatest informational content with the least likelihood of reversal. This is particularly

true of the primary cytogenetic changes that play key roles in neoplastic

transformation and upon the presence of which the neoplastic phenotype and cell

proliferation ultimately depend. Nevertheless, the usefulness of karyotype analysis for

the characterization of cell lines lies principally among those derived from tumors with

stronger associations with specific chromosome rearrangements (i.e., hematopoietic,

mesenchymal, and neuronal, rather than epithelial tumors) (Helgason, 2005).



Cytogenetic methods facilitate observations performed at the single-cell level,

thus allowing detection of intercellular differences. Accordingly, a second virtue of

cytogenetic data lies in the detection of distinct subclones and the monitoring of

stability therein. Except for doublings in their modal chromosome number from 2n to

4n “tetraploidization,” cell lines appear to be rather more stable than is commonly

supposed. Indeed, chromosomal rearrangement in cells of the immune system could

reach peak intensity in vivo during the various phases of lymphocyte development in

vivo. A further application of cytogenetic data is to minimize the risk of using false or

misidentified cell lines. At least 18% of new human tumor cell lines have been cross-

contaminated by older, mainly “classic,” cell lines, which tend to be widely circulated.

This problem, first publicized over 30 years ago but neglected of late, poses an

insidious threat to research using cell lines (Helgason, 2005).

In the event of cross-contamination with cells of other species, cytogenetic

analysis provides a ready means of detection. Although modal chromosome numbers

were formerly used to identify cell lines, their virtue as descriptors has declined along

with the remorseless increase in the numbers of different cell lines in circulation. Thus,

species identification necessarily rests on the ability to distinguish the chromosome

banding patterns of diverse species. Fortunately, cells of the most prolific mammalian

species represented in cell lines (primate, rodent, simian, as well as those of domestic

animals) are distinguishable by experienced operators (Helgason, 2005).

4.5 – METHODS TO INDUCE CELL CYCLE CHECKPOINTS

The way cells respond to radiation or chemical exposure that damages

deoxyribonucleic acid (DNA) is important because induced lesions left unrepaired, or

those that are misrepaired, can lead to mutation, cancer, or lethality. Prokaryotic and

eukaryotic cells have evolved mechanisms that repair damaged DNA directly, such as

nucleotide excision repair, base excision repair, homology-based recombinational

repair, or nonhomologous end joining, which promote survival and reduce potential

deleterious effects. However, at least eukaryotic cells also have cell cycle checkpoints

capable of sensing DNA damage or blocks in DNA replication, signaling the cell cycle

machinery, and causing transient delays in progression at specific phases of the cell



cycle. These delays are thought to provide cells with extra time for mending DNA

lesions before entry into critical phases of the cell cycle, such as S or M, events that

could be lethal with damaged DNA (Lieberman, 2004).

The precise mechanisms by which checkpoints function is under intensive

investigation and details of the molecular events involved are being pursued

vigorously. This owes not only to the complexity and the intellectually and technically

challenging aspects of the process but also to the relevance of these pathways to the

stabilization of the genome and carcinogenesis. Nevertheless, it is clear that

checkpoint mechanisms are very sensitive and can be induced by the presence of

relatively small amounts of DNA damage. For example, in the yeast Saccharomyces

cerevisiae, as little as a single double-strand break in DNA can cause a delay in cell

cycle progression. One important aspect of studying cell cycle checkpoint mechanisms

is an understanding of how to induce the process (Lieberman, 2004).

The application of radiations, such as gamma rays and ultraviolet (UV) light, are

capable of causing DNA damage, and thus leading to the induction of cell cycle

checkpoints. Certain chemicals or the use of temperature- sensitive mutants to disrupt

DNA replication, are also used routinely to induce checkpoints. Gamma rays cause

primarily single- and double-strand breaks in DNA but can infrequently induce

nitrogenous base damage as well. In contrast, UV light (i.e., 254 nm) causes a

preponderance of bulky lesions, such as pyrimidine dimers, although single-base

damage and strand breaks are a smaller part of the array of lesions that can be

produced. Regulation of cell cycle checkpoints induced by ionizing radiation versus UV

light is mediated by overlapping but not identical genetic elements (Lieberman, 2004).

4.6 – METHODS FOR SYNCHRONIZING MAMMALIAN CELLS

When studying cell cycle checkpoints, it is often very useful to have large

numbers of cells that are synchronized in various stages of the cell cycle. A variety of

methods have been developed to obtain synchronous (or partially synchronous) cells,

all of which have some drawbacks. Many cell types that attach to plastic culture dishes

round up in mitosis and can then be dislodged by agitation. This mitotic shake-off

method is useful for cells synchronized in metaphase, which on plating into culture



dishes move into G1 phase in a synchronous manner. A drawback to the mitotic shake-

off method is that only a low percentage (2–4%) of cells are in mitosis at any given

time, so the yield is very small. Also, cells rapidly become asynchronous as they

progress through G1 phase, so the synchronization in S phase and especially G2 phase is

not very good. The first limitation can be overcome by plating multiple T150 flasks with

cells, using roller bottles, or blocking cells in mitosis by inhibitors such as Colcemid or

nocodazole (Lieberman, 2004).

Mitotic cells that are collected can be held on ice for an hour or so while

multiple collections are done to obtain larger numbers of cells. To obtain more highly

synchronous populations of cells in S phase, the mitotic shake-off procedure can be

combined with the use of deoxyribonucleic acid (DNA) synthesis inhibitors, such as

hydroxyurea (HU) or aphidicolin (APH), to block cells at the G1/S border (but probably

past the G1 checkpoint). APH inhibits DNA polymerase α, whereas HU inhibits the

enzyme ribonucleotide reductase, though it may operate by other mechanisms also.

On release from the block, cells move in a highly synchronized fashion through S phase

and into G2 phase. In terms of the number of synchronized cells, this method has the

same limitation as discussed above, because the starting cell population derives from

the mitotic shake-off procedure. In addition, the block of cells with drugs can cause

unbalanced cell growth, so one cannot necessarily conclude that all biochemical

processes are also synchronized (Lieberman, 2004).

Large numbers of synchronous cells can be obtained using centrifugal

elutriation, Figure 4.2. This method requires the use of a special rotor in a large floor

centrifuge and separates cells into the cell cycle based on cell size. Cells may be

obtained in early or late G1 phase, or primarily in S phase. However, the cell

populations are not highly synchronous in S phase but instead have significant

populations of G1- and G2-phase cells included. Nevertheless, it is possible to

synchronize very large numbers of cells using this method, and biochemical processes

are not perturbed (Lieberman, 2004).



Figure 4.2 - Centrifugal elutriation (from (Wahl, 2001))

Another method that results in highly synchronous populations is based on

labeling cells with a viable dye for DNA (Hoechst 33342). Cells stained with this dye can

then be sorted by cell cycle phase. Sorted G1 cells will be distributed throughout G1,

cells in S phase can be sorted into a small window in S phase and thus will be highly

synchronized, but only a small number of cells can be obtained. G2 phase cells will be

contaminated with late S phase cells. Furthermore, some cell types do not stain well

with Hoechst 33342, so sufficiently good DNA histograms cannot be obtained Hoechst

33342 (Lieberman, 2004).

4.7 – ANALYSIS OF THE MAMMALIAN CELL CYCLE BY FLOW CYTOMETRY

One of the most common uses of flow cytometry is to analyze the cell cycle of

mammalian cells. Flow cytometry can measure the deoxyribonucleic acid (DNA)

content of individual cells at a rate of several thousand cells per second and thus

conveniently reveals the distribution of cells through the cell cycle (Lieberman, 2004).

The DNA-content distribution of a typical exponentially growing cell population

is composed of two peaks (cells in G1/G0 and G2/M phases) and a valley of cells in S

phase, Figure 4.3. G2/M-phase cells have twice the amount of DNA as G1/G0-phase

cells, and S-phase cells contain varying amounts of DNA between that found in G1 and



G2 cells. Most flow-cytometric methods of cell cycle analysis cannot distinguish

between G1 and G0 cells or G2 and M cells, so they are grouped together as G1/G0 and

G2/M. However, there are flow cytometric methods that can distinguish four or even

all five cell cycle subpopulations: G0, G1, S, G2, and M. Furthermore, each

subpopulation can be quantified. Obviously, flow cytometry with these unique

features is irreplaceable for monitoring the cell cycle status and its regulation

(Lieberman, 2004).

Figure 4.3 - A typical cell cycle distribution of DNA content (from (Cooper,2004))

Cell cycle checkpoint genes are key elements in cell cycle regulation.

Checkpoint gene mutation can lead to defects in one or more cell cycle checkpoint

controls, which can then result in cell death or cancer. Many of the cell cycle

checkpoint genes are tumor suppressors, such as p53, ataxia-telangiectasia mutant

(ATM), ataxia-telangiectasia and Rad3 (ATR), and BRCA1 (Lieberman, 2004).

In mammalian cells, the cell cycle checkpoint controls that can be analyzed by

flow cytometry are G1 arrest, suppression of DNA replication, and ATM dependent as

well as independent G2 arrest. Exposure to a genotoxic agent can activate some or all

the checkpoints (Lieberman, 2004).

4.8 – CONCLUSION

Effective in vitro maintenance and growth of animal cells requires culture

conditions similar to those found in vivo with respect to temperature, oxygen and



carbon dioxide concentrations, pH, osmolality, and nutrients. Within normal tissue in

vivo, animal cells receive nutrients through blood circulation. For growth in vitro,

animal cells require an equivalent supply of a complex combination of nutrients. For

this reason, the first attempts in animal cell culture were based on the use of biological

fluids such as plasma, lymph and serum, as well as on extracts from embryonic-derived

tissue (Castilho, 2008).

Medium composition is one of the most important factors in the culture of

animal cells. Its function is to provide appropriate pH and osmolality for cell survival

and multiplication, as well as to supply all chemical substances required by the cells

that they are unable to synthesize themselves. Some of these substances can be

provided by a culture medium consisting of low molecular weight compounds, known

as basal media. However, most basal media fail to promote successful cell growth by

themselves and require supplementation with more complex and chemically

undefined additives such as blood serum (Castilho, 2008).

Some cultivation processes are based on operational strategies that allow cells

to remain viable, but in a nonproliferative state, so as to prolong the productive phase

and to increase the productivity of the process. By these strategies cell proliferation

may be controlled by adding chemical additives that arrest the cell cycle, usually in the

G1 phase, increasing specific productivity. However, concomitantly undesirable effects

such as cytotoxicity may be observed, which result in a decrease in cell viability and in

the impossibility of maintaining the culture in a nonproliferative state for long periods

of time. Deprivation of specific nutrients and growth factors can also stop cell

proliferation, but in this case cell viability decreases and programmed cell death –

apoptosis – is activated. Currently, much research on the biochemical control of cell

cultures based on preventing the cell death mechanisms, to avoid cell death instead of

inhibiting cell growth, is being carried out with the aim of prolonging the productive

period of a cell culture process (Castilho, 2008).

Any process, industrial or laboratory-based, presents a series of important

variables that represent its state. In the case of cell culture, there are the variables

related to the environment to which the cells are exposed, such as temperature, pH,

dissolved oxygen, nutrients in the culture medium, and metabolite concentrations, as



well as those related to the cell itself, such as concentration, average size, or the

profile of intracellular enzyme activities (Castilho, 2008).

CHAPTER V

MATERIALS AND METHODS

CHAPTER V – MATERIALS AND METHODS



The main goal of this chapter is to perform a description of the materials and

methods of the following papers: “Lack of p53 function promotes radiation-induced

mitotic catastrophe in mouse embryonic fibroblast cells” of Fiorenza Ianzini,

Alessandro Bertoldo, Elizabeth A Kosmacek, Stacia L Philips and Michael A Mackey,

Cancer Cell International 2006 6:11, and “Nuclear accumulation and activation of p53

in embryonic stem cells after DNA damage” of Valeriya Solozobova, ALexandre

Rolletschek and Christine Blattner, BMC Cell Biology 2009 10:46.

The previous chapters enable the acquisition of theoretic knowledge regarding

the methods employed in the execution of the laboratorial work performed in the

mentioned papers.

The images resulting from the mentioned papers will be analyzed and

processed, as will be demonstrate in the next chapter.

5.2 – MATERIALS AND METHODS OF THE PAPER “LACK OF P53 FUNCTION PROMOTES RADIATION-

INDUCED MITOTIC CATASTROPHE IN MOUSE EMBRYONIC FIBROBLAST CELLS”

5.2.1 - CELL CULTURE

Mouse embryonic fibroblast (MEF) cells were grown in monolayer in Dulbecco's

modified eagle medium (DMEM) (GIBCO) containing 10% heat-inactivated fetal bovine

serum (Hyclone), non-essential aminoacids (GIBCO) and antibiotics (100 U/ml penicillin

and 100 μg/ml streptomycin) (GIBCO). Under these growth conditions, cells grew with

a doubling time of about 14 using an Olympus AX-70 microscope equipped with a

mercury lamp and ultraviolet filters. Only intact cells containing three or more

fragmented nuclei were scored for these experiments.

5.2.2 - Light microscopy

The same microscopic slides prepared for the cytology end point (described

above) were used to depict cell morphology changes post-irradiation. Images were

acquires using an epifluorescence Olympus BX51 microscope at a magnification of 10×.



5.2.3 - Bivariate BrdUrd-PI (bromodeoxiuridine-propidium iodide) flow cytometry

Analysis of cell cycle distribution during the post-irradiation interval was

determined using 10 μM BrdUrd pulse labeling techniques, followed by bivariate

analysis using anti-BrdUrd-PI staining, to monitor cells in G1, S, and G2 phases. Flow

cytometric analysis was performed using a FacSTAR, with excitation of fluorochromes

by an argon laser emitting at 488 nm with 300 mW power; red fluorescence (PI) was

detected using a 640 nm low-pass filter, and green fluorescence (FITC) using a 525 nm

band-pass filter.

5.2.4 - Bivariate cyclin B1-PI flow cytometry

Estimates of the relative intracellular levels of cyclin B1 were made using anti-

cyclin B1-PI analysis. Briefly, cells fixed in 95% ethanol were incubated (1 hour, room

temperature) with an anti-cyclin B1 monoclonal antibody, rinsed and incubated as

before with FITC-conjugated goat-anti-mouse IgG, treated with RNAse (1 mg/ml, 30

min, room temperature) after which 0.5 ml of 70 μg/ml of PI was added. The details

for flow cytometric data acquisition were the same as for the anti-BrdUrd-PI analysis

above. In all samples analyzed, cyclin fluorescence was detected only in cells with early

S DNA content.

5.2.5 - Western blotting

Western Blotting was performed to determine cyclin B1 protein expression in

the mutant p53 MEF 10 cells and in the wild-type p53 MEF 12 cells. Thirty μg total

proteins from whole cell extracts were boiled for 10 min in Laemmli sample buffer and

separated using 10–12% 1D SDS-PAGE. The separated proteins were transferred to

Immobilon-P membranes using a semi-dry blotting apparatus and probed with an anti-

cyclin B1 monoclonal antibody (anti-mouse), at a dilution of 1:500. Beta actin was

detected using goat anti-actin polyclonal antibody. Peroxidase-conjugated AffiniPure

goat anti-mouse IgG Fcy fragment specific and donkey anti-goat IgG HRP were used as



secondary antibodies at a dilution of 1:6000 and 1:2000, respectively. Detection was

performed using Western Lightning Chemiluminescence Reagent.

5.3 – MATERIALS OF THE PAPER “NUCLEAR ACCUMULATION AND ACTIVATION OF P53 IN EMBRYONIC

STEM CELLS AFTER DNA DAMAGE”

5.3.1 - Cell lines and their treatments

R1 and D3 ES cells were cultured in GlutaMAX™-I medium supplemented with

15% fetal bovine serum, 0.1 mM β-mercaptoethanol, 40 μg/ml gentamycin and 1000

units/ml LIF in culture dishes that had been coated with 0.1% gelatine. Mouse

embryonal fibroblasts that had been irradiated with 6.3 Gray served as feeder cells.

p53-/- ES cells were cultured in GlutaMAX™-I medium supplemented with 15% fetal

bovine serum, 0.1 mM β-mercaptoethanol, 1000 units/ml LIF and 300 μg/ml G418 in

culture dishes that had been coated with 0.1% gelatine.

SNL cells irradiated with 6.3 Gray served as feeder cells. CGR8 cells were grown

in Glasgow Minimum Essential Medium supplemented with 10% fetal bovine serum,

40 μg/ml gentamycin, 100 units/ml LIF, 0.05 mM β-mercaptoethanol, 2 mM L-

glutamine and 1 mM non-essential aminoacids without feeder cells. Culture dishes

were coated with 0.2% gelatine.

All ES cell lines were sub-cultured each day. 3T3 cells were grown in Dulbecco's

Modified Eagle Medium supplemented with 10% donor bovine serum and 1%

penicillin/streptomycin. All cells were cultured at 37°C and 6% CO2 in a humidified

atmosphere.

Cells were irradiated with a 60Co γ-ray source at a dose rate of 1 Gray per

minute in cell culture medium. For UV irradiation, the culture medium was removed

and saved. The cells were washed with PBS and irradiated with 30 J/m2. After

irradiation, the original culture medium was added back to the cells. Transfections

were performed using the mouse ES cell Nucleofector Kit according to the

manufacturer's recommendations.



5.3.2 - Immunofluorescence staining

ES cells were grown on feeder cells that had been grown on cover slips. After

washing twice with ice cold PBS, cells were fixed with ice-cold acetone/methanol (1:1)

for 8 min on ice. Cover slips were washed 3 times with PBS and cells were

permeabilized with 0.5% Triton-X-100 in PBS for 10 min. Cover slips were washed 3

times with PBS and incubated for 30 min in blocking buffer (1% bovine serum albumin;

1% goat serum in PBS). After blocking, cells were incubated overnight with an antibody

directed against p53 (Pab421) that had been diluted 1:200 in blocking buffer. Cover

slips were washed 3 times with PBS and incubated for 30 min at room temperature in

the dark with an antibody directed against mouse IgG coupled to Alexa-Fluor-488 and

Draq5 both diluted 1:1000 in blocking buffer. Cover slips were washed 3 times and

mounted with Hydromount on microscope slides.

5.3.3 - RT-PCR

Total RNA was prepared from cells using the RNeasy kit according to the

manufacturer's recommendation and treated with DNase I to remove residual genomic

DNA. RNA was transcribed into cDNA using random primers and RevertAidtm H

MinusM-MuLV reverse transcriptase. Real-time PCR was performed in duplicates with

a SYBR Green PCR mixture. The cDNA was denatured for 15 min at 95°C followed by 40

cycles of 95°C for 15 s and 50°C for 1 min using the 7000 ABI sequence detection

system and gene specific primers. The signals were normalized to the signals for the

housekeeping gene 34B4. Sequences of primers are available on request.

5.3.4 - Beta-Galactosidase staining

Cells were washed with PBS and fixed with 3.5% paraformaldehyde in PBS for

10 min on ice. Cells were washed 3 times with PBS, incubated with X-gal (0.25 mg/ml)

and solubilised in reaction buffer (5 mM potassium ferricyanide, 5 mM potassium

ferrocyanide, 2 mM magnesium chloride in PBS) for 16 h at 37°C.



5.3.5 - MTT-assay

106 D3 cells or p53-/- cells were plated on 0.1% gelatinecoated 6 cm-plates.

After attachment, cells were irradiated with 2 Gray or left unirradiated for control.

Each day, MTT (3-[4,5-Dimethylthiazol-2-yl]-2,5-diphenyltetrazolium Bromide) was

added to two plates at a final concentration of 2.5 mg/ml and incubated for 4 h. The

reaction was stopped by removal of the medium and solubilization of the MTT-

precipitate in 2 ml isopropanol. Absorbances were read at 595 nm.

5.3.6 - Colony assay

200 ES cells were plated in 3.5 cm dishes coated with 0.2% gelatine. Twenty-

four hours after plating α-pifithrin and μ-pifithrin were added to a final concentration

of 10 μM. Two hours after drug addition, cells were irradiated with 0.5 Gy, 1 Gy, 2 Gy

and 4 Gy. After additional two hours, the culture medium was replaced with fresh

medium without inhibitors and the cells were incubated for two weeks with a daily

change of culture medium. The cells were washed with PBS, fixed with methanol,

stained with 1% crystal violet in PBS and counted. For colony assays in the absence of

an inhibitor, cells were irradiated at four hours after plating and incubated for two

weeks with a daily change of culture medium.

5.3.7 - Apoptosis Assay by Annexin V staining

1 × 106 D3 or p53-/- ES cells were washed with ice-cold PBS and resuspended in

400 ml Ca-containing buffer (10 mM HEPES, pH 7.4, 140 mM NaCl, 5 mM CaCl2). 5 μl of

annexin V-FITC and 1 μg propidium iodide were added for 10 min and the cells were

immediately analyzed by flow cytometry.

CHAPTER VI

IMAGE PROCESSING AND ANALYSIS

CHAPTER VI – IMAGE PROCESSING AND ANALYSIS



Digital image processing is an area characterized by the need for extensive

experimental work to establish the viability of proposed solutions to a given problem.

An important characteristic underlying the design of an image processing system is the

significant level of testing and experimentation that normally is required before

arriving at an acceptable solution. This characteristic implies that the ability to

formulate approaches and quickly prototype candidate solutions generally plays a

major role in reducing the cost and time required to arrive at a viable system

implementation (González, 2004).

MATLAB is a high-performance language for technical computing. It integrates

computation, visualization, and programming in an easy-to-use environment where

problems and solutions are expressed in familiar mathematical notation. Typical uses

include the following:

Math and computation;

Algorithm development;

Data acquisition;

Modeling, simulation and prototyping;

Data analysis, exploration and visualization;

Scientific and engineering graphics;

Application development, including graphical user interface building.

MATLAB is an interactive system whose basic data element is an array that

does not require dimensioning. This allows formulating solutions to many technical

computing problems, especially that involving matrix representation, in a fraction of

the time it would take to write a program in a scalar non-interactive language such as C

or Fortran (González, 2004).

The name MATLAB stands for matrix laboratory and was written originally to

provide easy access to matrix software developed by the LINPACK (Linear System

Package) and EISPACK (Eigen System Package) projects (González, 2004).



The Image Processing Toolbox is a collection of MATLAB functions (called M-

functions or M-files) has extended the capability of the MATLAB environment for the

solution of digital image processing problems (González, 2004).

6.2 – IMAGE PROCESSING AND ANALYSIS

In this work, it is performed the image processing and analysis of the images

withdrawn from the following papers:

“Nuclear accumulation and activation of p53 in embryonic stem

cells after DNA damage”, of Valeriya Solozobova, ALexandre Rolletschek

and Christine Blattner, BMC Cell Biology 2009 10:46;

“Lack of p53 function promotes radiation-induced mitotic

catastrophe in mouse embryonic fibroblast cells”, of Fiorenza Ianzini,

Alessandro Bertoldo, Elizabeth A Kosmacek, Stacia L Philips and Michael A

Mackey, Cancer Cell International 2006 6:11.

The objective of that image processing and analyze was to improve, through

the image interpretation, the results achieved in the referred papers relating to the

cellular response to irradiation regarding the p53 gene.

For these propose, the images were enhanced and segmented through the

following procedure:

1. Read the input image;

2. Execute contrast adjustment to better understand the image

data, namely for images with low-contrast as it is the case. This is a fairly

low-contrast image, so I thought it might help;

3. Remove noise from the image and then overlay the perimeter on

the original image;

4. To separate the cells properly, one possible approach is the

marker-based watershed segmentation. With this method it is possible to

connect a partial group of connected pixels inside each object to be

segmented. The extended maxima operator is used to identify groups of



pixels that are significantly higher than their immediate surroundings and

then the image is overlaid again;

5. Remove noise from the result and then overlay it;

6. Complement the image so that the peaks become valleys.

7. Modify the image so that the background pixels and the

extended maxima pixels are forced to be the only local minima in the

image.

8. Trace the object boundary to better visualization of the

individual/group of cells.

These steps correspond to the following sequence of MATLAB commands, in

this case, of one of the images from the first paper mentioned earlier:

Read and show the image:

g1=imread('p53_IA.png'); g=rgb2gray(g1);

imshow (g)

Contrast adjustment, and here there is the representation of

both the histogram and the equalized histogram to a better

understanding of what is done:

g_eq = adapthisteq(g);

subplot (1,2,1), imhist (g), title('Histogram') subplot (1,2,2), imhist (g_eq), title ('Equalized Histogram')



o Cleaning the image and overlay the perimeter on the original image:

bw2=imfill(bw,'holes');

bw3=imopen(bw2,ones(5,5)); bw4=bwareaopen(bw3,40); bw4_perim=bwperim(bw4);

overlay1=imoverlay(g_eq,bw4_perim, [.3 1 .3]); imshow (overlay1)

o Extended-maxima transform and carrying out some morphological

operations followed by cleaning and image overlay:

mask_em=imextendedmax(g_eq,30);

mask_em=imclose(mask_em,ones(5,5)); mask_em=imfill(mask_em,'holes'); mask_em=bwareaopen(mask_em,40);

overlay2=imoverlay(g_eq,bw4_perim | mask_em, [.3 1 .3]); imshow (overlay2)

0

200

400

600

800

1000

1200

1400

1600

Histogram

0 50 100 150 200 250

0

200

400

600

800

1000

1200

Equalized Histogram

0 50 100 150 200 250

0

200

400

600

800

1000

1200

1400

1600

Histogram

0 50 100 150 200 250



o Complement the image and impose minima to the image:

g_eq_c=imcomplement(g_eq);

g_mod=imimposemin(g_eq_c,~bw4 | mask_em);

o Trace the object boundary:

dim = size (g_mod)

col = round(dim(2)/2)-90; row = min(find(g_mod(:,col)))

boundary = bwtraceboundary(g_mod,[row, col],'N'); imshow(g_mod)

hold on; plot(boundary(:,2),boundary(:,1),'b','LineWidth',2);

g_mod_filled = imfill(g_mod,'holes'); boundaries = bwboundaries(g_mod_filled);

for k=1:13 b = boundaries{k};

plot(b(:,2),b(:,1),'b','LineWidth',2); end

This procedure was executed for every image from both articles mentioned

before, and the result is:

1. “Nuclear accumulation and activation of p53 in embryonic stem cells after DNA

damage”:

a. Light microscopy photos of the mutant p53 cell line MEF 10(1) following

10 Gy γ-irradiation.



i. Morphology of MEF 10(1) cells at 32 hours post-irradiation:

ii. Morphology of sham irradiated MEF 10(1) cells, at time zero:

b. Light microscopy photos of wild-type p53 cell line MEF 12(1) following

10 Gy γ-irradiation:





2. “Lack of p53 function promotes radiation-induced mitotic catastrophe in

mouse embryonic fibroblast cells” of Fiorenza Ianzini, Alessandro Bertoldo,

Elizabeth A Kosmacek, Stacia L Philips and Michael A Mackey, Cancer Cell

International 2006 6:11:

a. Cytoplasmic localisation of p53 in proliferating ES cells. R1, D3 and CGR8

mouse embryonal stem cells were plated on cover slips (R1 and D3 in

the presence of feeders, CGR8 in the absence of feeders):

i. P53 in R1, D3 and CGR8:



ii. Cells were incubated in Draq5 (R1, D3 and CGR8):

b. p53 accumulates in the nucleus of irradiated ES cells:

i. R1 (p53 and Draq5) and D3 (p53 and Draq5) stem cells, grown on

feeders, were irradiated two days after plating with 7.5 Gray IR

(0, 1 and 2h first row, 4,8 and 24h second row):

1. R1 – p53



2. R1 – Draq5



3. D3 – p53

4. D3 – Draq5



ii. R1 stem cells, grown on feeders, were irradiated with 30 J/m2

UVC light (0, 1 and 2h first row, 4,8 and 24h second row):

1. R1 – p53

2. R1 – Draq5



6.3 - COMPARISON BETWEEN ORIGINAL AND PROCESSED IMAGES

In the paper “Nuclear accumulation and activation of p53 in embryonic stem

cells after DNA damage”, nuclear fragmentation is persistent during the time course of

the experiment and reaches values of about 80% at 48 hours post-irradiation.

Morphological changes in irradiated p53 mutant MEF 10(1) cells were detected. The

presence of fragmented nuclei and changes in cell shape are evident for the irradiated

population. Irradiated MEF 12(1) cells also confirm that the morphological features of

the irradiated cell population do not greatly differ from the control population, except

that a greater number of large cells is observed, probably reflecting the persistent G2

arrest.

In order to highlight these results, the light microscopy photos obtained with

this study were processed using MATLAB. The comparison between the images is the

following:





iii. Morphology of MEF 12(1) cells at 40 hours post-irradiation:

iv. Morphology of sham irradiated MEF 12(1) cells, at time zero:

The processed image allows a better visualization of points with higher

intensity in the cells as well as improved individualization of the cells/group of cells.

With these results the conclusion of the articles are highlighted since the

morphological changes due to irradiation are evident with the contoured cells, as well

as nuclear fragmentation in the MEF 10(1) cells at 32 hours post-irradiation (more

diffuse intracellular content).

In the paper “Lack of p53 function promotes radiation-induced mitotic

catastrophe in mouse embryonic fibroblast cells”, they tried to clarify the activity of

p53 in stem cells, determining the localization of the p53 protein in mouse ES cells,

investigating three different mouse ES cell lines, R1, D3 and CGR8 and determined p53

localization by immuno-fluorescence staining. R1 and D3 cells were cultured on feeder

cells while CGR8 stem cells do not require feeders for maintaining an undifferentiated

phenotype. They found the majority of p53 localized in the cytoplasm. However, these



results do not exclude the possibility that a small fraction of p53 exists in the nucleus.

As a transcription factor, p53 usually needs to be nuclear to be active.

One particularly important function of p53 is DNA damage signaling. Here, to

suppress tumorigenesis, p53 halts the cell cycle and induces apoptosis in primary cells

and in tumor cell lines. Since stem cells provide the pool of proliferative

pluri/toti/omni-potent cells within organisms, they are more likely to propagate DNA

lesions and mutations to daughter cells compared to differentiated cells.

Since p53 is primarily a transcription factor, nuclear localization of p53 should

be essential for its transcriptional activity and p53 accumulates in the nucleus of ES

cells after DNA damage.

In unstressed ES cells p53 was localized mainly to the cytoplasm. However, at

one hour after irradiation, p53 accumulated in the nucleus of irradiated ES cells. At

four hours after irradiation, p53 was still present in the nucleus of some cells, while in

others it had mostly disappeared. At eight hours after irradiation p53 had essentially

disappeared from the nucleus of all cells. Surprisingly, at 24 hours after IR, p53 was

again localized in the nucleus of most cells. Nevertheless, a minority of cells still

showed cytoplasmic localization of p53.

After UV-irradiation, p53 also accumulated in the nucleus of ES cells. In contrast

to IR-irradiated cells, p53 remained in the nucleus of UV-irradiated cells. At 24 hours,

most of the ES cells had died while the few remaining ones showed a very intensive

nuclear staining for p53.

Once again, to highlight these results, the light microscopy photos obtained

with this study were processed using MATLAB and here is the comparison between the

images:

i. Cytoplasmic localization of p53 in proliferating ES cells, R1, D3 and CGR8 mouse

embryonal stem cells:



- R1 – p53

- D3 – p53

-

-

-

- CGR8 – p53



– R1 – Draq5

-

-

-

- D3 - Draq5

- CGR8 - Draq5



- R1 - p53 - 0h

-

-

-

- R1 - p53 - 1h

-

-

- R1 - p53 - 2h



- R1 - p53 - 4h

-

-

-

- R1 - p53 - 8h

-

-

-

- R1 - p53 - 24h



- R1 – Draq5 - 0h

-

-

-

- R1 – Draq5 - 1h

-

-

- R1 – Draq5 - 2h



- R1 – Draq5 - 4h

-

-

-

- R1 – Draq5 - 8h

-

-

-

- R1 – Draq5 - 24h



- D3 - p53 – 0h

-

-

-

- D3 - p53 – 1h

-

-

-

-

- D3 - p53 – 2h



- D3 - p53 – 4h

-

-

-

- D3 - p53 – 8h

-

-

-

- D3 - p53 – 24h



- D3 - Draq5 – 0h

-

-

-

- D3 - Draq5 – 1h

-

-

-

- D3 - Draq5 – 2h



- D3 - Draq5 – 4h

-

-

-

- D3 - Draq5 – 8h

-

-

-

- D3 - Draq5 – 24h



- R1 – p53 – UV - 0h

-

-

-

- R1 - p53 – UV - 1h

-

-

-

- R1 - p53 – UV - 2h



- R1 - p53 – UV - 4h

-

-

-

- R1 - p53 – UV -8h

-

-

-

- R1 - p53 – UV - 24h



- R1 – Draq5 – UV - 0h

-

-

-

- R1 – Draq5 – UV - 1h

-

-

-

- R1 – Draq5 – UV - 2h



- R1 – Draq5 - UV - 4h

- R1 – Draq5 – UV - 8h

- R1 – Draq5 – UV - 24h

Once again, the processed images enable a superior visualization of points with

higher intensity in the nucleus (shown with Draq5) and in the cytoplasm. The results

described in the paper were better understood with the processed images.



6.4 – SUMMARY

It is possible that p53 functions to repress radiation-induced mitotic

catastrophe (MC) through its activity as a modulator of the G2 checkpoint mechanisms,

and the lack of p53 promotes MC as a mechanism for removing damaged mouse

embryonic fibroblast cells from populations (Ianzini, 2006).


demonstrates the morphological changes in irradiated p53 mutant MEF 10 (1) cell line.

The presence of fragmented nuclei and changes in cell shape are evident for irradiated

population and are hallmarks of MC. In the irradiated MEF 12 (1) cells there are a

greater number of larger cells probably due to the persistent G2 arrest, suggesting the

important role of p53 in the induction of MC following irradiation.

P53 is localized in the cytoplasm of embryonic stem (ES) cells and activates

transcription of a reporter gene in resting ES cells and of endogenous target genes in

response to DNA damage. The activity of p53 in resting stem cells shows that the p53

protein is also in ES cells in a latent state and can be activated when its activity is

required. However, after DNA damage, p53 did not activate all, but activates at least

some of its target genes in ES cells and the transcription of these genes was facilitated

by the presence of active p53 in the nucleus of irradiated cells (Solozobova, 2009).

After IR, p53 accumulated in two waves in the nucleus of ES cells. The first

nuclear accumulation occurred at one to two hours after irradiation and correlated

with an increase in the amount of the p53 protein. During the second wave of nuclear

accumulation of p53 at twenty-four hours after irradiation, there is not an increase in

p53 abundance suggesting that for the second wave of nuclear accumulation p53 was

translocated from the cytoplasm into the nucleus (Solozobova, 2009).


emphasize the changes in intracellular content due to irradiation of cells, and

emphasize the cytoplasmic localization of p53 in proliferating ES cells and the evidence

that p53 accumulates in the nucleus of irradiated ES cells.

Defining the mechanisms underlying the role of p53 in MC might lead to

strategies to improve clinical radiation response for those human tumors with defects

in p53, or p53-related pathways, and that avoid radiation induced apoptosis. If



radiation-induced MC is the predominant mode of cell death in p53-deficient cells,

clinical interventions designed to enhance its production might not affect surrounding

normal tissue, and thus lead to a therapeutic gain (Ianzini, 2006).

CHAPTER VII

CONCLUSIONS AND FUTURE WORKS

CHAPTER VII – CONCLUSIONS AND FUTURE WORKS


7.1 – FINAL CONCLUSIONS

Clearly that wild-type (WT) p53 not only has anti-proliferative and anti-

transforming activity but also possesses the ability to induce programmed cell death

(apoptosis) after exposure of cells to DNA-damaging agents, such as γ-irradiation or

anticancer drugs. The concept that p53 is a growth regulatory protein fits with its short

half-life, its nuclear location and transcription factor activity, and its increased

synthesis in DNA damaged cells. WT p53 regulates the transcription of a number of cell

replication associated genes (Ruddon, 2007).

Growth arrest induced by WT p53 blocks cells prior to or near the restriction

point in late G1 phase and produces a decrease in the mRNA levels for genes involved

in DNA replication and cell proliferation, such as histone H3, proliferating cell nuclear

antigen (PCNA), DNA polymerase α, and b-myb. To carry out these gene regulatory

events, WT p53 has to assume a certain conformational structure, apparently

modulated by its phosphorylation state, and oligomerize so that it can bind to DNA.

Mutant p53 cannot achieve the appropriate conformation and can block WT p53

function by forming oligomers with it. Even though a lot is known about the biological

actions of p53, e.g., the ability to induce G1 arrest, to induce apoptosis following DNA

damage, to inhibit tumor cell growth, and to preserve genetic stability, the way in

which it does all this is not totally clear (Ruddon, 2007).

The p53 network can be activated by at least three mechanisms. The first is

DNA strand breaks triggered by ionizing radiation or other DNA-damaging agents. This

mechanism is dependent on activation of the ATM (ataxia telangiectasia-mutated)

protein, Chk2, or other kinases. Interestingly, mice that are deficient in p53 function

and in the ability to repair DNA double-strand breaks because of a failure in non-

homologous end-joining (NHEJ) repair, develop highly aggressive pro-B cell

lymphomas. The second mechanism is overexpression or aberrant expression of

growth factor signals such as those turned on by oncogene proteins Ras or Myc.

Finally, cellular stress is induced by chemotherapeutic drugs, ultraviolet light, or

protein kinase inhibitors (Ruddon, 2007).

The role of p53 in maintaining genetic stability appears to involve induction of

genes that stimulate nucleotide excision repair, chromosomal recombination,

CHAPTER VII – CONCLUSIONS AND FUTURE WORKS


chromosome segregation and induction of the gene for ribonucleotide reductase. p53

also stimulates the expression of genes that inhibit angiogenesis (Ruddon, 2007).

The main scientific reports in which this work is based on are concerning the

role of p53 in the cellular behavior. The right and normal performance of these check

point mechanisms are very important to keep the equilibrium in the cellular turnover.

To be able to perform the adequate study of cells it is important to always have

in mind the proper behavior in laboratory, to ensure that there is no contamination in

the medium for culture cells (chapter IV).

To understand this kind of study, it is imperative to have knowledge regarding

the radiation effects in normal and neoplastic tissues (chapter III). It´s also important

to understand the mechanisms of cell cycle regulation and apoptosis (chapter II).

7.2 – FUTURE WORKS

The future prospect of this thesis is to continue the study with cells, performing

the analysis and image processing of breast cancer cells submitted to brachytherapy.

Hence, the study of morphological changes that occur in the irradiated cells, as well as

the modifications in the cellular environment to obtain the maximum information of

the electron microscopy images of these cells, will be done. Additionally,

computational algorithms will be developed to help that study in an automate and

robust manner.

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