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The biology of three novel natural product microtubule interacting agents : ceratamine A and B and dimethyl… Karjala, Geoffrey William George 2004

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THE BIOLOGY OF T H R E E N O V E L N A T U R A L PRODUCT M I C R O T U B U L E INTERACTING AGENTS: CERATAMINE A AND B AND DIMETHYL VARACIN  by  G E O F F R E Y WILLIAM G E O R G E K A R J A L A  B.Sc, The University of British Columbia, 2001  A THESIS SUBMITTED IN PARTIAL F U L F I L M E N T OF T H E REQUIREMENTS FOR T H E D E G R E E OF  M A S T E R OF SCIENCE  in  T H E F A C U L T Y OF G R A D U A T E STUDIES  (Department of Biochemistry and Molecular Biology)  T H E UNIVERSITY OF BRITISH C O L U M B I A December 2004  © Geoffrey William George Karjala, 2004  Abstract  Microtubules are dynamic polymers of the protein tubulin. The dynamic property of microtubules allows the network to breakdown and form new structures like the mitotic spindle. Microtubule associated proteins and small molecules can alter the dynamics. Changing the dynamics by the addition of small molecules can prevent the proper formation of the mitotic spindle and lead to mitotic arrest. Some of these antimitotic agents have had clinical success in cancer treatment with the vinca alkaloids inducing the underpolymerisation and paclitaxel (Taxol™) promoting the overpolymerisation of microtubules. In addition, small molecules that alter the microtubule network are gaining use in chemical genetics. A phenotypic antimitotic assay has been used to discover three novel compounds that interact with the microtubule network. The first two, ceratamine A and B are classic antimitotic agents that block cells in M-phase and prevent proliferation, probably by disrupting the microtubule network. They stimulate the over-polymerization of microtubules in vitro as determined by both microtubule polymerization assays and electron microscopy. In vivo, ceratamines induce the formation of tubulin-containing structures not previously described. These structures include pillars of tubulin in mitotic cells and a perinuclear microtubule network during interphase. Ceratamines do not compete with paclitaxel for binding to the microtubules, and are very structurally simple. The same phenotypic antimitotic assay was used to discover dimethyl varacin (DMV). Flow cytometry data indicated that D M V is not an antimitotic agent and does not block cells in any particular phase of the cell cycle, though it does inhibit proliferation. It  ii  induces strong GF-7 phospho-specific antibody binding (typical only in mitosis) from all phases of the cell cycle. Western blots and Kinexiis Kinetworks™ screens showed that a two hours D M V treatment at 5 uM leads to the strong activation of M A P K pathways (Erkl/2, p38 a M A P K , and JNK/SAPK) without a significant increase in Cdc2 activity or global phosphorylation. It was also found to strongly inhibit microtubule formation in vitro and in vivo. Though both D M V and the ceratamines were discovered in the same antimitotic screen and both target microtubules, they have dramatically different biological properties.  in  Table of Contents Section  Page  Abstract  ii  Table of Contents  iv  List of Figures  vii  List of Symbols, Nomenclature, and Abbreviations  viii  Acknowledgements  xiii  Chapter I - General Introduction  1  1.1- The Cytoskeleton  1  1.1.1 - Overview  1  1.1.2- Microtubules and Tubulin, Inseparable Partners  1  1.1.3- Microtubules and the Cell Cycle  7  1.1.4 - Microtubule Poisons  12  1.1.5- Microtubules and Microtubule-Associated Proteins  14  1.2- M A P Kinase Signaling  16  1.2.1- Mitogen Activated Protein Kinases (MAPK)  16  1.2.2 - M A P K and the Cytoskeleton  20  1.2.3 - M A P Kinase Phosphatases (MKP)  21  1.3- Goals  22  Chapter II - Materials and Methods  23  2.1-Materials  23  2.2 - Antimitotic Assay  23  2.3 - Antiproliferation Assay  23  iv  2.4 - In Vitro Tubulin/Microtubule Assays  23  2.4.1 - Promotion of Tubulin Polymerization Assay  23  2.4.2 - Inhibition of Tubulin Polymerization Assay  23  2.4.3 - Negative Staining Electron Microscopy of Microtubules  24  2.4.4 - H-Paclitaxel Competition Binding to Microtubules  24  3  2.5 - Flow Cytometry Analysis of the Cell Cycle  25  2.6 - Immunofluorescence Microscopy  25  _  2.6.1 - P-Tubulin Immunofluorescence Microscopy of D M V Treated Cells  25  2.6.2 - p-Tubulin Confocal Immunofluorescence Microscopy of Ceratamine Treated Cells _  25  2.6.3 - p-Tubulin Immunofluorescence Microscopy of D M V Treated Cells  26  2.7 - Kinase Assays  26  2.7.1 - Cdc2 Kinase Assays  26  2.7.2 - In-Gel Kinase Assay  26  2.8-Western Blots  26  2.9 - Kinexus Kinetwork™ Phosphosite Screen  27  2.10- Protein Phosphatase 2A Assay  27  Chapter III - The Ceratamines  29  3.1 - Ceratamine Specific Introduction  29  3.2-Results  31  3.2.1 - Effect of Ceratamine A and B on Cell Cycle Progression  31  3.2.2 - Antimitotic and Antiproliferative Assay  34  3.2.3 - Effect of Ceratamines on Microtubule Organization In Vivo  34  3.2.4 - Effect of Ceratamines on Microtubule Formation In Vitro  39  3.2.5 - Ceratamines do not Compete with Paclitaxel for Binding to Tubulin 3.3 - Discussion  ^ 45  Chapter IV - Dimethyl Varacin  52  4.1 - Dimethyl Varacin Specific Introduction  v  52  4.2 - Results  54  4.2.1 - Antimitotic Assay and Antiproliferation  54  4.2.2 - Dimethyl Varacin Inhibits Microtubule Polymerization  54  4.2.3 - Dimethyl Varacin Rapidly Breaks Down the Microtubule Network In Vivo 4.2.4 - Dimethyl Varacin Induces Mitotic-Like Phosphorylations from A l l Phases of the Cell Cycle  57 57  4.2.5 - Dimethyl Varacin does not Cause Global Activation of Cdc2  60  4.2.6 - In Gel Kinase Assays of Lysate from Dimethyl Varacin Treated Cells  62  4.2.7 - Kinexus Kinetworks™ Phosphosite Screens  65  4.2.8 - Confirmation Western Blots of Erkl/2, SAPK/JNK, and p38 M A P K Phosphorylation  75  4.2.9 - Protein Phosphatase 2A is not Inhibited by Dimethyl Varacin  78  4.3 - Discussion  79  References  89  vi  List of Figures Figure Title  Page  Figure 1 - Structure of a Microtubule  3  Figure 2 - The Cell Cycle  8  Figure 3 - Microtubule Dynamics and Polymer Mass During the Cell Cycle  10  Figure 4 - The Mitotic Spindle  11  Figure 5 - M A P Kinase Activation and Deactivation Pathways  18  Figure 6 - The Chemical Structure of Paclitaxel, Laulimalide, and Peloruside A  30  Figure 7 - The Chemical Structure of Ceratamine A and B  32  Figure 8 - Flow Cytometry of MCF-7 mp53 Cells Treated with DMSO, Nocodazole, Ceratamine A or Ceratamine B Figure 9 - Antimitotic and Cell Proliferation Assays with Ceratamine A andB Figure 10 - Confocal p-Tubulin Immunofluorescence Images of Ceratamine or Paclitaxel Treated Cells  33 35 37  Figure 11 - Cross Section P-Tubulin Staining in Mitotic Cells  38  Figure 12 - In vitro Tubulin Polymerization Assays with Ceratamine A and B  40  Figure 13 - Electron Microscopy Photos of Microtubules  41  Figure 14 - H-Paclitaxel Binding to Microtubule Assay  44  Figure 15 - The Chemical Structure of Varacin Compounds  53  Figure 16 - Antimitotic ELICA and Cell Proliferation Assays with Various Concentrations of Dimethyl Varacin and Different Cell Lines  55  Figure 17 - Dimethyl Varacin Inhibits Microtubule Formation In Vitro  56  Figure 18 - Dimethyl Varacin Disrupts Microtubules In Vivo  58  Figure 19 - Dimethyl Varacin's Effect on the Cell Cycle  59  Figure 20 - Dimethyl Varacin Does Not Activate Cdc2/cyclin B  61  Figure 21 - In-Gel Kinase Assays  63  3  Figure 22 - Summary of Data from Kinexus Kinetworks™ Phosphosite Screens Figure 23 - Western Blots of Phospo-Erkl/2, Phospho-JNK, Phosphop38 MAPK, and Total M A P K from D M V Treated Cell Lysates Figure 24 - Protein Phosphatase 2A Assay  66-67 76 77  vii  List of Symbols, Nomenclature, and Abbreviations  Abbreviation  Name  y-TuRC  y Tubulin Ring Complex  AMP-PK  A M P Dependent Protein Kinase  ASK1  Apoptosis Signal-Regulating Kinase 1  ATF1  Activating Transcription Factor 1  ATF2  Activating Transcription Factor 2  BSA  Bovine Serum Albumin  CaMK2  Ca /Calmodulin-Dependent Protein Kinase 2  Cdc  Cell Division Control  CDK  Cyclin Dependent Kinase  CREB  cAMP Responsive Element Binding Protein  CTKD  C-Terminal Kinase Domain  ddGTP  dideoxy Guanosine Triphosphate  DLK  Dual Leucine Zipper-Bearing Kinase  DMSO  Dimethyl Sulfoxide  DMV  Dimethyl Varacin  DNA  Deoxyribonucleic Acid  DSP  Dual Specificity Phosphatase  DTT  Dithiothreitol  E  Exchangeable Site of Tubulin  EGFR  Epidermal Growth Factor Receptor  2+  viii  gQ-p^  Ethylene glycol-bis(2-aminoethylether)-N,N,N',N'tetraacetic acid  eIF2Bs  Eukaryotic Initiation Factor 2B s Subunit  eIF2a  Eukaryotic Initiation Factor 2a  eIF4  Eukaryotic Initiation Factor 4  eIF4E-BPl  Eukaryotic Initiation Factor 4E-Binding Protein 1  ELICA  Enzyme Linked Immunocytochemistry Assay  ErbB2  Erb2/HER2 Receptor Tyrosine Kinase  Erk  Extracellular Regulated Kinase  FACS  Fluorescence-Activated Cell Sorting  FAK  Focal Adhesion Kinase  GDP  Guanosine Diphosphate  GDP-Tubulin  Tubulin with GDP at the Exchangeable Site  GRK2  G Protein-Coupled Receptor-Serine Kinase 2  GSK3  Glycogen Synthase Kinase 3  GTP  Guanosine Triphosphate  GTP-Tubulin  Tubulin with GTP at the Exchangeable Site  HPLC  High Performance Liquid Chromatography  IC50  Concentration of 50% Inhibition  IGFR  Insulin Growth Factor Receptor  IKK  I kappa B Kinase  IRS 1  Insulin Receptor Substrate 1  INK  c-Jun N-terminal Kinase  ix  K-fibre  Kinetochore Fibre  MAP  Microtubule Associated Protein  MAPK  Mitogen Activated Protein Kinase  MAPKAPK-2  M A P Kinase Associated Protein Kinase -2  MAPKK  M A P K Kinase  MAPKKK  M A P K K Kinase  MBP  Myelin Basic Protein  MEF2A  Myocyte Ehancer Factor 2A  MEK  MAP/Erk Kinase  MEKK  M E K Kinase  MK  M A P K Activated Protein Kinase  MKK  M A P Kinase Kinase  MKP  M A P Kinase Phosphatase  MLK  Mixed Linage Kinase  MNK  M A P K Interacting Kinases  MNK1  M A P K Interacting Kinase 1  MOPS  3-(N-Morpholino)propanesulfonic acid  MSG  Monosodium Glutamate  MSK  M A P K and Stress Activated Protein Kinases  MSK1  M A P K and Stress Activated Protein Kinase 1  MSK2  M A P K and Stress Activated Protein Kinase 2  MTOC  Microtubule Organizing Centre  x  mTOR  Mammalian Target of Rapamycin  J^IYY  3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium  bromide N  Nonexchangeable Site of Tubulin  NF-KB  Nuclear Factor K B  Op 18  Oncoprotein 18  PAGE  Polyacrylamide Gel Electrophoresis  PAK  p21 Activated Kinase  PDK1  Phospho-Inositide Dependent Kinase 1  PIPES  Piperazin-l,4-bis(2-ethanesulfonic  PKB  Protein Kinase B  PKC  Protein Kinase C  PKD  Protein Kinase D  PKN  Protein Kinase N  PKR  Double Stranded RNA-Dependent Protein Kinase  PPI a  Protein Phosphatase l a  PP2A  Protein Phosphatase 2A  PRK1  Protein Kinase C-Related Protein-Serine Kinase 1  PRK2  Protein Kinase C-Related Protein-Serine Kinase 2  PVDF  Polyvinyldifluoride  Rb  Retinoblastoma  RSK  Ribosomal S6 Kinase  RSK1  Ribosomal S6 Kinase 1  xi  acid)  RTK  Receptor Tyrosine Kinase  SAPK  Stress Activated Protein Kinase  SDS  Sodium Dodecyl Sulphate  SMAD  SMA- and M A D - Related Protein  STAT  Signal Transducer and Activator of Transcription  TAK1  Transforming Growth Factor-beta-Associated Kinase  TBS  Tris Buffered Saline  TBS-T  Tris Buffered Saline - 0.1% Tween 20  XKCM1  Xenopus Kinesin Catastrophe Modulator 1  XMAP  Xenopus Microtubule Associated Protein  Xll  Acknowledgements There are many people who I wish to thank. First and foremost is Diane, my wife, whose support during my Master's degree has been immense. I also wish to thank Dr. Roberge for welcoming me into his lab and providing guidance and assistance always with a smile. In addition to Dr. Roberge, the other two members of my supervisory committee, Dr. Andersen and Dr. Numata have been very helpful. The support from the Roberge lab has been wonderful. Along with the lab, there are agencies and people that without whose help the projects I worked on would not have been possible. NSERC provided the money for my stipend and CIHR supported the operation of the Roberge lab. The E7 monoclonal antibody developed by Michael Klymkowsky was obtained from the Developmental Studies Hybridoma Band developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA 52242. Radiolabeled paclitaxel was obtained from the Drug Chemistry and Synthesis Branch, National Cancer Institute. The U B C Bio-imaging facility was also very helpful in obtaining the confocal immunofluorescence images and the electron microscopy images. Andy Johnston at the U B C multi-user flow cytometry facility helped obtain the flow cytometry data. The dimethyl varacin project was greatly aided by two people. First, Lu Yang from the Andersen lab must be mentioned for isolating the compound. Second, Cristina Bigg of the Roberge lab deserves much credit for the project. She is responsible for the data used to prepare Figures 16, 19, 20, and 24. Also, Dr. Pelech and Dr. Zhang of Kinexus were very helpful.  xiii  The ceratamine project also benefited from the help of others. Emiliano Manzo of the Andersen lab is responsible for isolating the two compounds. Queenie Chan of the Roberge lab also contributed to the project and is responsible for Figures 9, 10, and 11.  xiv  Chapter I: General Introduction 1.1- The Cytoskeleton 1.1.1 - Overview Cells must exist in a three-dimensional world where external forces are always present and in some cases movement is required. In addition, cells must organize their internal structures, transport vesicles and segregate chromosomes at the appropriate time. In response, cells have evolved a system of filaments to provide a source of structural strength, to partition chromosomes during mitosis, to be "trails" for vesicles, to provide motility, and to organize the shape of the cell. These filaments are in fact, three distinct protein polymer classes that together form the cytoskeleton. The first polymer class is actin filaments (microfilaments), which are composed of actin subunits; the second is intermediate filaments composed of intermediate filament proteins; and the third, and the one to be discussed further, is microtubules composed of tubulin subunits. Microtubules partition the organelles in the cell and are responsible for the separation of chromosomes during mitosis. They also form cilia bodies and provide strength to the cell.  1.1.2 - Microtubules and Tubulin, Inseparable Partners Microtubules are polymers of the protein tubulin. The microtubules serve many functions within the cell such as providing strength, shape, and 'traffic' lanes. They also form the mitotic spindle that separates chromosomes during mitosis. To perform these functions tubulin and microtubules must be very adaptable. The innate dynamic instability of microtubules (Desai et al., 1997) allows the microtubules to rearrange and creates this adaptability.  1  Tubulin was initially discovered based on its irreversible binding to the microtubule poison colchicine (Weisenberg et al., 1968). Further work determined that tubulin was a heterodimer of a and p tubulin (Feit et al., 1971) and bound two molecules of guanine nucleotide per mole of tubulin dimer (Weisenberg et al., 1968). One of the guanine nucleotides is GTP bound to a non-exchangeable (N) site. The other guanine nucleotide is far more dynamic, and it is bound at an exchangeable (E) site on the tubulin heterodimer (Nogales et al., 1998). The GTP at the E site is crucial to the correct dynamics of the microtubule. Because tubulin is a heterodimer, it is asymmetrical. As such, the resulting microtubule is asymmetrical with a minus (-) and a plus (+) end. This polarity also plays a critical role in the dynamics of the microtubule as the two ends have different properties (Desai et al., 1997). The microtubule itself is a hollow tube of tubulin typically composed of 13 protofdaments in mammalian cells (Tilney et al., 1973; Evans et al., 1985). A protofilament is a linear polymer of tubulin, and a region of tubulin, the M-loop, is crucial for lateral contacts between protofilaments in the assembly of a microtubule (Li et al., 2002). A schematic of the microtubule structure is shown in Figure 1. It is possible for the number of protofilaments to vary from as few as 8 to as many as 17 (Pierson et al., 1978; Aamodt et al., 1986; Chretien et al., 1992) depending on temperature (Pierson et al., 1979), tubulin source (Aamodt et al., 1986), Microtubule Associated Proteins (MAPs), buffer conditions (Pierson et al., 1978), small molecules (Diaz et al., 1998) and site of nucleation (Evans et al., 1985). While it was initially thought that the number of protofilaments was determined at nucleation, it has now been shown that the number can vary within one microtubule (Chretien et al, 1992).  2  TJLC  : - • • n>~  nut En)  Figure 1 -  The Microtubule Structure  A microtubule is a hollow cylindrical polymer of tubulin. The cylinder is approximately 25 nm across and typically composed of 13 protofilaments in mammalian cells. A protofilament is a linear polymer of tubulin. Figure from Kline-Smith and Walczak (2004).  3  As mentioned previously, microtubules are very dynamic (Desai et al., 1997). They undergo periods of rapid growth where tubulin is added to the plus end before a 'catastrophe' occurs and the microtubule begins to shorten. After a period of shrinkage a 'rescue' can occur and the microtubule will begin to grow again (For an excellent review see Desai and Mitchison 1997). It is thought that these dynamics are in part regulated by the GTPase activity of tubulin. The microtubule is stabilized by a cap of tubulin with GTP at the E site. To this microtubule GTP-tubulin (tubulin with GTP at the E site) may be added. Once bound, the GTP is hydrolyzed to GDP. GDP-tubulin at the plus end of a microtubule, as is thought to occur when the rate of hydrolysis exceeds the rate of polymerization, is not stable and will lead to a catastrophe and shortening. This is thought to occur because the hydrolysis of the GTP to GDP has an allosteric effect on tubulin which introduces a curve into the protofilaments (Howard et al., 1986). This will initiate a period of shrinkage where the protofilaments turn inside out and dissociate from the microtubule (Tran et al., 1997). This occurs until a rescue event, after which the microtubule will enter a period of growth. This is the GTP-cap model first proposed in 1984 (Mitchison et al., 1984). Questions still being examined in this model include the rate of GTP hydrolysis and polymerization, and the amount of GTP-tubulin which must be bound at the plus (+) end of the microtubule to stabilize it. Between periods of growth and shrinkage the microtubule may appear static. However, it may be undergoing a treadmill action (this is also known as a flux state) (Margolis et al., 1998). It is thought this occurs because of the polarity introduced by the asymmetry of tubulin. The two ends have differing affinities for tubulin and consequently the critical concentration will vary (Walker et al., 1988). The critical concentration of  4  tubulin is the concentration above which the microtubule will grow, and below which the microtubule will shrink. Because the critical concentration values vary between ends, it is possible for tubulin to be added to one end, typically the plus, while being lost at the other. This leads to an apparent static microtubule, but it is really dynamic with tubulin dimers being added to one end and lost at the other. This state is typically observed in vitro and is called treadmilling (Margolis et al., 1998). It is believed that treadmilling plays a role in in vivo microtubule dynamics and work is ongoing to demonstrate this (Margolis et al., 1998). The structure of tubulin has been solved allowing a more in-depth explanation of earlier observations. The structure was initially solved by electron crystallography and has since been refined to a 3.5 A resolution (Nogales et al., 1998). a and (3 tubulin are not very similar at the primary level with 40% sequence identity (Nogales et al., 1998). However, at the tertiary level, the two subunits are nearly superimposable (Nogales et al., 1998). The differences are confined to the loops, slight offsets of secondary structures, and amino acid side chains (Nogales et al., 1998). The C-terminus is a region of hypervariability, and it is the main area of difference between the respective isotypes (6 for a, 7 for P) (Luduena 1998). The solution of the tubulin dimer structure led to the identification of three functionally distinct domains. The N-terminal domain is a Rossman fold, and it is where the guanine nucleotide binds. The middle domain is where the majority of microtubule drugs bind, while the relatively short C-terminal domain is where MAPs bind (Nogales et al., 1998). The solution of the structure unequivocally identified the plus end as having the p subunit exposed, while the minus end has the a subunit exposed. It also showed that the N site occurs at the interface between the a and  5  P subunit within one heterodimer while the E site occurs between the P subunit of one dimer and the a subunit of another (Nogales et al., 1998). This, in part, explains why the E site and the hydrolysis of GTP plays such a critical role in the dynamics of the microtubule as the E site is exposed at the plus end of the microtubule. Tubulin is far more complex than described above with multiple isotypes transcribed from different genes, often in a tissue-dependent manner (Luduena 1998). The function for these isotypes is presently under debate (Luduena 1993; Luduena 1998). It does appear that under some circumstances the different isotypes are interchangeable (Bond et al., 1986; Gu et al., 1988) though not in all circumstances (Hoyle et al., 1990; Renthal et al., 1993). More work will be required to determine the roles for each of the isotypes. In addition to the multiple isotypes tubulin undergoes significant post translational modification (Luduena 1998; Westermann et al., 2003). These modifications can include phosphorylation of serine and possibly tyrosine (Eipper 1974; Luduena 1998); acetylation of the epsilon amino group of lysine 40 of several alpha tubulin isotypes (L'Hernault et al., 1985; Piperno et al., 1987; Luduena 1998); tyrosination/detyrosination of the C-terminus of a tubulin (Luduena 1998); the removal of the final two amino acids (tyrosine and glutamate) are removed in a tubulin to form A2-Tubulin (Paturle-Lafanechere et al., 1991; Luduena 1998); polyglutamylation, the addition of multiple glutamate residues to the gamma carboxyl group of a C-terminus glutamate residue, in some isotypes of alpha and beta tubulin (Edde et al., 1990; Mary et al., 1994; Luduena 1998); and polyglycylation, the addition of multiple glycine residues to the gamma carboxyl group of a C-terminus glutamae residues, in a and P tubulin of axenomal microtubules (Rudiger et al., 1995; Luduena 1998). The outcome of these  6  modifications is still being investigated, though acetylation, polyglutamylation and polyglycylation appear to be involved in motility (Luduena 1998; Westermann et al., 2003). In some instances the enzymes responsible for these modifications are still being sought. Further, it is clear that the modifications do not occur to the same degree on both a and p tubulin, with acetylation and tyrosination/detyrosination being observed solely on a and phosphorylation, polyglycylation, and polyglutamylation occurring on various isotypes of both a and P tubulin (Luduena 1998; Westermann et al., 2003). The microtubule network does not occur haphazardly within the cell: the Microtubule Organizing Centre (MTOC) of the cell organizes it. In animal cells, the M T O C is typically the centrosome, which contains two protein structures called centrioles. Remarkably centrioles duplicate in a semi-conservative manner and form two centrosomes such that a bipolar spindle can form during mitosis and each daughter cell will have a centrosome (Wong et al., 2003). A key component of the M T O C is the y tubulin ring complex (y-TuRC) (Zheng et al., 1995), a protein complex that contains y tubulin and serves to nucleate the formation of a microtubule (Zheng et a l , 1995). The minus end of the microtubule is capped by the y-TuRC with the positive end extending into the cytoplasm (Zheng et al., 1995). During interphase, the M T O C in animal cells is typically near the nucleus. The MTOCs of the cell separate to form the poles of the mitotic spindle at the onset of mitosis.  1.1.3 - Microtubules and the Cell Cycle The cell cycle consists of four phases: G i , S, G2 and M phase (Figure 2). Collectively G i , S, and G2 form interphase during which the cell grows, replicates its  7  Figure 2 - T h e C e l l C y c l e  The cell cycle is composed of four phases: Gi, S, G2, and M. Gi, S, and G2 collectively make up interphase. M phase is further divided into mitosis and cytokinesis. Mitosis can then be separated into five distinct stages: prophase, prometaphase, metaphase, anaphase, and telophase. Figure taken from Karp (1999).  8  genome, and prepares to divide. During M phase the cell undergoes mitosis, itself consisting of 5 phases: prophase, prometaphase, metaphase, anaphase, and telophase. M phase results in the segregation of replicated chromosomes, followed by cytokinesis, the physical division of the cell. The main kinase responsible for entry into mitosis is thought to be the Cdc2/cyclin B complex (Cdc2 is also known as Cdkl) (Nigg 2001). It is generally accepted that this kinase triggers the hallmark events of mitosis such as the condensation of chromosomes and dissolution of the nuclear lamina (Nigg 2001). It may also play a role in the reorganization of the cytoskeleton (Nigg 2001). That said, it is possible to force cells to enter mitosis without Cdc2 phosphotransferase activity indicating more remains to be discovered (Gowdy et al., 1998). The microtubule network, as mentioned, plays a critical role in the cell cycle. During interphase, it is relatively stable with microtubules having a half-life of approximately 200 to 300 seconds (Walker et al., 1988; McNally 1996; Zhai et al., 1996). However, at the start of mitosis, the microtubule network breaks down with a dramatic loss of polymer mass (Zhai et al., 1996) and becomes much more dynamic as it forms the mitotic spindle (for review see (Kline-Smith et al., 2004)). This is reflected by a dramatic decrease in the half-life of microtubules to 15 to 70 seconds (Walker et al., 1988; McNally 1996; Zhai et al., 1996) (Figure 3). This increase in dynamics is thought to help the mitotic spindle find and attach to the chromosomes in a search and capture model. The mitotic spindle contains two types of microtubules (Figure 4) (Kline-Smith et al., 2004). The first are the kinetochore microtubules which bind to the kinetochore of the chromosome. Approximately 25 microtubules will bind to one mammalian kinetochore and associate to form K-fibres (McEwan et al., 1997) that separate the chromosomes  9  Gi  I  S  I  G  2  I  M  Cell Cycle Phase  Figure 3 - Microtubule Dynamics and Polymer Mass During the Cell Cycle During interphase, microtubules are very stable. However, at the start of M phase the microtubules become much more dynamic. Similarly, at the start of M phase the amount of microtubule polymer rapidly decreases before returning to the interphase level.  10  K-fttwr Astrai .'MT  • I ^ B i p o l a r MT  r«n*msnrm  Chmmnsnms  Figure 4 - The Mitotic Spindle The mitotic spindle is composed of kinetochore microtubules, approximately 25 of which associate to form a K-Fiber, and interpolar or overlap microtubules. Kinetochore microtubules are responsible for the separation of the chromosomes during anaphase. Interpolar microtubules separate the centrosome. Astral microtubules localize the centrosome. Figure from (Kline-Smith et al., 2004).  11  during anaphase (McEwan et al., 1997; Kline-Smith et al., 2004). The second are overlap microtubules that are critical for the formation of a bipolar spindle and separation of the centrosomes during mitosis. Overlap microtubules form when the plus end of microtubules that originate from each of the two centrosomes overlap at the midbody of the cell. These overlapped microtubules are cross-linked by a plus-end directed motor that forces the centrosomes apart. Finally, in the mitotic cell there are astral microtubules that orient the location of the spindle pole (Kline-Smith et al., 2004). The dynamics of the microtubules are critical for the proper progression of mitosis (Kline-Smith et a l , 2004). If these dynamics are perturbed by small molecule drugs the cells will typically block at the metaphase stage of M-phase with high Cdc2/cyclin B activity. Many chemotypes have been found that block cells in metaphase (Hamel 1996; Jordan et al., 1998). The small molecules that perturb microtubule dynamics and lead to mitotic arrest are termed microtubule poisons or antimitotic agents. While many are known in the laboratory, two families have been brought to the clinic as a very successful chemotherapeutic agents for cancer treatment, the vinca alkaloids and the taxanes (paclitaxel and taxotere) (Rowinsky et al., 2001).  1.1.4- Microtubule Poisons As discussed previously, the correct dynamics of the microtubule network is critical to the proper progression of the cell cycle (Kline-Smith et al., 2004). In fact, if a small molecule perturbs these dynamics, the cell will block in mitosis. This property has been exploited to produce two families of successful anti-cancer drugs: the vinca  12  alkaloids and the taxanes. To understand how these work it is necessary to first examine the effects of microtubule poisons on the microtubule network. There are two large classes of microtubule poisons (Hamel 1996; Jordan et al., 1998). The first, with over 100 members, comprises those that inhibit the polymerization of microtubules and/or produce a breakdown of the microtubule network. The vinca alkaloids are an example of such agents that are used in the clinic for treatment of testicular cancer, Wilms tumour, Acute Lymphocytic Leukemia, lymphoma, rhabdomyosarcoma, soft tissue sarcoma, neuroblastoma, and Non-Small-Cell Lung Cancer (Rowinsky et al., 2001). Nocodazole and colchicine are two examples that are commonly used in the laboratory. The second much smaller group, with approximately 10 members, comprises agents that promote the polymerization of tubulin into microtubules and stabilize existing microtubules. Here, the most well known example are the taxanes (paclitaxel and docetaxel) which are commonly used in the clinic for treatment of Ovarian, Breast, and Non-Small-Cell Lung Cancer as well as Kaposi's sarcoma (Rowinsky et al., 2001), as well as in the laboratory. There exist at least five drug-binding sites on tubulin that can disrupt the microtubule network (Hamel 1996; Altmann 2001). The first is the colchicine-binding site, the second is the vinca domain, the third is the rhizoxin/maytansine, the fourth is the taxoid domain, and the fifth is the sulfhydryl group of assembly critical cysteine residues. The majority of examined microtubule poisons bind at one of these sites. Those binding at the colchicine, vinca, rhizoxin/maytansine or cysteine domains typically inhibit polymerization of tubulin into microtubules, while those binding at the taxoid domain promote the polymerization of microtubules. There is also a peptide binding site on (3  13  tubulin that overlaps with the vinca domain (Mitra et al., 2004). Naturally occurring peptides such as hemiasterlin and doastatin 10 bind in the peptide binding site (Mitra et al., 2004). Other binding sites probably exist as evidenced by laulimalide and peloruside that promote microtubule polymerization but do not compete with paclitaxel for binding to the microtubule (Pryor et al., 2002; Gaitanos et al., 2004). The taxoid site is of particular interest. While still debated, it is thought that the taxoid binding site is only present once the microtubule has formed (Parness et al., 1981; Takoudju et al, 1988). The solution of the microtubule and tubulin structure has allowed for the determination of the location of the taxoid binding site on the interior of the microtubule (Nogales et al., 1998; Li et al., 2002). While diffusion from the ends is possible, this likely would occur slowly and may be blocked by MAPs associated with microtubule ends. Holes have been identified in the wall of the microtubule which might allow rapid diffusion of the taxanes into the interior of the microtubule (Li et al., 2002). While it was initially thought that all microtubule-disrupting agents worked by changing the overall amount of tubulin that was polymerized, a revolution in thinking has begun to occur. It is now thought that these agents work at doses lower than those that affect the mass of polymerized tubulin by decreasing the dynamics, particularly the rescue and catastrophe frequency (Jordan et al., 2004). This decrease in dynamics prevents the formation of the mitotic spindle leading to a mitotic arrest.  1.1.5 - Microtubules and MAPs Microtubules do not exist alone. They have many proteins associated with them that are, in general called MAPs. Some of these proteins, such as members of the kinesin  14  and dynein family, are motors that move along the microtubule and carry cargo with them (Hirokawa et al., 1998; Endow 2003; Vallee et al., 2003). Others, such as XMAP215 (Kinoshita et al., 2002), XMAP230 (Andersen 2000), Tau, MAP4 (Cassimeris 1999; Andersen 2000), MAP2, Opl8/Stathmin (Cassimeris 2002), X K C M 1 (Andersen 2000), and katanin (McNally et al., 1993) affect the stability of the microtubule. If microtubules are highways, then kinesin and dynein are the cargo carriers. These motors typically use ATP hydrolysis to power their movement and will move either toward the plus end, as is the case for most kinesin motors, or toward the minus end, in the case of dynein and at least one kinesin (Hirokawa et al., 1998). They carry vesicles and also play a role in the separation of the centrosomes during mitosis (Hirokawa et al., 1998). Motors attach to the overlap microtubules, and each is plus-end directed and will force the centrosomes, where the minus end is attached, to separate. A large number of MAPs, including XMAP215, XMAP230, Tau, MAP4, XKCM1 and katanin, regulate the dynamics of microtubules either in vitro or in vivo. On the one hand, XMAP215, XMAP230, Tau, and MAP4 all increase the length of the microtubules, although by different mechanisms. XMAP215 increases the rate of growth but does not change the rate of catastrophe or rescue (Andersen 2000). XMAP230, Tau and MAP4 stabilize the microtubule and reduce the frequency of catastrophe rather than increase the rate of growth (Andersen 2000). On the other hand, Opl8/stathmin and XKCM1 destabilize the microtubules. Op 18 has been shown to disrupt microtubules in two distinct manners. First, it is possible for Op 18 to sequester tubulin dimers which has the effect of lowering the concentration of tubulin below the critical concentration. It is  15  also possible for Op 18 to actively induce catastrophe (Cassimeris 2002). X K C M 1 has only been shown to induce catastrophe (Andersen 2000). Unlike the other MAPs that work at the microtubule ends, katanin activity occurs in the middle of the microtubule where it cleaves microtubules in an ATP-dependent manner (McNally et al., 1993). Katanin has been shown to be active during M phase in Xenopus extracts and is likely involved in the rapid breakdown of the microtubule network and the onset of mitosis (McNally et al., 1998). The MAPs are crucial for the correct dynamics of the microtubule network (Cassimeris 1999). In fact, the in vitro dynamics of purified tubulin are not similar to those observed in vivo. However, correct in vivo dynamics can be reconstituted if XMAP215 and X K C M 1 are added at the correct concentration to tubulin in vitro (Kinoshita et al., 2001). MAPs can be controlled via signaling pathways within the cell. They can be phosphorylated which in turn changes their effect on microtubule dynamics (Cassimeris 1999).  1 . 2 - M A P K Signaling 1.2.1 - Mitogen Activated Protein Kinase (MAPK) Protein phosphorylation plays a vital role in the regulation and signaling of the cell. Phosphorylation changes occur in response to a diverse range of stimuli from growth factors to stress. The outcomes of this phosphorylation can be just as varied ranging from proliferation to differentiation. One of the major family of kinases responsible for this signaling is the M A P K family (Roux et al., 2004). This family of kinases has at least five members: Erkl/2, Jnk/SAPK, p38 M A P K , Erk3/4 and Erk5 (Roux et al., 2004). There are  16  examples of MAPK-like families conserved in organisms from yeast to humans. These kinases are grouped as a family because they are activated following a kinase cascade that results in phosphorylations at two critical threonine and tyrosine residues in the consensus sequence T X Y on the M A P K (Roux et al., 2004). This consensus sequence occurs in the activation or T-loop of the MAPK. M A P K Kinase ( M A P K K / M K K ) , also known as a M A P K Kinase (MEK), is responsible for the addition of the activating phosphorylations on M A P K (Roux et al., 2004). M A P K K are dual specificity kinases that add phosphate to both the threonine and tyrosine residue of the activation loop. Similar to the M A P K , the M A P K K (MEK) must be phosphorylated to become activated. The kinase responsible for this phosphorylation is a M A P K Kinase Kinase (MAPKKK). M A P K K K s are typically activated at the plasma membrane by receptor tyrosine kinases (RTKs) or Gproteins (Roux et al., 2004), though there appears to be alternative methods of activation such as microtubule disruption (Stone et al., 2000; Zang et al., 2001). A schematic of M A P K activation is shown in Figure 5. Erkl/2 were the first M A P K to be discovered in mammals, and in fact were originally called MAP2K since it phosphorylated microtubule-associated protein 2 (Ray et al., 1987). Erkl/2 are kinases of approximately 44 and 42 kDa, respectively. The M A P K K for Erkl/2 is MEK1/2, while the M A P K K K for Erkl/2 is the Raf family of kinases (A-Raf, B-Raf, 1-Raf) (Roux et al., 2004). The Raf kinases are activated in response to phorbol esters or growth factors. The exact mechanism of activation is unknown but it does include a series of phosphorylations, the binding of Ras, and recruitment to the plasma membrane (Chong et al., 2003). Erkl/2 has many substrates including membrane proteins, nuclear proteins, cytoskeletal proteins and M A P K -  17  MAPKKK v.  )  1 MAPKK  •  •  MKP  Figure 5 - M A P Kinase Activation and Deactivation Pathways Phosphorylation of M A P K K by M A P K K K leads to the activation of M A P K K . In turn, M A P K K phosphorylates M A P K on a threonine and tyrosine residue in the consensus sequence of T X Y . This is the activation loop of the M A P K . In turn, the dual specificity MKP removes these phosphorylations resulting in an inactivation of M A P K .  18  Activated Protein Kinases (MKs) (Roux et al., 2004). These substrates often change transcription either directly or indirectly. Erkl/2 plays a role in activating the ribosomal S6 kinases (RSKs), the M A P K and stress activated protein kinases (MSKs), and the M A P K interacting kinases (MNKs). These activated kinases in turn affect many cellular processes including transcription (Roux et al., 2004). p38 M A P K has four isoforms: a, (3, y and 5. These isoforms are activated in response to stress rather than growth factors and phorbol esters as is the case with Erkl/2. The M A P K K for the p38 family is MKK3/6 while the number of M A P K K K for the p38 family is high and includes M E K K s 1 to 4, M L K 2 , M L K 3 , D L K , ASK1, Tpl2, and Tak2. Similar to Erkl/2, p38 M A P K has many substrates including phospholipase A2, Tau, ATF1, ATF2, M E F 2 A Sap-1, Elk-1, NF-KB, Ets-1 and p53. The MKs MSKs and MNKs are also activated. However, unlike Erkl/2, p38 M A P K do not phosphorylate RSKs (Roux et al., 2004). JNK/SAPK family contains ten isoforms that are activated in response to stress, similar to p38 M A P K (Davis 2000). The M A P K K s for JNK/SAPK include MEK4 and MEK7 (Davis 2000). M E K 4 is interesting, as it is known to activate p38 M A P K in vitro (Roux et al., 2004). This provides a possible point of cross talk and a possible explanation for how stress may lead to the activation of both p38 M A P K and JNK/SAPK. A n additional potential location for cross-talk is at the M A P K K K level. The known M A P K K K for JNK/SAPK include MEKK1-4, M L K 2 , M L K 3 , Tpl-2, D L K , Taol, Tao2, TAK1, ASK1, and ASK2 (Roux et al., 2004). Many of the known M A P K K K s for JNK/SAPK have also been shown to be M A P K K K s for the p38 M A P K pathway. There is one large difference between JNK/SAPK, and p38 M A P K and Erkl/2. There is no  19  known M K that JNK/SAPK activates. However, it does phosphorlyate and activate a number of transcription factors including the most well known, c-Jun (Roux et a l , 2004). MAPKs obviously play a large role in signaling. The length of the signal is crucial. It has been shown in the same cell line that the duration of activation can mean the difference between a signal to proliferate and a signal to differentiate (Marshall 1995).  1.2.2 - M A P K and the Cytoskeleton Erkl/2 in mammals was originally identified as a kinase capable of phosphorylating MAP2 (Ray et al., 1987). Despite early investigations, no association with microtubule was observed. However, a paper published in 1995 used multiple Erkl/2 antibodies to observe that upwards of 50% of Erkl/2 in a mammalian cell was associated with the microtubule network (Reszka et al., 1995). In fact, even after stimulation, nearly 50% of the active Erkl/2 remained associated with the microtubule (Reszka et a l , 1995). There is a body of evidence that hints that Erkl/2 signaling may be involved in microtubule stability (Reszka et al., 1995).Work has demonstrated a role for M A P K pathways in cell migration (Huang et al., 2004) which requires a rearrangement of the cytoskeleton. Recent work has hinted that it might be necessary to think of microtubules in a different manner. It has been shown that signaling can result in microtubule rearrangement. However, it has also been repeatedly shown that disruption of the dynamics of the microtubule can lead to the activation of the Erkl/2 and other M A P K pathways (Stone et al., 2000; Zang et al., 2001; Boldt et al., 2002). This activation  20  appears to be cell type specific, and the reason is not known. Nevertheless, it may be necessary to reconsider the microtubules not only as a target of signals but also as a 'receptor' capable of initiating a signaling response. In addition to being regulated by M A P K signaling, there is evidence that the microtubule network of the cell is controlled during the cell cycle. The evidence suggests that this control is via phosphorylation of regulatory proteins (Cassimeris 1999). Initially, it was found that active Cdc2/cyclin B could alter the lengths of microtubules in Xenopus egg extracts. Further, MAP4 phosphorylation deficient mutants block entry into mitosis and there are at least four required phosphorylations on Op 18 for mitosis to occur (Cassimeris 1999).  1.2.3 - M A P Kinase Phosphatase (MKP) As discussed, M A P Kinases must be phosphorylated at their T-loop in order to be activated (Pearson et al., 2001). In particular, there must be both a threonine and a tyrosine phosphorylated at the T X Y consensus sequence. To be deactivated, either the threonine or the tyrosine, or both residues must be dephosphorylated. As the length of the activation can have profound consequences it is probable that dephosphorylation is critical. In vitro it has been shown that a protein-serine/threonine (Alessi et al., 1995) or tyrosine (Pettiford et al., 2000) phosphatase can remove a phosphate from these residues in M A P K leading to its inactivation. However, it has been discovered that M A P K dephosphorylation is typically performed by dual specificity phosphatases (DSP) that dephosphorylate both threonine and tyrosine (Theodosiou et al., 2002; Farooq et al., 2004). These DSPs have come to be known as M A P Kinase Phosphatases (MKPs). There are now 10 known MKPs which have differing M A P K specificity (Theodosiou et al.,  21  2002). The majority of the activity assays have been conducted in vitro. More work will be required to determine which substrates are physiologically relevant to which MKPs. However, some interesting results have been obtained. It has been shown that the binding of Erkl/2 results in the activation of MKP-3 (Camps et al., 1998). Substrate binding at a location distinct from the catalytic residue leading to activation may be critical for the regulation of M A P K signaling. There has also been work on the catalytic residue of MKPs. It has been found that the sulfhydryl of a cysteine residue is the catalytic residue (Denu et al., 1995).  1.3 - Goals The goal of this thesis has been the characterization of the biology of three new microtubule poisons. The first two, ceratamine A and B, are structurally very related. They were discovered in an antimitotic screen and their effect on microtubules in vivo, microtubules in vitro, and the cell cycle has been examined. The third molecule is dimethyl varacin (DMV). D M V was also discovered in an antimitotic screen. Work with D M V has concentrated on its effect on microtubules, both in vivo and in vitro as well as its effect on the cell cycle, M A P K signaling, and cellular phosphorylations.  22  Chapter II: Materials and Methods  2.1 - Materials: Unless otherwise noted, all materials were purchased from SigmaAldrich Chemical Company, St. Louis, Missouri.  2.2 - Antimitotic Assay - The phenotypic antimitotic ELICA was performed as described previously (Roberge et al., 2000).  2.3 - Antiproliferation  Assay - The antiproliferation assay was performed as described  previously (Anderson et al., 1997).  2.4 - In Vitro Tubulin Polymerization  Assays  2.4.1 - Promotion of Tubulin Polymerization:  On ice, 35 uL of buffer A (80mM PIPES  pH 6.8, 1 mM M g C l , 1 mM EGTA) were added to each well of a half-area 96-well plate, 2  followed by 5 uL of 10 mg/mL bovine brain tubulin (TL238, Cytoskeleton Inc. Denver, Colorado) in buffer A containing 10% glycerol, 5 uL of a lOx solution of ceratamine A or B in buffer A, and 5 uL of 10 mM GTP in buffer A. The optical density at 340 nm was followed at 32°C over time.  2.4.2 - Inhibition of Tubulin Polymerization:  This assay was performed in a similar  manner to the promotion of polymerization assay but with the following differences. On ice, 25 uL of buffer A plus 10% glycerol were added to each well of a half-area 96-well plate, followed by 15 uL of 10 mg/mL tubulin in buffer A containing 10% glycerol, 5 uL of lOx ceratamine A or B solution in buffer A containing 10% glycerol and 5 uL of 10  23  mM GTP in buffer A plus 10% glycerol. The optical density at 340 nm was followed at 37°C over time.  2.4.3 - Negative Staining Electron Microscopy: The promotion of tubulin polymerization assay was performed as described above. Ten uL of the reaction (10 pg of protein) were removed and placed on a Formar coated copper grid and left for 1 minute at room temperature. The fluid was wicked off by Whatman filter paper and 10 uL of 2% uranyl acetate solution were added and immediately removed by Whatman filter paper wicking. The sample was analyzed by electron microscopy using a Hitachi H7600 Transmission Electron Microscope. (University of British Columbia, Bioimaging Facility).  2.4.4 - H-Paclitaxel 3  Competition Binding Assay: The paclitaxel binding assay was  conducted similar to that described previously (Pryor et al., 2002). Briefly, solutions of 10 u M H-paclitaxel (NCI) with a specific activity of 1600 dpm/pmol, various concentrations of trial compound [20 uM eletherobin, 100 u M or 250 u M ceratamine A], DMSO (20%o), and monosodium glutamate (0.75M) were prepared. A solution of microtubules was prepared by incubating tubulin (2.5 uM; 0.25 mg/mL), 2',3'dideoxyguanosine 5'-triphosphate (ddGTP; 25 uM) and monosodium glutamate (0.75 M). These solutions were incubated for 30 minutes at 37°C. Twenty five pL of drug solution were added to 100 uL of microtubule mixture for a final volume of 125 uL, and the mixture was incubated for 30 minutes at 37°C. Following incubation, 50 uL of reaction mixture were removed, centrifuged for 20 minutes at 13,000 rpm and the supernatant was counted in a liquid scintillation counter to determine the amount of unbound, or free, H-paclitaxel. Also, 50 uL of uncentrifuged reaction mixture were  24  counted to determine the total amount of H-paclitaxel. The statistical significance of the results was determined by a Student's t test.  2.5 - Flow Cytometry Analaysis of the Cell Cycle: Analaysis of mitosis-specific markers was conducted as described (Rundle et al., 2001) using a F A C S Calibur and GF-7 primary antibody, with the following minor modification: RNase was from Qiagen and used at a final concentration of 100 ug/mL. GF-7 primary antibody was raised using paired helical filaments from Alzheimer brains. While several bands are detected by Western blot (data not shown), the epitopes are typically only observed during mitosis. Data were analyzed using MDI 2.8.  2.6 - Immunofluorescence  2.6.1 - f-Tuhulin  Microscopy  Immunofluorescence  Microscopy of DMV Treated Cells: 1 mL of a  1/10 dilution of 80% confluent Swiss-3T3 cells was seeded onto glass coverslips in 12well microplates and incubated 48 hours to allow the cells to attach. The cells were treated with 10 uM D M V or 0.1% D M S O for either 30 or 120 minutes. The microtubules were visualized using the E7 monoclonal P-tubulin antibody (Hybridoma Development Bank, University of Iowa) as described in Anderson et al (Anderson et al., 1997). D N A was not stained.  2.6.2 - f-Tubulin  Confocal Immunofluorescence Microscopy  of Ceratamine Treated  Cells: MCF-7 cells cultured on glass coverslips were treated without or with ceratamine A and microtutubles were immunostained using the E7 monoclonal p-tubulin antibody. A  25  stack of 40 optical sections each 0.35 pm was collected using a Bio-Rad Radiance Plus confocal microscope and analysed using ImageJ software (NIH). 2.6.3 - P-Tubulin Immunofluorescence Microscopy of Ceratamine Treated Cells:  MCF-7  cells cultured on glass coverslips were treated without or with ceratamine and microtutubles were immunostained using the E7 monoclonal P-tubulin antibody (Anderson et al., 1997). D N A was stained using Hoechst 33248.  2.7 - Kinase Assays  2.7.1 - Cdc2 Kinase Assay: MCF-7 mp53 cells were plated in 100 mm dishes and grown to approximately 80% confluence. The cells were treated with 300 nM nocodazole, 0.1 % DMSO, and 10 uM D M V for 5 hours before harvesting by trypsinization. The assay was then carried out as described previously (Gowdy et a l , 1998).  2.7.2 - In Gel Kinase Assay: Following treatment by either D M V at 1, 5, or 10 uM; 300 nM nocodazole; or 0.1 % D M S O for 2 hours, an in gel kinase assay was carried out as described elsewhere (Gowdy et a l , 1998). The following changes were made. The lysis buffer contained neither DNase I nor RNase A. Further, the gel contained either no polymerised substrate to detect possible autophosphorylation, or 0.2 mg/mL myelin basic protein (MBP) from Invitrogen. Finally, the gel was visualized by autoradiography.  2.8 - Western Blots: Ten ug of protein from a whole cell lysate were separated on a 10% SDS-PAGE. A semi-dry transfer (Biorad; Transblot SD) was used to transfer to a PVDF membrane in Towbin buffer. The membrane was washed in dt^O prior to being blocked with a solution of 5% non-fat milk in TBS containing 0.1% Tween-20 (5%-TBS-T). The  26  membrane was incubated with primary antibody [Phospho-p38 M A P K T180/Y182; Cell Signaling Technology; Phospho-p44/42 M A P Kinase (T202/Y204), New England Bioloabs; p44/42 M A P K , Stressgen; Phospho-JNK/SAPK, Cell Signalling Technology] at the manufacturers' recommended dilutions in 5%-TBS-T for Phospho-p44/42 M A P K , Total Erkl/2 (p44/42 M A P K antibody), and phospho-JNK/SAPK or TBS containing 5% bovine serum albumin (BSA) and 0.1% Tween-20 for phospho-p38 M A P K . The blots were incubated overnight at 4°C. The membranes were washed in TBS-T prior to being incubated in the appropriate secondary antibody at the manufacturers' recommended dilution in 5%-TBS-T. The membrane was washed extensively before being developed using Enhanced Chemiluminesence (Pierce, SuperSignal West Pico).  2.9 - Kinexus Kinetworks™  Screens: Cells at approximately 80% confluency in three 100  mm tissue culture plates were treated with stimuli (0.05% D M S O or 5 uM D M V ) for 2 hours. Cells were collected, pooled and lysed as described in the Kinexus Bioinformatics Corp. (Vancouver, Canada) sample submission form. For each Kinetworks™ phosphosite screen (KPSS 1.3, KPSS 2.0, KPSS 3.0, KPSS 4.0), 500 uL of lysate at 1.0 mg/mL in loading buffer were submitted to Kinexus Co.  2.10 - Protein Phosphatase 2A Assay - Approximately 0.125 U (1 unit releases 1 nmol of phosphate per minute from 15 uM phosphorylase A at 30°C) of PP2A from Upstate Biotechnology were added to 10 uL of enzyme dilution buffer (20 m M MOPS, pH 7.5, 0.15 M NaCl, 60 m M p-mercaptoethanol, 1 mM M g C l , 1 mM DTT, 10% glycerol, 0.1 2  mg/mL BSA). To this solution, 10 uL of assay buffer (10% w/v glycerol, 0.1 mg/mL BSA) containing 25 uM or 2.5 uM D M V were added for a final volume of 20 uL. This  27  was left on ice for 5 minutes. Previously, 5 uL of peptide solution (0.75 mg/mL RRATPhosphate-VA-amine in phosphate free water) were added to 25 uL of assay buffer in a half-area 96 well plate and incubated at 37°C for 30 minutes. The enzyme solution containing the inhibitors was then added to the peptide solution for a final volume of 50 uL. This was incubated for 45 minutes at 37°C. Fifty uL of Malachite Green (1.05% w/v ammonium molybdate, 0.034% malachite green, 1 M HCI) was added and the absorbance measured at 595 nm. Controls were also performed with no enzyme present or no substrate present.  28  Chapter III: Identification of Ceratamines A and B as Microtubule-Stabilizing Agents With Unusual Biological Properties  3.1 - Specific Introduction: The number of microtubule-stabilizing chemotypes is on the order of 10 with the most well known example being the taxanes, including paclitaxel (Figure 6A) and its derivative taxotere (He et al., 2001). Other examples include the eleuthesides (Long et al., 1998) , laulimalides (Mooberry et al., 1999), GS-164 (Shintani et al., 1997), epothilones (Bollag et al., 1995), discodermolides (ter Haar et al., 1996), sarcodictyins (Hamel et al., 1999) , peloruside (Hood et al., 2002) and dictyostatin (Isbrucker et a l , 2003). Discodermolide, eleutherosides, sarcodictyins and epothilones compete with paclitaxel for binding at the taxoid binding site (Bollag et al., 1995; Kowalski et al., 1997; Hamel et al., 1999), but laulimalide (Figure 6B) and peloruside A (Figure 6C) have been unequivocally shown to bind at a site distinct from paclitaxel on the microtubule (Pryor et al., 2002; Gaitanos et al., 2004). Significantly, all of these microtubule-stabilizing agents except GS-164 have come from natural products and are very structurally complex (Figure 6). This is exemplified by paclitaxel which was isolated from the Pacific Yew tree (Taxus breviofolia) and has 11 chiral centers. This structural complexity led to large obstacles in the development of paclitaxel as a chemotherapeutic agent (Cragg et al., 1993). It is still made semi-synthetically though a total synthesis has been reported (Kingston 2000). As such, structurally simple antimitotic agents would be attractive lead  29  A  C  Figure 6 - The Chemical Structures of Paclitaxel (A), Laulimalide (B), and Peloruside A (C) Paclitaxel, laulimalide and peloruside A promote the polymerization of microtubules and microtubule bundling, but laulimalide and peloruside A cannot compete with paclitaxel for binding to microtubules.  30  compounds as they should be relatively simple to supply. There is also a pressing need to develop new antimitotic agents as cells can develop resistance to paclitaxel. Resistance may take the form of different tubulin isotypes (Verdier-Pinard et al., 2003), point mutations in tubulin (Verdier-Pinard et al., 2003), and expression of drug efflux pumps (Yusufet al., 2003). A cell-based phenotypic assay for antimitotic agents was used to screen extracts from natural sources. A n extract from the marine sponge Pseudoceratina  sp. collected in  Papua New Guinea showed activity in the assay. The active compounds were isolated by chromatographic steps using the assay to guide purification, and they were identified as ceratamines A and B (Figure 7) by analysis of their mass spectrometry and nuclear magnetic resonance data (Manzo et al., 2003).  3.2-RESULTS 3.2.1 - Effect of Ceratamines A and B on Cell Cycle  Progression:  The effect of ceratamines A and B on cell cycle progression was investigated using a dual labeling flow cytometry procedure that can identify the Gi, S, G and M cell 2  populations. Untreated cells showed a cell cycle profile typical of asynchronously proliferating cells, with large proportions of Gi, S and G2/M (Figure 8A). Exposure of cells to 21 u M ceratamine A or 22 u M ceratamine B for 16 h caused an almost complete disappearance of the Gi and S peaks with a corresponding large increase in the G2/M peak (Figure 8A). Staining with the GF-7 antibody, a phospho-specific antibody that is typically recognized only at mitosis, to distinguish between G and M showed the arrest 2  to be at M and not G : ceratamine A treatment caused 74% of the cells to arrest at M and 2  31  Figure 7 - The Chemical Structure of Ceratamine A and B  32  B  G1  DMSO  M  §  ^^2/M  Ul  G2 UO  GI  DMA (PI)  D N A (PI)  5 1 <  J8  Nocodazole  S  § m  LL  o DNA (PI)  Ceratamine A  J  I  Ul :  Ceratamine B  DMA (PI)  I  5  i 1  LL. *  i  o  DNA (PI)  D N A (PI)  Li  DNA (PI)  Figure 8 - Flow Cytometry of MCF-7 Ceratamine A or Ceratamine B  DNA (PI)  mp53 Cells Treated with  D M S O , Nocodazole,  (A) DNA profile of cells stained with propidium iodide. (B) GF-7 versus D N A dual labeling experiment. This permits the differentiation of G2 and M phase cells. The colours represent the number of events. As one moves through the spectrum from blue to red, the number of events increases.  33  ceratamine B caused 81% M arrest, (Figure 8B). The extent of M phase accumulation was very similar to that observed with the microtubule-targeting agent nocodazole (Figure 8B).  3.2.2 - Antimitotic and antiproliferative  activity of ceratamines.  To examine the concentration dependence of M arrest by ceratamines, cells were treated with different concentrations of compound and the proportion of cells in M was determined by staining the D N A and counting the number of cells with condensed mitotic chromosomes in the microscope. As shown in Figure 9A, M arrest was concentration-dependent, with half-maximal arrest at 12 u M for ceratamine A and 21 u M for ceratamine B. The IC50 results do not completely agree with those obtained with flow cytometry. The IC50 for ceratamine B is very similar to the concentration used in the flow cytometry experiment, where >80% of cells were blocked in mitosis. The reason for this discrepancy is unclear, but possible explanations may be differing batches. Also, IC50 values are approximations from the curve in Figure 9A. As expected, the compounds also inhibited cell proliferation, with IC50 of 2 u M for ceratamine A and 3 u M for ceratamine B (Figure 9B).  3.2.3 - Effect of ceratamines on microtubule organization in vivo. We next examined the effect of ceratamines on the morphology of interphase and mitotic microtubules using immunofluorescence microscopy with a P-tubulin antibody and counterstaining of DNA.  In ceratamine-treated cells arrested in M , the chromosomes  were condensed but failed to align at the cell equator (Figure 10B). Examination of microtubules showed that instead of containing a bipolar mitotic spindle as in control  34  Ceratamine (uM)  Ceratamine (uM)  Figure 9 - Antimitotic and C e l l Proliferation Assays with Ceratamine A and B  Antimitotic (A) and Cell Proliferation Assays (B) with various concentrations of either ceratamine A (open circles) or ceratamine B (closed circles). The percent of mitotic cells was determined by mitotic spreads after 16 hours with various concentrations of ceratamine. Cell proliferation was determined by M T T assays. Both assays used MCF-7 mp53 cells. Antimitotic assay was conducted once though the results are supported by flow cytometry. Cell Proliferation Assays were conducted 3 times; error bars represent the standard deveiation.  35  mitotic cells (Figure 10A), cells arrested in M by ceratamine treatment had no microtubules. Instead, multiple foci of P-tubulin staining were observed (Figure 10B and Figure 11). The number of foci varied from cell to cell from three to as high as about thirty. Not all foci had tubulin staining of equal intensity nearly all cells containing a few that were clearly brighter than the rest. three-dimensional  confocal  Examination of the structure of these foci by  microscopy showed  that they consisted  of pillar-like  structures that extend vertically from the glass surface upon which the cells were attached (Figure 1 IB and Figure 1 IC). The intensely staining foci spanned the entire thickness of the cell whereas less intensely staining foci had a similar structure but did not span the entire thickness of the cell. Focus formation occurred rapidly. Incubation with 21 uM ceratamine A for 30 min was sufficient to induce multiple centers of intense microtubule staining. Interestingly, some cells with apparently normal microtubules were observed but they were all at stages later than metaphase, indicating that ceratamines might only affect microtubules at early stages of mitosis. The focal tubulin staining persisted for more than ten hours in the presence of ceratamines. Longer incubation times caused a decrease in the number of tubulin foci, such that cells often had either two or four regions of heavy tubulin staining. The effect of ceratamines on mitotic microtubules was markedly different from that of the microtubule polymerizing agent paclitaxel, which caused the appearance of dense, "bushy" mitotic spindles (Figure 10C). Ceratamine A treatment also changed the organization of microtubules in some interphase cells.  In these cells, ceratamine A caused the appearance of a pronounced  perinuclear microtubule network with a distinct depletion of peripheral microtubules  36  Tubulin  DNA  No drug  Ceratamine A  Paclitaxel  Figure 10 - Confocal P-Tubulin Immunofluorescence Images of Ceratamine or Paclitaxel Treated Cells Images of P-tubulin staining and D N A (Hoescht 33248 staining) were collected of mitotic cells treated with (A) DMSO, (B) ceratamine A, or (C) paclitaxel. Images were collected of P-tubulin staining in interphase cells treated with (D) DMSO or (E) ceratamine A .  37  Figure 11 - Cross Section p-Tubulin Staining in Mitotic Cells P-Tubulin staining in mitotic cells treated with (A) DMSO or (B,C) ceratamine A were collected by confocal microscopy. Image J software was used to reconstitute crosssectional images.  38  (Figure 10E). In addition, the microtubule network of ceratamine-treated cells appears disorganized compared to that of control cells (Figure 10D).  3.2.4 - Effect of ceratamines on microtubule formation in vitro  Most antimitotic agents interfere with microtubule function by interacting directly with them and causing their depolymerization or their overpolymerization. To examine the mechanism of action of ceratamines, we examined the effect of these compounds on the polymerization of microtubules from pure tubulin in vitro. We first determined whether ceratamines inhibit microtubule polymerization. As shown in Figure 12A, 10 uM nocodazole prevented the assembly of microtubules but ceratamines showed no significant inhibition of tubulin polymerization at any concentration tested (up 110 uM). We next examined whether ceratamines can promote microtubule polymerization. As shown in Figure 12B, 10 u M paclitaxel stimulated microtubule polymerization, as expected. Ceratamine A (100 uM) also promoted microtubule polymerization (Figure 12B) but to a lesser extent than paclitaxel.  To determine whether the increase in  absorbance measured in this assay was indeed due to the formation of microtubules, a sample  was  removed  and analyzed by negative staining  transmission  electron  microscopy. As shown in Figure 13A and Figure 13B, ceratamine A caused the formation of microtubules, similar to those observed for the paclitaxel treated sample (Figure 13C). No microtubules were observed in samples treated with DMSO, the drug solvent (not shown).  39  A 0.3  0.2  0.1  0.0 0  5 10 Time (min)  15  B  0.12 0.10 0.08 0.06 0.04 0.02 0.00 -0.02  J  §  5 10 Time (min)  15  Figure 12 - In vitro Tubulin Polymerization Assays with Ceratamine A and B (A) Inhibition of tubulin polymerization with DMSO (open squares), nocodazole (open triangles), ceratamine A (open circles) and ceratamine B (closed circles). The final tubulin concentration was 3 mg/mL; the positive control was nocodazole. (B) Promotion of tubulin polymerization assay with DMSO (open squares), paclitaxel (closed triangles), ceratamine A (open circles), and ceratamine B (closed circles). The final concentration of tubulin was 1 mg/mL; the positive control was paclitaxel. Similar results for both (A) and (B) were obtained in a second experiment.  40  Ceratamine A - 30,000x  Paclitaxel - 120,000x  Figure 13 - Electron Microscopy Images of Microtubules A portion of the products from the promotion of tubulin polymerization assay was examined by negative staining electron microscopy. Paclitaxel was a positive control with clear microtubules present. The bar represents 500 nm. 41  3.2.5 - Ceratamines do not compete with Paclitaxel for binding to Tubulin: To probe the molecular mechanism by which the ceratamines promote the polymerization of tubulin into microtubules, a competition assay with radiolabeled paclitaxel was conducted. This was done to determine if the ceratamines bind to the taxoid binding site of tubulin. In the assay, microtubules are initially formed by incubation with ddGTP and a high concentration of monosodium glutamate (MSG). These conditions result in the almost complete polymerization of available tubulin. A solution containing the" radiolabeled paclitaxel and the compound in question, in this case ceratamine A , is then added to the microtubule solution. After 30 minutes to come to equilibrium a portion of the total reaction is counted in a scintillation counter. A second portion is centrifuged. This causes the microtubules and the small molecules bound to them to pellet at the bottom of the tube. The supernatant, containing unbound small molecules, is counted in a liquid scintillation counter. With both counts, it is possible to obtain a ratio of free to total paclitaxel. If the compound in question competes with paclitaxel binding, the amount of free paclitaxel will increase in the presence of the compound. This will be reflected by an increase in the ratio of free to total paclitaxel. Using this assay, developed by Hamel (Kowalski et al., 1997), it was discovered that ceratamine A does not compete with paclitaxel for binding to microtubules at concentrations up to 100 uM (Figure 14). As can be seen in Figure 14 when no drug is present in the binding assay, the ratio of free to total paclitaxel is 0.22±0.04. The ratio with 4 u M eleutherobin, a known competitor of paclitaxel binding (Hamel et al., 1999), is 0.40±0.06. However, with 20 uM ceratamine A the ratio is 0.19±0.02 and is not significantly different from that found  42  in the absence of drug. Even when the amount of ceratamine A is increased to 50 uM, a 25-fold excess over the amount of tubulin and paclitaxel, the ratio is 0.26±0.03, again a non-significant difference. This is clear evidence that ceratamine A is unable to compete with paclitaxel for binding to the microtubules at these concentrations.  43  o LU  Figure 14 - H-Paclitaxel Binding to Microtubule Assay 3  Microtubules were preformed and then incubated with no drug, eleutherobin, or one of two concentrations of ceratamine A. Each reaction contained 2 uM H-paclitaxel with a specific activity of 1600 dpm/pmol. The ratio of free to total paclitaxel was then determined by scintillation counting over multiple experiments. The bar represents the mean of 3 independent experiments with the error bars representing standard deviation. Eleutherobin is a positive control. The change between eleutherobin and D M S O is statistically significant. However, the changes between D M S O and either ceratamine treatement are not statistically significant. 3  44  3.3 - Discussion This work describes the characterization of the cellular effects of two novel chemically simple antimitotic agents, ceratamine A and ceratamine B. These compounds block the cell cycle in M phase and inhibit cellular proliferation. Ceratamine A disrupts the microtubule network in vivo in a manner that, to our knowledge, has not been previously described. In vitro, ceratamine A and ceratamine B promote the polymerization of tubulin similar to paclitaxel, but ceratamine A does not compete with paclitaxel for binding to microtubules. Ceratamine A and ceratamine B were discovered in a phenotypic antimitotic ELICA screen of natural products (Manzo et al., 2003). Not surprisingly, mitotic spreads and flow cytometry confirmed that ceratamine A and ceratamine B blocked the cells in M phase. This is similar to what has been reported with paclitaxel (Roberge et al., 2000). Significantly, the ceratamines had IC50 values of 12 and 21 uM, respectively, for antimitotic properties. These values are three orders of magnitude higher than the 3 nM reported for paclitaxel (Roberge et al., 2000). Ceratamine A and ceratamine B were also found to inhibit cellular proliferation in a dose-dependent manner. Similar to the case with their antimitotic property, ceratamine A has a lower IC50 value than ceratamine B (2 uM versus 3 uM). However, both ceratamine A and B have IC50 values significantly higher than the 5 nM reported for paclitaxel (Anderson et al., 1997). It must be mentioned that there is a 5- to 7-fold increase in the IC50 values of antimitotic activity versus antiproliferative activity for the ceratamines. While this was not observed by Anderson et al. (1997) for paclitaxel it was observed for vincristine where the reported antimitotic IC50 was 15 nM and the reported antiproliferative IC50 was 2 nM. This is a 7.5-fold  45  increase. The M T T assay used for the antiproliferative assay measures a slow down in the cell cycle, while the antimitotic assay measures a complete halt. It may be that at lower concentrations of antimitotic agents, the cells are not becoming blocked in mitosis, but the cell cycle is slowed resulting in an inhibition of proliferation. Although they are not very potent, the ceratamines may be useful lead compounds for drug development. Combinatorial chemistry using the structure of ceratamines as a starting point may lead to the development of more potent analogues. From these studies, there is evidence that changing a hydrogen to a methyl group can decrease the IC50 values of both the antimitotic and antiproliferation assay. Whether the experiments are conducted on more potent analogues or the ceratamines, only animal studies can address their potential for clinical use. While it had been found that the ceratamines block cells in mitosis and inhibit cellular proliferation, their mechanism of action was unknown. Since many antimitotic agents disrupt the microtubule network of the cell, this was investigated. Confocal immunofluoresence microscopy revealed that ceratamine A treatment of cells resulted in a disruption of the microtubule network. Only ceratamine A was examined as it was the more potent of the two, and the mechanism of action for ceratamine A and B is almost certainly the same given their close structural similarity. In interphase, not all of the cells were affected by ceratamine A treatment. However, those that were affected had intense perinuclear tubulin staining. Similar perinuclear staining has been described in response to aphidicolin treatment for 72 hours (Tanaka et al., 1998). What is more striking is the absence of microtubules in mitotic cells. Instead, pillars of tubulin are present. These structures are in sharp contrast to the bushy microtubule bundles observed following  46  treatment with paclitaxel. This hints that the ceratamines affect microtubules in a different manner than paclitaxel. The pillars are of particular interest. While it is clear that these structures are composed of tubulin, it is not clear whether they are groupings of microtubules or aggregates of tubulin. Electron microscopy could provide an answer to this question. It may be possible to visualize a cross section of the pillars and identify microtubules versus protein aggregates. These pillars raise several interesting questions: is their formation nucleated or not? If there is a site of nucleation, is it a single protein such as membrane bound tubulin that has been reported (Luduena 1998), or a protein complex? Why does the number of foci decrease with time? Do the pillars aggregate or do they breakdown and then reform on pre-existing ones? Are proteins other than tubulin found in the pillars and if so what are the roles of these proteins? With clear in vivo data that microtubules are affected by ceratamine A treatment, work shifted to an in vitro assay in an attempt to characterize the nature of this effect. As the vast majority of chemotypes that affect microtubule dynamics inhibit their formation, this was initially investigated. The ceratamines were found to not inhibit the polymerization of tubulin to microtubules in vitro. This is in direct contrast to nocodazole, which was used as a control and which effectively inhibited microtubule formation. This result clearly demonstrated that ceratamine A and ceratamine B did not 1  function as antimitotic agents through inhibition of microtubule polymerization. Next, the ceratamines were investigated for their ability to promote the polymerization of microtubules. In the in vitro assay designed to examine the ability of a compound to stimulate the polymerization of microtubules, the ceratamines were found  47  to be positive. Their ability to promote the overpolymerization was similar to that observed with paclitaxel, but they were much less potent. This is in line with earlier antimitotic and antiproliferative results which demonstrated that paclitaxel was more potent than ceratamine A and ceratamine B. Also consistent with earlier results was the relative potency of ceratamine A and B. At the same concentration, ceratamine A induced more microtubule polymerization than ceratamine B. This provides a possible explanation for why ceratamine A is more potent in the in vivo assays than ceratamine B. Ceratamine A may have a more dramatic effect on the microtubule network of cells. The microtubule polymerization assay measures the increase in light scattering as microtubules form. However, the assay does not differentiate between light scattering due to microtubule formation, precipitation of the drug or formation of protein aggregates. To determine whether ceratamine A was indeed promoting the formation of microtubules, the reaction mixture was analyzed by electron microscopy to visualize the products. It was found that similar to paclitaxel, ceratamine A was indeed promoting the formation of microtubules. This provided clear evidence that ceratamine A could promote the polymerization of microtubules. At this point in the investigation there was some conflicting data. From the in vitro work it was clear that the ceratamines promoted the polymerization of microtubules.. At this level they are similar to paclitaxel despite being less potent. However, from in vivo immunofluorescence microscopy it was evident that ceratamine affected microtubules in a manner distinct from paclitaxel. Paclitaxel treatment results in the formation of microtubule bundles in interphase and "bushy" spindles at mitosis. This was never observed with ceratamine treatment. Instead, the dramatic pillars of tubulin staining  48  were observed in mitotic cells, and perinuclear networks were observed in some interphase cells. We thus hypothesized that while both ceratamine A and paclitaxel promote microtubule polymerization, they might bind at different sites in the microtubule. This was examined by a paclitaxel binding competition assay. The paclitaxel binding assay demonstrated that ceratamine A could not compete with paclitaxel for binding to the microtubule polymer. There were three possible explanations for this. First, the concentration of ceratamine A may be insufficient to bind to the microtubule. This is unlikely as the highest concentration tested, 50 uM, was sufficient to induce polymerization of microtubules in vitro (data not shown). The second explanation is that the binding sites overlap but the binding affinity of ceratamine A is too low to compete with paclitaxel. This is a possibility that cannot be discounted. There was a 25-fold excess of ceratamine A over paclitaxel and still no competition was observed, but it is true, from earlier experiments, that ceratamine A was much less potent than paclitaxel at inducing microtubule polymerization. The third explanation is that ceratamine A binds to a region of the microtubule outside the taxoid binding site and does not disrupt it. This seems the most likely because it explains the results of the paclitaxel binding assay and provides an explanation for the in vivo results. While more evidence is required, intuitively it follows that binding to a site other than the taxoid binding site might produce different microtubule morphologies. Work remains to determine which of the possible three explanations for the competition assay results is correct. The most direct experiment to determine where the ceratamines bind on microtubules is to solve the structure of tubulin/microtubules with the ceratamines bound. Other methods that are theoretically possible include isothermal  49  titration calorimetry and HPLC. Both of these methods depend on prohibitively large quantities of both protein and drug. Another possible mechanism to resolve this question involves using a radiolabeled ceratamine molecule. If paclitaxel can compete with labeled ceratamine, then this would be evidence of binding to the same site. While this experiment does not require prohibitive amounts of either ceratamine or protein, it obviously requires that the labeled ceratamine be synthesized. This may not prove to be trivial. If the ceratamines do bind at a region on the microtubule distinct from the taxoid binding site, it places them in a very small group. To date, there are only two other molecules known to promote the stabilization of microtubules and bind outside the taxoid binding site. One is laulimalide (Pryor et al., 2002) and the other is peloruside (Gaitanos et al., 2004). In contrast to the ceratamines, laulimalide treatment of cells produces the characteristic microtubule bundling typical of paclitaxel treatment (Mooberry et al., 1999). On this basis alone, it would appear that the ceratamines bind to a site unique not only from paclitaxel, but laulimalide as well. The work to characterize the biology of the ceratamines has demonstrated that they are small molecules with traditional antimitotic and antiproliferative properties but unique cellular effects. The ceratamines affect the microtubule network both in interphase and mitosis, producing very unique tubulin staining, and promote the polymerization of microtubules in vitro. A simple model for the ceratamines mechanism would be that the drug binds to either tubulin or microtubules and inhibits the correct dynamics of the mitotic spindle. This leads to cells blocked in mitosis that in turn inhibits their  50  proliferation. Further, this model would include the ceratamines binding to a portion of the microtubule separate from the taxoid binding site. The goal of our antimitotic screening program is to discover novel antimitotic agents with potential for the treatment of cancer. The ceratamines' potential to become clinically relevant is unclear at this point. They are significantly less potent than paclitaxel, and hypothesizing from their structure their water solubility may be limited. These two factors may combine to prevent appropriate activity within either animal models or humans. These questions will have to be answered in animal studies with the ceratamines. Despite questions about their solubility and potency, the ceratamines do appear to have one strong asset that may be very useful for further work. They are structurally very simple, especially when compared to the taxanes. For a quick measure, this structural simplicity is reflected in the number of chiral centers present in the molecules. The ceratamines do not have any chiral centers, whereas the paclitaxel has eleven. Each one of these chiral centers creates significant problems when attempting to synthesize the drug. It is hoped that the absence of these chiral centers would allow the ceratamines to have none of the supply problems that so plagued the development of paclitaxel (Cragg et al., 1993). Even if not developed further, the ceratamines represent what is one of the simplest chemical structures of a microtubule-stabilizing agent yet described.  51  Chapter IV - The Biology of Dimethyl Varacin 4.1 - Dimethyl Varacin Specific Introduction Varacin, a benzopentathiepin or cyclic polysulfur, was initially isolated from the marine sponge Lissoclinum  vareau and found to be a cytotoxic and antifungal  agent (Davidson et al., 1991). Further work with varacin has shown that it can cleave D N A in vitro which may be the cause of its cytotoxic properties, though in vivo work is required to verify this observation (Chatterji et al., 1998). Other work has concentrated on the polysulfur ring of varacin. In silico calculations have shown that the polysulfur ring may open in a thiol dependent manner to release a triatomic sulfur molecule and a thiolate anion of the remaining varacin (Greer 2001). While this has not been demonstrated for varacin, some thiol reactive agents are known to affect the microtubule network of the cell (Jordan et a l , 1998). In particular, they usually inhibit the polymerization of tubulin in vitro and cause a breakdown in the microtubule network in vivo. Dimethyl varacin was isolated in our phenotypic antimitotic assay. Its structure was elucidated by Lu Yang using X-ray crystallography and is shown in Figure 15 A. The structure of varacin is shown in Figure 15B.  52  Figure 15 - The Chemical Structure of Varacin Compounds The structure of Dimethyl Varacin (A) and Varacin (B) is shown.  53  4.2 - RESULTS  4.2.1 - Antimitotic and antiproliferative  activity  The dose dependence of DMV's activity in the antimitotic assay was examined. It was found that D M V had an I C  50  value of 2 uM in HCT-116 p53 +/+ cells and 2 uM in  HCT-116 p53 -/- cells (Figure 16A). The presence or absence of p53 did not affect the antimitotic properties of D M V . The dose dependence of D M V ' s activity in the antimitotic assay with MCF-7 mp53 cells was also examined with an IC50 of 2 uM, similar to HCT-116 cells. It was next examined if D M V could prevent the proliferation of MCF-7 mp53 cells. D M V does inhibit proliferation with an IC50 value of approximately 50 nM (Figure 16B). The effect of D M V on the cellular proliferation of M D A 231 cells was examined and found to be very similar to that observed with MCF-7 mp53 cells (Figure 16B).  4.2.2 - Dimethyl Varacin inhibits microtubule  polymerization:  The effect of D M V on the polymerization of tubulin into microtubules was examined using an in vitro tubulin polymerization assay (Figure 17). It was found that, similar to 10 uM nocodazole, D M V at 50 uM was able to inhibit the polymerization of tubulin into microtubules. A sample with no drug (1% DMSO) did polymerize.  54  A  0.1  1  10  DMV Cone (uM)  B  D M V C o n e (uM) Figure 16 - Antimitotic E L I C A and Cell Proliferation Assays with Various Concentrations of Dimethyl Varacin and Different Cell Lines (A) Antimitotic ELICA assays were performed with MCF-7 mp53 (closed diamonds), HCT-116 p53 +/+ (open triangles) and HCT-116 p53 -/- (closed triangles) at various doses of D M V . (B) Cell Proliferation Assays were performed with MCF-7 mp53 (closed triangles) and MDA-231 (open inverted triangles) to examine D M V ' s effect of cellular proliferation. In both assays the error bars represent standard deviation of 3 measurements.  55  0.3  Time (min) Figure 17 - Dimethyl Varacin Inhibits Microtubule Formation In Vitro Inhibition of Tubulin Polymerization Assay with D M S O (open squares), nocodazole (open triangles), or D M V (closed circles). The assay was performed with a final tubulin concentration of 3 mg/mL.  56  4.2.3 — Dimethyl Varacin rapidly breaks down the microtubule network in vivo: Indirect immunofluorescence microscopy was used to analyze the microtubule network of cells treated with 10 uM D M V (Figure 18). It was found that D M V disrupts the microtubule network in the majority of cells within 30 minutes. Analysis of cells treated with 0.1% D M S O at 30 and 120 minutes revealed the presence of a defined microtubule network with a clear M T O C . However, in cells treated with 10 uM D M V for 30 and 120 minutes the microtubule network was clearly disrupted. The overall staining was far less intense and is distributed in the cytoplasm in an irregular manner. There did not appear to be any significant microtubule network present with no obvious M T O C . These results indicated that microtubules are disrupted in vivo by D M V treatment.  4.2.4 - Dimethyl Varacin induces mitotic-like phosphorylations from all phases of the cell cycle: Dimethyl varacin was discovered in a phenotypic antimitotic assay in which mitotic cells are identified indirectly, via the detection of a phosphorylation site on nucleolin (Roberge et al., 2000). To further examine the effect of D M V on the cell cycle, flow cytometry was utilized. Flow cyotometry by staining D N A with propidium iodide allows the detection of cells in G i , S, and G2/M phase based on differing amounts of DNA. Cells treated with 1 % D M S O showed a typical D N A profile for asynchronously cycling cells (Figure 19A). Treatment of cells with 300 nM nocodazole for 24 hours resulted in a strong accumulation of cells in G2/M phase (Figure 19A). This type of profile is typical for antimitotic agents. Treatment with either 5 or 25 u M D M V for 5 hours did not produce a mitotic arrest (Figure 19A). Instead, the D N A profile resembled  57  DMSO  10uMDMV  Figure 18 - Dimethyl Varacin Disrupts Microtubules In Vivo Indirect immunofluorescence of ^-tubulin in Swiss 3T3 fibroblasts treated with DMSO DMV.  58  DMSO  Nocodazole  5uM DMV  25uM DMV  Figure 19 - Dimethyl Varacin's Effect on the Cell Cycle Flow cytometry of MCF-7 mp53 cells treated with DMSO, nocodazole for 24 hours, 5 u M D M V or 25 u M D M V for 5 hours. (A) D N A profile of cells stained with propidium iodide. (B) GF-7 versus D N A content. The darkness of the spot represents the number of events.  that of DMSO for asynchronously cycling cells. To gain further information, it is possible to use an antibody with a phospho-epitope typically present only during M phase to distinguish between cells in G2 and M phase. Cells examined in this manner after exposure to 1 % D M S O revealed the expected pattern for asynchronously cycling cells with the majority of cells in interphase (Figure 19B) and a small proportion in mitosis. Cells treated with 300 nM nocodazole for 24 hours resulted in a profile characteristic of antimitotic agents with the majority of cells blocked in mitosis (Figure 19B). However, cells treated with 5 uM D M V for 5 hours yielded very startling results (Figure 19B): the phosphorylations that are typically only observed during mitosis are induced in all phases of the cell cycle including G] and S phase. Treatment with 25 uM D M V resulted in stronger induction of these phosphorylations (Figure 19B). Thus, D M V is not a traditional antimitotic agent, and the antimitotic activity describe in Figure 16 is not due to mitotic arrest but to an increased phosphorylation of proteins in all phases of the cell cycle.  4.2.5 - Dimethyl Varacin does not cause global activation of Cdc2: The primary kinase responsible for mitosis is thought to be Cdc2 (Nigg 2001). It was hypothesized that D M V induces the premature formation of mitotic phosphorylations by causing the overactivation of Cdc2 phoshpotransferase activity. A five hour treatment of MCF-7 mp53 cells with 10 uM D M V led only to an approximate 70% increase in Cdc2 phosphotransferase activity (Figure 20). However, a five hour treatment with nocodazole, an agent known to block cells in mitosis leading to an activation of Cdc2, led to an approximate 300% increase in Cdc2 phosphotransferase activity (Figure 20). This  60  Figure 20  - Dimethyl Varacin Does Not Activate Cdc2/cyclin B  Cdc2 Kinase Assays with DMSO, D M V , or nocodazole. The assay was conducted by an immunoprecipitation of Cdc2/cyclin B from a cell lysate following incubation with the indicated drugs. The assay was only conducted once, but the findings are supported by results from Kinexus Kinetwork™ screens.  61  result indicates that activation of Cdc2 kinase is not responsible for the extensive mitoticlike phosphorylations observed following D M V treatment.  4.2.6 - In gel kinase assays: We wished to further investigate the potential kinase activity changes following D M V treatment. To this end, an in-gel kinase assay was conducted either with or not myelin basic protein (MBP), a general kinase substrate, polymerized directly into the polyacrylamide gel. The gel is treated to renature kinases and the gel is incubated with [y32  P]ATP and active kinases can be detected by the resulting radioactive bands. The  identities of the kinases in question are not needed making this technique an excellent starting place. However, one must be mindful that heteromeric kinases, such as Cdks, and non-renaturable kinases will not be detected (Gowdy et al., 1998). A number of kinases were activated by treatment with D M V (Figure 21 A). Significantly, there was a large increase in overall kinase activity following treatment with 5 uM D M V compared with 1 uM D M V . Kinases of approximately 65, 52, and 44 kDa were strongly activated. The kinases of 65 and 52 kDa were not activated by nocodazole treatment, whereas the kinase at 44 kDa was activated by nocodazole. The 44 kDa kinase in cells treated with nocodazole might be activated by nocodazole induced disruption of the microtubules. Alternatively, if it is typically activated at mitosis increased activity may be observed because nocodazole is blocking the cells in mitosis. The similarities and differences between nocodazole and D M V indicate that while both depolymerize microtubules, the downstream effects are not necessarily the same. Significantly, the concentration at which D M V strongly activates the three kinases (5  62  A  B  *o  2=J. 5i  N  ^  LT>  > 2  > 2  O  "O  Q  Z  co 2  g o  Q  178 —114 —  Q  A _  2 > 2  Q  life-  82 — 61 — 47 — 36 — •mmm* 25 —  •  Figure 21 - In Gel Kinase Assays  Ten \ig of whole cell lysate from cells treated in the indicated manner were separated on a 10% SDS-PAGE containing either (A) 0.2 mg/mL MBP or (B) not. A kinase assay using y- P ATP was conducted directly in the gel. The ladder represents the distance traveled by prestained protein markers (kDa). 32  63  uM) is the same concentration at which D M V was observed to be active in the antimitotic assay and to induce microtubule depolymerization in vivo. It was possible that the active kinases were not phosphorylating MBP but were autophosphorylating. To examine this, an in-gel kinase assay was performed with no polymerized substrate (Figure 2IB). This would most likely examine autophosphorylation; however, it cannot be eliminated that the substrate is a different protein with a similar molecular weight to the kinase. There were similarities and differences observed between the assays performed with M B P present or not. Similar to the case with M B P polymerized into the gel, a band appeared at 44 kDa following both D M V treatment and nocodazole. It is therefore likely that a kinase of approximately this molecular mass was capable of autophosphorylation. However, the kinases at approximately 62 and 55 kDa that were very active with MBP present, did not appear when MBP was omitted. This would indicate that these kinases were incapable of autophosphorylation. Further, a kinase at approximately 34 kDa appeared to be inhibited or degraded following D M V treatment. This was not observed with M B P present. One possible explanation is the presence of a kinase of similar molecular mass capable of phosphorylating MBP but not autophosphorylation. When MBP was present in the gel, the activity of this second kinase masked any changes in the activity of the 34 kDa kinase. However, when there was no MBP present in the gel, no signal from the second kinase was observed allowing one to observe changes in the activity of the 34 kDa kinase. A second explanation is that there was only one kinase capable of both autophosphorylation and phosphorylating MBP at 34 kDa. However, D M V treatment may inhibit its ability to autophosphorylate but not to phosphorylate MBP.  64  4.2.7 - Kinexus Kinetwork ™ phospho-site screens: We had data from both flow cytometry and in-gel kinase assays that a number of kinases were becoming activated post-DMV treatment. However, we had no simple and efficient method of determining the identity of the kinases. To survey the widest possible number of possible substrates in the shortest time, four Kinexus Kinetwork™ phosphosite screens, 1.3, 2.0, 3.0, and 4.0 were conducted. Kinexus Kinetwork™ screens are conducted by multi-immunoblotting (Pelech 2004). In this technique 500 pg of protein from a cell lysate are separated in one broad lane on an SDS-PAGE. The proteins are transferred to a membrane such as nitrocellulose. A plastic manifold with approximately 20 different channels is then placed over the membrane effectively creating 20 equivalent membranes suitable for Western blotting. By conducting thorough testing beforehand to avoid cross-reactivity issues, a mixture approximately 3 antibodies may be added to each channel. The blot is then incubated with appropriate secondary antibodies and developed with E C L (Pelech 2004). This technique allows Kinexus to probe with approximately 35 primary phospho-specific antibodies per protein sample (Pelech 2004). The four screens conducted allowed us to determine changes in the phosphorylation status of 80 proteins at 111 independent sites within a three week turnaround time. This presented a significant time and cost savings over conducting the similar number of individual Western blots. Phosphorylations were detected at 72 of 111 sites (65%). Twenty of the sites investigated had changes between 100%, (or two-fold) and infinity; phosphorylation was not detected at 39 sites (35%). The results of this experiment are summarized in Figure 22. There are numerous observations emerging from these data:  65  Protein Phosphorylation (com)  im  m$  mm soao  mm  t2cco  I«OCO  Accv: 11« S72 M1P-PKTI72 COKi YlS COK I T W 1 & COKI TfC? <lf2aS52.gL[37 elR 82C& ERX1T202/Y204 EFB<2T?S5^I87  FAKY$7$.g.g;  FAK Y57? FAKS722 FAKS910 Hb& GRK2S670 G&-3a$21 fcl48 | CSK3<xV2?§ &SK3J}Y?I0 KKotSlSJ 47-47 KK|»St3l 4 62 SGFIRY1183 -B14 Insulnft Y 9 7 2 JWK p39 JNK p47 Lyn p a  Y5C7  Lyn p*5 Y507 MEK? S2 )/.»S22 MEK1 Y292 MEK! S2S3 MEK113K MEK2 T3S4 M.^a>5S!©?;S2a7 { j j j ^ j f g MKK0S2C7 MLK312? 7i?S2St  MSKU2p?SS37fi 451 MSKU2p858376 255 mTOftS2«H$|-12 tm\ S89S •  Figure 22 - Summary of Data from Kinexus K i n e t w o r k s ™ Phosphosite Screens (continued on next page) The combined results from the Kinetworks™ KPSS 1.3, 2.0, 3.0, and 4.0 screen analyses of the control (light bars) and those from the drug-treated (dark bars) cells were averaged and then graphed. The percentage difference between the drug-treated and control cells is indicated at the end of the bars. Only those target phosphoproteins that were detected with these screens are shown.  66  Protein Phospfsosylation (cpm) m  am  im  PAKpMSUl PAKpSOSUI  PDKi $241 PKBaS4J3 PKCa$£57 PKCS TSOS 4 0 l  PKC8 1538  PKft 1451 PP1aT32D -g^ PfiK1T77S PRX2 IBIS RE/I pSi S 2 » «sflp$9$2§9 -52RST3SS R8SC12 86 S7S0 R&S807ISS11 -1  RBT82I Rik! $380 R s i l T573 8£Kp70T«VH24 SSK o7C $5Kp?3 Sncl 546 Y23iMY240> Stel pGGY23iuY240 Src Y<tlS  im  ^H***  7  Src Y*35*J  STAli S7i? SMT3S72? ^  Figure 22 - S u m m a r y o f D a t a f r o m K i n e x u s K i n e t w o r k s ™ P h o s p h o s i t e S c r e e n s (continued from previous page) The combined results from the Kinetworks™ KPSS 1.3, 2.0, 3.0, and 4.0 screen analyses of the control (light bars) and those from the drug-treated (dark bars) cells were averaged and then graphed. The percentage difference between the drug-treated and control cells is indicated at the end of the bars. Only those target phosphoproteins that were detected with these screens are shown.  67  1. Cdc2 is not activated by treatment with 5 u M D M V for 2 hours. In fact, inhibitory phosphorylations at T14/Y15 increase approximately 30%, while activating phosphorylations at T161 decrease 22%. This is consistent with our earlier finding the Cdc2 is not significantly activated by D M V treatment. Other proteins important in cell cycle control whose phosphorylation was monitored include p53 S392. Consistent with the use of a cell line containing a dominant negative p53 mutation, no phosphorylation was observed in either sample. Phosphorylation changes in Rb were also monitored. Phosphorylation changes in Rb were increased at least 100% at 4 sites. Phosphorylation of T356 increased 148%. S612 increased 278%. S780 increased 36% while S807/S811 increased 66%. Finally, T821 and T825 increased 104% and 161%, respectively. The screen had examined seven phosphorylation sites on Rb. However, this is less than half of those that are known (Harbour et al., 2000). While the effect of phosphorylation at these seven sites on the cell cycle is unknown, it is accepted that hyperphosphorylation of Rb is associated with the release of E2F and the entry of the cell into S phase (Harbour et al., 2000). 2. The most significant phosphorylation increases are observed with kinases in the M A P K family. The phosphorylation status of the activation loop of Erkl/2, JNK/SAPK, and p38 M A P K are all significantly increased. These phosphorylations are typical of activated MAPK. Erkl activation loop phosphorylation (T202/Y204) increased 1560% while Erk2 T185/Y187 phosphorylation increased markedly because the control sample had no  68  observable signal. Two splice variants of JNK, p39 and p47, also increased markedly at T183/Y185. Finally, p38 a M A P K phosphorylation increased approximately 800% at T180/Y182 of its activation loop. Erk5 T218/Y220 phosphorylation was not observed, though it is unclear if this protein is expressed in MCF-7 mp53 cells. The activation loop phosphorylations on MAPKs are catalyzed by M A P K K s . Phosphorylation of the respective M A P K K s (MEK1/2 for Erkl/2, MKK3/6 for p38 a MAPK) was not observed to nearly the same extent. The phosphorylation change of M E K 1 depends on which site is examined. S217/S221 and T292 change only-1% and 1% respectively. S217/S221 phosphorylation is thought to be required for activation of MEK1 (Steelman et al., 2004). It is important to note that the phosphorylations were detected. S298 phosphorylation increases 82% while T386 phosphorylation increases 38%. S298 phosphorylation has been reported to be necessary for efficient phosphorylation of Erkl/2 by MEK1 by promoting M E K 1-Erkl/2 association (Eblen et al., 2004) while T386 and T292 phosphorylation is part of a negative feedback loop with Erkl/2 (Brunet et al., 1994). On MEK2 only one phosphorylation site, T394, was examined and found to decrease by 42%. The significance of the phosphorylation on T394 is unclear. S189/S207 phosphorylation required for the activation of MKK3/6 increased approximately 250% (Raingeaud et al., 1996). The results indicate that while MEK1/2 does not become significantly more activated, this is not the case for MKK3/6.  69  The phosphorylation changes on M A P K K K s vary. M L K 3 T277/S281 phosphorylation decreased by 8%. M L K 3 is a M A P K K K for the SAPK/JNK pathway and the phosphorylation of T277/S281 is required for activity (Leung et al., 2001). Rafl is a M A P K K K for Erkl/2 and its phosphorylation at S259 increased 73% in the p61 splice variant and 28% in the p69 splice variant. The phosphorylation of Rafl at S259 is thought to lead to inhibition of Rafl kinase activity (Pearson et al., 2001). Proteins downstream of the MAPKs are also phosphorylated. MSK1/2 phosphorylation increased significantly. At the S376 site, phosphorylation increased 250% in the p80 isoform and 450% in the p79 isoform. This site is phosphorylated by the C-terminal kinase domain (CTKD) of MSK1/2 (Roux et al, 2004). Erkl/2 or p38 activity is required for activation of the C T K D of MSK1 (Roux et al., 2004). Phosphorylations of RSK1 at S380 and T573 required for activation showed significant increases (Roux et al., 2004). S380 phosphorylation increased 103% while T573 phosphorylation increased markedly. S380 is phosphorylated by the C T K D of RSK1. T573 is thought to be phosphorylated by Erkl/2 leading to activation of the C T K D (Roux et al., 2004). RSK T360/S364 was not detected in either control or D M V treated samples. S364 appears to be essential for R S K activation whereas the significance of T360 phosphorylation is unknown (Roux et al., 2004). MNK1 T197/T202 phosphorylations involved in MNK1 activation (Waskiewicz et al., 1997) were not observed in either the control or D M V sample. It is unknown whether the protein was present in the cell and not phosphorylated or absent. MAPKAPK-2 T334, essential for M A P K A P K -  70  2 activity (Roux et al., 2004), was not detected. JUN S73 phosphorylation increased 80%. 3. The extent of Focal Adhesion Kinase phosphorylation change varied. At Y576 the change was -5%. At the adjacent Y577, phosphorylation increased 88%. These two phosphorylations are thought to be required for maximal activation (Parsons 2003). S722 and S910 increased 80% and 56% respectively. The role of these serine phosphorylations is not understood at present though they may be involved in protein interactions (Parsons 2003). 4.  Glycogen synthase kinase 3 phosphorylation increased 148% and 41% at the inhibitory S21 and at the activating Y279 respectively on the a isoform (Jope et al., 2004). On GSK3P, the activating phosphorylation of Y216 increased 39%, while the inhibitory phosphorylation S9 was not detected in either the control or D M V treated cells (Jope et al., 2004).  5. Phosphorylation at S141 of p21 Activated Protein Kinase (PAK) decreased approximately 25% in both the p54 and p56 isoform. The significance of this phosphorylation is currently unclear. It does reside within the autoinhibitory loop of PAK1 (Bokoch 2003). 6. Changes in P K C phosphorylation on the multiple isoforms. varied. Phosphorylation of S657 increased 44% on P K C a . PKCa/p phosphorylation decreased 18% at T638. S657 and T638 phosphorylations are thought to increase the stability of active P K C (Parekh et al., 2000). Phosphorylation of T505 on PKC5, the absence of which results in low activity (Parekh et al., 2000), increased 28%. PKC8 Y311 phosphorylation, apparently required for maximal activity in  71  response to oxidative stress (Jackson et al., 2004), was not detected in either the control or the D M V sample. PKCs and PKC9 had phosphorylation increases of 63 and 64%, respectively, at S729 and T538. Phosphorylation of S729 on PKCs is required for maximal activity (Parekh et al., 2000). T538 phosphorylation of PKC0 is required for full activity (Xu et al., 2004). Finally, phosphorylation at T410 on P K C ^ decreased 38%. The absence of this phosphorylation results in low activity of PKCC (Parekh et al., 2000). 7. Phosphorylation of adducin a S724 increased 295%. This phosphorylation is thought to inhibit adducin's ability to recruit spectrin and cap fast growing actin filaments (Matsuoka et al., 2000). 8. Phosphorylation of eukaryotic initiation factors also increased. eIF2oc S51 phosphorylation, which reduces protein synthesis (Pain 1996), increased 57%. eIF4 S209 phosphorylation that activates protein synthesis (Pain 1996) increased 29%. eIF2Bs S539 and eIF4E-BPl S65 phosphorylation were not detected. 9. IKKoc SI80 phosphorylation decreased 47% whereas IKK(3 SI81 phosphorylation decreased 62%. These phosphorylations result in the activation of IKK activity (Mercurio etal., 1997). 10. Shcl phosphorylation at Y239/Y240 changed. The amount of change depended on the isoform. p46 had a significant increase; the phosphorylation was undectable prior to treatment with D M V . p66 Y239/Y240 phosphorylation increased 55%. These phosphorylations increase c-myc signaling and may play a role in Erkl/2 signaling (Ravichandran 2001). She p52 Y239/Y240 was not  72  detected. It is unknown if the p52 isoform was not present in the cell or simply not phosphorylated. 11. The phosphorylation status of Src was monitored at two positions, Y418 and Y529, respectively. It was found that Y418 phosphorylation increased 67% while Y529 phosphorylation decreased 19%. Y418 phosphorylation is required for full activation of Src while Y529 phosphorylation is inhibitory (Yeatman 2004). 12. The phosphorylation of Lyn p44 and p46 was monitored at Y507. On p44, phosphorylation increased 22%; phosphorylation of p46 increased 56%. This phosphorylation is associated with inhibition of Lyn kinase activity (Hibbs et al., 1997). 13. PKB and PDK1 phosphorylation status was also monitored. PDK1 S241 phosphorylation, required for PDK1 activity (Casamayor et al., 1999), decreased 15%. PKBcx S473 and T308 phosphorylation increased 172 and 315% respectively. These phosphorylations are required for full activation of PKB (Hanada et al., 2004). 14. P R K l (also known as PKN) T778 phosphorylation increased 63% following D M V treatment. T778 phosphorylation of P R K l is thought to be associated with maintaining P R K l in an active conformation (Peng et al., 1996). PRK2 T816 phosphorylation increased 68%. 15. There were numerous phosphorylations that while detected did not change a significant amount. Insulin receptor Y972 decreased 11% whereas IGFR Y I 163 phosphorylation increased 14%. Mammalian target of rampamycin (mTOR) S2448 phosphorylation decreased only 12%. Phosphorylation of p70 S6 Kinase  73  was monitored at two locations, T421/T424 and T389 where phosphorylation increased 20% and decreased 22%, respectively. STAT1 S727 phosphorylation decreased 15% and STAT3 S727 phosphorylation decreased 9%. A M P - P K phosphorylation on T172 decreased 23% while GRK2 S670 phosphorylation decreased 6%. P P l a T320 phosphorylation decreased 3%. PKR T451 phosphorylation increased 18%. It is important to note that while these phosphorylations did not significantly change they were detected. 16. Numerous phosphorylations were not detected in the phospho-site screens. These include adducin y S662; CREB S133; STAT1 Y701; STAT5 Y694; SMAD1 S463/S465; c-Kit Y730 and Y936; c-Met Y1003 and Y1230/Y1234/Y1235; EGFR Y1068 and Y I 148; ErbB2 Y1139; IRS1 Y I 179 and Y612; Lck S158, Y192, and Y505; Paxillin Y118 and Y31; Pyk2 Y579; Btk Y223; CaMK2 T286; p85 S6K T412 and T444/S447; PKD S916; Syk Y352; and Zap70 Y319. It is impossible to say if these sites were not phosphorylated or if these proteins are not transcribed or translated in this cell line. The phosphorylation of several sites was measured repeatedly in two or more of the different screens. Typically, while the absolute numbers would vary, the trend would be the same. This is true for Erkl T202/Y204, Erk2 T185/Y187, p38 a M A P K T180/Y182, P K B a S473, and CDK1 T14/Y15 amongst others. Notably, some sites including PCK5 T505, JNK p47 T183/Y185, Rb S612, She p46 Y239/Y240 and She p66 Y239/Y240 were undetected in one of the screens but observed in another. The reason for this is completely unknown as the cell lysate sent to Kinexus was homogenous.  74  4.2.8 - Verification Western blots of Erkl/2, SAPK/JNK, and p38 MAPK: To verify the data from the Kinexus screen, Western blots were performed using phospho-specific antibodies for the activation loop of Erkl/2 (pT202/pY204), SAPK/JNK (pT183/pY185), and p38 a M A P K (pT180/pY182) as well as total Erkl/2 (Figure 23). As expected, it was found that the activation loop phosphorylation of Erkl/2, SAPK/JNK, and p38 M A P K greatly increased following D M V treatment. Further, the total levels of Erkl/2 did not change in response to D M V treatment.  4.2.9 - Protein Phosphatase 2A is not inhibited by dimethyl varacin: The phosphorylation status of any protein is the result of a balance between phosphorylation by kinases and dephosphorylation by phosphatases. The results from the Kinexus screens and follow-up Western blots had shown that the phosphorylation state of the activation loop of Erkl/2, SAPK/JNK, and p38 a M A P K greatly increased following a two hour treatment with D M V . However, it was not clear whether this increase was due to an increase in the upstream kinase activity or a decrease in the phosphatase activity. DMV's effect on in vitro protein phosphatase 2A (PP2A) activity was examined. PP2A is inhibited by okadaic acid (Garcia et al., 2003), and okadaic acid can promote the condensation of chromosomes, a hallmark of mitosis, in the absence of Cdc2 activity (Gowdy et al., 1998). Further PP2A has been reported to play a role in the downreguation of Erk signaling (Lechward et al., 2001). No inhibition was found at concentrations up to 10 uM (Figure 24).  75  o  o  o  CO  N 03 T3 O O O  z  11 Ii Q  O  o Phospho J N K / S A P K  —,  Phospho p38 M A P K Phospho Erk1/2  Figure 23 Western Blots of Phospo-Erkl/2, Phospho-JNK, Phospho-p38 a M A P K , and total Erkl/2 from D M V Treated Cell Lysates  Western blots with the indicated antibody were conducted. Ten pg whole cell lysate protein from cells treated in the indicated manner for 2 hours were separated on a 10% SDS-PAGE. The proteins were transferred to PVDF and probed with the indicated antibody. The phospho-specific antibodies for the various M A P Kinases required that both the threonine and tyrosine of the consensus sequence T X Y in the activation loop be phosphorylated. These phosphorylations are required for activation.  76  Figure 24 - Protein Phosphatase 2A Assay Protein phosphatase 2A activity was assayed by using a phosphate sensitive dye that absorbed light at 595 nm when phosphate is released from phospho-peptide substrate. Controls include no substrate and no enzyme with low A595 values. The bars represent the mean of 3 experiments; error bars represent the standard deviation.  77  4.3 - Discussion The investigation of the biology of D M V has revealed a molecule with very interesting effects. D M V was found to inhibit cellular proliferation and affect the microtubule network in vivo and in vitro. The in vivo work demonstrated that the microtubule network was rapidly broken down in response to D M V treatment. This work was confirmed by the in vitro inhibition of tubulin polymerization. Though discovered in an antimitotic screen, D M V is not an antimitotic agent. D M V was found to induce mitotic phosphorylations from all phases of the cell cycle but not by activating Cdc2/cyclin B. D M V has heretofore undescribed effects on the phosphorylation status of the MCF-7 mp53 cell line. Of 111 total phospho-sites examined, 72 were observed to have phosphorylations. Only 20 had changes greater than 100% and 12 were on proteins within M A P K pathways, 4 were on Rb, 2 were on PKBa/p, 1 was on GSK3a, and 1 was on adducin a. D M V is unlikely to be a general phosphatase inhibitor as only 18% of phosphorylation sites change more than 100% in response to D M V treatment nor does D M V inhibit PP2A, a member of the phospho-protein phosphatase (PPP) family (Garcia et al., 2003). It appears that D M V has some degree of specificity as treatment leads to strong activation of all members of the M A P K family investigated (Erkl/2, p38 a M A P K and JNK/SAPK), through an unknown mechanism, with 12 of 20 sites with changes greater than 100% linked to activation of M A P K pathways. D M V was discovered in a phenotypic antimitotic screen and was later found not to be an antimitotic agent nor block cells in any particular phase of the cell cycle. To fully explain this result, it is first necessary to explore the screen's design. The antimitotic screen developed by Roberge in 2000 is an enzyme linked immunocytochemistry assay  78  (ELICA). Key to the antimitotic ELICA's success is the epitope for the TG-3 antibody. While raised against paired helical filamentous proteins from the brains of Alzheimer patients (Vincent et al., 1996), this antibody binds to phosphorylated nucleolin typically present only during mitosis (Dranovsky et al., 2001). Over time cells treated with an antimitotic agent accumulate in mitosis leading to an increase in signal. What is being measured is not an increase in mitotic cells but an increase in the phosphorylation of this epitope. While it is typically only present during mitosis, D M V induces its formation from all phases of the cell cycle leading to a false positive signal in the antimitotic ELICA. GF-7 antibody was also raised against paired helical filaments from brains of Alzheimer patients, but in MCF-7 mp53 cells is typically recognized only during mitosis. However, while the proteins that GF-7 binds to during mitosis are unknown there are at least three bands on a Western blot (unpublished data). Despite D M V not being an antimitotic agent, it inhibits cellular proliferation at concentrations a 1000-fold lower than those that lead to a signal in the E L I C A assay. This is a clear indication that D M V has targets necessary for proliferation besides those that lead to mitotic phosphorylations. D M V causes a rapid breakdown of the microtubules of all cells. The P-tubulin staining is discontinuously spread throughout the cytoplasm. There are remnants of a microtubule network visible in some cells though it is clearly not similar to control cells in both organization and intensity. At 30 minutes the majority of cells treated with D M V has detached from the cover slip and therefore, cannot be stained and observed. Given the rapid nature of the breakdown it is likely that the mechanism of action is distinct from that of nocodazole. At doses that block cells in mitosis, nocodazole has little effect on the  79  microtubule polymer mass of interphase cells (Vasquez et al., 1997). While the microtubule network does not appear completely normal, microtubules are not broken down in response to nocodazole treatment. The finding that D M V causes the breakdown of microtubules in vivo was supported by in vitro experiments showing that D M V can inhibit the polymerization of microtubules. Only excess amounts of drug to the amount of tubulin heterodimer were used. A future experiment to investigate the minimum ratio of D M V to tubulin heterodimer required to inhibit microtubule polymerization is worth consideration. On the other hand, if the ratio of D M V to tubulin heterodimer was near stoichiometric, it would indicate that D M V acts by sequestering tubulin heterodimers. On the other hand, substoichiometric ratios would indicate a different mechanism. The molecular mechanism by which D M V inhibits the polymerization is of interest. There has a report suggesting that the polysulfur ring of benzopentathiepins may be able to form a thiolate anion (Greer 2001). This may attack sulfhydryl groups of cysteine residues, and assembly critical cysteine residues such as C239 in tubulin are known to be sensitive to attack by other cysteine directed reagents (Luduena et al., 1991). One plausible hypothesis is that D M V ' s polysulfur ring opens up and attacks an assembly critical cysteine leading to a covalent modification of cysteine and inhibiting tubulin's polymerization. The most elegant proof of this would be from mass spectrometry experiments in the absence of reducing agents. This experiment, in theory, should readily demonstrate the appearance of a new peak reflecting a covalent tubulin-DMV product following incubation of tubulin with D M V . However, this experiment is complicated by several factors. First are the isotypes of tubulin, at least several of which are present in the purification. Further, these isotypes may have several post-translational modifications  80  leading to increased sample heterogeneity (Luduena 1998). These factors combine to create a very heterogeneous sample significantly complicating the mass spectrometry experiment. Less elegant proofs may include pre-treating D M V with a thiol reducing agent such as p-mercaptoethanol or dithiothreitol and examining for loss of microtubule polymerization inhibition. Evidence of covalent modification may also come from an electrophoretic mobility shift in a non-denaturing gel electrophoresis. Again, this may be complicated by multiple species of close yet differing masses. There is one outstanding question with the hypothesis of D M V attacking a cysteine residue of tubulin. Would D M V bind only to cysteines in tubulin or would any cysteine in any protein be reactive? Presently there is no answer. It is possible that D M V is binding only to a tubulin cysteinyl residue. In this model, D M V would probably initially interact noncovalently with tubulin. This would create a favorable orientation for D M V to attack the cysteinyl residue. Additionally, the possible cysteine residue's environment may make the residue very reactive (Britto et al., 2002). Another possibility is that any cysteine residue in any protein may suffice. This may explain the severe toxicity and differing effects between D M V and other tubulin directed antimitotic agents. It may be that D M V does interact with tubulin leading to its breakdown, yet there may be other targets within the cell that D M V is interacting with. As discussed previously, D M V is not an antimitotic agent but it induces mitotic like phosphorylations. The kinase responsible for these phosphorylations was of particular interest. While no final answer was found, there were some interesting results. The first kinase investigated was the Cdc2/cyclin B complex. As it is thought to be the main kinase responsible for the onset of mitosis (Nigg 2001), it is a logical choice for an  81  activated kinase producing mitotic-like phosphorylations. Cdc2/cyclin B complex was not activated in response to D M V treatment. This was later confirmed with the Kinetwork™ phospho-site screens that indicated an increase in inhibitory phosphorylations and a decrease in activating phosphorylations. The activity status of Cdc2/cyclin B proved another point of difference between nocodazole and D M V . Nocodazole, along with other antimitotic agents, typically causes a large increase in Cdc2/cyclin B activity over time. The finding that Cdc2/cyclin B is not activated by D M V left an important question: If cdc2/cyclin B is not responsible for these mitotic-like phosphorylations, what is? The kinase activity status of the cells following DMSO, D M V , and nocodazole treatment was investigated using an in-gel-kinase assay in an attempt to reach an answer. These two assays, with and without MBP present, provided some interesting results. It was readily seen that a number of kinases were strongly activated two hours after D M V treatment both when MBP was present and absent. Further, they were activated only at a D M V dose of between 1 and 5 uM. This is the same dose that disrupts the microtubules in vivo, suggesting a link between the microtubule network disruption and the activation of the kinases. Significantly, while a number of kinases were activated, the result produced distinct bands. This implies that the phospho-epitope formation was not a byproduct of a global cellular phosphorylation increase. It was evident that some of the kinases were activated in response to D M V treatment alone, whereas others were activated by D M V and nocodazole. This is further evidence that D M V is not a typical antimitotic or microtubule poison. The results with the in-gel-kinase assay were intriguing but provided no identification of the kinases activated.  82  At this point, lysates from D M V and DMSO treated cells were sent to Kinexus for four phospho-site Kinetwork™ screens. These screens provided significant insight into the phosphorylation status of the cell following D M V treatment and were analogous to performing numerous phospho-specific Western blots. O f 111 sites monitored, 20 had changes greater than 100%. The majority of proteins with major changes are involved in M A P K signaling. Four members of the M A P K family were investigated: Erkl/2, p38 a MAPK, JNK/SAPK, and Erk5. M A P Kinase activity is regulated by threonine and tyrosine phosphorylation of the consensus sequence T X Y (Roux et al., 2004). Of the four members examined only Erk5 was not found to have a significant increase in these activating phosphorylations. Phosphorylation of Erk5 was not observed though it is possible that Erk5 is not expressed. The upstream kinases for the Erkl/2 and p38a M A P K pathways were investigated. As expected, activating phosphorylations were observed to increase greater than 100% in MKK3/6 the upstream kinase in the p38 a M A P K pathway. This would suggest that p38 a M A P K is being activated in a traditional manner. The upstream kinase for the Erkl/2 pathway, M E K 1/2, is regulated by multiple phosphorylations. None of the observed phosphorylations increased greater than 100%. In fact phosphorylations at S217/S221 required for activation (Steelman et al., 2004) decreased by 1%. This is indicative that the increase in phosphorylation observed on Erkl/2 does not occur in a traditional manner and another mechanism may be involved. No monitored M A P K K K s had phosphorylation changes greater than 100%. She Y239/Y240 phosphorylation increased markedly. This phosphorylation has been reported to activate the Erkl/2 pathway (Ravichandran 2001).  83  Erkl/2 and p38 a M A P K have many targets downstream including numerous protein kinases (Roux et al., 2004). Of the downstream targets examined, MSK1 had an activating phosphorylation increase of at least 250%. Interestingly, MSK1 can phosphorylate SerlO and Ser28 of Histone H3 (Davie 2003). This phosphorylation is known to be present during mitosis (Nowak et al., 2004). RSK is also downstream of Erkl/2 (Roux et al., 2004) and four phospho-sites were examined. Two of these sites were observed to increase over 100%. However, phosphorylations at T360/T364, required for activation (Roux et al., 2004) were not detected. This suggests that RSK was not active in D M V treated cells. These results also provide a possible answer to a question posed earlier. If Cdc2 is not responsible for the observed mitotic phosphorylations, what is? Erkl/2, p38 a M A P K and JNK/SAPK activating phosphorylations markedly increase and are likely responsible for increasing the TG-3 and GF-7 phosphorylations either directly or indirectly. TG-3 detects a phosphorylation at a proline directed site and members of the M A P K family are proline directed kinases (Roux et al., 2004) similar to Cdc2 (Norbury 1995). Phosphorylations detected by GF-7 may be added either by M A P K members directly, or through downstream kinases such as MSK1. In addition to the M A P K pathways, elements of the PKBa pathway appear to be activated. PKBa phosphorylation increases more than 100% at two activating sites, T308 and S473 (Hanada et al., 2004). GSK3a S21, downstream of PKBa (Hanada et al., 2004), had an increase in phosphorylation greater than 100%. GSK3a phosphorylation highlights contradictory signaling events observed following D M V treatment. S21  84  phosphorylation increases as does Y279 phosphorylation. These two phosphorylations are antagonistic as S21 increases activity while Y279 inhibits activity (Jope et al., 2004). While kinase assays and Kinexus Kinetwork™ screens indicate that Cdc2 was not activated, at least one cell cycle protein was significantly affected by D M V treatment. Four phosphorylation sites on Rb were increased at least 100%. Hyperphosphorylation of Rb is associated with entry into S-phase (Harbour et al., 2000). The last protein whose phosphorylation increased greater than 100% was adducin a. Adducin a is associated with the actin cytoskeleton (Matsuoka et al., 2000). While D M V targets the microtubule cytoskeleton network, as the cell changes shape the actin cytoskeleton may be disturbed. This may lead to the changes in adducin a phosphorylation and warrants further study. The results from the M A P K family are very interesting, particularly Erkl/2 where MEK1/2 does not appear to significantly increase in activity. The role of MEK1/2 may be addressed via U0126 and PD98059, two chemical inhibitors of MEK1/2. Monitoring Erkl/2 activation loop phosphorylation after the addition of either of these inhibitors with D M V may provide insight into the role of MEK1/2 and upstream signaling. If increased upstream kinase activity does not play a role in the M A P K phosphorylations than the presence of U0126 or PD98059 should have little effect. If the activity of MEK1/2 is not increasing, there must be another explanation. The phosphorylation status at any site is a balance of phosphate addition by kinases and removal by phosphatases. Typically M A P K K s add phosphorylations to the activation loop of MAPKs. In turn, MKPs remove these phosphorylations. It is clear that D M V treatment leads to significant increases in the activation loop phosphorylations of  85  Erkl/2, p38 a M A P K , and SAPK/JNK. Further, results from Kinexus indicate that, at least in the case of Erkl/2, the upstream M A P K K was not significantly increased. In that case a possible explanation of the increased activation loop phosphorylations is the inhibition of MKPs (Vogt et al., 2003). The effect of D M V on in vitro M K P activity should be examined (Vogt et al., 2003). Throughout this project there exist two seemingly distinct branches: disruption of the microtubule network and induction of mitotic-like phosphorylations. It is tantalizing to think of possible models to explain them. The first is that they are separate occurrences. That is, D M V directly attacks the microtubules leading to their disruption and directly attacks another target leading to signaling events. It has been established that D M V can inhibit microtubule formation by acting directly on tubulin. It is also hypothesized that inhibiting MKPs would lead to the increase in M A P K signaling (Vogt et a l , 2003). MKPs' catalytic residue is a cysteine (Denu et al., 1995). If D M V can attack a cysteine residue in tubulin, it may equally well attack a cysteine residue in an M K P . The plausibility of this model should be answered by assays to determine D M V ' s effect on M K P activity in vitro (Vogt et al., 2003). A second model is that D M V leads to an increase in signaling, which in turn leads to a rearrangement of the microtubule network. This is the least likely model. There is clear evidence that in vitro D M V can act directly on tubulin and that D M V treatment leads to microtubule breakdown in vivo. Though probable that in vivo D M V is disrupting microtubules directly, this has not been demonstrated unequivocally. There is evidence that M A P K play a role in cell migration and the cytoskeleton (Huang et al., 2004), but activation of M A P K does not typically lead to a dramatic breakdown of microtubules. D M V may activate/deactivate the correct  86  combination of signaling events to lead to the breakdown. The third and perhaps most intriguing model is that D M V is causing a breakdown of the microtubules resulting in strong mitotic phosphorylations. As with the other models, this one lacks clear evidence though there are tantalizing hints. As mentioned repeatedly, D M V can act directly on tubulin in vitro. The glaring problem with this final model is why does D M V breakdown of microtubules produce this signaling, while all other examined microtubule poisons do not? There is no easy answer to that question. Perhaps D M V is attacking microtubules in a manner not previously seen. D M V may attack the middle section of the microtubule leading to its cleavage. While there is no evidence for this, it would be a unique method for a drug-microtubule interaction and may be investigated using cleavage assays initially used to isolate and characterize katanin (McNally et al., 1993). Additional supporting evidence is provided by the effect other microtubule poisons on M A P K signaling (Stone et al., 2000; Zang et al., 2001; Boldt et al., 2002). Nocodazole has been shown to induce Raf-1 activity, which is assayed by monitoring activation of MEK1 and phosphorylation of Erk2 (Zang et al., 2001). In KB-3 cells paclitaxel, vinca alkaloids, and colchicine have been shown to induce INK activity. However, maximal activation occurs 8-12 hours after addition of the drug with minimal activation 2 hours after addition (Stone et al., 2000). In contrast D M V has very strong activation two hours after addition. Dissimilar to D M V treatment, vincristine, paclitaxel, and colchicines all lead to a decrease in activation loop phosphorylations on Erkl/2 and p38 in KB-3 cells (Stone et al., 2000). It has also been described that activating phosphorylations on Erkl/2 and INK occur in response to paclitaxel treatment in HeLa, A341, and MCF-7 cells. 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