The Chemistry and pharmacology of Taxol® and its derivatives

PHARMACOCHEMISTRY LIBRARY

ADVISORY BOARD

T. Fujita E. Mutschler N.J. de Souza D.T. Witiak F.J. Zeelen

Department of Agricultural Chemistry, Kyoto University, Kyoto, Japan Department of Pharmacology, University of Frankfurt, F.R.G. Research Centre, Hoechst lndia Ltd., Bombay, lndia College of Pharmacy, The Ohio State University, Columbus, OH, U.S.A Organon Research Centre, Oss, The Netherlands

VII

LIST OF CONTRIBUTORS Dr. G. Appendino Dipartimento di Scienza e Tecnologia del Farmaco via P. Giuria 9 10125 Torino ITALY Dr. S.H. Chen Bristol Myers Squibb Pharmaceutical Research Institute P.O. Box 5100 Wallingford, CT 06492-7660 U.S.A.

Dr. L. Landino Chemistry Department University of Virginia Charlottesville, VA 22901 U.S.A. Dr. T. MacDonald Chemistry Department University of Virginia Charlottesville, VA 22901 U.S.A.

Dr. T. Cresteil INSERM U75 Universite Rene Descartes 75730 Paris Cedex 15 FRANCE

Dr. B. Monsarrat Laboratoire de Pharmacologie et Toxicologie Fondamentales CNRS 205 Route de Narbonne 31400 Toulouse FRANCE

Dr. R.C. Donehower Division of Pharmacology and Experimental Therapeutics Johns Hopkins Qncology Center Baltimore, MD 21287 U.S.A.

Dr. E.K. Rowinsky Div. of Pharmacology and Experimental Therapeutics Johns Hopkins Oncology Center Baltimore, MD 21287 U.S.A.

Dr. V. Farina Department of Medicinal Chemistry Boehringer Ingelheim Pharmaceuticals 900 Ridgebury Road Ridgefield, CT 06877 U.S.A.

Dr. I. Royer Laboratoire de Pharmacologie et Toxicologie Fondamentales CNRS 205 Route de Narbonne 31400 Toulouse FRANCE

Dr. D. Gu6nard Institut de Chimie des Substances Naturelles CNRS 91190 Gif-sur-Yvette FRANCE Dr. J. Kant Bristol Myers Squibb Pharmaceutical Research Institute P.O. Box 5100 Wallingford, CT 06492-7660 U.S.A.

Dr. D.M. Was Bristol Myers Squibb Pharmaceutical Research Institute 5, Research Parkway Wallingford, CT 06492-7660 U.S.A. Dr. M. Wright Laboratoire de Pharmacologie et Toxicologie Fondamentales CNRS 205 Route de Narbonne 31400 Toulouse FRANCE

The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B.V. All rights reserved

PREFACE Taxol |

a naturally occurring diterpenoid marketed by the Bristol-

Myers Squibb Company, is one of the most exciting antitumor drugs available today. Its current indications (refractory ovarian and metastatic beast cancer) may soon be expanded since the drug is showing activity against lung and head-and-neck cancers. Taxotere |

a closely related analog, is being developed

by Rhone-Poulenc Rorer, and may receive approval this year. Although there are many reasons to be excited about Taxol | there are also reasons to think that chemists can improve upon it. Its low water solubility and the difficulties in formulating it, its lack of activity against certain types of cancer, its numerous clinical side-effects, this drug, all combine to make This book is therefore stimulate further research in

and the rapid emergence of resistance against it a less than optimal chemotherapeutic agent. written for medicinal chemists, in order to this area and to provide the reader with the

necessary background information to start a research program in the area. There are already many reviews on specialized aspects of Taxol | research, as well as two books that cover essentially every possible aspect of research in the field. Therefore, I feel compelled to justify the publication of yet another comprehensive review of the subject, and explain how this work differs from all the other ones. As mentioned above, the book is written mainly for the medicinal chemist, although it should provide useful background to any researcher in the field. I have chosen specific topics that will be relevant in medicinal chemistry research. I will begin by listing the topics that I have not included and why. No historical accounts on the discovery of Taxol | is given here, not because it is not of interest, but because it has been extensively described elsewhere. 1 The "supply problem" is not directly addressed here. It has been widely publicized, especially in the lay press, that supplying all ovarian cancer 1 Wall, M.E.; Wani, M.C. In Taxane Anticancer Agents: Basic Science and Current Status; Georg, G.I.; Chen, T.T.; Ojima, I.; Vyas, D.; ACS Symposium Series; Washington, 1994, p. 18.

patients with Taxol |

would have catastrophic environmental consequences.

At present, the "patient vs. tree" dilemma is no longer an issue, since the core component (a baccatin derivative) from which Taxol | is made can be found in the leaves of the yew tree in high abundance. Other promising fields of research addressing the supply problem, such as plant tissue culture or even fermentation, have been discussed at length elsewhere. 2 For the practicing chemist, the good news is that the baccatins are available commercially, mostly owing to the extensive efforts by Bristol-Myers Squibb and RhonePoulenc Rorer to secure a steady and reliable supply of Taxol | and Taxotere | respectively. With Taxol | soon becoming generic, there will be no lack of companies that will market one or more of the biologically relevant taxanes. Therefore, although the search for alternative methods of taxane production is fascinating and important, these compounds are now becoming available in quantity to researchers. Clinical results are, of course, the endpoint of all pharmaceutical research, and implications obtained from clinical work are discussed throughout the book, especially in chapters 3 and 7. Nevertheless, a detailed account of Taxol | clinical research is not included here, partially because many good accounts have been presented lately, 3 and partially because much of the information contained in these accounts does not help the medicinal chemist in his design of better taxanes. Finally, although scientifically exciting, no account of total synthetic approaches to the taxanes is reported here, for the same reasons given above. Many accounts have been published of the many elegant approaches to Taxol |

and there is no point in duplicating all that large body of information

here. Also, total synthetic efforts have played essentially no role in clarifying the issues of interest to the medicinal chemist, i.e. S t r u c t u r e - A c t i v i t y Relationships (SAR), and will probably contribute very little in the near future, due the structural complexity of these molecules. The present book opens with a review of the naturally occurring taxoids (Chapter 1), written by Prof. Giovanni Appendino. The chapter is not a comprehensive list of all taxoids isolated to date, but attempts a systematic 2 Taxol: Science and Applications; Suffness, M., Ed.; CRC: Boca Raton (in press). 3 Holmes, F.A.; Kudelka, A.P.; Kavanagh, J.J.; Huber, M.H.; Ajani, J.A.; Valero, V. In Taxane Anticancer Agents: Basic Science and Current Status; Georg, G.I.; Chen, T.T.; Ojima, I.; Vyas, D.; ACS Symposium Series; Washington, 1994, p.31.

approach at describing the different classes of taxoids, with particular reference to all skeletal types and the various functionality patterns. Biosynthetic studies are also discussed, as well as some of the basic chemistry and common functionalities of the taxoidic skeleton. Although Taxol | and the baccatins have been the starting materials for the preparation of analogs for SAR studies, this does not have to be true in general and, with a knowledge of the various taxanes available in nature, one can plan the synthesis of compounds that are not readily accessible from the baccatins. In turn, the search for novel compounds of this family may ultimately lead to new antitumor substances. Chapter 2, also by Prof. Appendino, deals with the structural identification of taxoids, mostly by spectroscopic means. The section on NMR spectroscopy contains the first detailed analysis of the influence of structural factors on proton and carbon chemical shifts in taxoids, and therefore should be of extreme utility to workers in the field. The pictorial analysis of 19 1H and 13C spectra of a number of representative taxoids should provide instant help to chemists who are attempting to identify new taxanes (natural or synthetic). Chapter 3, by Dr. Dolatrai Vyas, discusses the formulation of taxanes. After a detailed discussion of the various Taxol |

formulations of possible

clinical relevance, the chapter explores the concept of prodrugs for the purpose of achieving water solubility and bioequivalence with Taxol| The chapter will be useful to all medicinal chemists working on these drugs, as well as on second-generation analogs. Some of the concepts discussed are general enough to be of interest to all medicinal chemists. Chapter 4, By Prof. Michel Wright et al., deals with the metabolism and pharmacokinetics of Taxol| and Taxotere | Knowledge of the fate of a drug in vivo, and specifically its biodistribution, plasma concentration and half-life, as well as inactivation by metabolic transformations and excretion, is extremely important in planning the synthesis of new analogs. New compounds could be specifically prepared to avoid known metabolic processes that lead to inactivation, like side-chain cleavage and oxygenation (hydroxylation) reactions. In Chapter 5, Dr. Shu-Hui Chen and myself give a comprehensive account on the chemistry of taxanes in relation to SAR studies. The SAR field has undergone an explosive growth. In the early 80's, the scarcity of material

as well as of interest in the drug conspired to keep our knowledge of Taxol | SAR to a minimum. The last 4 years have witnessed a frenzy of publications, as academic and industrial laboratories compete to solve problems of chemoselectivity, in order to modify the functional array of the taxanes around the molecule's core. From these efforts, a clearer picture of the role of the various functionalities on the mode of action of the taxanes is emerging. Some issues remain to be resolved, but the next obvious step is to try to design new, easily accessible, scaffolding systems that may hold the essential binding elements found in Taxol |

in the right spatial configuration for

proper binding to microtubules. In addition it is obvious that, among the hundreds of synthetic analogs of Taxol | that have been prepared, some had to have better binding properties or cytotoxicity than Taxol | itself, and this is being reported with increasing frequency. A fuller evaluation of these analogs in vivo is, of course, needed before unrealistic claims are made. Analogs

endowed with better potency or lower toxicity will probably emerge. In addition, since the primary mechanism of resistance to the taxanes appears to be of the mdr type, it seems reasonable to assume that a taxane within a tumor cell will be available for binding with microtubule structures and with the P-glycoprotein (the export p u m p ) in a competitive fashion. While decreased affinity for the promiscuous export p u m p seems hard to engineer, higher affinity for the target (already precedented) may have the overall effect of increasing intracellular drug concentration by shifting the equilibrium in favor of the tubulin-drug complex, and therefore may reduce or suppress resistance

in vitro and perhaps even in vivo. The answers to all these

questions have not emerged yet, but will likely be the subject of future investigations. In C h a p t e r

6, Dr. Joydeep Kant discusses

SAR aspects of the

phenylisoserine side chain. The C-13 side chain is an important element in the binding of Taxol |

to its biological target, and it has become necessary to

devote a whole chapter to this topic in view of the many chiral approaches to phenylisoserines for SAR studies and the m a n y analogs prepared. With baccatins becoming readily available, the easiest modifications that can be made are the ones that incorporate new side chains, and there is no doubt that medicinal chemists will be active in this area for years to come. The issue of how the side chain folds both in solution and at the binding site has piqued the interest of many workers and is also discussed in this chapter.

Finally, in Chapter 7 Prof. Timothy Macdonald and Lisa Landino discuss the mode of action of the taxanes and the mechanisms of resistance. The dynamics of microtubules and the many sophisticated controls for the formation of these interesting and important structures from soluble tubulin are given special emphasis. Learning the precise binding site of Taxol | within its target should help design better analogs, or even attempt

de n o v o

design of Taxol | mimics with completely different structures. In addition, the understanding of how cancer cells become resistant to taxanes may help develop new strategies and modalities in cancer chemotherapy. Research in the Taxol|

area is proceeding at record pace. About half of

what we know about this drug and its analogs, at least measured in terms of number of papers, has been learned in the last two years. An analysis of publications in this area (Figure 1) shows, after the slow 70's, a more rapid phase in the 80's, with

ca.

100 papers/year, followed by an exponential phase

in the 90's coinciding with the clinical development of Taxol |

In 4 years the

number of papers published yearly has increased almost ten-fold, although the relative growth seems to be slowing and perhaps ready to plateau. 1000 900

800 700 600 o

500 400

~ aoo 200 100

0 oO O~

~cO O~

~ o13 O~

r cO O~

~I" oO O~

~ I~0 O~

r4D oO O~

~ o13 O~

~ ~ O~

O~ o13 O~

0 O~ O~

~-O~ O~

04 O~ O~

03 O~ O~

year Figure 1: Publication trend in the taxane area over the last 14 years (searched through Current Contents and Medline)

O~ O~

I surmise therefore that an update of the field in 1995 is especially timely and justifiable. Finally, one w o r d about names: taxol was the name given by its discoverers to the active principle of Taxus brevifolia. The Bristol-Myers Squibb Company, on registration of this c o m p o u n d as its own brand, has made the unusual and perhaps unfortunate choice of registering the trivial name as its own trademark. The c o m p o u n d should then be referred to as Taxol |

Since the generic name has been withdrawn, the company had to

choose a new generic name for the c o m p o u n d , and picked "paclitaxel", certainly not an attractive choice. This has generated confusion in the literature, and in general workers in the field, with the exception of BMS workers, have obvioulsy not paid any attention to the name change, and continue to use the term "taxol" instead of "paclitaxel". In this book, we use the name Taxol |

and paclitaxel, but not the old trivial name. Rhone-Poulenc

Rorer has followed suit, registering the trivial name of their clinical candidate as their o w n trade name (Taxotere |

and baptizing the generic version

"docetaxel". Whatever their names, I am certain that these derivatives will continue to be at the center of the attention in the field of cancer chemotherapeutics, and I hope this book will help workers to advance the field with newer, even more exciting results. A c k n o w l e d g e m e n t s : I wish to thank Dr. Helen Oen (Boehringer Ingelheim, Scientific Information) for keeping me up to date on the copious literature in this area over the last two years, and Mary Feron for extensive retyping of parts of the manuscript. I dedicate this book to my parents, Renato and Wanda Farina.

Vittorio Farina Boehringer Ingelheim Pharmaceuticals 900 Ridgebury Rd Ridgefield CT 06877

Ridgefield, 2/6/95

The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B.V. All rights reserved

1 NATURALLY OCCURRING TAXOIDS Giovanni Appendino Dipartimento di Scienza e Tecnologia del Farmaco, via Giuria 9, 10125 Torino, Italy

1.1. I N T R O D U C T I O N

Taxoids (taxane diterpenoids) are a structurally homogeneous class of compounds that occur in two genera of the yew family (Taxus and Austrotaxus). The very limited distribution of taxoids within the plant kingdom is the result of the peculiar taxonomical position of the yew tree, which stands relatively apart from the other seed plants. Furthermore, fossils of ancient yews (Paleotaxus rediviva, T. jurassica, T. grandis) show a close similarity to the modern yews, suggesting a limited evolution through the ages. Part of the remarkable adaptability and evolutionary longevity of the yew tree is presumably related to its complex and peculiar secondary metabolism. Indeed, ancient trees like the yew and the gingko are a storehouse of biologically active compounds, whose complex and unique molecular frameworks give us a glimpse of the biochemical virtuosity of early plant chemistry. The most famous yew (T. baccata L.) is considered a dying-out species. It has become almost extinguished in natural plant communities, and only survives as an important element of green areas (parks, gardens). With the current rate of plant extinction, one may wonder how many taxols and gingkolides will be missed by future generations of plant chemists.

1.2. S Y S T E M A T I C S OF THE YEW T R E E

The yew family ( T a x a c e a e ) has only five g e n e r a ( A m e n t o t a x u s , Austrotaxus, Pseudotaxus, Taxus and Torreya) [1, 2]. Other genera t h a t used to be included in the Taxaceae family (Cephalotaxus,

P o d o c a r p u s ) a r e now

considered part of independent families. Owing to the absence of seed cones, the yew family has sometimes been excluded from the conifer order, raising its taxonomic state to a new order or even class. The systematics of the genus T a x u s are controversial. The yew tree is distributed t h r o u g h o u t the n o r t h e r n hemisphere, and occurs in eight distinct geographical regions. Little, if any, overlap between these enclaves exists, and yews are commonly named from their area of distribution (Table 1). Yews look very much alike, and the presence of only one collective species (T. baccata L.) is often assumed. A classification of this type was proposed by Pilger, who divided the genus into seven subspecies (see Table 1) [1]. However, more recent dendrological work recognized the infraspecific taxa as independent species (e.g. Krfissmann [2], Table 1), and this opinion prevails nowadays. In addition to the n a t u r a l species, two infraspecific hybrids are shown. They originated from cultivations in North America, on breeding the Japanese yew with the European yew (T. x media Rehd.) and with the Canadian yew (T. x hunnewelliana Rehd.). The needles of one cultivar of T. x media Rehd. (T. x media Rehd. cv. Hicksii) contain Taxol| and Taxol-related compounds in amounts comparable to those reported from the bark of the Pacific yew, and might become an i m p o r t a n t and renewable source of Taxol |

[3]. The genus Austrotaxus is monotypic, and its

only species (A. spicata Compt.) is endemic to the New Caledonian rain forest. Confusion exists in the phytochemical literature as to the identity of the Himalayan and the Chinese yew. The former is often referred to as T. baccata L., following Pilger's classification [1] and the Index Kewensis, whereas the Chinese yew, referred to as T. mairei Hu ex Liu, T. chinensis Rehd., T. y u n n a n e n s i s Cheng et L.K. Fu or T. celebica (Warburg) Li., might actually comprise more t h a n one species. As a result of this taxonomic shuffling, one has to source information on a specific yew under several different Latin names. Especially confusing is the fact t h a t the name T. baccata L. has also been applied to some Asian yews. The distinction between the various yews is difficult, and mainly based on three morphological characters: the length of the needles (10-30 mm), the way

they are attached to the twigs (straight or bent), and the shape of the bud scales (crenate and pointed or non crenate and blunt) [1, 2]. In the absence of fruits, the distinction between the yews and some species of Cephalotaxus and Torreya can be difficult even for the best trained eyes. This is evidenced by the name cephalomannine given to a Taxol| analog isolated from a plant identified as Cephalotaxus mannii Hook at the time of collection and chemical analysis [4], but later recognized as a yew species (T. waUichiana Zucc.) [5]. Table 1. The Systematics of the Genus Taxus

Trivial name

Pilger classification [1]

Krtissmanm classification [2]

European yew

T. baccata subsp, eubaccata Pilger

T. baccata L.

Himalayan yew

T. baccata subsp. wallichiana (Zucc.) Pilger

T. wallichiana Zucc.

Chinese yew

T. celebica (Warburg) Li. T. cuspidata Sieb. et Zucc.

Japanese yew

T. baccata subsp, cuspidata (Sieb. et Zucc.) Pilger

Pacific yew

T. baccata subsp, brevifolia (Nutt.) Pilger T. baccata subsp, globosa (Schlechtd.) Pilger

T. brevifolia Nutt.

Florida yew

T. baccata subsp, floridana (Nutt.) Pilger

T. floridana Nutt.

Canadian yew

T. baccata subsp. canadensis (Marsh.) Pilger

T. canadensis Marsh.

Mexican yew

T. globosa Schlechtd.

T. x media Rehd. T. x hunneweUiana Rehd. Many varieties and cultivars of yew have been developed for ornamental purposes. Krtissmann lists 139 of them, giving clues to their identification [2]. No comprehensive survey on the phytochemical pattern of the various yew species, hybrids and cultivars exists. However, some general trends are

l0 emerging, at least at the species level, from the wealth of data published in the last few years (see section 1.10). 1.3. HISTORICAL P E R S P E C T I V E

The first studies that correctly established the constitution of the taxane nucleus appeared in 1963, when three groups (Lythgoe's [6], Nakanishi's [7] and Uyeo's [8]) independently reported their conclusions regarding the carbon skeleton of some Taxus constituents. However, interest in the chemistry of the yew tree dates from the mid-nineteenth century, since a mixture of taxoids was obtained by the German pharmacist Lucas as early as in 1856 [9]. Reading a report by a French veterinarian on the poisonous properties of the yew, Lucas remembered a fact occurred in his own town (Arnstadt) two decades earlier, when a flock of sheep had been placed in a fenced yard landscaped with a few large yew trees. The following day five or six sheep (he could not remember exactly) had died, and he was contacted by a veterinarian, who asked him to analyze the stomach of the dead animals. Poisoning from heavy metals was suspected, but Lucas could not find any evidence for this. Inspection of the yard where the sheep had been kept revealed that the yews had been stripped of leaves as high as the animals could reach. Poisoning from the yew seemed the most plausible explanation for the death of the animals, and the yard's owner had all the yews eradicated from his property. The report by the French veterinarian and his own experience made Lucas suspect the presence of alkaloids in the yew. Many compounds of this type had already been isolated, and some of them were highly poisonous (strichnine, nicotine, coniine). After a laborious extraction, Lucas obtained an amorphous white powder showing basic properties. He named the material taxine, -ine being the ending given at that time to alkaloids. An improved isolation procedure was worked out by Marm~ twenty years later [10]. His preparation gave a crystalline and apparently purer material, but this claim was not substantiated by later workers, who always described taxine as an amorphous powder giving amorphous salts. The first systematic investigation on the biological properties of taxine was carried out by Borchers in 1876 [11]. He recognized the high toxicity of this alkaloid and described its action on the respiratory system and the heart.

ll The structural characterization of taxine was extremely slow. The early studies were unsuccessful, and the first clue came only in 1923, w h e n Winterstein showed that taxine is the ester of a polyalcohol esterified with acetic acid and (L)-~-dimethylamino-~-phenylpropionic acid [12]. Divergent physical constants were reported for taxine (mp 82-124~

[a] D +35-96~ and these early

studies could not dispell the obvious suspicion that taxine was actually a mixture of compounds. A breakthrough came in 1956, a hundred years after the isolation of taxine, when Graf showed t h a t this alkaloid is a mixture of at least seven compounds [13]. G r a f w a s able to obtain three of them in pure form (taxine A, B and C, Figure 1), but the structural elucidation of these compounds was achieved only recently [14-16]. HO AcO'"

I

O

~ ~ sIlI ~

0

O

NMe2

RO 3.1.1 3.1.2

Taxine A R=Ac T a x i n e C R=H

O

.O.

NMe 2

OH 3.1.3 Taxine B Figure 1: The Taxines A different approach was followed by Lythgoe, who discovered t h a t chemical modification of taxine can afford pure compounds. Thus, after acetylation and conversion of the Winterstein esters into cinnamic esters, two products were obtained (5-cinnamoyltriacetyltaxicin I and II), whose constitution was established in 1963 [6], and stereochemistry three years later [17]. The same

12 approach

was

followed

by

Nakanishi

[7]

and

Uyeo

[8].

5-

Cinnamoyltriacetyltaxicin II turned out to be identical to taxinine, a compound obtained in 1925 by Kondo and Takahashi from the needles of the Japanese yew [18]. Taxinine is thus the first natural taxoid to be obtained in pure form and structurally elucidated (Figure 2). The fact t h a t taxine was characterized well after the advent of chromatographic techniques is surprising, since the toxicological relevance of the yew tree has not diminished, and cases of human and animal poisoning are still reported on a regular basis [19]. One has to consider, however, that taxine is unstable, being decomposed by acids and light, and that many of its constituents are prone to isomerization during the purification procedure (see section 1.8). Furthermore, two seminal discoveries shifted the attention of the scientific community towards other constituents of the yew tree. In the late 1960s, interest in taxine was overshadowed by the discovery of the outstanding antitumor properties of Taxol | [20] and by the detection of large amounts of ecdysones in yew tissues [21]. The isolation and structural elucidation of Taxol| reported by Wani and Wall in 1971 [21], was a remarkable accomplishment, because of the low concentration (ca. 0.02% of dry bark weight) and structural complexity of this compound. AcQ

OAc O ,,,,O~jl

0 R 3.1.4 3.1.5

OAc

R=OH (5-Cinnamoyltriacetyltaxicin I) R=H (5-Cinnamoyltriacetyltaxicin II, Taxinine) Figure 2

In those years, systematic studies on the non-alkaloidal constituents of the yew were undertaken by the groups of Nakanishi in Japan and Halsall in England. These groups discovered new members of the taxane group of diterpenoids. Halsall in particular is responsible for the numbering of the taxane skeleton used today, and for the isolation of the baccatins. His entire work was published as a series of six short notes [22-27], and details on the isolation of

13 these important compounds were never reported. In the 1980s, Potier's group in France isolated several new taxanes, including a structural type characterized by a C-12, C-17 oxygen bridge [28], and several analogs of Taxol|

[29]. Other

major achievements by the French group were the first partial synthesis of Taxol| described in 1988 [30], and the discovery of the excellent antitumor properties of Taxotere | a semisynthetic taxane now in advanced clinical study [31]. The partial synthesis of Taxol|

from 10-deacetylbaccatin III, a taxoid

available in relatively large amounts (up to 0.1%) [30] from a renewable source (needles and clippings of several yew species) solved the supply problem and paved the way for the commercialization of Taxol| as a drug. The literature on naturally occurring taxoids was reviewed by Kingston et al. in 1993 [32]. Their work updated a series of previous reviews, summarizing

what was known on the occurrence and the reactivity of taxoids up to March 1992. This work covers relevant literature up to, and including, August 1994.

1.4. REPRESENTATIONS, NUMBERING AND TRIVIAL NAMES Taxoids are compounds having a [9.3.1.03, 8] tricyclopentadecane ring system or closely related skeleta. The bidimensional representation of the taxane skeleton offers the opportunity for a number of drawing modes, but gives no clue as to the actual shape of the molecule (see Figure 10 in chapter 2). Two planar representations are in use (Figure 3): the linear one, by Lythgoe (A) [6], and the angular one, by Miller and Kingston, (B) [32,33]. Both have merits and drawbacks. In both representations, the orientation of the substituents at the tetrahedral ring carbons of rings B and C is represented by the conventional stereochemical symbols (thickened and broken lines) with reference to the a and faces of the molecule. These are defined as in steroids (a=lower face, ~=upper face), observing the molecule with the methyl group at C-8 (C-19) placed in the "northern hemisphere" and pointing toward the observer. The absolute configuration of the natural taxoids is such that, if the molecule is oriented in this way, ring A is to the left and ring C to the right side of the observer. If the taxane ring system is drawn according to Lythgoe (A), ring B is not in its most expanded form, and C-15 is the apex of a reentrant angle. Its substituents must thus be drawn inside ring B. Since reentrant and vertex angles are related by a C2-rotation along an axis passing through the Cn-1 and C n + l atoms, the meaning of thickened and broken lines is reversed, and the actual orientation of

14 a substituent is the opposite of what is intuitively expected. Therefore, the C-16 methyl group, cis to the C-19 methyl and ~ according to the steroid convention, is represented here by a broken line, and the opposite is true for C-17. 18

10

13 / a

9 19(17)

1~.~'(20) r I

"'

2

A

16

6

m(16)

H

B

C

Figure 3: Bidimensional representation of the taxane stereoparent (old methyl numbering in parentheses). In the Miller-Kingston representation (B), ring B is drawn in its most expanded form, and the actual orientation of the gem-dimethyl groups is straightforward from their stereochemical symbols. However, this representation is difficult to draw rigorously, since the perimeter of ring A is too small to contain, in graphical terms, the gem-dimethyl groups. Thus, these are placed into different rings: C-16 (~, thickened line) into ring B, and C-17 (a, broken line) into ring A. Another point of concern is the way the orientation of the substituents on ring A and on the bridgehead carbon C-1 are indicated. Lythgoe considered C-1 as a cyclooctane ring B carbon joined by a three-carbon a-chain (C-12 through C14) to C-11 [17]. The substituent at C-1 (hydrogen of hydroxyl) is thus ~ (cis to the methyl at C-8), but descriptors different from a/~ are needed for the other ring A substituents, since the fragment C-12 through C-14 is considered a t r a n s a n n u l a r bridge and not part of the main ring system. For these substituents Lythgoe used an exo/endo notation [17]. The stereochemical descriptors for the Lythgoe and Miller-Kingston representations are based on the B-C ring system, and the C-1 to C-14 bond should be drawn using a broken line. In practice, the perimeter of the ring system is drawn using a line of normal thickness for this bond as well, and this conventional representation is well established. It must be emphasized, however, that in the current bidimensional representation of taxoids, the stereochemistry at

C-1 is actually not indicated, since only one stereochemical descriptor is

15 employed for the substituents of this stereogenic carbon. This is unambiguous in fused systems, but not in bridged systems. Furthermore, the a/~ notation is applied to taxoids in a rather peculiar way, not fully consistent with the steroid rules. In all n a t u r a l l y occurring taxoids, the non-nuclear C-1 substituent (hydrogen or hydroxyl) is cis to the C-19 methyl, and ~ according to the Lythgoe convention. The most correct and practical representation of the taxane skeleton would be one that considers it as a cyclodecacyclohexane with a methano bridge, and the Chemical Abstract name of taxanes is based on this ring system (Figure 3, C). As with the gem-dimethyl groups of camphor, no stereochemical descriptor would be necessary for C-16 and C-17, which could be written with lines of normal thickness but in different rings (cf. the Miller-Kingston representation [22, 32]), and defined not as a/~, but as syn or anti to a certain element (e.g. the C-19 methyl group). Furthermore, all substituents along the periphery of the ring system would be defined by the same descriptors (a/~), avoiding the cumbersome use of syn / anti for those at C- 12, C- 13 and C- 14. The stereochemistry at C-1 could also be clearly indicated, making the course of some reactions obvious from simple inspection of the bidimensional r e p r e s e n t a t i o n (e.g. the syn allylic epoxidation t h a t established the C-1 functionality in Nicolaou's synthesis of Taxol | [34]). The numbering of the taxane skeleton is based on the IUPAC name of the parent ring system (4,8,12,15,15-pentamethyltricyclo[9.3.1.03,S]pentadecane), and was proposed in 1964 by the leaders of the three major groups working on this class of compounds (Lythgoe, Nakanishi and Uyeo) [35]. In 1969, Halsall proposed a different numbering for the methyl groups [23], similar to the one used in cembranoids and other diterpenoids (tiglianes, daphnanes, ingenanes), where the gem-dimethyl groups are C-16 and C-17. Both systems were used during the following decade, but in 1978 the IUPAC blue book adopted Halsall's numbering [36], which has been the only one used since then. It must be emphasized, however, that in 2(3->20)abeotaxanes (taxine A-type compounds) biogenetic and structural numberings are different (see section 1.6). Many taxanes have been assigned trivial names, derived from the botanical name [baccatins (Figure 4), austrospicatine, brevifoliol] or the geographical location (taiwanxan) of the yew tree where they were first found. A combination of both has also been employed (taxagifine). Many trivial names of taxoids begin with the syllable tax, resulting in confusing proliferation of similar

15 names, often with additional suffix numbers (taxagifines) or letters (taxinines, taxchinins, taxchins, taxuyunnanines). Particularly frustrating is the situation with the taxinines, since only a few scattered letters have been employed (A,B,E,H,J,K,L,M, Figure 5), a situation reminiscent of that of gingkolides (A,B,C,M). The structure of baccatin II is not known. Based on the molecular formula [22], this compound may be 1-hydroxybaccatin I, which generally cooccurs with baccatin I. AcO

OACoAc

AcO

A c O ....

O OH

H O .... R"

v

H

iiii

O

OAc

OAc

HO

4.1.1 R=H, Baccatin I 4.1.2 R=OH, Baccatin II (?)

OBz

OAc

4.1.3 7~-OH, Baccatin III 4.1.4 7(z-OH, Baccatin V OAc " OAc

AcQ

A c O .... O HO

OR

OAc

4 . 1 . 5 R=Ac, Baccatin IV 4.1.6 R=Bz, Baccatin VI 4 . 1 . 7 R=n-Hexanoyl, Baccatin VII

Figure 4: The baccatins Further confusion arises from the fact that also non-taxoidic compounds isolated from the yew (taxicatine) and even synthetic products (taxilan, taxylone) begin with the syllable tax. To complicate matters even more thoroughly, spelling problems exist. Thus, in the English literature, the alkaloidal mixture from the yew has been referred to both as taxin or taxine, but baccatins are taxoids and baccatine is a triterpenoid [37]. Furthermore, baccatin and baccatin I are the same compound [25], but taxinine [7, 8] and taxinine A-M (Figure 5),

17 referred to as taxinin and taxinins A-M by Chemical Abstract, are different compounds. AcO

o

OAc

AcO ~,

OAc ,~ OAc

_

Y

OAc

.... OOinn

OAc

4.1.10 Taxinine B

3.1.5 R=Cinn, Taxinine 4.1.8 R=H, Taxinine A 4.1.9 R=Ac, Taxinine H AcO OAc R

AcO ~,

OAc ~ R

AcO ....

O

....OCi an

"'OR OAc

OAc

4.1.13 R=H, Taxinine K 4.1.14 R=Ac, Taxinine L

4.1.11 R=H, Taxinine E 4.1.12 R=OAc, Taxinine J AcO AcO j / O n

0

" ' " ""

I

Bz

" /OAc

, .... Iss I

OH

OAc

4.1.15 Taxinine M Figure 5: The taxinines

More t h a n one trivial name has been assigned to the same compound, and taxchinin A [38] and 2a-acetoxybrevifoliol [39] are the same compound, as are t a x a c u l t i n [40] a n d taxol D [41]. To avoid this i n t o l e r a b l e confusion, a nomenclature system based on the names of only a few basic taxane structures would be highly desirable. Figures 4 and 5 show the formulas of the baccatins

18 and taxinines. Most of these compounds were structurally elucidated in the sixties and seventies, and their names are rooted in the literature. Whenever possible, plant chemists should try to name new taxoids as derivatives of these compounds. Taxols are baccatin III derivatives esterified at C-13 with phenylisoserines bearing various N-acyl and N-alkyl groups. An alphabet system was proposed by Potier [29], using suffix letters to distinguish between compounds bearing different N-acyl groups (Figure 6). New letters are introduced in alphabetical order whenever a new taxol is isolated [40-42a]. This system allows a rational naming of closely related compounds, and deserves widespread use. Also, it avoids the misleading name cephalomannine for N-debenzoyl-N-tigloyltaxol (now taxol B). Chemical modification within the diterpenoid core can occur naturally at C-7 (epimerization [32, 42c], xylosidation [29]), C-9 (reduction [42b]) and C-10 (deacetylation, oxidation [32, 42c], esterification with ~-hydroxybutyric acid [29]). Overall, twenty-two natural taxols are known to date.

RIN'R20 phil"_

AcO

O OH

0 ....

oH

HO

-

BzO

OAc

4.1.16 RI=BZ, R2=H, Taxol A (Taxol) 4.1.17 Rl=Tigloyl , R2=H , Taxol B (Cephalomannine) 4.1.18 Rl=n-Hexanoyl , R2=H , Taxol C 4.1.19 Rl=n-Hexanoyl, R2= Me, N-Methyltaxol C 4.1.20 Rl=n-Butanoyl , R2=H , Taxol D (Taxacultin) Figure 6: The taxol alphabet 2.5. CHEMODIVERSITY AND STRUCTURAL TYPES 2.5.1 Skeletal types Natural taxoids are rather homogeneous in functional complexity. Indeed, compared to other classes of terpenoids, the structural variations within taxoids

19 are limited, and many compounds only differ in their esterification pattern (e.g. baccatins IV, VI and VII, see Figure 4). The main structural diversity was found within taxanes from the tropical species Austrotaxus spicata Compt. [43, 44], and interesting new structural types might be present in other yews that cross the equatorial line (Indonesian and Malayan yews). However, these plants have not yet been investigated from the botanical or the chemical point of view. Besides taxanes, three other skeletal types of natural taxoids are known (Figure 7), resulting from closure of an extra ring between C-3 and C-11 (3,11cyclotaxanes), or from rearrangement of ring A and ring B [(11(15->1)- and 2(3>20)abeotaxanes, respectively)]. Taxanes are by far the most widespread skeletal types of taxoids, accounting for 96 out of 101 natural taxoids listed in Kingston's review [32, 39], but recent studies have highlighted the importance and widespread distribution of the other minor skeletal types [16, 39]. The four skeletal types of taxoids are exemplified by taxinine [18], brevifoliol [39, 45], taxine A [14], and taxinine K [46], the first compounds of each type to be isolated (Figure 7). Under mild conditions (see section 1.8), certain 13- or 9-oxo-All-taxoids can be turned into 3,11-cyclotaxanes [46, 47], and C-1 hydroxylated All-taxenes can rearrange to 11(15-> 1)abeotaxanes [48]. Taxanes and 2(3->20)abeotaxanes actually have a different biogenesis (see section 1.6). Many other skeletal types have been obtained by radical, cationic or anionic rearrangements of taxanes (see chapter 5), but none of them has so far been encountered in nature. 1.5.2. Functionalization of the terpenoid core Taxanes: The site of main structural variation in the terpenoid core is the C-4/C-20/C-5 moiety. According to its functionalization, taxanes can be divided into five different structural types (Figure 8): 5a-hydroxy-A4,2~ (olefin-type, A), 5a-hydroxy-4~,20-ether type (epoxide-type, B), 4r (oxetane-type, C), 4(z,5a, 20-triol-type (D) and 5~,20-diol-type (E). The tertiary hydroxyl at C-4 is generally esterified, but the secondary hydroxyl group at C-5 and the primary one at C-20 can occur both in free and esterified form. Oxo bridges are always ~, an important observation in regard to the biogenesis of these structural types.

20 AcO ~,

OAc -

0

....0 . ~ ~

Ph

OAc 3.1.5 Taxinine, a taxane OBz OAc " OAc

AcO 11

HO ....

O -

OAc

"r "'.. sssS

s~ 0

OH

OAc

HO 5.1.1 Brevifoliol, a 11(15-> 1)abeotaxane

4.1.13 Taxinine K, a 3,11-cyclotaxane

O NMe 2 ~ , ~ ....0 Ph

AcO . . . . . . . . ..-" AcO

2o

OH

3.1.1 Taxine A, a 2(3->20)abeotaxane Figure 7: Skeletal types of natural taxoids Another i m p o r t a n t point of variation is the oxidation state of C-9 and C13, where both a hydroxy or a keto group can occur (Figures 4 and 5). However, four compounds with unfunctionalized C-13 [23, 49, 50], three compounds with both C-9 and C-13 unfunctionalized [23, 50, 51], and one compound with unfunctionalized C-9 [50] have been described. C-l, C-2, C-7 and C-14 can be oxygenated or not, w h e r e a s C-6 is always unfunctionalized. The t e r t i a r y hydroxyl at C-4 is generally esterified, but the secondary hydroxyl group at C-5 and the p r i m a r y one at C-20 can occur both in free and esterified form.Oxo

21 bridges are always ~, an important observation in regard to the biogenesis of these structural types.

5 '"OH(OCOR)

_ "OH(OCOR)

20 A

B

~

C

"OH(OAc)

'"OAc

1

OH(OAc) D

O OAc(OH)

OAc E

Figure 8:C-4/C-20/C-5 Functionalization types All of the natural taxanes, with the exception of taxagifine and its derivatives, have a double bond at C-11/C-12. This plays an important role in stabilizing the twist-boat conformation of the cyclooctane moiety of the taxanes and in preventing transannular interaction between the substituents at C-3a and C-12a [53]. Indeed, A ll-taxenes are an important example of "hyperstable olefins", where the linear and the angular strain associated with the introduction of a bridgehead double bond are overridden by the decrease in t r a n s a n n u l a r interactions and I-strain caused by the conversion of tetrahedral carbons into trigonal ones [54]. Various combinations of functionalities can occur, as shown in Figures 4 and 5. However, important associations of functional groups exist, possibly due to the presence of "gene cassettes". For example, all oxetane-type taxoids are also oxygenated at C-2 and C-7 (cf. baccatins II-VII, figure 4), whereas all taxoids with a C-13 keto group bear a double bond or an epoxide at C-4/C-20 (cf. taxinines A, B, H, K, L and M, Figure 5).

22 AcO AcO

I

"

OAc

0

0 ....O " j ~ " H

Ph

: OAc 5.2.1

Figure 9: Taxagifine, on oxo-bridged taxane 3,11-Cyclotaxanes: All the non-alkaloidal compounds of this type isolated to date are phototaxicin I and II derivatives [46, 55, 56], and have the same acylation pattern as their corresponding and co-occurring taxicines. However, the taxane corresponding to the 3,11-cyclotaxane pseudoalkaloid spicaledonine (2a-acetoxycomptonine) is unknown as a natural product [44]. AcO

~,

0

OAc

-

"i, "-.. ....OR 2

OAc 5.2.2 RI=OH, R2=Cinn , Triacetyl-5-cinnamoylphototaxicin I 5.2.3 RI=H, R2=Cinn , Triacetyl-5-cinnamoylphototaxicin II 5.2.4 RI=H , R2=COCH(OH)CH(NMe2)Ph , Spicaledonine Figure 10: Some naturally occurring 3,11-cyclotaxanes 11(15->1)Abeotaxane~: All natural compounds of this class (see 5.1.1, Figure 7), bear an oxygen function at C-15 [39], whereas acidic treatment of C-1 hydroxylated taxanes gives mostly rearranged products of this class, but with a C-15/C-16 double bond [48]. Oxygen bridges can form between C-15 and C-10 [57] and between C-15 and C-13 [58], presumably by intramolecular nucleophilic quenching of cations centered at C-15 and C-13, respectively. No 11(15>1)abeotaxanes with a C-13 keto group have been reported.

23

2(3->20)Abeo.taxanes: All compounds of this type are derivatives of taxine A [14] (3.1.1, Figure 7), and only differ in their acylation pattern [16, 59]. 1.5.3. Acylation patterns All hydroxyl groups of taxoids, with the exception of that at C-1, can be found esterified with various acids and aminoacids (Table 2) [60, 61]. As a result, taxoids can be classified as pseudoalkaloidal or non-nitrogenous. The aminoacid can be nicotinic acid or a series of phenylpropanoid ~-amino acids (Winterstein acid, phenylisoserines, variously O- and N-acylated or N-alkylated). Nicotinyl esters of terpenoids are relatively rare as natural products, and occur mainly in the dihydroagarofuran-type sesquiterpenoids from Celestraceae. Phenylpropanoid ~-aminoacids are instead typical building blocks of yew constituents. Nicotinyl residues are found at O(9), and phenylpropanoid aminoacids at 0(5) and O(13). Aminoacids at 0(5) are N-alkylated, and those at O(13) N-acylated [42]. Two exceptions are known: an O(5)-pseudoalkaloid from A. spicata Compt. with a free amino group [44] and the O(13)-pseudoalkaloid N-methyltaxol C, where the aminoacid nitrogen is both alkylated and acylated [40, 42a]. The aminoacid nitrogen of phenylisoserine can be acylated with benzoic, tiglic, capronic (hexanoic) or butyric acid (cf. the taxol A-D series, Figure 6). The non-nitrogenous acids esterifying the hydroxyl groups of the diterpenoid core can be acetic, cinnamic, benzoic, capronic, a-methylbutyric, [5hydroxybutyric or (z-methyl ~-hydroxybutyric acid. Acetyl residues can be found at the tertiary hydroxyl group at C-4 and at all the secondary hydroxyl groups with the exception of C-14. The other acyl residues have instead a more specific location, suggesting the involvement of selective acylases. Interestingly, the acylation pattern of taxanes and ll(15->l)abeotaxanes is generally different, and benzoyl residues have a wider distribution in abeotaxanes, being found not only at C-2, but also at C-7, C-9 and C-10 [39, 58]. Non-enzymatic acyl migrations have been observed between the hydroxyl groups at C-7, C-9 and C-10 [62-64] and between those at C-2 and C-14 [65] (see section 1.8). It is therefore likely that the acylation pattern observed in some taxoids is the result of both enzymatic and non-enzymatic pathways.

24 Table 2. Common Side Chains of Taxoids Structure

Name

Abbreviation

-OCOCH3

Acetate

Ac

-OCO(CH2)4-CH3

Hexanoate

-OCO(CH2)2-CH3 O OH

Butyrate ~-Hydroxybutanoate

0 a-Methylbutanoate

l-oJ

MeBu

a-Methyl-~Hydroxybutanoate

0 Cinnamate

~- 0

v

-Ph

-ok@h 0

Winterstein acid

NN e

ON' -OCOPh 0

l-o

Cinn

Phenylisoserinate Benzoate

Bz

Tiglate

Tigl

Nicotinate

Also, the n a t u r e of the ester group can be affected by non-enzymatic reactions, since Winterstein acid esters are converted into E-cinnamic esters under mild acidic conditions. This conversion can be sometimes useful for the

25 characterization of alkaloidal taxoids. For preparative purposes, Hoffmann elimination after nitrogen quaternization has been used [66], but the more straightforward method is the Cope elimination of the corresponding N-oxides [67]. Treatment with m-chloroperbenzoic acid in THF turns Winterstein acid esters into E-cinnamates. In alkaloidal taxoids, formation of the N-oxides is much faster than double bond epoxidation, and if only a moderate excess of peracid is employed, quantitative yields of the corresponding cinnamates can be obtained [16]. The N-oxides of phenylisoserine derivatives are more stable and can be isolated, as shown by the preparation of the N-oxide of taxine A [16]. 1.5.4. Glycosidation Patterns Taxol| and some analogs have also been isolated in glycosidic form from woody tissues of T. baccata L. (trunk, roots) [29]. In all cases, the residue was Dxylose and the glycosidic bond was at the C-7 hydroxyl group. 1.6. B I O G E N E S I S

1.6.1. Carbocyclic skeleton The taxane skeleton is a terpenoid, and many authors have speculated upon its origin, but very few biosynthetic studies have been carried out. The early suggestion that taxanes are degraded triterpenoids like quassinoids [8] has been dismissed, and the current view is that taxoids are diterpenoids from both the structural and the biogenetic point of view. Lythgoe was the first to propose a reasonable biogenetic derivation for the taxane skeleton [6b]. His scheme (Figure 11) involves the head-to-tail cyclization of E,E,E-geranylgeranyl pyrophosphate (6.1.1) to a C-15 (taxane numbering) macrocyclic cembrene cation (6.1.2). Quenching of the positive charge by the C-11/C-12 double bond and loss of the C-11 proton might afford a verticillane derivative (6.1.4). A transannular cyclization of the 1,5-diene system of this bicyclic intermediate would eventually give the tricyclic taxane skeleton. The transannular cyclization of a 1,5-diene system is a common leitmotiv in the biosynthesis of isoprenoids, and the relationship between verticillenes and taxanes is the same existing in sesquiterpenoids between germacradienes and eudesmanes. The 2(3->20)abeotaxanes could be derived from a modification of this scheme, mediated by double bond isomerization prior to the 1,5-diene cyclization (for the numbering of 2(3->20)abeotaxanes, see section 1.4), and the

25 3,11-cyclotaxane skeleton is probably the result of the photocyclization of suitable taxane precursors. The ll(15->l)abeotaxanes might derive from C-1 hydroxylated taxanes via a Wagner-Meerwein rearrangement, or, alternatively, from 1,15-epoxycembrene derivative 6.1.5, via a transannular epoxide cyclization.

6.1.1 OPP

6.1.5

6.1.4 Verticillene ~ , ~

6.1.2

Q H+

6.1.6

~ H

H

t

H+

H

OH 6.1.9

11( 15-> 1)Abeotaxane

6.1.7 Taxane I

~2 " 6.1.8

HQ

2

6.1 10

o

2(3->20)Abeotaxane

6.1.11

3,11-Cyclotaxane

Figure 11: Biogenesisof the taxoidic skeleta

27 Although this scheme for the derivation of the taxane skeleta seems plausible, it should be noticed that the configuration at C-1 of most cembranes and all verticillanes isolated from plants is different from that of the taxanes [68, 69], and t h a t attempted cyclization of cembrane and verticillane derivatives failed to give any detectable amounts of compounds with the taxane skeleton [7O]. These observations do not disprove Lythgoe's proposal, but may point to a more subtle mechanism. On the other hand, the course of the cyclization of verticilladienes might be steered toward taxane derivatives by a conformational bias induced by the oxygen functions. Indeed, all naturally occurring taxoids are heavily functionalized, with a number of oxygenated sites spanning from three (taxuyunnanine D, 6.1.12 , Figure 12 [50]) to eleven (taxagifine III,6.1.13 [71]), and no simple taxane hydrocarbon has ever been isolated. Furthermore, the only report of the transformation of a verticillane-type diterpenoid into a compound

friedo-verticillane

with a taxane-like skeleton is the one in the trioxygenated clemeolide, 6.1.14 [72]. 0 \\ Aco

....

HO ... O I "..li_L

, ....

H O

p'.,.... H OH

__Ac 6.1.12, Taxuyunnanine D

O OAc

6.1.13, Taxagifine III

O

AcO

OAc " OAc

~

0

OAc

....

%

~

0 //

AcO,,, "OAc OH

6.1.14, Clemeolide

OAc

6.1.15, Taxchin A

Figure 12: Examples of biogenetically related skeleta

28 A survey of the various synthetic procedures employed to assemble the taxane skeleton [73] highlights the importance of radical and anionic processes, but reactions of this type have rarely been employed by Nature for the assembly of terpenoid skeleta. 1.6.2. C-4/C-20/C-5 Functionalization The transannular cyclization of the 1,5-diene system of verticillanes gives a C-4 taxane cation. This might evolve into a 4(20)-double bond, present as such or in vestigial form (epoxide, diol, oxetane) in all but one (taxchin A, 6.1.15 [74]) of the natural taxoids. The further oxidative elaboration of the double bond, following or concomitant to C-5 oxygenation, has been the subject of much speculation, especially with regard to the formation of the oxetane ring (Figure 13). Potier's group has elaborated a synthetic procedure for the transformation of the 4a,5(z,20 triol system (A) into a 4-hydroxy-5~(20)-oxetane (B) [75], and a similar approach was also used by Danishefsky on a model compound [76]. Both schemes are based on an SN2-type displacement of a 5a-leaving group by the C20 hydroxyl moiety. The sequence triol->oxetane is biogenetically plausible and supported not only by the results of studies on model compounds, but also by the occurrence of 4,20,5-trioxygenated taxoids with the correct configuration at C-4 and C-5 [77]. The role of the 4~(20)-epoxides (C) and their synthetic relationship with the 4(20) olefins are instead not obvious. Halsall postulated the conversion of 4~(20)-epoxides into 4a,20-diols and then into oxetanes via anchimerically assisted opening of the epoxide by a C-5 hydroxyl or C-5 ester carbonyl group [26]. This sequence is plausible, but no study reporting a reaction of this type in taxoids or model compounds has appeared. Direct formation of the 5~(20)-oxetane from natural 4~(20)-epoxides via an oxabicyclobutonium ion (D, Figure 13) was proposed by Swindell [89], but studies on model compounds did not support the idea. Furthermore, in all natural taxoids, the 4,20-epoxide is ~ [41], whereas the epoxidation of A4(20)taxenes afforded exclusively the a epoxides, due to the steric effect of the C-8 ~methyl group [41]. Thus, it is not clear whether the natural 4~(20)-epoxides are the precursors of the 5~(20)-oxetanes, or whether they derive instead from the 4(z,20 diols via an alternative ether bridge formation. Another possibility that should not be discounted is the occurrence of a convergent process, involving more than one path, for the formation of the oxetane ring. In all schemes for the

29 oxetane ring formation, the final step is an SN2-type reaction at C-5, where the leaving group may be a phenylpropanoid aminoacid migrating to C-13, a fascinating hypothesis proposed by Potier [79].

"'OH 20 4,20-dihydroxylation

r 4

20,5-ether formatio~/~

5

,s,

',0~ H

m

H

A

r

"•20,4-ether ormation

r

5

~

4

0

_

OH

,5 "OH

s

s SS

F ~~

4

5

| D

Figure 13: Plausible biogenesis of the oxetane ring of taxoids

1.6.3. Phenylpropanoid Aminoacids Many taxoids have the C-5 or the C-13 hydroxyl groups esterified with phenylpropanoid ~-aminoacids. According to the definition of Hegnauer [80], these taxoids are pseudoalkaloids, since they contain nitrogen, but their cyclic carbon skeleton is not derived from an aminoacid. Winterstein acid and N-alkyl and N-acyl phenylisoserines are typical constituents of yew pseudoalkaloids.

30 Their derivation from phenylalanine has been confirmed by feeding experiments [81-84], but exact details of the various steps are not known, nor have the relevant enzymes been characterized. The observation that cinnamic acid is not incorporated into T a x u s ~aminoacids suggests the involvement of a dyotropic rearrangement, catalyzed by an aminomutase, that turns phenylalanine into its corresponding ~-aminoacid (Figure 14) [84]. This step is reminiscent of the phenyllactate-> tropate rearrangement in the biosyntheis of atropine and related alkaloids [85]. The formation of Winterstein acid from phenylalanine takes place with the exclusive loss of the p ro-R C-3 hydrogen [82], an observation that is consistent with the finding that cinnamic acid is not incorporated into yew ~aminoacids [84]. Indeed, the enzyme phenylalanine:ammonia lyase transforms phenylalanine into E-cinnamic acid by removing the C-3 pro-S hydrogen, and the biosynthesis of cinnamic acid and the yew ~-aminoacids are distinct. Hydroxylation at C-2 and N-acylation or N-alkylation would then complete the biosynthesis of the side chain of taxol- and taxine A-type pseudoalkaloids. Feeding experiments have shown that, during the biosynthesis of Taxol| the side chain is not attached in its final form, but as phenylisoserine, whose acylation takes place after C-13 esterification [86]. As discussed in section 1.5, the yew ~-aminoacids generally occur at 0(5) when N-alkylated and at O(13) when N-acylated [42]. Winterstein acid esters are often accompanied by the corresponding cinnamates, and cinnamoyl groups are thus generally found at 0(5). One exception exists, i.e. the ll(15->l)abeotaxane taxchinin B, 6.1.16(Figure 15), where the cinnamate is at O(13) [87]. It has been suggested t h a t 0(5)- and O(13)-esters are related by the intramolecular migration of the aminoacid side chain from 0(5) to O(13) [79]. Although no experimental evidence for a reaction of this type is known, inspection of models shows the plausibility of this hypothesis, since the taxane skeleton has an inverted cup shape, and the oxygen function at C-13 and C-5 are in close spatial proximity. Doubts on this hypothesis were cast by feeding experiments, since labeled baccatin III could be incorporated into Taxol | even though its functionalization pattern precludes O(5)->O(13)-transesterification [86]. If a simple biogenetic relationship between 0(5) N-alkylated and O(13)-Nacylated alkaloids does exist, its manipulation could allow one to switch the metabolism of yew needles from the abundant (1%) N-alkylated esters like taxine B to the much rarer (ca. 0.004-0.01%) N-acylated esters like Taxol|

31 phJ~1

CO 2H mutase NH2 r p hH ' ~ " v - -CO2 NH 2

(L)-Phenylalanine

N-Methylation = Winterstein acid

1. Hydroxylation 2. Esterificationwith baccatin(ROH) _NH2 _NHBz p h j ~ z / _ CO2R gzCoA ph~:~,~,_ CO2 R _

OH Phenylisoserine

OH Taxol

Figure 14: Biosynthesis of Taxus ~-aminoacids and of Taxol|

BzO

_OACoAc

CinnO .... O HO

OAc

6.1.16 Figure 15: The structure of Taxchinin B

Minute amounts of taxoids are also produced by Taxomyces andreanae, a fungal endophyte of the Pacific yew [88], whose ability to synthesize taxanes is a remarkable example of horizontal transfer of genetic material [89]. The fungal and the plant biosynthesis of Taxol|

could be distinguished with labeled

precursors, since the aminoacid leucine is a precursor of Taxol| in T. brevifolia Nutt. but not in Taxomyces andreanae, where formation of ~-hydroxy-~methylglutaryl CoA (and thus isopentenyl pyrophosphate) from this aminoacid is negligible compared with its formation from acetate [88]. Further details on the fungal biosynthesis of Taxol| are not known. 1.7. C H E M I C A L R E A C T I V I T Y OF T A X O I D S

Taxoids are relatively unstable compounds, being sensitive to acidic, basic and oxidizing conditions. Furthermore, the inverted cup shape of these skeleta

32 and the high density of functional groups make a variety of intramolecular reactions

possible.

We briefly

summarize

here

the

major

types

of

transformations that are somewhat general for taxoids. For a thorough review on the chemistry of

Taxol|

and specifically the synthesis of analogs for

Structure-Activity Relationship (SAR) studies, the reader is referred to chapter 5 and 6. 1.7.1. Skeletal rearrangements Modifications of the carbon connectivity of all three rings of the taxane system have been reported. Most of these rearrangements have been observed in baccatin III derivatives, and are discussed in chapter 5. The focus here is on on rearrangements observed in other structural types of taxoids. Ring A rearrangements: Formation of a cationic species at C-1 triggers the Wagner-Meerwein rearrangement of ring A, resulting in contraction of this ring and formation of a tertiary cationic species at C-15 (Figure 16) [39, 48]. The latter can eliminate an a-proton (7.1.3) or be quenched by a nearby hydroxyl (7.1.4) or a water molecule (7.1.5) [57]. The formation of a cationic species at C13 can trigger two different rearrangements of ring A, depending on the substituent present at C-1 (Figure 17). In C-1 hydroxylated taxanes, the C-1/C15 bond fragments, generating a hydroxylated cation at C-1 and eventually affording cyclodecene derivative 7.1.8 [90]. When a hydrogen is present at C-l, the C-1/C-15 bond fragments in a different way, generating ring-contracted C-15 cation 7.1.10 [22]. Contraction of ring A was observed during the irradiation of certain taxinine derivatives (Figure 18) [91]. In these compounds, the usual photochemical reactivity of 13-oxoA4(2O),ll-taxoids (hydrogen migration from C-3 to C-12 and bond formation between C-3 and C-11) was shut down by the saturation of the 4(20)-double bond or by the formation of the C-9/C-10 acetonide (as in 7.1.14, Figure 18), and the formation of cyclopropanated derivatives 7.1.17 was observed instead, via C-C bond migration between C-1 and C-12. Removal of allylic activation for the C-3 hydrogen, introduction of steric constraints due to the presence of the acetonide methyls, or a longer C-4 to C-11 distance may be responsible for this behavior.

33 HO 1

HO"'

7.1.1

HO

HO ....

7.1.2

Figure 16: Cationic rearrangements of ring A: C-1 trigger T r a n s a n n u l a r cyclizations:

Reactions of this type have been observed under

radical and photochemical conditions. The radical cyclization of baccatin III derivatives is discussed in chapter 5. The irradiation of taxinine-type compounds causes an unusual photochemical reaction, involving hydrogen transfer from C-3 to C-12, the enone (z-carbon (7.1.13, Figure 18). In reactions of this type, hydrogen transfer to the enone [~-carbon or to the ketone carbonyl is generally observed. This r e m a r k a b l e cyclization was discovered by Nakanishi [46], and proceeds via a ~,~* triplet or a C-11/C-12 diradicaloid [89]. The inverted-cup shape of the taxane skeleton makes the hydrogen transfer to the C-11 or to the enone oxygen impossible, whereas H-3 and C-12 are spatially dose and can form a bond.

34

0

7.1.6

7.1.7

7.1.8

7.1.9

7.1.10

7.1.11

H+

Figure 17" Cationic rearrangements of ring A: C-13 trigger Ring B rearrangements" The rearrangement of taxinine to anhydrotaxininol (Figure 19) holds a venerable position in taxoid chemistry, since it was discovered as early as 1931 [92], and represents the first skeletal rearrangement observed in this class of compounds. The rearrangement takes place in a basic madium, v i a a vinylogous retro-aldol fragmentation, followed by aldol condensation between the C-14 enolate and the C-9 formyl, and by SN2-type displacement of the C-2 acetate by the C-14 enolate. The stereochemistry of anhydrotaxininol is not known. 1.7.2. Functional group modifications Hydroxyl groups: The very low reactivity of the 5a-hydroxyl group toward acylation and silylation seems general in all skeletal types. In reactions of this type, the 13-hydroxyl generally shows a lower reactivity than the C-2, C-7 and C9 hydroxyls, and in oxetane-type compounds this is enhanced by the formation of hydrogen bonding with the C-4 ester carbonyl [30]. The relative reactivity of the other oxygen functions depends on the functionalization pattern, and no general rule can be given as to their relative reactivity in esterification or hydrolysis reactions. Furthermore, the reactivity order of the various hydroxyl groups in oxidation reaction does not parallel that observed in acylation and silylation reactions.

3.5

AcO .~

AcO

~'

:~

hv

0

=

0

H-3

, %

OH : OAc

Cinn

3.1.4

H.I AcO~ .,~ O

OCinn migr.

OIH'~_. H ~ OAc 7.1.12

OAc F;~,

AcO

"" " ~ ~ ~ =L_I..., 9 .&P %

OIH"~__ - I OAc

C-3/C-ll bond

OCinn

formation

?Ac

H ....

~ O

",~, "-.-

% "

,,

"OCinn

OAc

7.1.13

5.2.2

oXo .-~

oXo ~ .~ "OAc

H

OAc

Ac

:

OAc 0

0

0

C-l, C-12bond=,O~

~

migration

J, bondform.: O [1 "c~

: 7.1.16

OAc

C-1,C-11

0

X

"OAc

II ~Ac

7.1.17

OAc

Figure 18: Photochemistryofthe 13-oxo-All-enone systemin taxoids

36 AcO

~OAc

HO

(OH

/_,J O

KOH =

O

"OCinn

.... H

OAc

"OH

:

OAc 7.1.18

3.1.5

O I

O Z

"-"~ 0

O I ~

H 7.1.19

OAc O

9

0

H

-

OH

"OH

--7.1.20

OH

0

"OH

OH

OH

%'O

7.1.21, Anhydrotaxininol Figure 19: The rearrangement of Taxinine to Anhydrotaxininol Thus, the allylic 13-hydroxyl, which is difficult to acylate, can be oxidized very easily even with reagents not normally used for the oxidation of alcohols to ketones (MCPBA, OsO4, DDQ, NBS) [93]. The oxidation of the side-chain hydroxyl of phenylisoserine taxane esters leads to a-ketoesters that are stable in the Taxol |

(N-acylation)

series

[94], but are easily degraded to the

corresponding enones, probably via a cyclic mechanism, in the taxine A (Nalkylation) series [16]. Vicinal hydroxyls (C-9/C-10; C-1/C-2 or C-1/C-14) can be protected as cyclic acetals [6b], carbonates [95], or orthoesters [96]. The C-l, C-2 carbonate served as a benzoate precursor in Holton's [97] and Nicolaou's [34] total syntheses of Taxol |

37 Keto groups: In natural taxoids, keto groups occur at C-9, C-10 and C-13. The C9 carbonyl is sterically hindered and unreactive toward reducing agents and other nucleophiles. The C-13 and C-10 keto groups do not react with hydrazinetype reagents [6b], but can be reduced with borane [98] or hydrides [34]. Formation of only the a-alcohol at C-13 was reported in a baccatin III derivative (Figure 20 [34]), but closely related compounds gave mixtures of both epimers as well as the product of overreduction [16, 75]. The unusual electronic structure of All-13-oxo and/or 9-oxo-taxanes is discussed in section 2.3. It is worth noting t h a t the regioselectivity of the reduction of 13-oxo-All-taxanes is rather capricious, and sometimes opposite to what expected. The formation of taxininol (7.2.1, Figure 20) upon t r e a t m e n t of taxinine with lithium aluminum hydride involves a remarkable conjugate reduction of the enone system to a ketone [99]. Furthermore, the 13-oxo group of taxinine-type compounds is totally unreactive under conditions in which baccatin-type compounds are reduced [59]. Double bonds: Reactions of the 4(20)-double bond of taxanes yields predictable products. Thus, this double bond is cleaved by ozone [6b], whereas catalytic hydrogenation [6b], osmylation [75], and epoxidation [41] occur with exclusive attack from the less hindered a-face. The catalytic hydrogenation is plagued by the hydrogenolysis of the C-5 oxygen function, especially in polar solvents [100]. On account of a more caged and h i n d e r e d structure, A4,10-3,11-cyclotaxanes and their hydrogenolysis products are inert toward hydrogenation [100]. The C-11/C-12 double bond is quite unreactive toward electrophilic and nucleophilic addition as well as toward catalytic hydrogenation. Some reactions have occasionally been described, but appear to be associated with a specific functionalization pattern of the taxane skeleton [41]. Thus, the epoxidation of some A4(20),ll-taxadienes occurred preferentially at the endocyclic double bond [41], but in closely related compounds where the hydroxyl groups allylic to the l 1-double bond are acylated, peracid attack occurred mainly at the exocyclic double bond (7.2.7 v s . 7.2.8, Figure 21) [41], whereas under these conditions baccatin III derivatives were oxidized to the corresponding 13-enones (see 7.2.10, Figure 21) [16].

38 1.8. I S O L A T I O N ARTIFACTS

PROCEDURES,

ANALYSIS

AND

EXTRACTION

Two taxoids (Taxol| and 10-deacetylbaccatin) are commercially available. Their isolation from various plant parts has been thoroughly investigated and optimized, but details have not been disclosed for obvious proprietary reasons. AcO ~,

OAc .:

HO

LiA1H4, THF H

:

"OCinn

~ 7

~ ..,

ref. 99 H

3.1.5, Taxinine

OH

OH

OAc

AcO

OH

7.2.1, Taxininol

O OTES

AcO

O~

O OTES

NaBH4, MeOH, 84%~HO .... O HO

: OBz

ref. 34

O

OAc

HO OBz

7.2.2

HO

OAc 7.2.3

20TES

,,

HO ~,

0 / / OTES

% P

O

NaBH4, MeOH, 34% HO" _-

O HO

: OBz 7.2.4

OAc

ref. 16

""

O

OBz 7.2.5

Figure 20: Hydride reductions of the 13-oxo-A11-taxenesystem On a laboratory scale, early studies on the yew focused on the basic alkaloidal constituents, and used aqueous acids for the extraction of the plant material [66]. Neutral and polar solvents (CH2C12, CHC13, methanol, ethanol,

39 acetone) are used nowadays for the extraction of the whole taxoidic fraction. Nonpolar solvents (i.e. petroleum ether) have occasionally been used for the selective extraction of relatively nonpolar taxoids, like the baccatins from the heartwood of T. baccata L. [101]. Extracts from the needles are generally purified by partition between aqueous methanol and petroleum ether. In this way, part of the pigments, i.e. simple phenolics like 3,5-dimethoxyphenol as well as lipids are removed.

RO

OR

OR

\

RO ....

MCPBA _ "O R

H

RO ~, /

RO,,,,~~,.

OR " OR *

CH2C12,

H

ref. 41

0

"O R

7.2.7 7.2.6

+

RO ,

R

7.2.7

H Ac

traces major 80% 20%

O

OR J,

"

OR

7.2.8

Ro

,

"oR 7.2.8

,,,', .o

HO ~,

O

HO

/70_H

0

MCPBA

0

_

O

OBz

7.2.9 R=H 7.2.10 R=OH

CH2CI2, ref. 16

O

OBz 7.2.11 R=H (69%) 7.2.12 R=OH (62%)

Figure 21: Reactions of All-taxenes with peracids

An alternative technique to remove these compounds is the treatment of a 1:1 water-ethanol solution of the extract with lead(II) acetate. A procedure of this type can also be applied to seed extracts for the removal of the abundant lipid

40 fraction [102]. The purified extracts are then separated using conventional chromatographic techniques, open column chromatography on SiO2 still being the most extensively used one. HPLC, both on normal and reverse phase, can be used for f u r t h e r purification of the c h r o m a t o g r a p h i c fractions. More sophisticated techniques (high speed planetary countercurrent chromatography) have also been used [103]. Most taxoids are crystalline compounds, and crystallization is very useful as a purification method. Unfortunately, mixed crystals can sometimes form, especially with taxols [16]. The qualitative and quantitative analysis of yew extracts is routinely carried out by HPLC on reverse phase columns (phenyl, cyano or C-18) [3]. Since the polarity of taxoids varies considerably and the extracts can contain up to 5060 different taxoids in detectable amounts, no universal solvent for the resolution of these mixtures exists. The analysis generally focuses on taxoids having a defined range of polarity (e.g. taxols, baccatin III and deacetylbaccatin III), and selectivity is achieved using UV detection, thus avoiding interference from non UV-active taxoids (brevifoliol, baccatin IV). The speed of the analysis is an important parameter when a large number of samples have to be analyzed, and MS/MS techniques for the detection of Taxol| requiring less t h a n five minutes per sample have been developed [104]. Extensive surveys on taxoid distribution in yew extracts must have been carried out by companies that commercialize taxoids (Indena, Hauser), but these results have not been published for proprietary reasons. This, and the difficulty of obtaining many taxoid standards, is responsible for the paucity of studies devoted to the study of taxoids other t h a n taxols and taxol-equivalent compounds (baccatin III and its 10-deacetyl derivative) in yew extracts. Methods for the q u a n t i t a t i v e analysis of taxoids in plant m a t e r i a l s using mass spectroscopy (FAB, MALD, spray ionization and MS/MS techniques) [104], or immunochemical methods [105] have also been reported. The major problem plaguing the isolation of taxoids is the formation of artifacts, since several reactions of taxanes are possible under conditions similar to those encountered during the preparation of plant extracts and their separation. Acyl .migrat..i.on: Reversible acyl migrations involving the hydroxyl groups at C-7, C-9 and C-10 have been documented in CDC13 solution (Figure 22) [62-64]. In spite of the topological similarity, no reaction of this type was observed in partially deacylated baccatin VI and abeobaccatin VI derivatives [39], suggesting t h a t the a c t i v a t i o n energy for the acyl m i g r a t i o n d e p e n d s on the

41 functionalization pattern of the taxoidic core. However, it should be noted that CDC13 used for NMR spectroscopy does not contain soluble stabilizers, and formation of HC1 may have been responsible for the observed migrations. Indeed, solutions of monoacylated cinnamoyl taxicins in stabilized chloroform are stable for weeks, and this solvent could even be used for their separation [62].

AcO

OH

O ==~

OH

.~ n ,0' ' NMe2 O / u0 ' v ~ ~,;OCin I1 Ph 16 . -NHMe2 ()H ref.

-

HO

AcO

(3H

"-

3.1.3

HO

8.1.1

~OAc

O

~l . HO OH 8.1.2

OAc

HO Q

NMe2

./91,,,O,..Jl,,.~ph

_NHMe20

OH

"OCin

8.1.3

Figure 22: Isomerization/degradation reactions observed in CDC13solution Loss of Dimethylamine from Winterstein Acid Esters: It was recognized very early that taxine is unstable and loses activity upon storage, presumably as a consequence of its loss of dimethylamine and formation of the corresponding cinnamates (Figure 22). This was observed, along with O(10)->O(9) acyl migration, with CDC13 solutions of taxine B [16]. The derivation of cinnamic esters of taxoids from their corresponding Winterstein acid esters seems highly plausible, although it is not clear to what extent this derivation may be biogenetic or simply chemical. At room temperature, prolonged (months) contact with chloroform and silica gel is required for elimination [43], and it may well be that the reaction is also under enzymatic control.

42 For unobvious reasons, Winterstein acid esters with an acetoxyl at C-13 are more stable than those with a keto group at this position, like taxine B [106]. C-7 Epimerization: In taxoids bearing a keto group at C-9, the C-7 ~-hydroxyl can reversibly epimerize via a retro-aldol/aldol mechanism (Figure 23). The reaction has been reported to occur under mildly basic or acidic conditions (basic alumina or silica gel) [57] and even on concentration of HPLC chromatographic fractions [107]. In the a-epimer, the hydroxyl group is pseudoaxial, and this arrangement is stabilized by the formation of a hydrogen bond with the C-4 acetate carbonyl group [108].

H

~HO,,,

Ho

_ HO

-Bz O 7.2.9

,__Y,.;o OAc

L/H_-Y".,J ~ OAc 8.1.6

HO"- r ' HO

O OAc OBz 8.1.7

Figure 23: Retroaldol equilibration of baccatin derivatives [57] Ring A Contraction: In acidic medium, C-1 hydroxylated taxanes can rearrange to the corresponding ll(15->l)abeo derivatives (Figure 16). A mixture of C-15 hydroxy and A15,16-derivatives is formed, in a ratio depending on the reaction conditions [48]. However, all ll(15->l)abeotaxanes isolated to date have a C-15 oxygen function, and their dehydration products have never been detected in yew extracts [39]. Furthermore, the acylation pattern of l l(15->l)abeotaxanes and taxanes is generally different [59]. Thus, baccatin VI is widespread, but its corresponding ll(15->l)abeotaxane (abeobaccatin VI) is unknown as a natural product, since only deacetylated [59] or isomeric (O(10) vs. 0(2) benzoate [39]) derivatives have been isolated. It is therefore likely that 11(15-> 1)abeotaxanes are natural constituents of the yew. More details about this rearrangement are discussed in chapter 5. Photocyclization: Taxinine-type compounds are converted by UV light to their corresponding 3,11-cyclotaxanes (Figure 18) [46]. The reaction is often quantitative, and is accompanied by the E->Z photoisomerization of the cinnamoyl residues [55], and by the reductive deamination of Winterstein acid

43 residues to the corresponding ~-phenylpropionates [100]. The fact that no taxoid with a Z-cinnamoyl residue has ever been isolated points to an enzymatic origin for 3,11-cyclotaxanes. Further details on the photocyclization reaction of taxanes can be found in chapter 5. Reactions During Plant Drying and Storage:

Taxoid fractions obtained from

fresh and dried plant samples can show remarkable differences [16], and the Taxol | content of the needles decreases substantially during drying and storage [109]. The nature of the reactions responsible for this is presently unknown. 1.9. BIOLOGICAL A C T M T Y Interest in the biological activity of taxoids has centered around the tubulin-mediated antitumor activity of Taxol | (see chapter 7 for details). However, cytotoxicity has been reported also for taxoids structurally unrelated to taxols, and yew extracts display various activities, some of which may be due to taxoids and may be of relevance for medicine or toxicology. 1.9.1. Non Tubulin-Mediated Toxicity Taxol | was initially isolated because of his remarkable cytotoxicity to KB cells [20], but yew extracts also contain less cytotoxic compounds, both of taxoidic and non-taxoidic structure (lignans). In particular, some non-alkaloidal taxoids can inhibit DNA replication and protein synthesis in tumor cells [110]. The mechanism for this activity is presently unknown, and only a limited number of compounds have been tested in these systems. The most active one was a A4(2~ [110], but the significance of this finding is difficult to assess without additional information. 1.9.2. Cardiotoxicity The yew has a remarkable history as a poison [111], and the plant is still relevant today in human and veterinarian toxicology. The toxicity of the yew has been ascribed to taxine, whose administration can reproduce the cardiac and respiratory disturbances typical of yew poisoning [112]. Taxine is a heart poison that seems to act on ion channels, blocking sodium and calcium currents [113]. The mechanism of this activity is not known at the molecular level. The study of taxine has been hampered by the instability and the complexity of this alkaloidal mixture. Its LD50 is known only

44 approximately, since no established criteria of purity exist, and the composition depends on the method of preparation and the yew species from which it derives [112]. F u r t h e r m o r e , m a n y studies were done on soluble taxine salts (hydrochloride, sulfate), whose purity and stability may have been different from those of crude taxine. Only one study has been carried out on purified constituents (taxine A and B), showing that taxine B is toxic (LD50 4.5 mg/Kg in mice by intravenous administration), whereas comparable amounts of taxine A are not [114]. Taxine B is the major (20-30%) component of taxine, and should also represent its poisonous constituent. Other components of taxine have been reported recently, but they were isolated in insufficient amount for a detailed in vivo investigation of their cardiotoxicity [106]. Taxol| shows only modest cardiotoxicity [115], but the presence of alkaloidal impurities, and in particular of taxine B, may be clinically relevant, since Taxol | is administered at relatively high doses, and taxine B has been reported to give cardiac anomalies at doses as low as 0.5 mg/Kg [114]. No analytical method for the characterization of taxine has been reported to date. 1.9.3. Other Activities Yew extracts have hormonal activity in insects and mammals. Indeed, the needles of the European yew are a good source of the moulting hormone ecdysones [21]. Block of ovulation and anti-implantation activity have been reported in rabbits and albino rats [116, 117]. Several studies have evidenced a sedative activity in yew extracts [118], and their cosmetic use is also discussed in the patent literature [119]. Apart from the ecdysones, the compounds responsible for these activities have not been identified, but the sedative activity may be due to dimeric flavonoids and not to taxoids. Amentoflavone binds to the BDZ site of the GABA receptor [120], and this compound occurs, along with several other derivatives, in the yew [121]. Among the phenolic constituents of the yew, the phenylbutanoid glucoside rhododendrin also shows hepatoprotective activity [122]. The severe allergic reactions reported upon administration of Taxol| have been ascribed to the vehicle Cremophore EL. However, an almost fatal allergic reaction was reported in a child who had chewed a few needles of yew [123], suggesting that the plant may contain powerful allergens. Contact dermatitis from yew wood has also been reported [124].

45 1.10. TAXONOMICAL R E L E V A N C E A N D RAISON D'ETRE

The distribution of taxoids within the Taxaceae family has taxonomical value, allowing a clearcut distinction between morphologically related genera. Taxoids are so far the only non-steroidal (triterpene) isoprenoids isolated from the genera Taxus and Austrotaxus, and this sets them apart from the genus Torreya, which contains labdane diterpenoids and furanosesquiterpenoids. Curiously, the seeds of some torreyas are edible [125], provided that the aril is removed, whereas the fleshy aril surrounding the seed is the only part of the yew that lacks taxoids and can be eaten. Comparison of the taxoid pattern within the genus Taxus might also help in unraveling the taxonomical relationship between the various species, varieties and cultivars of the yew. However, as usual with secondary metabolites, the production of taxoids is under a complex control mechanism, including genetic and epigenetic factors (environment, season, cultivation practices, plant part). As a result, clearcut conclusions are difficult to draw. The longevity of the yew tree, the presence of sexually distinct plants and the ease of hybridization are additional complications. The greater infraspecific variation of the taxoid pattern regards the basic alkaloids. Indeed, it was recognized very early that taxine from various yew species is different, and that some yews (e.g.T. brevifolia Nutt.) contain only small amounts of it [126]. Hegnauer suggested that the smallest amounts of alkaloids are contained in the species t h a t lacks cyanogenetic glycosides [127], but this correlation has not been investigated further. Some general trends are also emerging in relation to other types of taxoids. Thus, in the European yew, taxicin-type compounds are generally hydroxylated at C-l, whereas in the Japanese yew compounds of this type are generally unfunctionalized at C-1. Furthermore, C-14 hydroxylation seems typical of taxoids from the Himalayan and the Chinese yew, but 2(3->20) abeotaxanes have so far been detected only in the European yew. The production and storage of secondary metabolites is probably the most i m p o r t a n t s t r a t e g y adopted by plants to defend t h e m s e l v e s a g a i n s t microorganisms, herbivores and other plants. Very little is known about the ecophysiological role of taxoids, but the observation that the yew is poisonous to most vertebrates and insects suggests that its secondary metabolites have evolved as part of a defense strategy.

46 Furthermore, the various yew species produce not just a single taxoid, but complex mixtures of a limited amount of major metabolites and a larger number of minor derivatives, as expected for compounds designed to confer a broadspectrum resistance against predators. Thus, the poisonous properties of taxine and the anticancer activity of Taxol | might not be the result of an accidental affinity of these compounds for receptor sites on animal proteins, but may be part of a more subtle strategy whose exact details are elusive. The production of secondary metabolites is believed to develop during evolution by natural selection, and the long evolutionary life of the yew tree indicates that associations seen today may not be those in which the chemical interaction originally evolved. Taxoids may well be important and multitask "fitness factors" for the yew tree, but the plant must also have evolved strategies to avoid autotoxicity, since Taxol| interacts with a cellular structure (the microtubule) present in all eucaryotic cells, and plant tubulin is sensitive to this drug. As the age of molecular medicine dawns, plants represent a still largely untapped source of chemodiversity. The yew tree has provided us with Taxol | but other drugs or pharmaceutical leads may emerge in the future among the unique and fascinating secondary metabolites of this plant.

Acknowledgements I am very grateful to my wife Enrica for her patience and understanding during the time I dedicated to this chapter and not to her. I t h a n k Prof. P. Gariboldi (University of Camerino), Dr. B. Gabetta and Dr. E. Bombardelli (Indena, Milano) for their useful suggestions and comments. I am grateful to all members of my research group for showing me every day that scientists are happy people, and that the essence of science is independent thinking and hard work, not equipment.

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The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B.V. All rights reserved

55

2 THE STRUCTURAL ELUCIDATION OF T A X O I D S Giovanni Appendino Dipartimento di Scienza e Tecnologia del Farmaco, Via di Giuria 9, 10125 Torino, Italy

2.1. I N T R O D U C T I O N

The taxane skeleton can r e a r r a n g e under a variety of experimental conditions (acids, bases, light), and the early work based on degradative chemistry gave no indication on its actual constitution. Puzzling [YV features together with the difficulties in assessing purity concurred to delay the structure elucidation of taxoids until the advent of NMR spectroscopy. The application of powerful two-dimensional techniques (COSY, NOESY, HMBC etc.) and the availability of a wealth of published information have made the structure elucidation of new taxoids almost trivial, provided the compound is available in sufficient quantity and purity for the application of these techniques. 2.2. M A S S S P E C T R O S C O P Y

Mass spectroscopy (MS) of taxoids has been used to obtain information on the molecular weight and to identify the acyl groups bound to the diterpenoid

55 core. Soft ionization techniques (FAB, DCI, TSP) have been used [1]. Thus, the chemical ionization spectra (NH3) of taxols shows a parent ion (M + NH4)+, and a prominent peak at m/z 586 [M + NH4 - side chainH] +, resulting from the loss of the aminoacid side chain [2]. The constitution of the latter can be elucidated by the analysis of the fragmentation pattern [3], but MS had so far played only a marginal role in the analysis of the diterpenoid core of taxoids. The most prominent peaks in the spectrum of the semisynthetic taxane tetraacetyltaxinol (Figure 1) have been tentatively identified [4], but study of closely related taxanes failed to confirm the generalization of the proposed fragmentation pattern.

0

AcO

OAc

H

-

_

"OAt_,

OAc 2.1.1 Tetraacetyltaxinol Figure 1

2.3. UV AND CD (ORD) S P E C T R O S C O P Y

The most important chromophore in the diterpenoid moiety of taxoids is the enone system found in compounds of the 10- and 13-oxo-A 11 type. In taxanes, these systems can be distinguished on the basis of UV data (Figure 2). The 13oxo-A 11 system shows an anomalous n->~* absorption band at 275-285 nm (a ca. 5,000-6,000) [5], rather different from the value calculated from Woodward's rules (Xmax ca. 255 nm). Comparison with related compounds highlights the role of ring strain and of the g e m - d i m e t h y l group at C-15 bathochromic shift (ca. 7 nm) by the C-1 hydroxyl groups was The 10-oxo-A 11 system, on the other hand, does not properties, since its maximum (Xmax 240-250 nm, a c a .

for this effect. A also observed [5]. show unusual UV 3,000) is in good

agreement with the value calculated from Woodward's rules [6]. In compounds of this type, the chair-boat conformation of the cyclooctane B ring allows little overlap between the bridgehead double bond and the C-10 carbonyl, since the

57 corresponding u orbitals are almost orthogonal to each other [7]. Thus, conjugation is attained only if the conformation of ring B is significantly changed in the fragment C-8/C-11. Taxanes and 2(3->20)abeotaxanes of the 13-hydroxy (acyloxy)- A l l type show an anomalous n->u* absorption (~ ca. 10,000) at 210-230 nm (expected value ca. 190-200) [5], showing t h a t the bridgehead double bond has an anomalous electronic distribution, possibly r e l a t e d to the gem-dimethyl substitution at C-15 and to its rigid trans-cyclodecene (dodecene) nature. This anomalous absorption is responsible for an intense and positive Cotton effect, amenable to octant analysis [8]. The results of this configurational assignment are supported by comparison with the CD spectra of chiral olefins (transcyclodecene and trans-cyclooctene) and with the results obtained on taxoids with other techniques (NMR, X-rays, Horeau methods). The A4,2~ is responsible for a weak negative Cotton effect, which is of no practical use, being overshadowed by the strong positive Cotton effect ot the All system [8]. The C-9 keto group of baccatin III gives rise to a Cotton effect at 304 nm, whose observed negative sign is predicted by the carbonyl octant rule [8]. The benzoate sector rule and the dibenzoate chirality rules were applied to taxanes of the A4(20)-5a benzoyloxy type and 0(9), O(10)-dibenzoyl type, respectively [9, 10], and these results were used for the structure elucidation of taxinine [9].

HO

OH

H

OH 3.1.1 4(20)-Dihydro-5-deoxytaxicin I )~m~x = 283 nm (e = 5,700) [5]

"

H

3.1.2 Taxuyunnanine D ~ m a x -- 248 nm (e = 2,275) [6b]

Figure 2: UV differences between 13-oxo and ll-oxo All-enones The absolute configuration at C-1 of taxoids is opposite to the one found in their alleged biogenetic precursors (verticilloids) isolated so far from plants (see section 1.6.1). However, all evidence from CD studies (olefin and carbonyl octant

58 analysis, benzoate sector and dibenzoate chirality rules) fully confirms such absolute stereochemistry. 2.4. IR S P E C T R O S C O P Y

The IR spectra of taxoids do not show any unusual absorption band. Owing to extensive inter- and intramolecular hydrogen bonding, the hydroxyl and carbonyl regions generally show broad bands, especially in the solid state. 2.5. N M R S P E C T R O S C O P Y

Despite the importance of NMR spectroscopy for the structure elucidation of taxoids and the impressive technical advances made in the last decade, no recent review on the NMR features of taxoids is available. In 1963, Nakanishi discussed in detail the 1H-NMR spectrum of taxinine and some of its derivatives [11]. Although based on a wrong stereostructure (C3~H), most of these assignments are correct. In 1967, Lythgoe made extensive use of 1H-NMR data to correctly assign the stereochemistry of taxicin I and II [12], confirming most of the assignments and observations made by Nakanishi [11]. In 1980, Miller reviewed the available 1H-NMR data on taxoids [13], and in 1983 Rojas discussed the general features of the 13C-NMR spectra of several structural types of taxanes [14, 15]. In the last few years, many papers have discussed in detail the NMR spectra of Taxol| and related compounds [16-23]. All resonances were assigned, and solvent-related conformational changes at the side chain were noted [20-23]. Until the advent of two-dimensional techniques, most assignments in the 13C-NMR spectra were carried out by analogy or left uncertain. Reference will be made here mainly to data derived from fully assigned spectra. In order to better appreciate the stereochemical information derived from the NMR spectra, a section on the general conformational features of taxoids is presented next, before the full discussion of the NMR data. 2.5.1. Conformational Aspects of the Taxoidic Skeleta The shape of the diterpenoid core of taxoids is dictated mainly by the conformation of ring B. The latter is a cyclooctane in taxanes, a cycloheptane in

ll(15->l)abeotaxane, a cyclodecane in 2(3->20)abeotaxanes, and a bicyclo[3.3.0]

59 octane in 3,11-cyclotaxanes (see Figure 7 in chapter 1). More than one conformation is possible for ring B, except in 3,11-cyclotaxanes, where the presence of a t r a n s a n n u l a r bond prevents conformational mobility. Several conformations are possible for cycloheptane, cyclooctane and cyclodecane, but the junction with ring A and ring C makes only a few of them possible for ring B of taxoids. The major variation occurs along the C-9/C-10 bond, the only one between carbons not shared with ring A or ring C. Among the staggered conformations around C-9/C-10, two conformers are possible, and they are characterized by a syn or an anti relationship between the oxygen functions at C9 and C-10 (Figure 3, A and B). An eclipsed conformation with the oxygen syn (C) is also possible. The staggered, oxygen syn conformation around C-9/C-10 (A) is typical of taxanes, with the notable exception of taxagifine and its derivatives, which adopt the eclipsed conformation C [24]. Both staggered conformations A and B have been found in ll(15->l)abeotaxanes, whereas 2(3->20)abeotaxanes adopt the staggered/oxygen-syn conformation (A) typical of taxanes. All ananchomeric (i.e. conformationally locked) taxoids with the C-9/C-10 staggered, oxygen-anti conformation (B) have an extra oxygen bridge, between C-10 and C15 or C-13 and C-15 (Figure 4) [25, 26].

H9

'R

OR

R

H9Ho

Hlo

~(c-9/c4o)=6oo A

OR

Hlo ~(c-9/c-1 o)=-oo o B

~(c-9/c-~o)=12o o C

Figure 3: Newman projections along the C-9/C-10 bond in the three major conformations of taxoids.

No universally accepted denomination for the topological forms of medium-sized rings exists, and pictorial stereoviews of taxoids are difficult to draw and to interpret. The arrangements A and B correspond to a chair-boat and a chair-chair conformation for ring B in taxoids, and to a twist-boat and a twistchair conformation for ring B in ll(15->l)abeotaxanes. The Bucourt notation [27] makes reference to the succession of the endocyclic torsion angles, and seems more meaningful in this context [26].

60 Ring C is relatively rigid in oxetane-type taxoids, and adopts a sofa conformation with C-7 as the flap (Figure 5, A). The c h a i r and the boat c o n f o r m a t i o n are i n s t e a d

possible in A4(20)-taxoids, d e p e n d i n g on the

conformation of ring B or the presence of an additional C-3/C-11 bond (Figure 5, B and C) [28, 29]. Ring A adopts a distorted boat conformation in taxanes and

2(3->20)abeotaxanes, and an envelope conformation in ll(15->l)abeotaxanes.

~Ac

10 ~ - - "/'~O

AcO

_ ~ 0 OBz OAc

OAc

0

0 OBz

5.1.1

5.1.2

Figure 4: Ananchomeric taxoids with a staggered, oxygen-anticonformation around the C-9/C-10 bond. 2.5.2. Proton and Carbon NMR Assignments of the Diterpenoid Core For the sake of clarity and practicality, the NMR features of the proton(s) and carbon referring to the same position of the taxoidic skeleton will be discussed under the same heading. Throughout this section, bold numbers refer to compounds whose spectroscopic data are presented in section 2.5.4.

H H

O 02 Me

Ac

02 H713

H5

HFo~ A

02

Me H71~ ~

OR

HFa

//

B

Me H7~

~m

H7c~

C

Figure 5: Conformation of ring C of taxoids C-l:

D e p e n d i n g on the s t r u c t u r a l type of taxoids, C-1 can be a

h y d r o x y l a t e d q u a t e r n a r y carbon, an aliphatic m e t h i n e , or an aliphatic quaternary carbon, resulting in diagnostic resonances at 5 75-80, 40-50 or 60-70,

61 respectively. C-1 is an aliphatic quaternary carbon in 11(15->1)abeotaxanes (13 and 14), and its downfield resonance (5 60-70) is diagnostic for this type of taxoids [29b]. This chemical shift, unusual for a non-oxygenated carbon, might be the result of linear strain, since in these compounds some of the carboncarbon bonds centered at C-1 are significantly longer than normal [29b]. Owing to slow conformational equilibration, the signal of C-1 in ll(15->l)abeotaxanes is sometimes very broad or even undetectable at room temperature [26]. The C-1 hydroxyl is not acylated u n d e r s t a n d a r d or even forcing conditions, but acylation can be attained in situ (i.e. in the NMR probe) with the powerful acylating reagent trichloroacetyl isocyanate (TAI) [29b]. Several hours are required for the reaction, which causes a dramatic downfield shift on H-1413 (>1 ppm), and smaller shifts on the gem-dimethyls at C-15, H-14a and H-2 (Figure 6). This p a t t e r n of A5 allows a straightforward distinction between C-1 hydroxylated taxanes and C-15 hydroxylated 11(15->1)abeotaxanes. These two classes of compounds show similar 1H NMR spectra, and the rigorous establishment of the carbon connectivity via long-range 1H-13C-correlations can be difficult because of the fluxional behavior of 11(15->1)abeotaxanes [29]. However, acylation of the tertiary hydroxyl of these compounds has only a small effect on the resonance of H-14[~, but causes a large paramagnetic shift on the geminal methyls, a diamagnetic shift on H-913, and has a negligible effect on H-2 (Figure 6).

AcO

OAc

~ AcO .... +1.44

H

+0.29 H

L ~

-

:

c

L

~

OH ,~'~ n OAc BzO H +0.38

5.2.1 Baccatin VI

BzO

H -0.20 OAc " OAc

AcO,,. 0

H ~ ~ ' ~ ~ ~ / _ H~ OAc OAc +o.46H0~ +0.58

O

+0.14

5.2.2 Isoabeobaccatin VI

Figure 6: A (TAI) observed upon acylation of the tertiary hydroxyls of baccatin VI and isoabeobaccatin VI [29b].

62 H-1 is almost orthogonal to H-14a, and J l,14a is ca. 0 Hz. Coupling is instead observed with H-14[~ (ca. 7 Hz) and H-2~ (ca. 2 Hz). C-2: In C-2 oxygenated taxoids, the assignment of H-2 is straightforward, owing to a peculiar multiplicity pattern (J1,2 c a . 2 Hz; J2,3 ca. 7 Hz), easily distinguished from that of the other oxymethines. In taxoids of the 4,20-epoxide type, J2,3 is smaller (3-4 Hz) (1), but the splitting pattern of H-2 is still diagnostic, as is in taxagifine-type taxoids, where J2,3 is c a . 10 Hz (15). In taxanes having a C-2, C-9, C-10, C-13 hydroxylation (acyloxylation) pattern, H-2 is generally the most shielded of these oxymethines, regardless of the acyl residue at C-2 (5.2.1, Figure 7). In the corresponding l l ( 1 5 - > l ) a b e o t a x a n e s , H-2 is instead the most deshielded of these signals (5.2.3, Figure 7) [26, 30]. This observation has diagnostic value, although exceptions exist in taxagifine-type taxoids (15).

AcO

OAc

AcO ~,

OAc ~ OAc

6.31/ \lib ,/~.~ 6.13'~,,~ AcO'"

4

HO

% ,,

O

AcO....~

=.

6Bz 5.2.1 Baccatin VI [26]

Hd \

O ~6.36 OAc OBz

5.2.3 Abeobaccatin VI [30]

Figure 7: Ring A/B oxymethine resonances in baccatin VI and abeobaccatin VI

In C-2, C-10 di-oxygenated taxoids, the a s s i g n m e n t of C-2 is not straightforward, since its chemical shift (ca. 70-75 ppm, depending on its acylation state and that of C-l) is similar to that of C-10. However, since the signals of H-2 and H-10 are very well separated, two-dimensional techniques or selective decouplings allow an unambiguous assignment. When C-2 is a methylene, the C-1 hydroxyl has a strong deshielding effect (ca. 10 ppm, cf 9 vs. 11 ) on its resonance. Only a moderate effect ( ca. 2 ppm, cf 6 vs. 7) is instead observed when C-2 is an oxymethine, presumably because of a shielding 7-gauche interaction between the vicinal oxygens.

63 C-3: In 2(3->20)abeotaxanes, C-3 is a methylene, and the appearance of an isolated AB system with lines at 8 2.60-2.80 and 1.60-1.80 is a diagnostic

feature of taxine A derivatives (see 17). The corresponding 13C resonance is found at 8 ca. 35, close to the C-6 triplet. In taxanes and 11(15->1)abeotaxanes, C-3 is an aliphatic methine and H-3 is a doublet, resonating at relatively low field in compounds of the A4(20)-type (8 c a . 3.5) and 9-oxo-4a-acyloxy-5,20oxetane type (baccatin III/V derivatives) (8 ca. 3.5-4.0). In oxetane-type taxoids lacking a 9-oxo group (baccatin IV,VI,VII derivatives) or a 4-acetyl (see 4), H-3 resonates at 8 2.5-3.2, and the same chemical shift range is observed in taxoids of the 4(20)-epoxide type (see 1). The resonance of C-3 is found at 8 35-45 in compounds of the A4(20)-type, and at 8 45-50 in compounds of the oxetane type. An unusual value for C-3 (8 ca. 59 !) was found in A4(20)-taxoids bearing an amethylbutyrate ester at C-2 [15]. This remarkable effect was rationalized in terms of a conformational change in ring B that moves C-3 away from the 11-12 double bond [15]. However, the oxygenation pattern of these compounds should be confirmed by modern NMR techniques, since several other resonances are quite unusual for the proposed structure. In 3,11-cyclotaxanes (16), C-3 is a quaternary carbon, resonating at relatively low field (8 ca. 60) for a nonoxygenated tetrahedral carbon. As with C-1 in 11(15->1)abeotaxanes, linear strain may be responsible for this unusual chemical shii~ value. C-4: In all but one of the naturally occurring taxoids, C-4 bears no hydrogens. In compounds of the A4(20)-type, C-4 resonates at 8 140-150,

depending on the oxygenation state of C-2 and C-5. The corresponding resonance in compounds of the oxetane- and epoxide-type are 8 ca. 80 and 60 respectively. Hydrolysis of the 4-acetoxy group causes an expected upfield shift on C-4 (ca. 6 ppm, cf 4 and 5). C-5: In all natural taxoids C-5 is oxygenated, and the splitting pattern of

H-5 can give important information on the stereochemistry at this carbon as well as on the conformation of ring C. In taxoids of the A4(20)-type, H-5 is always ~, and its splitting pattern depends on the conformation of ring C. In A4(20)_ taxenes, ring C has a chair conformation (B, Figure 5), and H-5 is equatorial (J5,6a=J5,6~<3 Hz) [11]. In 11(15->1)abeotaxanes both the chair (B) and the boat conformation (C) have been detected [28, 29]. In the latter, Js,6a=J5,6~=ca. 5 Hz. In A4(20)-3,11-cyclotaxenes (see 16), ring C has a twist-boat conformation, and J5,6a=J5,6~=ca. 9Hz [31]. In taxoids of the 4,20 epoxide type, H-5 resonates at unusually high fields, yielding misleading information on the acylation state of

54 the C-5 hydroxyl (see 1). In oxetane-type taxoids, H-5 is a and ring C has a sofa conformation (A, Figure 5). In these compounds, H-5 resonates as a doublet of doublets (J5,6a= ca. 9 Hz, J5,6~=ca. 2 Hz). Opening of the oxetane via anchimeric assistance from the 4-acetyl inverts the stereochemistry at C-5, and H-5 turns into a narrow triplet, with J5,6a and J5,6~<3 Hz [32], a value t h a t is diagnostic of a chair conformation for ring C. In oxetane-type taxoids, C-5 resonates at lower field t h a n in other structural types of taxoids (5 83-85 vs. 70-78).

C - 6 : C - 6 is a methylene in all natural taxoids, and resonates at 5 30-35. The signals of H-6a and H-6~ are generally well differentiated, especially when C-7 is oxygenated. H-6a is more deshielded t h a n H-6~, but the chemical shift of the former can be dramatically affected by the presence of an aminoacidic side chain at C-5. Thus, H-6a resonates at negative values of 5 (-0.21 I) in taxine A (17), and at values o f ca. 1 ppm in other O(5)-taxane esters of phenylisoserine [33, 34]. The remarkable chemical shift of H-6a in taxine A is due to the fact that this hydrogen lies in the shielding cone of the aromatic ring of the side chain (see section 2.5.3) [35]. Thus, in peracetyl taxine A, H-6a resonates at 5 1.67, a value very close to the one observed in deaminoacyl taxine A [35]. This, and the decrease of J2',3' from 10 to 4.5 Hz, show t h a t the conformation of the side chain has changed, resulting in an a r r a n g e m e n t with H-2' and H-3' synclinal and the phenyl away from the concave face of the taxoid moiety. C - 7 : C - 7 can be an aliphatic methylene or an oxymethine, resonating at 5 25-30 and 68-75, respectively. In baccatin III derivatives, the 7-hydroxyl can epimerize via a retroaldol reaction (see section 1.8). This causes remarkable changes in the chemical shift and the multiplicity of H-7, which t u r n s from a downfield doublet of doublets (5 c a . 4.40 in CDC13) into a narrower and more upfield signal (5 c a .

3.70 in CDC13, cf. 1 8 and 19). This upfield shift is

presumably due to removal of the deshielding effect of the acetate carbonyl at C4. Epimerization from the n a t u r a l (7~-OH) to the u n n a t u r a l (7r affects the chemical shift of C-19, C-20 and all the ring C carbons. Especially diagnostic are the shift of C-19 (downfield ca. 9 ppm) and C-3 (upfield c a 5 ppm; cf. 18 and 19). "Long-range" effects on ring A and B carbons are also observed (vide infra).

C-8: In all natural taxoids, C-8 is an aliphatic quaternary carbon, and its chemical shift is mainly affected by the oxidation state at C-9. When C-9 bears a free or an esterified hydroxyl, C-8 resonates around 5 45, but the presence of a keto group at C-9 shifts this resonance downfield to 5 55-60.

65 C - 9 : C - 9 can be an oxymethine, a carbonyl or an aliphatic methylene, resonating at 8 75-82, ca. 210 and ca. 58, respectively. When C-9 and C-10 are both hydroxylated or acyloxylated, C-9 resonates at lower field than C-10 (see 411), but in O(9), O(10)-monoesters the most downfield carbon is the one bearing the ester group (see 12). In 3,11-cyclotaxanes both C-9 and C-10 resonate at 8>80. In 9,10-dihydroxylated taxanes, H-9 resonates at higher field than H-10 (8 ca. 4.3 and 5.0, respectively), and a similar trend is observed in 11(15>l)abeotaxanes

(14). This observation is useful in establishing the acylation

pattern of O(9),O(10)-monoesters [36]. In 9,10-dihydroxylated taxoids, the vicinal hydroxyls are always t r a n s (~ and ~, respectively), but the value of J9,1o depends on the conformation of ring B, which dictates the sign of the torsion angle C-8/C9/C-10/C-11. When this fragment is staggered and the angle is positive, H-9 and H-10 are t r a n s - d i a x i a l , and J9,1o is around 10 Hz (Figure 3, A). Rotation around the C-9, C-10 bond and the attainment of a negative value for the endocyclic torsion angle around these carbons makes H-9 and H-10 t r a n s - d i e q u a t o r i a l , and therefore J9,10 decreases to ca. 4 Hz (Figure 3, B), as observed in some 11(15->1) abeotaxanes

[28, 29a]. Small values of J9,10 (ca. 3 Hz) are also typical of

taxagifine-type compounds (oxygen bridge between C-17 and C-12, see 15), where the fragment C-8/C-9/C-10/C-11 is in an eclipsed conformation (~ ca. 120~ and H-9 and H-10 are anticlinal (Figure 3, C). C-10: In all natural taxoids C-10 is an oxymethine, whose chemical shift (5 65-75) is little affected by acylation or by the presence of a keto group at C-9. A change in hybridization of C-11, as in 3,11-cyclotaxanes (see 16), shifts C-10 downfield (8 80-82). Epimerization at C-7 of baccatin III to baccatin V causes downfield shifts at H-10 (ca. 0.5 ppm) and C-10 (ca. 3 ppm) (cf. 18 and 19) and a similar, but opposite, effect on C-12. The long-range effects are difficult to rationalize, but probably reflect subtle conformational differences and/or hydrogen bonding patterns. An even more marked downfield shift (AS ca. 5 ppm) on the chemical shift of C-10 was observed between 10-oxo-derivatives of the baccatin III and baccatin V series, suggesting better conjugation between the C10 carbonyl and the 11-12 double bond in baccatin V derivatives [37]. C - 1 1 : C - 1 1 is a tertiary alcohol in taxagifine derivatives (8 c a . 80), a quaternary aliphatic carbon in 3,11-cyclotaxanes (8 ca. 55) and an olefinic carbon in all the other taxoids, resonating at 8 130-135 (150-155 when a 13-oxo group is present). In 1 3 - h y d r o x y - 1 1 ( 1 5 - > 1 ) a b e o t a x a n e s , C-11 is slightly more deshielded than in the corresponding taxanes (AS ca. 2 ppm, cf. 5 and 14). The chemical shift

55 of C-11 is affected by esterification of the allylic hydroxyls at C-10 and C-13, which causes an upfield shift (ca. 3 and 1 ppm, respectively). C-12: C-12 is an aliphatic methine (5 ca. 50) in 3,11-cyclotaxanes, a quaternary oxygenated carbon in taxagifine derivatives (5 ca. 90) and an olefinic carbon (5 135-140) in all the other structural types oftaxoids. C-13: Like C-9, C-13 can be an oxymethine, a carbonyl, or an aliphatic methylene. In 13-hydroxy (acyloxy) taxoids, the chemical shift of C-13 depends on the structural type, and in l l ( 1 5 - > l ) a b e o t a x a n e s the resonance for this carbon is found at lower field than in taxanes (5 77-80 vs. 67-72, cf. 14 and 5). The carbonyl of All-13-oxotaxanes resonates around 199 ppm, as expected for a 2-cyclohexenone carbonyl, but the chemical shift of the carbonyl of 13-oxo-3,11cyclotaxanes (5 214-216, see 16) is rather downfield for a cyclohexanone carbonyl. In natural taxoids, H-13 is almost always ~, and its signal is rather broad, due to allylic coupling with H-18 (J ca.1 Hz). In taxanes, J13,14~ is ca. 9 Hz, whereas the value of J13,14a varies considerably (3-9 Hz), depending on the conformation of ring A. In l l ( 1 5 - > l ) a b e o t a x a n e s , J13,14~ is generally smaller (ca. 7 Hz) than in the corresponding taxanes [26]. Only one example of natural taxoid with a 13~oxygen function has been reported to date. In this compound, values of ca. 0 and 1.8 Hz were observed for J13,14a and J13,14~, respectively [26]. C-14: C-14 can be an aliphatic methylene (5 35-45, depending on the functionalization of its a-carbons)or an oxymethine (5 ca. 70). The geminal H-14 protons are generally well separated, and H-14a, pointing toward the concave face of the molecule, is the most shielded. A C-13 oxo group deshields H-14a more t h a n H-14~, and consequently, in some taxicin I derivatives, the resonances of H-14a and H-!4~ are close to each other. H-14a is also deshielded by an oxygen function at C-1 and, in oxetane-type taxoids, H-14a,~ are sometimes almost isochronous, or their chemical shift values can sometimes be reversed (see 4). C-15:C-15 is an aliphatic quaternary carbon (5 37-43, depending on the presence or absence of a hydroxyl group at C-l) in all taxoids, except in 11(15> l ) a b e o t a x a n e s , where this carbon is oxygenated (5 75-79). In these compounds, a strong intramolecular hydrogen bond between the hydroxyl at C-15 and the oxygen function at C-10 exists, and the C-15 hydroxyl proton resonates as a singlet, unaffected by dilution, at 5 2.40-2.50 [29b]. In taxagifin-type taxoids, C15 is aliphatic, but the oxygen functions on the a-carbons (C-11 and C-17) shift its resonance downfield to ca. 50 ppm (see 15).

67

C-16, C-17: In taxanes, the proton and carbon signals of the g e m dimethyls are often well differentiated. The C-17 methyl, facing ring A, is generally the most shielded in the 1H NMR spectrum, but an opposite relationship is observed in the 13C NMR spectrum. A5 of up to 0.60 ppm (1H NMR) and 15 ppm (13C NMR) have been reported (see 6 and 7). A keto group at C-9 has a shielding effect on H-16, but has an opposite effect on C-16. This upfield shiit of H-16 is responsible for the small A5 between the geminal methyls in the 1H NMR spectra of baccatin III/V derivatives. Furthermore, the phenylisoserine residue at C-13 has a deshielding effect on H-17, and consequently in taxols H-17 resonates more downfield than H-16, in contrast with what observed in the other types of taxoids (see 18 and 19). The C-1 hydroxyl shifts H-16 upfield and H-17 downfield, and both C-16 and C-17 upfield, as a result of a 7-gauche shielding interaction (see 6 vs. 7 and 9 vs. 11). X-ray analysis has shown that in taxanes the OH/C-I/C-2 bond angle is smaller than the tetrahedral value (100-105 ~ [14]. As a result, the hydroxyl bisecting the C-16/C-15/C-17 angle is closer to C-16 than C-17, and the T-gauche effect is larger for C-16 than C-17. This might also explain why, upon acylation of the C-1 hydroxyl, H-16 suffers a larger downfield shift than H-17 (see Figure 6). The singlets for H-16 and H-17 are generally broader than the H-19 singlet. To explain this observation, Nakanishi suggested the presence of a weak coupling between the geminal methyls [11], but two-dimensional techniques (COSY) show little, if any, coupling between these hydrogens. The singlet for H16 is often taller than the one for H-17, especially when a C-1 hydroxyl is present. It is likely than both gem-dimethyls of taxanes are subject to long-range couplings but, apart from H-l, the protons involved have not yet been identified. C-18: In the 1H NMR spectra of taxoids of the All type, the 18-methyl is the one generally resonating at the lowest field. The presence of a keto group at C-10 or C-13 does not cause an appreciable downfield shift, but acylation of the C-10 hydroxyl causes a downfield shift (0.10-0.15 ppm) that has diagnostic value for the location of the acylation site in 0(9), O(10)-monoesters [36]. In oxetanetype taxoids, the presence of the aminoacid side chain at C-13 causes a marked upfield shift on H-18 (5 = 2.01 in baccatin III, but 1.79 in Taxol| which resonates close to H-19 (see 18 and 19). In compounds of this type, a change in solvent polarity has little effect on the chemical shirt of H-18. This is surprising, since changes are observed within the hydrogens of the side chain, which is spatially close (NOE effect) to H-18.

58 Allylic coupling is observed between the 13~ hydrogen and the 18-methyl (4J13,18 c a . 1.0 Hz). In 3,11-cyclotaxanes (see 16), the 18-methyl resonates as a doublet (J ca. 7 Hz) and this is a diagnostic feature for this class of taxoids, since l l , 1 2 - d i h y d r o t a x a n e s are u n k n o w n as n a t u r a l products. In A ll-taxenes, the resonance of C-18 has a fairly narrow chemical shift range (12-15 ppm). The oxidation state of C-13 has little effect, b u t contraction of ring A from a cyclohexene to a cyclopentene, as in l l ( 1 5 - > l ) a b e o t a x a n e s , shifts the allylic methyl upfield (AS ca. 3 ppm), as shown by comparison of the 13C NMR spectra of 5 and 14. A similar, but not so large, effect is also observed in the 1H NMR spectra (5 =2.01 for H-18 in baccatin VI, 5.2.1, but 1.73 in abeobaccatin VI, 5.2.3, Figure 7)[30]. C-19. The chemical shift of the 19-methyl is affected by oxygen functions at C-7, C-9 and C-20, and its 1H and 13C resonances vary over a wider range than the ones for the other methyl groups. An a or ~ C-7 hydroxyl has little effect on H-19, but a 7~-hydroxyl causes a large upfield T - g a u c h e shii~ (up to 7 ppm) on C-19 (see 18 and 19). This allows a clearcut distinction between baccatin III and baccatin V derivatives, since C-19 resonates at ca. 10 ppm when the 7-OH is ~, and at c a . 15 ppm when the 7-OH is (~ (cf 18 and 19 ). Comparison of the resonance of C-19 for baccatin III (5 9.5) and baccatin VI (5 12.7) shows that, "when compared to a C-9 acetoxyl, the C-9 keto group has a shielding effect on C19. The presence of a 5,20-oxetane ring has little effect on the chemical shii~ of C-19, but causes a marked downfield shift (ca. 0.50 ppm) on H-19. Since also a 9-oxo group has a deshielding effect on H-19, in baccatin III derivatives H-19 is the most deshielded of the three non-allylic methyls (5 ca. 1.60, see 2, 3, 18 and 19). On the contrary, in A4(20)-taxoids, H-19 is generally the most shielded methyl, and resonates at fields as high as 0.80-1 ppm (see 6-12). A 4,20~-epoxide has also a deshielding effect on H-19, but not so large as a 5,20-oxetane ring (cf. 1 and 10). In 19-oxygenated taxoids, 2j 19a,~ is ca. 12 Hz (see 15). The AB system of H19a,~ is easily distinguished from the AB system of the oxymethylene protons at C-17 (taxagifine derivatives) or at C-19 (oxetane-type taxoids). Indeed, the presence of a cyclic structure in these latter compounds decreases the geminal coupling to ca. 8 Hz. C-20:

C-20 is an olefinic m e t h i n e in 2 ( 3 - > 2 0 ) a b e o t a x a n e s ,

and a

functionalized methylene, resonating as an AB system, in all the other structural types of taxoids. The separation of the AB system can yield information on the

59 stereochemistry at C-4 (4,20-epoxides), at C-7 (5,20-oxetanes) or on the functionalization of C-2 and C-5 (A4,20-taxoids). Thus, in the n a t u r a l 4~,20 epoxides, H-20a,~ are well separated (AS > 1 ppm), due to the downfield shift of the hydrogen facing ring B (5 ca. 3.50). In the isomeric and u n n a t u r a l 4a,20 epoxides, H-20cz,~ are instead much closer together (AS <0.40) or even isochronous, since the oxirane hydrogen facing ring B is more shielded (5<3) [34]. In taxols, the signals of the oxetane protons are generally well separated in CDC13 (AS 0.10-0.20 ppm), but epimerization at C-7 brings these resonances much closer to each other (AS 0.00-0.10, cf 18 and 19). Interestingly, in oxetan~ type taxanes H-20cz is generally more deshielded than H-20~, whereas in the corresponding 15(ll->l)abeotaxanes the opposite is true (cf 5 and 14). The geminal olefinic hydrogens of A4(20)-taxoids are subject to anisotropic effects from the oxygen functions and/or the ester groups at C-2 and C-5. In C-2, C-5-dihydroxylated compounds, H-20a,~ are close to each other (see 8), but a large difference (up to 0.60 ppm, see 7) is observed when one or both the hydroxyls are esterified (see 6, 7, 10, 11) and in 5-hydroxylated taxoids. However, H-20a,~ can be close together when the 0(5) ester is Winterstein acid (see 12) [33]. In cases where the geminal olefinic hydrogens have been assigned, the more upfield one is usually the one pointing toward ring B, but an opposite relationship was found for 3,11-cyclotaxanes (e.g. 16 ) and in 2,6-dihydroxylated A4(20)-taxoids. 2.5.3. Conformation and NMR Assignments for the Side Chain. The conformation of aminoalcohols of the type Ar-CHNH2-CHOH-X has been extensively investigated by NMR spectroscopy [38]. The results showed that rotation around the sp3carbons is restricted, and that the conformation is determined not only by purely steric factors, but also by the tendency to form a net of intramolecular hydrogen bonds between the two functional groups and the residue X. In phenylpropanoid esters of taxoids, hydrophobic interactions might also play a role, owing to the presence of lipophilic ester groups (benzoate, nicotinate, hexanoate) bound to the diterpenoid core. However, only certain conformations of the aminoacid side chain allow interactions between the groups on the diterpenoid core and the aminoacid side chain [20-23]. Simple esters of N-acyl phenylisoserine adopt the staggered conformation A (Figure 8) both in solution and in the solid state (see section 2.6). This conformation is also adopted by the aminoacid side chain of Taxol| in CDC13

70 [20-23], but the alternative staggered conformations

B and C have been found in

other taxoid esters (taxine A [35] and N-methyltaxol C [39], respectively).

H.

H

I -.

0/

Bz/,;~

Ph

OR H3'

H3"~

\

OH

NMe2

Ro c-y-H ' Ph

A

B

Ph,~

H3'

Ro c- "r" N ROC" "Me C

Figure 8: Proposed conformation of phenylisoserine derivatives in: A) Taxol (CDC13); B) Taxine; C) N-Methyltaxol C The type of conformation can be assessed by 1H NMR spectroscopy. Large values (9-10 Hz) of J2',3' are typical of conformation B [35], where these protons are a n t i p e r i p l a n a r .

Smaller values

(2-5 Hz) are i n s t e a d

observed in

conformations A and C, where these protons are synclinal [35, 39]. Conformations A and C can be differentiated by inspection of NOE effects: a strong NOE between H-3' and the C-2' hydroxyl proton being typical of conformation C [39]. The conformation adopted depends on several factors, the most important being the n a t u r e of the aminoacid nitrogen and its hydrogen bonding donor/acceptor properties. The aminoacid nitrogen can act as a hydrogen bond acceptor when alkylated (taxine A, B in Figure 8), whereas N-acylation makes it a hydrogen bonding donor (Taxol|

A in Figure 8), and a combination of N-

alkylation and N-acylation renders the aminoacid nitrogen inert toward hydrogen bonding (N-methyltaxol C, C in Figure 8). 0-2' Acylation has little effect on the conformation of N-acyl esters, but it changes the conformation of Nalkyl esters, as shown by the marked decrease (4-5 Hz) in the J2',3' (9.5 Hz in taxine A and 4.5 in its peracetyl derivative) [35]. In O(5)-esters, acylation of the C-2' hydroxyl has also a dramatic effect on the chemical shift of H-6a. These changes can be rationalized in terms of a shift from a conformation of type B, where H-2' and H-3' are antiperiplanatr and H-6a is within the shielding cone of the side-chain phenyl, to conformations of type A and C, where these hydrogens

71 are synclinal and the aminoacid phenyl group is away from the concave face of the taxoidic core [35]. The conformation of the phenylisoserine moiety of pseudoalkaloidal taxoids can also be affected by the solvent. Indeed, differences in J2',3' have been observed in N-acyl and N-alkyl esters when the spectra are taken in lipophilic (CDC13, benzene-d6) and hydrophilic solvents (MeOH-d4, DMSO-d6, DMSOd6/D20 mixtures). Interestingly, the changes are opposite for the two types of esters. Thus, J2',3' increases with the polarity of the medium in N-acyl esters [2023], but decreases in N - a l k y l esters [35]. In N-acyl esters like taxols, conformation A is preferred in apolar media, but conformations like B are preferred in aqueous media due to hydrophobic clustering of the side chain phenyl, 2-benzoate and 4-acetyl groups [22]. In N-alkyl esters, the contribution of the hydrogen-bonded conformation B is less important in polar media, due to competitive intramolecular hydrogen bonding with the solvent. N-Methyl and N,N-dimethyl derivatives of t a x u s ~-aminoacids can easily be distinguished by 13C NMR spectroscopy (5 N-Me c a . 35 and 42, respectively). A discussion of the conformation of Taxol| in relation to its bioactivity also appears in Chapter 6. Non-nitrogenous side chains generally show normal chemical shifts, but certain acyl groups can display peculiar NMR features. Thus, in O(9), O(10)diesters, when one of the acyl groups is an acetate and the other a benzoate, the 1H NMR resonance of the acetate methyl is shifted to high field (5 1.70-1.80, see 13). In oxetane-type taxoids, the 0(4) acetyl resonates at low field both in the 1H(5 c a . 2.20-2.40) and in the 13C NMR spectra (5 ca. 22). Epimerization at C-7, with formation of a strong intramolecular hydrogen bond between the 7-OH and the 4-acetyl, shifts the acetyl resonance to even lower fields (5 2.30 in Taxol| and 2.50 in 7-epi-taxol, see 18 and 19). The resonances of the cinnamate ~ and o r t h o - h y d r o g e n s

differ in 13-oxo-

and 13-hydroxy(acycloxy)-taxoids, presumably because the cinnamoyl residue is in the concave face of the molecule and is subject to the anisotropic effect of the C-13 carbonyl. The detection of NOE effects between the cinnamate hydrogens and H-18 is in agreement with this [11]. Thus, in taxinine derivatives the more downfield signal of the cinnamoyl residue is H - o r t h o (Figure 9), and H(~) resonates at 5 c a . 7.60. In taxinine J derivatives [40], the resonances of the cinnamate moiety are instead normal, and H(~) is the more downfield signal (5 7.80), whereas the resonance of the o r t h o - h y d r o g e n s other aromatic hydrogens (5 ca. 7.50) [40].

ca.

is close to that of the

72

AcO

OAc

O

OAc

71~,,6

H 7.76

6.43

5.3.1 Taxinine [36]

~

AcO

OAc

C

7.80 O

H

H 7.50

AcO ....

6.60

5.3.2 2-Deacetoxytaxinine J [40] Figure 9: Resonances of the cinnamate hydrogens in taxinine and 2-deacetoxytaxinine J

2.5.4. Selected Examples Formulas 1-19 represent a selection of fully assigned 1H and 13C NMR spectra of taxoids and are representative of all the major naturally occurring structural types and functionalization patterns. All spectra were taken in CDC13 operating at 300 MHz for 1H and 75 MHz for 13C, except for 2 and 3 (DMSO-d6) and 15, 18 and 19 (500 MHz and 125 MHz for 1H and 13C, respectively). Multiplicities in the 1H NMR spectra are given after D 2 0 exchange, and longrange couplings have not been considered. Thus, the resonance of H-18 is given as a singlet, even though under optimized conditions the signal is a narrow doublet (J

ca.

1 Hz).

73

1-HYDROXYBACCATIN I (1) [ ref.41] OAc

AcO 2.22 s

"

6.21 d

1.24 s

OAc

6.03 d

H

1.23 s

AcO,,,,

5.48 dd

6.08 dd

' " ' H 2.17m

4.21 t

eessss 1.64s 2.53

1.74m

1 4,

ddH

'OAc

,....

H HO

5.47 d :

1.87 dd

~Ac

H 3.53 d H

2.31 d

Ac: 8 2.21" 2.11"2.08, 2.05; 2.04; 1.99 (s)

J(Hz): 2,3=3.6; 5,6(z=5,6~=3.0; 7,6(x=4.5; 7,6~=12.2; 9,10=10.8" 13,14(x=6.6; 13,141~=9.5; 14oq14~=15.0; 200~,20~=5.1.

OAc

AcO 15.4

"

140.4

13.7

OAc

28.4

AcO,, ,, < 7,1

31.1 46.6

43.3 77.7 i i#

41.3 21.8

"OAc

38.5

HO

72.2" m

49.9

~Ac Ac: 170.1" 169.9; 169.8;169.4; 169.2; 169.2 (s); 21.9; 21.8; 21.6; 21.4; 20.9; 20.6 (q).

74

10-DEACETYLBACCATIN [ref.42] HO 1.90s 9.

(DMSOd6) (2)

III

O

5.13s

I

J

J

1.51s

OH H

0.93 s 4.62 m

H O'"'

s99

""H

2.28m

|

t# t 0.94 s 2.40m

4.09 dd

1.64m

,[.

4.90 dd

H

"O H HO

,,oo.-

3.81 d

oAc-

H

2.15m

1":t~40~s

OBz

Ac: 8 2.20 (s). Bz: 8.08 (d, J=7.8 Hz); 7.75 (t, J=7.8 Hz); 7.61 (t, J=7.8 Hz).

J(Hz): 2,3=6.9; 5,6o~=8.9; 5,613=2.0; 7,60~=7.9; 7,6B=9.9; 13,140~=13,14B=8.4.

HO

O

14.8

9.7 74.3

OH

210.2

141.5

26.7

36.5 57.0

H O""

66.0

####9

46.5

! 83.7

20.1

HO

~4~:.

OAc 7~4

ogz Ac: 169.5 (s); 22.2 (q). Bz: 165.2 (s); 130.2 (s, i) 129.4 (d, o) 128.6 (d, m) 133.1 (d, p).

?5

1413-HYDROXY-10-DEACETYLBACCATIN

III

(DMSO de) (3)[ref.421 HO 1.91 s

_0 OH

1.53 s

5.13s

H 0.92 s

HO,,,,

4.07 dd

""H

4.41 m

_

"

~

%

H HO

-

0

i

5.54d --

3.79 d

2.28m

4.90 dd

0.97 s

HO

1.64m

3.75d

(~Ac

OBz

H 3.99d H ~.~7d

Ac: 8 2.24 (s). Bz: 8.09 (d, J=7.8 Hz); 7.73 (t, J=7.8 Hz); 7.61 (t, J=7.8 Hz). J(Hz): 2,3=7.1 5,6o~=9.1 5,613=2.0; 7,6o~=7.9; 7,6~=10.1 13,14=5.5; 200q20~=8.2

HO

O

~,

14.9

//~

OH.

21 139.5

26.6

H O,,,, ( 74.6

36.6 57.3

42.: II I

46.0

II I

| 83.6

21.4

O HO

HO

74.0i

H OAc

75.4

6Bz Ac: 169.8 (s); 22.3 (q). Bz: 165.4 (s); 129.7 (s, i); 129.8 (d, o)" 128.7 (d, m); 133.3 (d, p).

76

4-DEACETYLBACCATIN IV (4)[ref.34]

AcO 2.03 s

OAc 9

6.O5 d

OAc

1.45 s

5.84 d

H

1.08 s

AcO,,,,

5.25 t

' " ' H 2.35m

5.72 dd 4.80 dd

1.65 s 2.25

1.90m

ddH

o

H HO

5.64

2.40 dd

d -

0

2

d

H

OAc

"

..

H4.36d

H 4.41

d

5 2.12; 2.10; 2.08; 2.01 (s) J (Hz)" 2,3=5.5; 5,6(z=8.1" 5,613=2.6; 7,6o~=7,6~=9.2; 9,10=11.0; 13,14(z=9.7; 13,1413=3.7; 140~,1413=15.5; 20~,2013=7.7.

Ac:

AcO

OAc

16.4

9

70.7

13.2

OAc

74.8

139.7

29.8

33.9 45.1

AcO,,,,

71.0

42.1 86.3

50.5 20.3

O

35.8

HO

72.5" 1

H

OH

79.7

OAc Ac:

171.0; 170.0; 170.0;170.0; 169.1 (s); 21.2; 21.2; 21.1-20.9 20.6 (q).

7"7

BACCATIN Vl (5)[ref.26]

AcO 2.01 s

OAc 9

6.20 d

1.58 s

OAc

5.98d

H

1.21 s

AcO,,,,

5.50 dd

'"'H

6.14t

2.48m

4.95 d

1.76 s 2.20 m

1.84m

H H

HO

H

~8~- -

2.20 m

3.16 d

~

(5 AC "

(~Bz

H

4.10 d

H 4.31 d

8 2.27; 2.17; 2.09; 2.08; 1.98 (s). Bz: 8.09 (d, J=7.7 Hz); 7.46 (t, J=7.7 Hz)" 7.59 (t, J=7.7 Hz). J (Hz)" 2,3=6.2; 5,6(z=8.9;7,60~=7.8; 7,613=9.7; 9,10=11.5; 13,14(z=13,1413=8.6; 20o~,2013=8.3.

Ac:

AcO

OAc

14.9

9

OAc

75.0

133 141.2

AcO,,,, (

12.7

28.2

34.5 45.7

69.6

42.7

47.2

II 83.8

22.2

O

35.1

HO

73.2" m

H O A c ~~

~Bz 170.4; 170.1" 169.8; 169.1- 168.8 (s); 22.7; 21.3; 21.2; 20.9; 20.7 (q). Bz: 166.9 (s)- 129.2 (s,i)- 130.1 (d,o); 128.6 (d,m); 133.6 (d,p).

Ac:

78

TAXININE (6)[ref.36]

AcO 2.28 s

~.

OAc 9

6.04 d

L 5.89d~

0.93s

1.70-1.80 m '

9

,,. 9 9

.H'N

1.15s

IIII H

0

O

## 5.35 t

ss S 1.76 s

2.84ddH

"

"

H-" H'

~4~,

H e.43d

5.55 dd :

~"

(~A

3

C

O

I

d

/ \

H 4.84s

Hs.3s s

H~

7.76 d

7.66d

Ac: 8 2.07; 2.06; 2.05 (s). J (Hz)" 1,2=2.1" 2,3=6.6; 5,60~=5,6~=2.5; 9,10=10.3; 14~,14~=20.0; 1413,1=7.0; 2',3'=16.1" o,m=ca 8.5.

AcO

7.43 m

OAc

14.0

17.5 73.5

27.5 28.4

44.5

O

7.43 m

O

199.4 #####

78.2

25.2

s#s O

H

117.9

=.=

117.2

145.8

134.6

OAc 128.

Ac: 169.9; 169.7; 169.4 (s)" 21.4; 20.9; 20.7 (q).

130.4

79

5-01N N AM OYLTR IA CETY LTAXI CI N i (7) [ref. 36] AcO 2.29 s

OAc 9

6.12d

0.96

I

5.94 d 1.24 s

IIII H

O 2.90d

~

##### 1.71 s

5.35d

O

###

H

H

H HO

~~

~~d"-

6.43d

3.55 d

/\

OAc

H,.,o, a5.37s

H

~

7.76d

7.67 d

Ac: 8 2.14; 2.10; 2.08 (s).

J (Hz)" 2,3--6.8; 5,6o~=5,6~=2.5; 9,10=10.0; 14oq14~=19.9" 2',3'=16.1" o,m=ca 8.5.

AcO

OAc

13.8

17.5

73.1

75.

27.6

34.4

O

7.44 m

199.2

#####19.8

HO

28.5

44.5

42.8

m

O

78.2 #

45

117.8

se 0

A 117.o

14

.

34.6

OAc

Ac: 171.7; 169.9; 169.7 (s) 21.2; 20.9; 20.7 (q).

130.4

80

2-DEACETYLDEClNNAMOYLTAXININE

E (8)

[ref. 35] AcO 2.11s

OAc

9 -

H 1.89m

0.88 s

6.05 d

____4,'

1.60 m

5.71 d

9 9

r"

A c O ' ' ' ' ( 5.76dd

1.59 m

"

#l I

H

..... 18om

I

""'OH

~.,, ddd H H 1.40 dd

(~H

H s.~

Ha.sos

8 2.08; 2.04; 2.00 (s). J (Hz): 1,2=2.1" 2,3=6.2; 5,6(z=5,6~=2.7; 9,10=10.4; 13,14(z=5.0; 13,14~=10.4;14~,14~=15.7" 1,14~=8.6.

Ac:

OAc

AcO 15.8

,~ 133.9

17.4

72.7

26.7 32.5

30.7

44.9

A c O , , , , ('""

~"

76.7

stssss26.1 43

I #I

"OH H

B

116.1

oH Ac:

170.3; 170.0; 169.8 (s)- 21.0; 20.9; 20.8 (q).

81

2-DEACETOXYDECINNAMOYLTAXININE J (9) [ref. 35] AcO 2.18s

OAc 9

6.25 d

OAc

0.79 s

5.81 d

H

0.98 s

5.67 dd

" " H 1.95m

A c O ' ' ' ' ( ,.7~dd 4.28 t

sssse S 1.55s

1.65m

2.77 ddd H

I #I

"OH H 1.08 dd

H

9,

3.20 d

1.78 m

H

H

1.88 m

H 4.83 s

1.72 m

H5.16s

Ac: 8 2.05; 2.04; 2.01" 1.96 (s).

J (Hz)" 213,3=5.4; 5,6o~=5,6~=3.0; 7,60~=5.4; 7,613=11.6; 9,10=10.8; 13,14o~=4.7; 13,1413=10.7;14(z,1413=15.7;1,1413=8.5.

AcO

OAc

15.9

.~ 135.9

12.6

OAc

72.1

137.

32.1

36.0

46.7 AcOllll

(70.0

38.9 sssss

$26.2

73.3

35

#

32.

I #

"OH

H

26.9 112.6

Ac: 170.3; 170.0; 169.6; 169.2 (s) 21.4; 21.0; 20.9; 20.8 (q).

82

1 - H Y D R O X Y D E C l N N A M O Y L T A X l N I N E J (10) [ref. 35] AcO

OAc

2.20 s

9

6.25 d

OAc

0.96 s

5.85 d

H1.59

1.10s

5.67 dd

' " ' H 1.95m

A c O ' ' ' ' ( ~-~"d 4.23 t

sssse S 1.63s

m

I #

2.34 dd

i

"OH 5.55 d -

H HO

3.66 d

1.94 dd

6Ac

H 4~s

H5.29 s

8 2.09; 2.08; 2.03 2.02 1.98 (s). J (Hz)" 2,3=6.2; 5,6o~=5,613=2.5; 7,6(~=5.4; 7,6~=11.5" 9,10=10.9" 13,14o~=5.3"13,14~=10.1"14ot,14~=15.6.

Ac:

AcO

OAc

15.9

9

134.5

71.3

13.0

OAc

75.,

140. 29.0

37.1

47.9

AcO,,,,

70.6

42.0 ssstes20. 8 43

I

75.2 ##

"OH HO

m

(3Ac Ac:

116.2

171.4; 169.9; 169.8; 169.6; 169.1 (s)- 21.3; 21.1 20.9; 20.9; 20.7 (q).

83

1 -HYDR OXY-2-D EAC ETOXYD E Ci N NAMOY L TAXININE J (11)[ref. 41] AcO 2.18s

OAc 9

6.27 d

0.81 s

OAc

5.83 d

H

1.66m

1.06 s

" " H 1.95m

AcO,,,, <,~ s s s t t S 1.55s

4.30 t

2.57 dd H

I

'#,#

"OH H HO

3.15d

1.55 dd

H

H

2.02 dd

1.77 d

H 4.89 s

H5.20 s

8 2.06; 2.05; 2.02; 1.98 (s). J (Hz) 20c,213=15.4;213,3=5.7; 5,6(x=5,613=3.0; 7,6o~=5.0; 7,613=11.4; 9,10=10.7; 13,14(z=4.6; 13,14~=10.3; 14~, 1413=15.3.

Ac:

AcO

OAc

15.8

.~ 135.5

71.7

12.6

OAc

76.

139

28.2

35.8

47.0

AcO,,,,

71.0

43.1

lllll

l

73.1 I I#

20.4 41

"OH

HO

36.6 112.8

Ac:

170.2; 169.9; 169.7; 169.2 (s)" 21.4; 21.0; 20.9; 20.8 (q).

84

TAXINE B (12) [ref.43]

OH

AcO 9

2.15s

I

1.78 m

H

1.13 sH_

5.85 d 4.28 d

1.37m

:

~

1.25 s

0

##### 1.52 s

~.,,

-"

4

H HO

"

,H~.

3.98 d ..

5.10t

II

'"0

H ~~3d~

H

2.92 dd

3.17d

m

~t

/\

oH

Hs3ss

Me2N ' " '

5.4os

//

~

7.32 m

Ac: 8 2.22 (s). J (Hz)" 2,3=7.0; 5,6o~=5,6~=2.5" 9,10=9.5; 2'oq3'=7.0; 2'~,3'=7.0" 2'0q2'~=16.0.

AcO

OH %

13.9.

17.9 76.6

139.1

34.2

199.9

0

26.4 29.2

45.6

42.3

78.2

46

HO

O U

m

118.1

o.

42.3

| 66.5 .. ~ , ~ .

M"~ 2 N '

\1~"~ 127.6

Ac:

170.2 (s); 21.2 (q).

8.5

B R E V I F O L I O L (13) [ref. 29b, 41] BzO 1.99 s

OAc 9

6.51 d

OAc

0.87 s

6.02 d

H1.81 5.55 dd

HO'"'

4.36 t

m

' " ' H 1.97m

4.39 t S

##,

~.~ ddH ~ 1.28 dd

"OH

H

~ f ~ s A

1.32

H 2

~.~d

2.7~4 d

Hl~d

H ,,7~s H ~ s "

"

Ac: 8 2.05; 1.72 (s). Bz: 7.84 (d, J=7.6 Hz); 7.53 (t, J=7.6 Hz)" 7.41(t, J=7.6 Hz).

J (Hz): 2r 2~,3=8.9; 5,60~=5,613=2.5; 7,6o~=5.3; 7,613=11.2; 9,10=10.5 13,14e~=13,14~=7.3; 14e~,1413=14.0.

BzO

OAc

12.0

.~

OAc

70.4

133.9

151

12.9

36.1

45.0

HO,,"

77.2 72.4

37.9

#

62.5

SI

47.

'OH 76.0

27.0

29.1 112.0

HO

24.8

Ac: 170.5" 170.0 (s) 21.4; 20.7 (q). Bz" 166.4 (s); 129.3 (s,i)" 129.4 (d,o); 128.7 (d,m); 133.2 (d,p).

86

7,9,10-TRIDEACETYLABEOBACCATIN (14) [ref. 30] OH

HO 1.84 s

Vl

OH

1.86 s

4.56 d 4.35 d

H 4.23 t

1.80m

" " H 2.57m

A c O , , , , { 5.72t 4.92

~.~dd , H""a

: Y~O

%

1.87 dd

H

-

1.10

HO

1,3s

2H6

(~Bz

-

" OAc H H 4.08d

4.40d

Ac: 8 2.15; 2.13 (s). Bz: 7.96 (d, J=7.6 Hz); 7.59 (t, J=7.6 Hz)" 7.45(t, J=7.6 Hz). J (Hz): 2,3=7.3; 5,6(x=8.2; 7,6(x=7,6~=8.3; 9,10=10.0; 13,14(x=13,1413=7.3;14o~,1413=14.0;20(x,2013=8.1

HO

OH

11.3

,9

143.~

12.2

OH

68.3

139.0

37.2

42.4

AcO,,,,

79.4 84.9 67.8 36

O 76.4

27.9

HO

k24.7

"

74.4

oAc

~~)az

Ac: 170.8; 169.1 (s); 22.1" 21.2 (q). Bz: 166.2 (s) 129.7 (s,i); 129.6 (d,o); 128.6 (d,m); 133.6 (d,p).

87

TAXININE M (15) [ref. 44] OBz

AcO

AcO

1

OAc

i,

1.17s

.....

5.31 d

HO

"

~.~d\ I ..... I 4.08 d 3.63 d

~11 . ~ " ~5"51

H~orn "~ dd

' " ' H 2.23m

O 4.45 t

1.29 s

#

3.00 dd H

"OH H

H

2.75 d

,,,~d-9-

2.48 ddd

OAc

3.71 d

/

H 4.68s

H 5.41s

Ac: 8 2.15; 2.11" 2.03; 2.03 (s). Bz: 8.16 (d, J=7.9 Hz); 7.61 (t, J=7.9 Hz) 7.51 (t,J=7.9 Hz).

J (Hz) 1,2=2.4; 2,3=10.4" 5,6o~=5,6~=2.5; 7,6o~=6.2;6eL,6~=14.2"7,613=10.7; 9,10=3.0; 143,1413=19.2; 14~,1=0.7; 1413,1=11.6; 17o~,1713=8.1" 19c~,1913=12.2.

OBz

AcO

AcO

1

OAc

1

12.0 ssss

1

HO

1

64.0 91.3

70.C 82.2

39.0

50.0

O 72.6 # I#

15.5 38.

"OH

33.7

H

i

(]Ac 172.7; 170.0; 168.7; 168.4 (s)" 21.2; 21.2; 20.7; 20.7 (q). Bz: 166.8 (s)" 129.2 (s,i); 130.0 (d,o); 128.6 (d,m); 133.6 (d,p).

Ac:

113.3

88

9,10-DIACETYL-5-CINNAMOYLPHOTOTAXICIN I (16)[ref 31] OAc

AcO 1.26 d

9

5.64 d

1.36

s

1.76 m H1.26

_

5.64 d

iiS

m

H 2.18m

,1.08 s

O 2.36 d

O IIIm, H 1.76 m

%

H

5.60 t

"~ 1.50 s

H -"

6.38 d

'O

4 4.82 d .. m

H HO 3.03 d

H sa, s H5.61s

oH

7.54 d

H ~j,.7.65 d

Ac: 8 2.04;2.04 (s).

J (Hz)" 2,3=1.8; 5,6ct=5,613=9.2; 9,10=9.8; 14oq14~=20.0; 2',3'=16.4; o,m=ca 8.5.

7.38 m

OAc

AcO 15.8

9

25.9

79.7 51.3

25.8 23.5

31.0 44.9

O

214.4

#1

%

O

76.5

9

"0

117.0

HO 128.8

14

.

34.1

oH

Ac: 171.0; 170.0 (s)" 21.1" 21.0 (q).

130.4

89

TAXlNE A (17) [ref. 35, 46]

HO 1.93 s

O

OH

.17s

5.17s

H

1.11 s 2.64 AcOIII!

1.16s

~.74..H'--"

1.68 d

H

H

1.62 dd

-0.20 d ~

"~

s S

1.82 dd

'"'H 0

H_

~'5.43h,dd

1.77 m

5.43 d

9

H

",

"O"

3.81 d

2.25 s

5.30 d

~ 5.61 dd

(3H ,,,'---"~ 46~d

Me2N '""

AcO

8 2.29; 1.98 (s). J (Hz): 1,2=2.5; 2,20=10.0; 3(z,313= 15.5; 5,613=6.4; 60q613=15.0; 7,6o~=3.6; 7,613=13.6; 13,14o~= 3.1; 13,1413=11.0; 14o~,14~=16.5; 1413,1=7.5; 2',3'=10.0.

7.38 m

Ac:

HO

O

16.7

7.38 m

OH

18.3

76.9

7.26 m

213.~ 132.4 24.5

AcO,,,,

53.2

70.5

37~se

9 34.9

27.4

H

\

,,,OH

' O" 1~"" 1%6

69.8

AcO

O

~o;~~

#

18.3

34.4

123.4

40.8

Ue2N"-

~ l ~ k 71"9

,,

12 Ac:

170.7; 169.9 (s) 21.4; 21.2 (q).

90

TAXOL (18)[ref.16] H

Bz \

AcO

7.01 d

/

N = "

1.79d

O 5.78 dd

OH -

1.88 ddd

H

II

1.24 s

_

~'4. d .~8

Illl H

U ' " ' s ~.~'

OH

#l l

2.54 ddd

494 dd

## 1.14 s

O

~--9

7.,~ m

7.35 m

.68 s

6.27 s

~'~ ~

1 5.67 d -

H HO

3.79 dd

i

SAc"

'H

H

OBz

4.19 dd

4.30 ddd

Ac" 8 2.38 (s, C-4)" 2.23 (s, C-10). O-Bz: 8.13 (dd, J=8.4,1.3 Hz); 7.61 (tt,J=8.4,1.3); 7.51 (m). N-Bz 7.74 (dd,J=9.0,1.5); 7.49 (m); 7.40 (m). J (Hz): 2,3=7.1" 5,6(z=9.6; 5,613=2.3; 6oq613=14.8; 7,6o~=6.7; 7,613=10.9; 13,14o~=13,1413=9.0; 140q14~=15.3; 20cz,2013=8.5" 2',3'=2.7; N-H=8.9.

Bz

H N

AcO O

1~8

~.~

133. " 55.0 .,.

O

172.7

1

,~

b,

I mmI (72.3

43.2

75.5

203.(

A, 26.9

138.0 .73.2

1~7.OOH

O

Ill

ll

i .. 58.6 45.6 t

21.8 35.7 V

128.3

129.o

HO

OH

74.9i

OBz Ac: 171.2 (s,C-10);170.4( s,C-4); 22.6 (q,C-4)" 20.8 (q,C-10). O-Bz 167.0 ( s ) 129.1 (s,i); 130.2 (d,o)" 128.7 (d,m); 133.7 (d,p). N-Bz: 167.0 (s); 133.6 (s,i); 127.0 (d,o)" 128.7 (d,m); 131.9 (d,p).

35.6

~4.4

91

7-EPITAXOL Bz

\

/

H

AcO

7.00 d

N

1.79 d

O

i

O

6~0s~ ) ~ . ~ ,

67s O._ H

/

m n n

(19)[ref.16]

5.81 dd

\1

~7 ddd

:

H

1.19s

lilt H Bill

.. 4.81 d

( 6.23t

##

i

8H

7.34-7.56 m

~.~,,,

ss 1.15s

H ~

~.4~d H

.[

2.33 ddd

494dd

0

i

HO

5.76d -

d

8Aci

II

'H 4.39

OBz

H

S

4.39 s

Ac: 8 2.50 (s, C-4); 2.19 (s, C-10). O-Bz: 8.18 (dd, J=8.5,1.3 Hz); 7.62 (tt,J=7.4,1.6); 7.52 (m). N-Bz: 7.72 (dd,J=8.4,1.3); 7.56-7.34 (m). J (Hz): 2,3=7.5; 5,6o~=9.0; 5,613=3.5; 6oq613=16.0; 7,6(x=2.1; 7,613=5.0; 13,140~=13,14J3=9.0; 14oq1413=15.4; 2',3'=2.6;3', N-H=9.0.

Bz

H N

AcO O

_: ~4.8 "

II 172.9

O

14.7

1

133.~

7~.1 ~o7.~\

~

25.9

" 36.1

138.1

57.6 9. 73.1

I I I I { 72.3

i

42.6

I #

# #

i

1~7.oO H

1~,8

82.8

40.3

21.2

i

1289

OH_

16.1

35.3

HO

O 75.3 -

H

i

Ogz Ac: 169.5 (s,C-10);172.4 (s,C-4); 22.5 (q,C-4) 20.8 (q,C-10). O-Bz: 167.3 ( s ) 129.4 (s,i); 127.2 (d,o) 129.1(128.4, 128.8)(d,m) 132.0 (d,p). N-Bz: 167.2 (s); 133.7 (s,i); 130.0 (d,o) 128.4 (129.1,128.8) (d,m); 133.8 (d,p).

SAc 7-6

92 2.6. X-RAY A N A L Y S I S

Many natural taxoids have been analyzed by X-ray diffraction methods (Figures 10, 11), and data are available for all the major structural types. F u r t h e r m o r e , several derivatives of n a t u r a l taxoids and m a n y synthetic compounds related to taxoids have also been subjected to X-ray analysis. X-ray data on Taxol| have never been published, but data are available on its semisynthetic analog Taxotere| [47], as well as on the side chain of Taxol| (both as methyl ester [48] and as the p-bromobenzoate of the methyl ester [49]) and of cephalomannine (taxol B) [50]. Baccatin III, corresponding to the diterpenoid core of Taxol| has also been analyzed by X-ray [51], and Figure 12 depicts an ORTEP view of the crystal structure. In taxoids and some of their rearrangement products, certain bonds are u n u s u a l l y long, especially those at the ring junctions (C-1 in 11(15>l)abeotaxanes [29b]; C-3 in 3,11-cyclotaxanes [62]). Given the crowded and compact topology of taxoids, several bond angles deviate significantly from the ideal values. However, few non-bonded interactions involving non-hydrogen atoms are present. The most common conformational types are exemplified by the torsion angle sequences observed in the solid-state conformations of baccatin III [51], taxagifine [24], 5-cinnamoyltriacetyltaxicin I [55], and 13-deacetylisoabeobaccatin VI [29b] (Figures 13-16). 2.7. C O N C L U S I O N S The structural elucidation of taxoids can be carried out using a variety of spectral methods, especially NMR techniques. Some spectroscopic features of taxoids are very unusual, and their rationalization provides unique challenges to spectroscopists.

93 AcO H ~ O

OAc "~

eeteee

AcO

OAc

O

ss

"OAc

"OCinn

OAc

OAc

14~-Bromotaxinol acetate [52] AcO

AcO ....

Triacetyl-5-cinnamoyltaxicin I [53] AcO ~,

OAc

,.....

O

OAc -

susoS

"O H

"OAc OAc

Taxusin [54] AcO AcO ,("

T a i w a n x a n [55]

R

R20,, .

.%

O

~,

II

,.., ~. 1

HO . . . . . . . .

O

R3

. -

OAc R=H Taxagifine [241 R=OBz Taxacin [56]

OBz

O OAc

RI=~-OH , R 2 =Ac, R 3 =H,Baccatin III[53] RI=~-OH, R2 =H, R3=OH,14~-HydroxyDAB[42] RI=~- I OAc,R 2 =IOAc,R 3 =H, DAB-7,10-bisiodoacetate [49] RI=(~-OH,R 2 =Ac, R 3 =H, Baccatin V [57]

Figure 10: Natural taxoids and simple derivatives investigated by X-ray diffraction (DAB=10deacetylbaccatin III). References are given in brackets for each entry.

94 AcO

OHHo

AcO

OAc OAc "

AcO'"'

AcO" HO

OBz

OAc

O

_

HO~

'"OAc

Taxuchin [61]

7,9-Bisdeacetylbaccatin VI [58]

R40,,, R2

RI=OAc,R2 =H,R3 =Bz,R4=H Taxchinin A [59] RI=H,R 2 =Ac,R3 =Bz,R4=H Brevifoliol 5-acetate [60] RI=Ac,R2 =H,R3 =Bz,R4=Ac Taxchinin D [28] RI=Ac,R2 =R3 =H,R4=Ac Taxchinin G [28]

-OR~A c R40'"'~ O OAc

HO OR1 RI=Ac,R2 =Ac,R3 =Bz,R4=H, 13-Deacety]isoabeobaccatin VI [29b] RI=Bz,R2=Bz,R3 =H,R4=Ac, 9,10-Deacety]-9-benzoy]abeobaccatin VI [26] HO AcO O // OTES _

~

AcO,,,

O HO

= OBz -

O

AcO ....

OAc

7-TES- 13-Acetylphotobaccatin III [62]

NMe 2

'"O H . AcO Taxine A [46]

Ph OH

Figure 11: Natural taxoids and simple derivatives investigated by X-ray diffraction.

95

C(041l C(042)

0{023) 0(009)

0(006) C(017)

0(005)

C(027) C(010)

C(033)

[016}

C(024)

C(019)

~_C(037)

~,030)C(021 4) 01002)

~,

C(029) C(036)

0(007)

1

0(0031 C(034) 0(004)

0{008}

0(0011

1035}

C(025) C(026) C(039} C(0401~,~1~ C(038) 0(031)

Figure 12: ORTEP drawing of baccatin III

95

AcO

OAc %

59.9 58.9

O::~~

49.7

' -15.4 ~ , J -21.6 57.1

T-54.9

" -49.3

-46.al 61.4 ,14.3

-59 ",j

1

99

-75.2

HO

~,,166.8

7-

ou.~

99

"'0Cinn

H

m

OAc Figure 13: Endocyclic torsion angles in the solid-state conformation of triacetyl-5-cinnamoyltaxicin I [53]

AcO

OAc OAc

%

HO 122.8 9-35.0

46.6

-51.

-55.5

-51.9

O

I I I I I I

-36ok

.~n ~

/84.8

-5:

100.,

-86.0

H

60.0

.,,,~-7o.7

I

"OCinn

m

OAc Figure 14: Endocyclic torsion angles in the solid-state conformation of taxagifine [24]

97

O

AcO

OH 55.5

55.2 -34.7

HO

40.4

-43.5

54.3

-6.0

'lll

t 29.8

"53"'~"

-58.! 57.9

.7

,0"~,.

t9

-70.7

. . . . 0.3

,os.~,-

O

,,..

i

HO

OAc

OBz Figure 15: Endocyclic torsion angles in the solid-state conformation of baccatin III [51]

OAc

BzO

OAc 59.0 27.6

-50.6 ~, I

/

~

39.4

-66.5

H O,,,,

-11.1

|

| -49.3

17.8

-28.3

"

HO

-2.7

-44.1|60.7 -2.6

6Ac

OAc

Figure 16: Endocyclic torsion angles in the solid-state conformation of 13-deacetylisoabeobaccatin VI [29b]

98 REFERENCES

o

.

o

.

~

~

~

.

10. 11. 12. 13. 14. 15. 16.

17.

Griffini, A.; Peterlongo, F.; De Bellis, P.; Pace, R. Fitoterapia 1993, 64, 53. Barboni, L.; Gariboldi, P.; Torregiani, E.; Appendino, G.; Gabetta, B.; Bombardelli, E. Phytochemistry 1994, 36, 987. Bitsch, F.; Shackleton, C.; Ma, W.; Park, G.; Nieder, M. Rapid Commun. Mass Spectrom. 1993, 7, 891. Kurono, M.; Nakadaira, Y.; Onuma, S.; Sasaki, K.; Nakanishi, K. Tetrahedron Lett. 1963, 7, 2153. Lythgoe, B. in The Alkaloids; Manske, R.H.F., Ed.; Academic Press, New York, 1968, vol.10, p.597. (a) Ettouati, L.; Ahond, A.; Convert, O.; Poupat, C.; Potier, P. Bull. Soc. Chim. Fr. 1989, 687. (b) Zhang, H.; Takeda, Y.; Minami, Y.; Yoshida, K.; Matsumoto, T.; Xiang, W.; Mu, O.; Sun, H. Chem. Lett. 1994, 957. Harrison, J.W.; Scrowston, R.M.; Lythgoe, B. J. Chem. Soc. (C) 1966, 1933. Della Casa de Marcano, D.; Halsall, T.G.; Scott, A.I.; Wrixon, A.D.J. Chem. Soc. Chem. Commun. 1970, 582. Harada, N.; Ohashi, M.; Nakanishi, K. J. Am. Chem. Soc. 1968, 90, 7349. Harada, N.; Nakanishi, K. J. Am. Chem. Soc. 1969, 91, 3989. Woods, M.C.; Nakanishi, K.; Bhacca, N.S. Tetrahedron 1966,22,243. Eyre, D.H.; Harrison, J.W.; Lythgoe, B. J. Chem. Soc. (C) 1967, 452. Miller, R.W.J. Nat. Prod. 1980, 43,425. Rojas, A.C.; de Marcano, D.; M~ndez, B.; De M~ndez, J. Org. Magn. Res. 1983,21, 57. De Marcano, D.; M~ndez, B.; De M~ndez, J.; Monasterios, J.; Rojas, A.C.; Halsall, T.G. Org. Magn. Res. 1983, 21, 524. Chmurny, G.N.; Hilton, B.D.; Brobst, S.; Look, S.A.; Witherup, K.M.; Beutler, J.A.J. Nat. Prod. 1992,55, 414. For the revised assignments of the 13C NMR resonances of the N-benzoyl and the aminoacid phenyl, see" Harris, J.W.; Katki, A.; Anderson, L.W.; Chmurny, G.N.; Paukstelis, J.V.; Collins, J . M . J . Med. Chem. 1994, 25, 706 (footnote 11). Baker, J.K. Spectr. Lett. 1992,25, 31.

99 18. 19. 20. 21. 22. 23. 24. 25. 26.

27. 28. 29.

30. 31. 32. 33. 34.

Falzone, C.J.; Benesi, A.J.; Lecomte, J.T.J. Tetrahedron Lett. 1992, 33, 1169. Hilton, B.D.; Chmurny, G.N.; Muschik, G.M.J. Nat. Prod. 1992, 55, 1157. Dubois, J.; Gu~nard, D.; Gu~ritte-Voegelein, F.; Guedire, N.; Potier, P.; Gillet, B.; Beloeil, J.C. Tetrahedron 1993, 49, 6533. Williams, H.J.; Scott, A.I.; Dieden, R.A.; Swindell, C.S.; Chirlian, L.E.; Francl, M.M.; Heerding, J.M.; Krauss, N.E. Tetrahedron 1993, 49, 6545. Vander Velde, D.G.; Georg, G.I.; Grunewald, G.L.; Gunn, C.W.; Mitscher, L.A.J. Am. Chem. Soc. 1993, 115, 11650. Williams, H.J.; Scott, A.I.; Dieden, R.A.; Swindell, C.S.; Chirlian, L.E.; Francl, M.M.; Heerding, J.N.; Krauss, N.E. Can. J. Chem. 1994, 72,252. Chauvi~re, G.; Gu~nard, D.; Pascard, C.; Picot, F.; Potier, P.; Prang~, T. J. Chem. Soc. Chem. Commun. 1982, 495. Appendino, G.; Ozen, H.~.; Gariboldi, P.; Torregiani, E.; Gabetta, B.; Nizzola, R.; Bombardelli, E. J. Chem. Soc. Perkin Trans. I 1993, 1563. Barboni, L.; Gariboldi, P.; Torregiani, E.; Appendino, G.; Cravotto, G.; Bombardelli, E.; Gabetta, B.; Viterbo, D. J. Chem. Soc. Perkin Trans. I 1994, 3233. Toromanoff, E. Tetrahedron 1980, 36, 2809. Li, B.; Tanaka, K.; Fuji, K.; Sun, H.; Taga, T. Chem. Pharm. Bull. 1993, 41, 1672. (a) Appendino, G.; Tagliapietra, S.; Ozen, H.~.; Gariboldi, P.; Gabetta, B.; Bombardelli, E. J. Nat. Prod. 1993,56, 514. (b) Revised structure: Appendino, G.; Barboni, L.; Gariboldi, P.; Bombardelli, E.; Gabetta, B.; Viterbo, D. J. Chem. Soc. Chem. Commun. 1993, 1587. Appendino, G.; Cravotto, G.; Enrifi, R.; Jakupovic, J.; Gariboldi, P.; Gabetta, B.; Bombardelli, E. Phytochemistry 1994~ 36, 407. Appendino, G.; Lusso, P.; Gariboldi, P.; Bombardelli, E.; Gabetta, B. Phytochemistry 1992, 31, 4259. Chen, S.-H.; Huang, S.; Wei, J.; Farina, V. Tetrahedron 1993, 49, 2805. Appendino, G.; 0zen, H.~.; Fenoglio, I.; Gariboldi, P.; Gabetta, B.; Bombardelli, E. Phytochemistry 1993, 33, 1521. Appendino, G.; Cravotto, G.; Enrifi, R.; Gariboldi, P.; Barboni, L.; Torregiani, E.; Gabetta, B.; Zini, G.; Bombardelli, E. J. Nat. Prod. 1994, 57, 607.

100 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54.

Barboni, L.; Gariboldi, P.; Appendino, G.; Enrifl, R.; Gabetta, B.; Bombardelli, E. Liebigs Ann. Chem., in press. Appendino, G.; Gariboldi, P.; Pisetta, A.; Bombardelli, E.; Gabetta, B. Phytochemistry 1992, 31, 4253. Appendino, G.; Fenoglio, I.; Cravotto, G.; Varese, M.; Gariboldi, P.; Gabetta, B. Gazz. Chim. hal., 124, 253. Schmid, G.H. Can. J. Chem. 1968, 46, 3415. See also: Potapov, V.M. Stereochemistry; 1979, Mir Publisher, Moscow, p.256. Barboni, L.; Gariboldi, P.; Torregiani, E.; Appendino, G.; Gabetta, B.; Bombardelli, E. Phytochemistry 1994, 36, 987. Min, Z.-D.; Jiang, H.; Liang, J.-Y. Acta Pharm. Sin. 1989, 24, 673. Barboni, L.; Gariboldi, P.; Torregiani, E.; Appendino, G.; Gabetta, B.; Zini, G.; Bombardelli, E. Phytochemistry 1993, 33, 145. Appendino, G.; Gariboldi, P.; Gabetta, B.; Pace, R.; Bombardelli, E.; Viterbo, D. J. Chem. Soc. Perkin Trans. I 1992, 2925. Ettouati, L.; Ahond, A.; Poupat, C.; Potier, P. J. Nat. Prod. 1991, 54, 1455. Beutler, J.A.; Chmurny, G.M.; Look, S.A.; Witherup, K.M.J. Nat. Prod. 1991, 54, 893. Appendino, G.; Lusso, P.; Gariboldi, P.; Bombardelli, E.; Gabetta, B. Phytochemistry 1992, 31, 4259. Graf, E.; Kirfel, A.; Wolff, G.-J.; Breitmaier, E. Liebigs Ann. Chem. 1982, 376. Gu~ritte-Voegelein, F.; Gu~nard, D.; Mangatal, L.; Potier, P.; Guilhelm, J.; Cesario, M.; Pascard, C. Acta Cryst. 1990, C46, 781. Peterson, J.R.; Do, H.D.; Rogers, R.D. Pharm. Res. 1991 8, 908. Wani, M.C.; Taylor,H.L.; Wall, M.E.; Coggon, P.; McPhail, A.T.J. Am. Chem. Soc. 1971, 93, 2325. Miller, R.W.; Powell, R.G.; Smith Jr., C.R.; Arnold, E.; Clardy, J. J. Org. Chem. 1981, 46, 1469. Viterbo, D.; Appendino, G., unpublished results. Shiro, M.; Koyama, H. J. Chem. Soc. (C) 1971, 1342. Begley, M.J.; Freckmall, E.A.; Pattenden, G. Acta Cryst. 1984, C40, 1745. Ho, T.-L.; Lee, G.-H.; Peng, S.-M.; Yeh, M.-K.; Chen, F.-C.; Yang, W.L. Acta Cryst. 1987, C43, 1378.

101 55. 56. 57. 58. 59. 60.

61. 62.

Ho, T.-Y.; Lin, Y.-C.; Lee, G.-H.; Peng, S.-M.; Yeh, M.-K.; Chen, F.-C. Acta Cryst. 1987, C43, 1380. Yoshizaki, F.; Fukuda, M.; Hisamichi, S.; Ishida, T.; In, Y. Chem. Pharm. Bull. 1988, 36, 2098. Castellano, E.E.; Hodder, O.J.R. Acta Cryst. 1973, B29, 2566. Gunawardana, G.P.; Premachandran, U.; Burres, N.S.; Whittern, D.N.; Henry, R.; Spanton, S.; McAlpine, J.B.J. Nat. Prod. 1992, 55, 1686. Fuji, K.; Tanaka, K.; Li, B.; Shingu,T.; Sun, H.; Taga, T. Tetrahedron Lett. 1992, 31, 7915. Chu, A.; Furlan, M.; Davin, L.B.; Zajicek, J.; Towers, G.N.H.; SoucyBreau, C.M.; Rettig, S.J.; Croteau, R.; Lewis, N.G. Phytochemistry 1994, 36, 975. Zhang, S.; Lee, C.T.-L.; Chen, K.; Kashiwada, Y.; Zhang, D.-C.; McPhail, A.T.; Lee, K.-H. J. Chem. Soc. Chem. Commun. 1994, 1561. Chen, S.-H.; Farina, V.; Huang, S.; Gao, Q.; Golik, J.; Doyle, T.W. Tetrahedron 1994, 50, 8633.

The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B.V. All rights reserved

103

3 PACLITAXEL (TAXOL| F O R M U L A T I O N AND P R O D R U G S Dolatrai M. Vyas Bristol-Myers Squibb Company, Pharmaceutical Research Institute 5 Research Parkway, Wallingford CT 06492-7660 U.S.A.

3.1.

INTRODUCTION

Paclitaxel (1.1.1, Figure 1), 1 a natural diterpene isolated from the bark of

Taxus brevifolia (Pacific yew) [1] has been heralded as the antitumor agent of the 1990's because of its promising clinical activity against a variety of human solid tumors such as ovary, breast, lung, head and neck, and melanoma [2-4]. Interest in various aspects of this drug has mushroomed due to the clinical significance of its antitumor profile in treating common h u m a n cancers [5, 6]. Currently, paclitaxel is an FDA-approved cancer agent in the United States for second-line t r e a t m e n t against cis-platinum-refractory ovarian cancer and metastatic breast cancer which is refractory to anthracycline treatment. It has also been approved in other countries for similar indications. On the basis of broader ongoing clinical trials, there is a good likelihood of paclitaxel becoming a first-line chemotherapeutic agent in the very near future [7] . Within the current arsenal of cancer chemotherapeutics [8], paclitaxel is a unique tubulin-interacting agent. Unlike other clinical antimitotic agents such as the vinca alkaloids [9, 10], which inhibit the microtubule assembly process, 1 Taxol | is a registered trademark of the Bristol-Myers Squibb corporation. The generic name "paclitaxer' is used throughout this chapter.

104 paclitaxel promotes tubulin polymerization and stabilizes the resulting microtubules toward depolymerization [11]. This shift in the normal dynamics in the cellular tubulin-microtubule system by paclitaxel is currently widely recognized as its mode of action for cell cytotoxicity. Owing in part to the discovery by Susan Horwitz and coworkers in 1979 [11] of paclitaxel's mechanism of action, the National Cancer Institute (NCI) decided to accelerate the human clinical trials for this drug. Eventual marketing of paclitaxel for oncology clinical use was facilitated by a Cooperative Research and Development Agreement (CRADA) between the NCI and the Bristol Myers Squibb (BMS) Company in 1989 and the subsequent massive efforts by BMS to secure ample clinical supplies of paclitaxel [12].

BzNH O

P. O, -

AcO !II

2

OH

iSSSss

B~

O

Ph O -

HO I I I I

BzO 1.1.1. Paclitaxel

(Taxol)

O

iisiss

BzO 1.1.2. Docetaxel (Taxotere)

Figure 1: Clinicallyrelevant taxanes In spite of paclitaxel's promising clinical antitumor profile and the ongoing efforts to optimize its utility in multimodality treatment for curative cancer therapy, the drug has presented problems during its intravenous (i.v.) administration to patients [13]. These are formulation-related problems stemming from the use of the excipient Cremophor EL | [14] as a solubilizing detergent for i.v. administration. Interestingly, a close analog of paclitaxel, namely, docetaxel, 1.1.2(Taxotere | ) [15] (Figure 1) which is currently awaiting FDA approval for clinical use, is devoid of such problems. Docetaxel possesses a slightly better aqueous solubility than paclitaxel and is administered as a Tween-80 (polyoxyethylene sorbitan monooleate)/ethanol formulation. Several of paclitaxel's alleged formulation-related patient-care issues, such as hypersensitivity reactions, have been successfully addressed and managed in the clinic [16]. However, the pharmaceutical issues [17] related to Cremophor EL's use still persist during intravenous administration and demand precautions

105 and adherence to strict pharmacy protocols. The main intent of this chapter is to document and highlight all of the formulation-related issues stemming from the current clinical formulation and briefly discuss possible solutions sought to alleviate or circumvent them. The quest toward the development of a widely acceptable and safe intravenous formulation devoid of Cremophor EL is continuing in several research laboratories and is discussed herein. 3.2.

PHARMACEUTICAL DEVELOPMENT SUMMARY

Paclitaxel's early pharmaceutical development history [12, 18, 19] is mainly confined to the NCI laboratories, since it was through a joint NCI-USDA (United States Department of Agriculture) initiative in the early 1960's to screen plant material for novel cytotoxic agents that paclitaxel was first discovered. The isolation and identification of this agent were accomplished in 1966 at the NCI contract laboratories of the Research Triangle Institute, North Carolina, by M. Wani and M. Wall, approximately four years after the initial collection of Taxus Brevifolia plant material [18].

The isolation of pure paclitaxel was facilitated through a bioassay-guided fractionation protocol following in vitro cytotoxicity in KB cells and in vivo antitumor activity against murine tumor models such as leukemia P1534 and carcinosarcoma Walker 256. During the next few years, the preclinical antitumor profile of paclitaxel in several of the NCI's murine hematological tumor models, namely leukemias L1210, P388, and P1354, was established. Also, activity against the Walker 256 sarcoma model was demonstrated. These efficacy evaluations were carried out in mice by administering paclitaxel as a suspension in the peritoneal (i.p.) cavity against i.p.-implanted tumors. Due to the extremely hydrophobic character of paclitaxel [20], it was difficult to obtain a purely water-based formulation for i.v. or i.p. administration. Consequently, the vehicles employed for i.p. administration included steroidal suspensions, vegetable oils (peanut, sesame, olive), normal saline and carbomethoxy cellulose (CMC). It was not until 1974 that the first evidence of paclitaxel's efficacy in an i.p./i.p, murine solid tumor, B 16 melanoma, was obtained. Until this time, paclitaxel was considered an unexciting cytotoxic agent with solubility problems and in vivo efficacy confined mainly to i.p./i.p. localized tumor models. Consequently, there was little enthusiasm in pushing this drug further toward h u m a n clinical trials. However, by 1980 several new findings had made paclitaxel a prime candidate for h u m a n clinical trials. In

106 1978, after the introduction of h u m a n t u m o r xenograft models to the NCI screening program, paclitaxel was shown for the first time to possess distal tumor efficacy against the LX-1 lung, the MX-1 breast, and the CX-1 colon xenografts [18]. In all these t u m o r models the drug was a d m i n i s t e r e d subcutaneously to the h u m a n tumor implanted in the sub-renal capsule (kidney) of mice.

More importantly, the excitement and interest about this drug was

heightened in 1979 when a report by S.Horwitz and coworkers [11] disclosed that paclitaxel acted v i a a novel tubulin-interacting mechanism. Unfortunately, at this juncture a suitable formulation for intravenous a d m i n i s t r a t i o n of the drug was lacking and remained to be developed.

At the outset [21], it was

determined at the NCI t h a t paclitaxel was totally devoid of any activity when administered orally to mice up to a dosage of 160 mg/Kg. 3.2.1. Early Formulation Studies Since 1978, a major effort at the NCI was directed at developing an intravenous formulation for paclitaxel in order to initiate h u m a n clinical studies. Paclitaxel's extremely low w a t e r solubility (< 0.01mg/ml), coupled with the absence of a suitable chemical functionality (amine or carboxylic acid) for salt formation, led to the evaluation of cosolvents and excipients as the first strategy for developing an i.v. formulation. Much of this effort is described in detail in a recent report [21]. The approximate solubility of paclitaxel in aqueous vehicles and certain organic solvents are reported in Table 1. Table 1: Solubility of Paclitaxel in Various Solvents a

Solvent

Solubility (mg/ml)

Methylene Chloride

> 19

Ethanol 75% Propylene glycol

39 <1.4

ca.

75% Polyethylene glycol 400 (PEG 400)

31

35% PEG 400

0.03

Soybean oil

0.3

Triacetin

75

(a) adapted from ref. 21.

107 Paclitaxel has substantial solubility in organic solvents such as ethanol and methylene chloride. Highly concentrated (millimolar) paclitaxel solutions in some of these solvents were attainable. However, diluting paclitaxel solutions of water-miscible solvents (such as ethanol) with water presented problems of precipitation. The solubility of paclitaxel in lipids such as soybean oil (intralipid) was also not quite adequate for formulation considerations. The two intravenous formulations which received in-depth evaluation at the NCI were those involving cosolvent PEG(polyethylene glycol) 400 (75%) and surfactant Cremophor EL| [19, 21]. The 75% PEG 400 solution in water containing 16 mg/ml paclitaxel was found to be chemically stable by HPLC analysis and free of particulate material for up to 14 days at 25~ However, upon dilution for infusion, precipitation ( c l o u d i n e s s ) w a s discernible. Formulation with solubilizing surfactant Cremophor EL| (5%) in ethanol (5%) and 0.9% saline under equilibrium conditions gave a solution with paclitaxel concentration of about 0.1 mg/ml. With this formulation, concentrations greater than 0.6 mg/ml were achievable by dilution of a 6 mg/ml solution of paclitaxel in 1:1 Cremophor EL:EtOH. Fortunately, this solution had adequate physical and chemical stability over periods as long as 24 h; >96% of drug was found in solution over this time period. Thus, the Cremophor formulation provided some assurance against the possibility of drug precipitating during the infusion period. Also, this formulation was the best on an efficacy basis, since paclitaxel administered i.p. as an aqueous suspension containing Cremophor EL | was found to be more efficacious against an i.p.-implanted B 16 melanoma tumor than paclitaxel administered i.p. in PEG 400. Due to all these considerations, in 1980 the Cremophor formulation was selected for clinical trials. The current clinical dosage form of paclitaxel consists of a 5ml size vial containing 30 mg of paclitaxel, 2.64 g of Cremophor EL|

and 49.7% EtOH (1:1 v/v).

This

concentrated solution required further dilution with injectable fluids such as 5% dextrose, 0.9% sodium chloride, and 5% dextrose in Ringers solution. The intact vial shelf-life is estimated to be 5 years under refrigeration [21]. 3.2.2. Formulation Issues With the l a u n c h i n g of paclitaxel into clinical trials, several pharmaceutical and patient-care issues surfaced at the very outset. These apparently stemmed from the use of Cremophor EL|

as an excipient in the

intravenous formulation. Cremophor EL| is a chemically and physiologically

108 active surfactant [14]. Thus, the formulation is associated with potential medical liabilities depending upon the amount of Cremophor EL| present and the route by which this formulation is administered to patients. As it currently stands, the paclitaxel clinical formulation has the highest amount of this surfactant among all currently marketed drugs [14]. Other drugs with Cremophor in their formulations are teniposide [22] and cyclosporine [23]. The potential for harmful effects of Cremophor to patients is further augmented by the fact that large quantities of this solubilizing surfactant are required for intravenous delivery of therapeutic doses of paclitaxel ranging from 150 to 300 mg/m 2. P h a r m a c e u t i c a l Issues: P h a r m a c e u t i c a l issues [17] s t e m m i n g from the intravenous administration of the Cremophor EL| formulation have mainly affected hospital health care professionals such as pharmacists and nurses who are required to exercise caution and care during i.v. infusion. Chemical stability studies have shown that the paclitaxel formulation diluted with standard infusion fluids is stable for up to 27 hours at room temperature. This has facilitated the preparation and storage of i.v. solutions in hospital pharmacies. However, due to the leaching properties of Cremophor toward phthalate plasticizers from PVC infusion bags and intravenous administrationset tubings, use of polyolefin plastic containers, glass bottles or vinyl acetatetype bags has been mandated. Also, to guard against the perils of drug precipitation during infusion, paclitaxel is administered with an in-line filtration device containing a 0.22 ~nn membrane filter (e.g. IVEX-2 and IVEX-HP). The issue of paclitaxel compatibility with other medications has also been studied in some detail [24, 25]. An examination of visual and turbidimetric compatibility of paclitaxel with 59 other drugs was carried out. The list of drugs included antimicrobials, anticancer agents, analgesics, antiemetics, and antiallergy medications. The experiment involved the mixing of 1.2 mg/ml solution (4 ml) of paclitaxel in D5W with a 4 ml sample of the test drug at clinically relevant concentrations. All of the drugs but four were compatible with paclitaxel based on the turbidity m e a s u r e m e n t criteria. These included two cancer drugs, mitoxantrone hydrochloride and doxorubicin. In the case of doxorubicin, a loss of 12% paclitaxel in 96 hours was determined by HPLC measurements. It was stressed in this study that the absence of turbidity effects does not automatically imply chemical compatibility with paclitaxel, and caution should be exercised, particularly when need arises for combination therapy with other drugs.

109 Patient Care Issues: In clinical trials paclitaxel has manifested some of the classical drug-related toxicities associated with an anticancer agent [16]. These include neutropenia, neurotoxicity, mucositis, GI toxicities, and alopecia. The major, dose-limiting, toxicity was demonstrated to be neutropenia, which was usually severe at doses >200 mg/m 2. The incidence of severe neutropenia was similar at all dose levels and recovery was complete after 5-10 days. Since hematopoietic growth factors

(e.g. G-CSF) are now commonly used to ameliorate

neutropenia, neurotoxicity remains a major concern in high single and cumulative dose paclitaxel therapy. The neurotoxicity manifested is sensory neuropathy, which is common when paclitaxel is administered by a 24 h infusion and exceeds a dose of 200 mg/m 2. Hypersensitivity reactions, p a r t i c u l a r l y anaphylactoid signs and symptoms, remain a major medical liability and patient-care issue in the clinical use of paclitaxel [26]. The hypersensitivity reaction is alleged to be due to Cremophor EL | contained in the formulation. The symptoms include rapid onset of hypotension, respiratory distress

(e.g. bronchospasms), urticaria, and

rash. These appear to be classical histamine release-mediated reactions, which have also been observed in dogs treated with Cremophor EL| alone [27]. These allergic reactions due to the Cremophor threatened the continuation of earlier paclitaxel clinical trials, due to the death of a patient [7]. However, they were controlled by discontinuation of paclitaxel infusion and emergency treatment of patients with antihistamines, H2-blockers, and oral steroids [13]. Currently, in order to minimize the occurrence and the severity of hypersensitivity reactions in patients, two routine measures have been instituted. These include the extension of the infusion duration time to 6 or 24 h and the prophylactic use of antihistamines and corticosteroids prior to paclitaxel administration [7]. These measures do not completely eliminate the histamine release effects of paclitaxel administration, but they certainly reduce the episodes and severity of hypersensitivity reactions. In spite of the introduction of these safeguards, working with paclitaxel in the hospital setting remains a challenge. A non-Cremophor based intravenous formulation should alleviate the above pharmaceutical and patient-care concerns. However, a suitable clinical substitute has not yet been identified. Several NCI-funded labs and independent research groups are currently actively engaged in research to either develop a safer parenteral administration of paclitaxel through water-soluble prodrugs or to identify a suitable carrier system for i.v. administration.

ll0 3.3. P R O D R U G S OF PACLITAXEL

A thoroughly explored preclinical strategy for i.v. administration of paclitaxel has involved the use of prodrugs in a non-Cremophor-containing, preferably 100% aqueous, formulation [28]. Initially, this effort has involved the synthesis and evaluation of prodrugs with moieties carrying a solubilizing group. In this context, an ideal handle for prodrug synthesis has been the C-2' hydroxyl functionality, since derivatives masked at this position are devoid of activity until unmasked in vivo through hydrolysis by enzymatic or chemical means. A free hydroxyl group at C-2' seems to be a prerequisite for tubulin polymerization and consequently for cytotoxicity [29]. In contrast, the C-7 hydroxyl functionality is not such an ideal site for prodrug design, since masked derivatives (e.g. esters) at this position are usually much more stable than their C-2' counterparts to in vivo cleavage [30]. Consequently, only special C-7 derivatives with a rapid in vivo unmasking pathway have the potential to serve adequately as a prodrug. Pro-moieties which are slow to unmask in vivo may simply act as paclitaxel analogs [31]. 3.3.1. Acyl Derivatives as Prodrugs In 1984 Mellado et al. [32], during their investigation on the biological activity of paclitaxel acetates, established that the C-2' hydroxyl can be readily and selectively acetylated over the C-7 hydroxyl functionality. Furthermore, they found the C-2' acetate to be more labile toward hydrolysis t h a n its C-7 counterpart. These earlier observations formed the basis for considering the C-2' hydroxyl group as the preferred functionality for future research efforts on water-soluble prodrugs [28]. Paclitaxel C-2' Esters: The basic strategy in designing C-2' esters as watersoluble prodrugs has been the incorporation of ionizable groups such as amines, amino acids, and sulfonic acid groups. In this vein, a plethora of C-2' derivatives have been synthesized and evaluated for their suitability as water-soluble prodrugs of paclitaxel. These include (Figure 2) succinate (e.g. 3.1.1 ) and glutarate derivatives (e.g. 3.1.2 ) [33, 34]; sulfonic acid derivatives (e.g. 3.1.3 and 3.1.4 ) [35] and amino acid derivatives (e.g. 3.1.5 and 3.1.6 ) [30, 34]. Although several of these derivatives (e.g. 3.1.1 and3.1.2 ) possess adequate water solubility (up to 1%) for an i.v. formulation and good biological activity in vivo,

lll they were found unsuitable as prodrugs of paclitaxel.

This was attributed to

their chemical instability in aqueous solution, a property deemed unacceptable for intravenous administration of a highly insoluble agent such as paclitaxel. To date, among this class of prodrugs, 2'-[3-(NJV-diethylamino)propionyl]paclitaxel, 3.1.6 , prepared by Stella and coworkers [30], appears to be best suited for prodrug delivery of paclitaxel.

It has acceptable solution stability at pH 3.5

(half life >400 h). However, the half life at physiological pH (7.4) was <30 min. In h u m a n p l a s m a the h a l f life was even shorter (< 5 min).

The aqueous

solubility of this prodrug was determined to be >10 mg/ml. In vivo, 3.1.6 showed complete tumor remission in the MX-1 m a m m a r y tumor model.

Ac~~,

BZ~NH 0 Ph

-

/jO OH

0 ....

OR

HO

-: AcO OBz

3.1.1 R= CO-(CH2)2-COOH (HOCH2CH2)3N R= CO-(CH2)3-COONa 3.1.3 R= CO-(CH2)2-SO3Na 3.1.4 R= CO-(CH2)2-CONH-CH2-SO3Na 3.1.5 R= CO-(CH2)2-NH2-HCO2H 3.1.6 R= CO-(CH2)2-NEt2. CH3SO3H 3.1.7 R= COCH2OCH2CO2H 3.1.8 R= COCH2SCH2CO2H 3.1.9 R= COCH2S(O2)CH2CO2H 3.1.10 R= CO-(CH2)2-CONH(PEG) 3.1.2

Figure 2: Water-soluble C-2' esters as paclitaxel prodrugs More recently, a Scripps group led by Nicolaou has reported another series of C-2' esters n a m e d 'protaxols' [36]. These are monoesters (3.1.7, 3.1.8 and 3.1.9) of dicarboxylic acids incorporating a heteroatom functionality (oxygen or sulfur) for the purpose of imparting further water solubility. These compounds have been reported to have acceptable water solubility (ca. 1 mg/ml) and stability at neutral pH and room t e m p e r a t u r e .

Under basic conditions, 3.1.9 rapidly

112 generated paclitaxel.

The authors envisage that such a mechanism may be

relevant in vivo in the basic microenvironment of certain tumor cells. However, in vivo antitumor activity of these protaxols in murine tumor models have not yet been reported by the Scripps group.

Thus far all of the above approaches have only yielded prodrugs with moderate (up to 10 mg/ml) water solubility. However, conjugates of hydrophobic molecules with polyethylene glycol (PEG) of molecular weight of 2kD or greater are known to impart even greater water solubility. In this context, Greenwald and co-workers [37] have recently reported the synthesis and limited biological evaluation of a variety of C-2' polyethyleneglycol esters as highly water-soluble prodrugs of paclitaxel (3.1.10). The PEG esters reported are claimed to have water solubility of >666 mg/ml at ambient temperature. The half life of 3.3.10 was determined to be approximately 4.0 h and 1.1 h in pH 7.4 buffer and rat plasma, respectively. The authors have not reported any in vivo antitumor activity, but the approach seems very promising. Paclitaxel C-7 Esters: In general, C-7 esters have found little utility as watersoluble prodrugs of paclitaxel. To date, several C-7 derivatives have been reported [30] and in most cases they have been the counterparts of their corresponding C-2' derivatives. Their poor prodrug properties have been attributed to enhanced in vivo stability cleavage.

toward esterases and hydrolytic

For example, the cationic water-soluble derivative 3.1.11(Figure 3) had a half life of 378 h at pH 3.8 compared with a half life of of 96 h for its corresponding C-2' ester derivative. A similar trend was observed at physiological pH (7.4), where 3.1.11 had a half life of 34 h compared to 6 h for its C-2' counterpart. Half lives in human plasma for 3.1.11 and its C-2' counterpart were reported to be 3 h and <30 min, respectively. The enhanced hydrolytic (chemical and enzymatic) stability of C-7 esters over their C-2' counterparts can be ascribed to the steric congestion surrounding this position. Biologically, unlike the C-2' esters, cationic water-soluble C-7 esters such as 3.1.11 were shown to promote microtubule assembly as effectively as paclitaxel, but have been found to be poorly bioactive in whole cell assays. This was attributed to poor cell membrane penetration.

113

BZ"NH O P h ~ O

()H

AcO

O OCOCH2NMe2 MeSO3H

....

3.1.11 OBz

Figure 3: Typical C-7 ester prodrug of paclitaxel

Paclitaxel C-2' Carbonates: Recently, the BMS group and the Scripps group have demonstrated that C-2' carbonates, like their ester counterparts, behave as paclitaxel prodrugs in vivo. Carbonates and carbamates have so far found little utility as prodrugs because they are more stable than the corresponding esters toward hydrolytic and enzymatic cleavage. Notable exceptions in this area are carbamate derivatives of drugs such as 10-hydroxy-camptothecin [38] and CC 1065 [39], which have been shown to be effective prodrugs in vivo. The BMS group has recently disclosed the synthesis and in vivo biological profile of C-2' carbonates derivatives 3.1.12-17 (Figure 4)[40].

BZ.NH

Ph

O

AcO

O

OH

3.1.12 R= OH3

O ....

o),= RO

O

HO

: AcO OBz

o

3.1.13 3.1.14 3.1.15 3.1.16 3.1.17

R= CH2CH3 R= CH(CH3)2 R=CH2Cl R=CH2Ph R=CH=CH2

Figure 4: C-2' carbonate derivatives synthesized by the BMS group When evaluated in the cytotoxicity assay, paclitaxel-2'-carbonates were 210 times less cytotoxic than paclitaxel against human colon cancer cell line (HCT 116). Also, they were found to be inactive in the tubulin polymerization assay. However, after incubation in rat plasma at 37 ~ for 18 hrs, some of these carbonate derivatives, particularly 3.1.12 and 3.1.13, were found to promote microtubule assembly, indicating the generation of the parent compound, paclitaxel, in rat plasma. All of the above observations were fully substantiated

114 by t h e i r in vivo a n t i t u m o r activity in the Madison 109 m u r i n e lung carcinoma (M109) t u m o r model [41], as s u m m a r i z e d in Table 2. In this t u m o r model both the p r o d r u g and the t u m o r were localized i n t r a p e r i t o n e a l l y (i.p.), a n d as such r e p r e s e n t e d an 'in vivo test-tube' model. All carbonates were effective (%T/C > 125% is considered to be active) in increasing life span. The m e t h y l carbonate 3.1.12 was less active t h a n paclitaxel, b u t all the o t h e r c a r b o n a t e s exhibited comparable in vivo a n t i t u m o r activity w i t h respect to paclitaxel. These in vivo results indicated t h a t paclitaxel-2'-carbonates are converted to the p a r e n t drug u n d e r the in vivo conditions and act as true prodrugs of paclitaxel. Table 2: In vivo A n t i t u m o r Activity of Paclitaxel-2'-Carbonates 3.1.12-17 In vivo A n t i t u m o r Activity a %T/C (mg/K~inj ection) Compound

Carbonate Deriv.

Paclitaxel

3.1.12

162% (90 mg/Kg) b

276% (75 mg/Kg) b

3.1.13

>475% (60) c

275% (30) c

3.1.14

247% (100) b

197% (50) b

3.1.15

275% (60) c

275% (30) c

3.1.16

310% (50) c

270% (50) c

3.1.17

>475% (60) c

275% (30) c

a Madison 109 murine lung carcinoma (M109) i.p.implant model. Drugs administered i.p. in 10% Tween 80 in saline (paclitaxel), in 10% DMSO/saline (3.1.13-17), or in 10% DMSO/H20 plus a few drops of Tween 80 (3.1.12). %T/C refers to the percentage of the median survival time of drug-treated mice (six per dose) vs. saline-treated control. The %T/C values are determined at the maximum tolerated dose (shown in brackets). %T/C > 125% is defined as active in this tumor model, b Dose administered i.p. on days 5 and 8. c Dose administered i.p. on days 1, 5, and 9. One obvious drawback of the above carbonate derivatives is their extreme insolubility in w a t e r .

As will be discussed later,

a water-soluble prodrug

s t r a t e g y for one of these c a r b o n a t e s was successfully developed by a t t a c h i n g soluble pro-moieties at the C-7 h y d r o x y l functionality.

In a s i m i l a r vein,

Nicolaou a n d c o w o r k e r s h a v e r e c e n t l y r e p o r t e d novel w a t e r - s o l u b l e C-2' arylsulfonyl e t h y l c a r b o n a t e s 3.1.18-20(Figure 5) [36].

These compounds were

p r e p a r e d in order to evaluate a novel m e c h a n i s m of paclitaxel release in vivo. Accordingly, they were conceived to generate paclitaxel in vivo t h r o u g h a base-

115 induced ~-elimination reaction. However, these authors have not yet reported whether these putative prodrugs are actually effective in vivo.

AcO ~,

BZ~NH O Ph

0 ....

~

RO

o

O .. // .uH

,

7~O

0

/2

~

a.1.19 R -

\\ //

NO2

0

OBz

3.1.20R= ~~

\\~ ' ~//

NH2

O Figure 5: Paclitaxel 2'-carbonates prepared by the Scripps group 3.3.2. Phosphate Esters Derivatives as Prodrugs The Bristol Myers Squibb (BMS) group elected, for reasons of novelty, to focus on the synthesis and evaluation of water-soluble phosphatase-cleavable prodrugs of paclitaxel [28]. This strategy mandated the synthesis and evaluation of hitherto unknown phosphate esters of paclitaxel. Use of phosphate derivatives as water soluble-prodrugs of clinically useful drugs such as etoposide [42], clindamycin [43], and mustards [44] has been well documented. The rationale behind their use as prodrugs stems from the ubiquitous nature of phosphatase enzymes in mammalian systems [45]. Interestingly, certain tumors are shown to express high levels of membrane-bound alkaline phosphatases and thus provide an opportunity for selective cleavage and accumulation of these derivatives at the tumor site [46]. In the BMS prodrug program, the initial targets synthesized were the obvious prototypic C-2' and C-7 phosphate derivatives 3.2.1 and 3.2.2 (Figure 6) [47]. Although the sodium salts of 3.2.1 and 3.2.2 were endowed with impressive water solubility (~10 mg/ml), their in vitro and in vivo evaluation indicated that they were poor prodrugs of paclitaxel. The first indication of this was evident when both derivatives failed to generate paclitaxel in vitro upon treatment with purified preparations of bovine intestinal alkaline phosphatase.

116

gZ'NH

AcO ~'

O

O / / OR2

3.2.1 RI=PO(OH)2; R2=H 3.2.2 RI=H; R2=PO(OH)2

....

OBz Figure 6: Prototypical paclitaxel phosphates This was further corroborated by their extreme stability and failure to generate paclitaxel in r a t plasma. Also, both were inactive in promoting microtubule assembly in vitro. I n vivo evaluation of sodium salts of 3.2.1 and 3.2.2 against the i n t r a p e r i t o n e a l (i.p.) Madison-109 m u r i n e (M109) lung t u m o r model in comparison with paclitaxel d e m o n s t r a t e d (see Table 3) t h a t these analogs possessed m a r g i n a l a n t i t u m o r activity at best (in this model %T/C > 125 is considered active). Table 3: In vivo Activity of Sodium Salts of 3.2.1 and 3.2.2 in the M109 Tumor Modela

Maximum T/C b (mg/kg/inj.) Experiment

Compound

Phosphate

PaclitaxelC

id

3.2.1 (Na salt)

140(25)

270(50) e

2d

3.2.2 (Na salt)

123 (30)

190(10)

aMurine lung carcinoma i.p. implant model, bT/C refers to the percent of the median survival time of drug-treated mice (6 per dose) vs. saline-treated controls, at the maximum tolerated dose (in brackets). CAdministered in 10% Tween 80 in saline, dDose administered i.p. on days 1, 5 and 9. eAlso, a dose of 25 mg/kg/inj, achieved a T/C of 240 with 1/6 mice cured. The lack of in vitro enzymatic cleavage and inferior in vivo activity of 3.2.1 and 3.2.2 indicated t h a t these p h o s p h a t e s are poor s u b s t r a t e s for alkaline phosphatase. Sterically, both the C-2' phosphate and the C-7 phosphate moieties in 3.2.1 and 3.2.2 are quite close to the congested taxane core. Consequently, it is likely t h a t the p h o s p h a t a s e enzymes are not able to easily bind these functionalities and process them effectively to generate paclitaxel. In order to

117 address the steric barrier encountered in prototypes 3.2.1 and 3.2.2 toward phosphatase cleavage, the BMS chemists pursued a more complex strategy, i.e. the synthesis of pro-prodrugs of paclitaxel which can be activated by phosphatase (Figure 7).

0 Aoo

o

o

P h ~ O

....~

..... ~

phosphatase

l o'Oo

OBz 3.2.3

BZ'NH O spont.

phil'__

3.2.4

AcO~ /jO OH

~

= self- immolating linker

0 ....

ON OBz 1.1.1

Figure 7: Pro-prodrug strategy based on phosphatase enzymes. Pro-prodrug strategies have been sucessfully applied in medicinal chemistry [48]. The two most widely employed concepts in the design of pro-prodrugs have been the exploitation of the fragmentation cascade [49] and of the lactonization reaction [50]. Adaptation of these strategies to paclitaxel entails the design and use of appropriate self-immolating linkers at either C-2' or C-7 (as depicted in Figure 7). The concept involves dephosphorylation of the pro-prodrug 3.2.3 by phosphatases in vivo, followed by the rapid collapse (through a fragmentation cascade reaction or lactonization step) of transient dephosphorylated intermediate 3.2.4 to yield paclitaxel 1.1.1. The lactone approach has been previously exploited by medicinal chemists in prodrug design, since the factors

118 that influence the rates of lactonization reactions are generally well understood [51].

~

0

N

NH2

0

OMe

i

+

Esterase OMe

3.2.5

3.2.6

3.2.7

Figure 8: A self-immolative linker approach to drug delivery A prototypic pro-prodrug design to demonstrate the 'proof of principle' was based on the 'trimethyl lock' linker recently reported by Borchardt and coworkers [52] in the design of an esterase cleavable pro-prodrug of an amine (Figure 8). It was elegantly demonstrated by this group through in vitro experiments with isolated enzyme preparations and plasma studies that 3.2.5 was an effective pro-prodrug of p-methoxy aniline 3.2.7. The success of this approach was ascribed to the phenomenal lactonization rate enhancement (of the order of > 105) achieved by the presence of methyl substituents on the alkyl chain and the aromatic ring. The estimated half life of lactonization for the desacetyl analog of 3.2.5 is of the order of 1 min. Extension of this strategy to phosphate derivatives required the synthesis of acid linker 3.2.8 (Figure 9).

This was then utilized in the synthesis of the

desired targets 3.2.9 and 3.2.1{} [53]. The sodium salts of 3.2.9 and 3.2.10 were found to possess acceptable water solubility (ca. 10 mg/ml) for i.v. evaluation of these prodrugs in murine tumor models. Solution stability determinations in pH 7.4 Tris buffer (50mM) at 37 ~ were substantially different. T90 values, defined as the time required for the compound to undergo 10% degradation, were determined to be ca. 20 h for 3.2.9 and > 100 h for 3.2.10. Thus, 3.2.9 may lack the stability necessary for intravenous administration without encountering precipitation problems during i.v. infusion. An in-depth in vitro evaluation of these two pro-prodrugs in biological fluids and with pure enzyme was undertaken

to establish

all the

necessary p a r a m e t e r s .

Also, kinetic

investigations of the cleavage process using bovine intestinal AP and p-

119 nitrophenol phosphate (PNP) as a reference standard were performed. The Vmax/Km ratio for the prodrugs was determined and compared with that of PNP as a measure of enzyme efficiency with these substrates. It was found that 3.2.10 had Vmax]Km = 0.6 (vs. PNP=I), whereas Vmax]Km for 3.2.9 was only 0.13. This may indicate that the enzyme is confronted with lesser steric congestion around C-7 than C-2'. Interestingly, the self-immolation process for the linker to generate paclitaxel after dephosphorylation by the AP in both cases was rapid (ty2<5 min), in agreement with literature precedents.

AcO

O ...

BZ'NH O (OH)

O

ph/~g~2' O.... -

ttSs S

O

H

Ac()~ / O OBz

3.2.8

3.2.9 (HO)2OP

\ O ~/ BZ'NH O P h @ O

~ PO(OH)2

AcO~ 2 0 ~ ~ ' / / ~ ....

OH

10 OBz

Figure 9: Phosphate pro-prodrugs of paclitaxel The above in vitro observations for the two phosphate derivatives were further substantiated by their in vivo antitumor performance. When evaluated against the i.p/i.p. M109 tumor model, the C-2' ester/phosphate 3.2.9 was found to be marginally active, whereas the C-7 ester/phosphate 3.2.10 was as active as paclitaxel (Table 4). The above results led to the exploration of other self-immolating linkers, with the aim of establishing a relationship between rates of release of paclitaxel

120

in vitro a n d p r o - p r o d r u g s p e r f o m a n c e in vivo. C o n s e q u e n t l y , acids 3.2.11 - 13, c a r r y i n g a p r o t e c t e d p h o s p h a t e functionality, were s y n t h e s i z e d a n d studied for t h e i r s u i t a b i l i t y as self-immolating linkers in p r o - p r o d r u g s of paclitaxel (Figure 10) [28]. Table 4 : In vivo A n t i t u m o r Activity of C o m p o u n d s 3.2.9 a n d 3 . 2 . 1 0 i n the M109 T u m o r Model. a %T/C b (mg/Kg/inj .) Experiment

Compound

Prodrug

Paclitaxel e

Ic 2d

3.2.9 (Na salt)

144 % (100)

275 % (30)

3.2.10 (Na salt)

156 % (140)

144 % (40)

153 % (70) a Murine lung carcinoma, i.p. implant model, b T/C refers to the percentage of the median survival time of drug-treated mice (six per dose) vs. saline-treated controls, c Dose administered i.p. on days 1,5 and 9. Vehicle: water, d Dose administered i.p. on days 5 and 8. Vehicle: water. e Administered in 10 % Tween 80 in saline.

OPO(OBn)2 0 (OBn)2OPO-~~~O H

H

3.2.11 3.2.12 PO(OBn)2

COOH 3.2.13

Figure 10: New self-immolating linkers for paclitaxel delivery

Gem-dimethyl y-hydroxybutyric acid l i n k e r 3.2.11 w a s selected b a s e d on the k n o w n fact t h a t the r a t e of lactonization is e n h a n c e d by gem-dialkyl groups

121 because of the Thorpe-Ingold effect [51]. The choice of o-hydroxyphenylacetic acid linker 3.2.12 was based on the published study on the rates of lactonization of o-substituted phenylacetic acids [54]. The o-hydroxymethylbenzoic acid linker 3.2.13 was also considered for the study because, according to Fife and Benjamin [55], its rate of lactonization was estimated to be intermediate between that of the 'trimethyl lock' linker 3.2.8 and that of the 3.2.12.

AcO

O ...-.

BZ'NH O p h ~ ~ Z-

0,,, i

ss#sss

OBz

3.2.14

R I = I " ~ ~....~ A ... OPO(OH)2 R2=H ,, / \ O

3.2.15

RI=H

R2= l ~ O P O ( O H ) 2 O OPO(OH)2

OPO(OH)2 3.2.17

RI=H

3.2.18

RI= l

R2= ~ [ ~ ~ " y _ ~,%j

O

u

t

R2=H OPO(OH)2

3.2.19

RI=CO2Et R2= OPO(OH)2

Figure 11: Potential pro-prodrugs of paclitaxel containing self-immolating linkers

122 The potential pro-prodrugs of paclitaxel t h a t were synthesized and evaluated in vitro and in vivo are shown in Figure 11 [28].

Evaluation of in vitro p a r a m e t e r s for pro prodrugs 3.2.14-19 included m e a s u r e m e n t of their w a t e r solubility, solution stability m e a s u r e d as time required (t90) at 37 ~ and pH =7.4 for 10% decomposition of the prodrug, relative Vmax/Km ratios determined from enzyme kinetics (Bovine AP) and rates of linker self-cleavage (i.e. lactonization rates). Table 5 lists these properties for the pro-prodrugs evaluated. Table 5: In vitro Parameters for Pro-Prodrugs 3.2.14-19.

Compound

Water-solubility (mg/ml)a

Stability (t90)b

Linker SelfCleavage (tl/2) c

Relative Vmax/Km e

3.2.14

>10

8.3h

<5min

0.21

3.2.15

>10

23h

-80h

0.17

3.2.16

> 10

4 lh

<5min

0.91

3.2.17

5.5

15.2h

>200h

1.47

3.2.18

5.0 d

20.5h

<5min

0.50

3.2.19

2.5 d

60h

ND

ND

a Determined on the sodium salts by HPLC of saturated solutions after filtration through 0.2 ~m Nylon filters, bDetermined in 50mM Tris buffer, pH=7.4. CAt pH=7.4 at 37 ~ dMono-potassium salt was used. eBovine alkaline phosphatase (Sigma) in 50mM Tris buffer containing 1% DMSO, pH=7.4 at 37 ~ The Vmax/km ratio for reference PNP is set =1. The in vivo evaluation of these putative pro-prodrugs was carried out by a more 'realistic' protocol, [56] in comparison to t h a t employed in evaluating prodrugs 3.2.9 and 3.2.10, i.e. a distal tumor model was employed. The t u m o r was i m p l a n t e d s u b c u t a n e o u s l y (s.c.) in CDF1 mice.

The

animals received their drug treatments via intravenous (i.v.) injections on a five day consecutive daily t r e a t m e n t schedule, beginning on either day 4 or day 5 post-tumor implant. levels.

Each derivative was evaluated at a m i n i m u m of three dose

Antitumor activity was judged by the relative median time for tumors to

reach 1 g in drug-treated as compared to control groups (i.e. T- C values, in days).

123 A drug was considered active by this p a r a m e t e r if it produced a T- C value of > 4 days. Results are summarized in Table 6. Table 6: I n v i v o Antitumor Activity of Paclitaxel Phosphates 3.2.14-19 Against S.c.-Implanted Madison 109 Lung Carcinoma. Delay in Tumor Compounda

Optimal dose b

3.2.14

61

Growth-(T-C) days c Prodrug Paclitaxel 9.0

14.5

3.2.15

31

8.3

14.0

3.2.16

34

16.5

15.8

3.2.17

61

8.0

15.8

3.2.18

23

6.0

14.0

3.2.19

40

17.0

10.8

aConsecutive daily treatment schedule beginning on day 4 post-tumor implant, bHighest nontoxic dose (as sodium or potassium salt) given intravenously (in mg/kg/inj.). CRelative median time for tumors to reach 1 g in drug treated vs. control groups (8 mice per dose). All of the pro-prodrugs, in the form of salts, were endowed with acceptable water solubility for i.v. administration in water. Based on a stability criterion of t90 >24 h, only derivatives 3.2.16 and 3.2.19 qualify as suitable candidates for f u r t h e r evaluation, since t h e y are least likely to p r e s e n t with precipitation problems during i.v. infusion. Interestingly, these two derivatives were endowed with antitumor activity comparable to t h a t of paclitaxel under the same protocol. All the other derivatives tested produced delays in t u m o r growth t h a t were inferior to those caused by paclitaxel.

Compounds showing comparable

activity to paclitaxel reflect the fact t h a t self-immolating linkers associated with these derivatives were effective (see self-cleavage r a t e s in Table 5).

In this

context it is noteworthy t h a t o-hydroxymethyl benzoic acid linker 3.2.13 is a viable linker for pro-prodrug design even though its rate of lactonization (selfimmolation) is slower t h a n t h a t of the ' t r i m e t h y l lock' l i n k e r 3 . 2 . 8 .

On

comparison of the self-cleavage h a l f lives of linkers for the pairs of prodrugs 3.2.14 vs. 3.2.15, and 3.2.16 v s . 3.2.17, it appears t h a t linkers at C-2' are much more labile t h a n their C-7 counterparts. The i n v i v o profile of 3.2.1{} and 3.2.19 raises i n t e r e s t i n g speculations as to their m e c h a n i s m of activation.

In 3.2.1{}

124 either activation by an esterase or phosphatase may be responsible for generation of paclitaxel in vivo. The activity seen in 3.2.19, however, is mainly due to activation by two different enzymes; an esterase for cleavage of the C-2' carbonate moiety and a phosphatase for the cleavage cascade at C-7. The above limited examples pave the way for further development of phosphatase-cleavable pro-prodrugs of paclitaxel and/or other sophisticated studies in prodrug design. The clinical efficacy of either esterase- or phosphatase-based prodrugs remains to be established. Both approaches appear promising and will require further pharmaceutical effort and thorough biological optimization (i.e. dose and schedule variations) to select a candidate worthy of clinical evaluation. 3.4. M I S C E L L A N E O U S P R O D R U G S

As evident from the above discussions, the major efforts toward the development of water-soluble prodrugs of paclitaxel have focused on the study of either acyl or phosphate derivatives. The intended in vivo activation mode for these prodrugs is enzymatic (hydrolysis). Recently, Nicolaou and co-workers have reported a novel class of C-2' derivatives, e.g. paclitaxel-2' methyl pyridinium acetate 4.1.1 as water-soluble prodrugs of paclitaxel [57]. These preliminary studies have shown that this compound has high water solubility (>20mM) and the desired paclitaxel-releasing properties. For example, in human plasma at 37 ~ conversion of 4.1.1to paclitaxel is reported to be complete in <10 min. The u n m a s k i n g of paclitaxel in vivo is envisaged as a result of a nucleophilic attack by water or any bio-nucleophile at the 2-position of the pyridinium moiety. In water and buffered solutions (pH 6.0-7.4) at ambient temperature 4.1.1 is stable for several days and displays aggregation properties resulting in colloidal solutions. In vivo experiment (subcutaneously implanted PC-3 prostate carcinoma) showed that 4.1.1 (injected intraperitoneally) exhibits tumor growthinhibitory properties. For example, at the maximum tolerated dose (MTD) 4.1.1 was reported to be effective (4/8 animals were tumor-free), whereas paclitaxel recorded deaths (4/8) due to toxicity. In a separate report [58] by the same group, a paclitaxel C-7 methyl pyridinium salt analogous to 4.1.1was disclosed as a highly stable and water-

125 soluble derivative.

This derivative, due to its extreme stability in vivo, was

deemed unsuitable as a prodrug of paclitaxel.

Bz~NH O

Ph~O

AcQ

O

OH

....

o

[~[/+

AcO-

~ , . ~ N2Me

4.1.1 Figure 12: Methylpyridinium ether prodrug of paclitaxel 3.5. L I P O S O M E F O R M U I ~ T I O N S In recent years liposomes have received great attention as drug delivery vehicles for a variety of pharmaceutically important drugs [59]. In 1987, Tarr and co-workers were the first to report the formation of stable emulsions of paclitaxel in 50% triacetin as an attempt to develop alternative non-Cremophorbased formulations for i.v. administration of paclitaxel [60]. Although stable emulsions were reported at paclitaxel concentrations of 10-15 mg/ml of emulsion, triacetin itself was toxic to mice when administered i.v. in concentrations required to deliver therapeutic doses of paclitaxel. Subsequently, in 1990 Bartoli et al. reported in vitro and in vivo antitumor experiments with paclitaxel encapsulated both in liposomes and in nanocapsules [61]. In vivo experiments in P388 m u r i n e l e u k e m i a (i.p. implanted) model were carried out by administering paclitaxel i.p. in the carrier system under study. Paclitaxel encapsulated in liposomes was found to be as effective as free paclitaxel, whereas nanocapsules proved to be toxic. The real significance of the above formulations for clinical development has been difficult to gauge, due to the absence of in vivo results in realistic distal site tumor models. Most recently, the group led by Straubinger has reported significant progress towards developing a liposome-based i.v. formulation for delivery of paclitaxel [62]. This group has systematically studied over 300 sets of liposome

126 formulations to arrive at the optimal one for clinical studies. The formulations with a molar ratio of 1:33 of paclitaxel to phospholipid prepared from phosphatidylglycerol (PG) and phosphatidylcholine (PC) (1:9 molar ratio) were found to possess suitable chemical and physical stabilities for in vivo evaluation. A murine colon tumor (C-26) model which was highly resistant to paclitaxel in vitro was developed for in vivo evaluation of these formulations. Notably, the in vivo tumor model was a distal tumor model where the tumor was implanted

subcutaneously and the liposomal formulations administered intravenously. The results in this model established that the liposomes were equipotent or slightly more potent than the current clinical formulation containing Cremophor EL. The MTD for the liposome formulation was greater (2 to 7 fold) vs. that of free paclitaxel. This formulation allowed the observation of an antitumor effect at dose levels ca. 30% higher than those which would have been lethal using free paclitaxel. Further study of this promising liposome-based formulation in clinical trials will be a logical step in the evaluation of these alternative delivery systems. 3.6. C O N C L U S I O N S

In summarizing the progress to date, it appears that preclinical research efforts at developing a safer intravenous delivery of paclitaxel has continued unabated and in many directions, and several promising alternatives to the current formulation are available for evaluation in h u m a n clinical trials. Whether removal of Cremophor in the newer delivery systems will provide the oncologist with the opportunity to fully explore the dose-response profile of paclitaxel remains to be seen. Certainly, the presence of Cremophor has hindered the clinical efforts to develop optimal schedules and dose regimens for paclitaxel adminstration to patients. Several infusion schedules (3, 24 and 96 h) have been investigated randomly to maximize benefit and minimize adverse effects of Cremophor to patients [7, 63]. The role of Cremophor paclitaxel cytotoxicity has also been recently discussed and Paclitaxel has been shown to be an inducer and substrate resistance (MDR) in cultured cells [64-66]. There is considerable

in enhancing investigated. of multidrug debate on the

clinical contribution of Cremophor in the current formulation for enhancing the cytotoxicity of paclitaxel through the inhibition of MDR (see also chapter 7 for a full discussion). Unfortunately, the clinical relevance of this mechanism of

127 resistance for paclitaxel is unclear and is only now being probed [67]. The cytotoxicity of paclitaxel against resistant cell lines has been enhanced by concurrent exposure with agents known to reverse the effects of MDR [68]. Cremophor EL| is a well known MDR-reversing agent and its application in reversing MDR has been demonstrated in vitro for paclitaxel [69]. Also, a recent publication has reported that levels of Cremophor administered to patients through the current formulation are sufficient to achieve levels found to be efficacious for the reversal of MDR in vitro [70]. It is premature to predict whether or not any of the approaches reviewed here will ultimately succeed in supplanting the current paclitaxel formulation. It is clear that the development of second-generation analogs of paclitaxel, which may be inherently endowed with the same water-solubility problems as paclitaxel, will certainly benefit immensely from the present advances in the art. REFERENCES

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The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B. V. All rights reserved

131

4 METABOLISM AND P H A R M A C O L O G Y OF T A X O I D S M. Wright, B. Monsarrat, I. Royer Laboratoire de Pharmacologie et Toxicologie Fondamentales, CNRS, 205 route de Narbonne, 31400, Toulouse, France

E.K. Rowinsky, R.C. Donehower Division of Pharmacology and Experimental Therapeutics, Johns Hopkins Oncology Center, Baltimore, MD 21287, U.S.A.

T. Cresteil INSERM U75, Universit6 Ren6 Descartes, 75730 Paris cedex 15, France

D. Gu6nard Institut de Chimie des Substances Naturelles, CNRS, 91190 Gif-sur-Yvette, France 4.1. INTRODUCTION Among taxoids, paclitaxel 1 (Taxol | and more recently docetaxel (Taxotere | (Figure 1) have been submitted to clinical trials. The 1Taxol is a registered trademark of the Bristol-Myers Squibb corporation. The generic name paclitaxel is used throughout this chapter. Taxotere is a registered trademark of Rhone-Poulenc Rorer. The generic name docetaxel is used in this chapter.

132 u n d e r s t a n d i n g of t h e i r a n t i n e o p l a s t i c activity in h u m a n s and the i m p r o v e m e n t of their use in the clinic require knowledge of the main characteristics of their disposition. During early clinical trials, the pharmacokinetics of paclitaxel and docetaxel have been studied [1, 2]. In contrast, metabolic investigations have been slow due to the necessity to purify taxoid derivatives in sufficient quantities from the bile and/or from the feces for chemical characterization by mass and NMR spectroscopy. This time-consuming step was required for the description of possible metabolic processes, such as epimerization, hydrolysis, hydroxylation, sulfation and glucuroconjugation. Moreover, the purification of the metabolites in sufficient quantities [3, 4] and/or their synthesis [5] were necessary in order to determine the cytotoxicity. Once these questions were answered, it became possible both to characterize the enzymes involved in the metabolism of paclitaxel and docetaxel and to determine whether studies aimed at further modulating the effects of metabolism on clinical activities had to be restricted to humans or could be performed on convenient animal models.

AcO

O

BzNH O ph~.~'-O ~ -

Boc.

HO

O

NH O ,,

i i

oo

ph/~~O...

ee

-

H

BzO

so S s s

AcO" ~ 0

1.1.1. Paclitaxel

(Taxol)

i

BzO 1.1.2. Docetaxel (Taxotere)

Figure 1. Chemical structures of paclitaxel and docetaxel.

4.2. TAXOID DERIVATIVES O B S E R V E D IN BIOLOGICAL F L U I D S

Three main modifications of taxoids have been shown to occur when they are i n t r o d u c e d in the organism: epimerization, hydrolysis, and hydroxylation (Figures 2 and 3).

133 4.2.1. E p i m e r i c Derivatives The e p i m e r i z a t i o n of paclitaxel to 7-epi-paclitaxel (2.1.1, F i g u r e 2) h a s been r e p o r t e d by Ringel a n d Horwitz [6] in cell culture, w h e r e it c o n s t i t u t e d the major paclitaxel derivative. It w a s detected b o t h in the m e d i u m a n d inside the cell. After 72 h of i n c u b a t i o n , the ratio of 7-epi-paclitaxel to paclitaxel v a r i e d from 0.36 to 0.48 in the cell a n d from 0.25 to 0.32 in the m e d i u m , d e p e n d i n g on the cell line a n d on the initial concentration of paclitaxel.

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10-Deacetyltaxol

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Figure 2 : Chemical structures of paclitaxel metabolites. Metabolites I, V, VI, VII are observed in rat bile. Metabolites VI', VII', VIII' in human bile.

134 The epimerization of paclitaxel occurred in the medium in the presence or absence of cells. Moreover, this process was reversible, since 7-epi-paclitaxel was partially epimerized to paclitaxel when incubated in the medium. The presence of 7-epipaclitaxel in the cells may be the result of the similar affinities of 7-epi-paclitaxel and paclitaxel for microtubules, as demonstrated by displacement experiments [6]. 0

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Figure 3: The metabolic oxidation of docetaxel Surprisingly, 7-epi-paclitaxel was not detected in the bile and in the urine of either rats or h u m a n patients treated with paclitaxel [3, 4, 7]. Using the HPLC/APCI-MS method, M o n s a r r a t et al. recently characterized 7-epipaclitaxel in both plasma and urine of patients receiving a 3 h infusion of paclitaxel [8]. The quantity of 7-epi-paclitaxel in urine accounted for 0.7% of

135 total urinary excretion of paclitaxel. In plasma, the maximal concentrations were observed at the end of the infusion period, reaching 2.3-6.0 pM for paclitaxel and 0.11 ~M for 7-epi-paclitaxel. Thus, the plasma concentration of 7-epi-paclitaxel remained 20- to 55-fold lower t h a n the concentration of paclitaxel. The absence of 7-epi-paclitaxel in h u m a n bile and its low concentration in plasma and urine might suggest little epimerization of paclitaxel in vivo. However, using the urine collected from patients 16-24 h after the infusion period, Monsarrat et al. observed that 7-epi-paclitaxel may correspond to 2% of the paclitaxel derivatives excreted in the urine [8]. The effect of microtubules on the epimerization of paclitaxel has been studied by Ringel and Horwitz [6]. They incubated 10 ~M paclitaxel for 24 h in minimal essential cell culture medium with or without 10% fetal bovine calf serum in the absence and in the presence of calf brain microtubule proteins (2 mg/ml). The ratio of 7-epi-paclitaxel to paclitaxel was higher in the absence of microtubule proteins (0.15 and 0.16, respectively) than in their presence (0.04 and 0.07, respectively), both with and without serum. Since paclitaxel binds poorly to unassembled tubulin [9, 10], it is likely that paclitaxel was bound to uncharacterized tubulin assemblies [10-12] and was less susceptible to the epimerization process. Thus, the ratio of 7-epi-paclitaxel to paclitaxel resulted from complex equilibria involving both the reversible epimerization of paclitaxel and 7-epi-paclitaxel and the protection of both compounds from epimerization by reversible binding to intracellualr microtubules. Total elimination of 7-epi-paclitaxel in urine is low, and accounts for 5% of the total amount of paclitaxel derivatives excreted by this route. Paclitaxel and 7-epi-paclitaxel show almost identical cytotoxic potencies. Ringel and Horwitz [6] reported an EDso (drug concentration inhibiting cell division by 50% after 72 h) of 0.08 ~M and 0.12 ~M with mouse J744.2 cells, and 0.32 ~tM and 0.31 ~M with CHO cells, respectively. However, using KB cells, Kingston reported that EDso of 7-epi-paclitaxel was three times higher than the EDso of paclitaxel [13]. Similar epimerization has been shown to occur with docetaxel. Less than 2% of the total drug detected in the feces of patients treated with [14C] docetaxel was in the form of 7-epi-docetaxel [14]. A small amount of 7-epidocetaxel was also found in plasma extracts. The proportion of 7-epi-docetaxel reached 2.6% in rat bile [15, 16]. 7-Epi-docetaxel was less cytotoxic than docetaxel in P-388 leukemia (ICso 60 and 7 ng/ml, respectively). Moreover,

136 besides the major docetaxel biliary metabolite (VI) which is hydroxylated at the t-butyl group of the side chain at C-13, Vuilhorgne e t al. observed the corresponding 7-epimer (XII) with a ratio VIDCII of 0.15 [14, 16]. Similarly, 7epi isomers of three other hydroxylated metabolites of docetaxel have been characterized by 600 MHz NMR among minor docetaxel metabolites found in mice, rabbits, rats and dogs [14]. In contrast, the 7-epimer derivatives of paclitaxel and paclitaxel metabolites have not been observed in rat and human bile, and 7-epi-paclitaxel is barely detectable in h u m a n plasma and urine. Thus, docetaxel is a p p a r e n t l y more susceptible to 7-epimerization t h a n paclitaxel, leading to an apparent larger number of 7-epimeric metabolites and to more HPLC peaks than in the case of paclitaxel. 4.2.2. Hydrolytic Derivatives Two derivatives resulting from the hydrolysis of paclitaxel have been observed in h u m a n patients [8]. Cleavage of the side chain at C-13 of the taxane ring results in the formation of baccatin III, while 10-deacetyl paclitaxel results from the cleavage of the acetyl group at the C-10 of the taxane ring (Figure 2). Baccatin III was first reported by Ringel and Horwitz [6] in the culture medium of Chinese ovary cells (CHO) and mouse macrophage-like cells (J774.2) treated with 1-5 ~M paclitaxel for 72 h. It was detected in small quantities t h a t never exceeded 10% of the concentration of paclitaxel in the medium. In addition, it was barely detected in cells. The unequal distribution of baccatin III could result from its very low affinity for microtubules [17, 18]. Baccatin III has been chemically characterized in the bile of rats treated with paclitaxel [4]. It accounted for about 2% of the paclitaxel excreted by this route and 1-2% of the total administered dose. In initial studies it remained undetected in the urine of patients treated with paclitaxel [1]. Monsarrat et al. failed to detect it in the bile of a patient treated with paclitaxel (135 mg/m 2, administered as a continuous i.v. infusion over 24 h) [3]. Recently, using a new interface HPLC coupled to Atmospheric Pressure Chemical Ionization Mass Spectrometry (HPLC/APCI-MS), Monsarrat et al. observed both baccatin III and 10-deacetyl paclitaxel in the urine of h u m a n patients who received 135 m g / m 2 paclitaxel during a 3 h i.v. infusion [8]. Baccatin III is devoid of cytotoxicity (194-fold less active than paclitaxel vs. L1210 leukemia [4]) and 10deacetyl paclitaxel in 23-fold less active than paclitaxel v s . KB cells (human

137 nasopharyngeal carcinoma cells) [19].

These two hydrolytic derivatives of

paclitaxel were observed primarily in urine collected 16 to 24 h after the end of the infusion period, where their amount corresponded to 0.3% and 1%, respectively, of the total amount of paclitaxel recovered in the urine (5% of the total administered drug). It is likely that both the reduced infusion period (3 h vs. 24 h in the initial studies) and the higher analytical sensitivity of the HPLC/APCI-MS method could account for the apparent discrepancies between the first publications reporting the absence of baccatin III and 10-deacetyl paclitaxel in the urine and their recent characterization. A similar cleavage of the side chain at C-13 is likely to occur in the case of docetaxel, since Gaillard and Vuilhorgne [14] identified a very small amount of the side chain as the carboxylic acid in rabbit feces. 4.2.3. Hydroxylated Derivatives. Hydroxylated derivatives of paclitaxel were first reported by Monsarrat et al. in the bile of rats treated with paclitaxel [4, 7]. These observations were

extended to human subjects [3, 20] and recently to docetaxel in both animals and humans [14, 16]. Several metabolites of paclitaxel have been detected by HPLC in the bile of rats and a human subject, respectively [3, 4, 20, 21]. These compounds were subsequently purified and identified by mass spectrometry.

In all cases the

DCI mass spectra showed some of the prominent fragmentations at m/z 286 (side chain at C-13), 447 (taxane ring), and 569 (taxane ring with the side chain at C-2). Of the nine metabolites found in rat bile, mass and NMR spectroscopy analysis permitted the characterization of three monohydroxylated and a dihydroxylated one. Similarly, two of the five metabolites found in human bile are monohydroxylated and one is dihydroxylated. In both rats and humans, the principal biliary metabolites are hydroxylated compounds. However, interspecies differences were apparent. Only one of the four hydroxylated metabolites identified in rat bile was also observed in h u m a n bile (Figure 2). This metabolite (metabolite VII' in human and metabolite V in rat, 2.1.3) is hydroxylated at the p a r a - p o s i t i o n the phenyl group at C-3'.

of

It was the major biliary metabolite in rats,

accounting for 13 _+ 1.5% of total drug disposition (i.e. 33% of paclitaxel excreted

138 in rat bile), but it was a minor biliary metabolite in humans, accounting for only 2% of total drug disposition (i.e. 10% of paclitaxel recovered in the bile). The two other hydroxylated paclitaxel derivatives characterized in rat bile were minor metabolites and were not detected in the bile of the h u m a n patient (Figure 2). Metabolite VI, 2.1.4, is hydroxylated at the m e t a - p o s i t i o n of the benzoate moiety at C-2 and accounts for 5 _+ 1% of the injected paclitaxel, while metabolite VII, 2.1.5, is hydroxylated at the C-19 methyl group and accounts for 1-2% of the administered dose. The chemical characterization of metabolites V and VI was further confirmed by Walle et al., who studied paclitaxel metabolism in rat hepatocytes [22]. Monsarrat et al. [3, 7, 8, 20] characterized 6-hydroxypaclitaxel (2.1.7, Figure 2) as the major paclitaxel metabolite from the h u m a n bile. The hydroxylation at C-6 of the taxane ring was somewhat surprising because 6hydroxypaclitaxel had not been previously reported and would not have been predicted based on the chemical reactivity of this portion of the molecule. 2.1.7 accounted for 12% of total drug disposition in h u m a n (i.e. 60% of paclitaxel excreted by the bile route) but was not recovered in the rat bile [3]. In addition to the hydroxylation at C-6 (the major h u m a n metabolism) and the hydroxylation at the phenyl group at C-3' (minor h u m a n metabolism), Monsarrat et al. described a minor h u m a n metabolite, 2.1.6, resulting from the hydroxylation at both sites and accounting for 2.5% of total drug disposition (i.e. 12.5% of paclitaxel excreted in the bile) [3, 7]. The h u m a n biliary metabolism of paclitaxel was further confirmed by Harris et al. [23] who used the bile of another patient, and by Cresteil et al. [24], Kumar et al. [25] and Harris et al. [26], who studied the metabolites of paclitaxel produced in vitro in the presence of h u m a n liver microsomes. All of these investigators detected 6hydroxypaclitaxel. The h y d r o x y l a t e d paclitaxel m e t a b o l i t e VII' was characterized by Cresteil et al. [24] and Harris et al. [26], who also detected 2.1.6. In contrast to the initial studies which failed to observe hydroxylated paclitaxel metabolites in plasma and urine, M o n s a r r a t et al., using the HPLC/APCI-MS method, identified 6-hydroxypaclitaxel both in urine and plasma [8, 27]. Although 6-hydroxypaclitaxel accounted for only 1.5% of the urinary clearance of paclitaxel, it accounted for 11% of the paclitaxel present in plasma. At the end of the infusion period (135 mg/m 2 for 3 h) the average plasma concentrations of 6-hydroxypaclitaxel and paclitaxel were 0.44 ~M and

139 3.9 BM, respectively. A similar observation has been made by Gianni et al., who reported 6-OH paclitaxel in the plasma of human patients [28]. Similarly, Huizing et al. reported putative paclitaxel metabolites in the plasma of human patients [29]. Besides docetaxel, 18 docetaxel derivatives were detected by HPLC and mass spectrometry in the bile of rats treated with this drug [14]. Docetaxel metabolites corresponding to the four major peaks have been purified and their chemical structure has been determined by mass spectrometry and NMR spectroscopy (Figure 3). Similarly, Vuilhorgne et al. have isolated docetaxel metabolites from the feces of different species [14]. They observed four major metabolites and several minor ones. The simplest docetaxel metabolite (2.1.9, V I ) r e s u l t s from the hydroxylation of the t-butyl group of the C-13 side chain, whereas metabolite XII corresponded to its C-7 epimer. The two metabolites V and VII correspond to two isomeric hydroxy-oxazolidinones 2.1.10, while metabolites XI and XII corresponded to their C-7 epimers (Figure 3). Finally, an additional metabolite (XVI), which corresponded to oxazolidinedione 2.1.11, was also isolated. These metabolites were present in all species studied to date including rats, mice, rabbits, dogs and humans [14]. However, the relative quantities of the metabolites were species-dependent. It is likely that metabolites V/VII and XVI resulted from further modification at the same site of the molecule (Figure 3) from the initially hydroxylated docetaxel derivative 2.1.9, mediated by cytochrome P450 enzymes [14, 16]. Thus, this hydroxylation process is likely to be responsible for the major hepatic metabolism of docetaxel, accounting for 36.5%, 64% and 45% of docetaxel clearance in rats, mice and humans, respectively. An additional site of hydroxylation, characterized on the minor docetaxel metabolite VIII, was localized on the phenyl group at C-3' [16]. This site of hydroxylation was already known to result in the major biliary paclitaxel metabolite in rats (VI) [3, 4]. Typical HPLC chromatograms of biliary extracts for paclitaxel and docetaxel are shown in Figure 4 and 5. To summarize, the principal metabolites of paclitaxel and docetaxel result from hydroxylation. In both cases, hydroxylation processes result in metabolic deactivation with respect to cytotoxicity. Hydroxylation of paclitaxel

140 at the C-6 of the taxane ring, on the phenyl group at C-3', and on the C-2 benzoate reduced the cytotoxicity 31-, 10-, and 41-fold, respectively [1, 4, 23]. V

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Figure 4. Biliary metabolites of paclitaxel. Unextracted bile samples were injected directly onto the HPLC column..4~ Rat bile samples: (a) Chromatogram of a bile sampled 1 h after paclitaxel treatment. Unmetabolized paclitaxel corresponds to peak X. Paclitaxel metabolites correspond to peaks labeled I to IX. (b) Chromatogram of a pre-treatment bile sample. (c) Chromatogram of a bile sample obtained 1 h after i.v. administration of the paclitaxel formulation alone. B. Human bile samples: (a) Chromatogram of bile collected during the first 0-6 h following 24 h infusion of paclitaxel. Unmetabolized paclitaxel corresponds to peak IX'. Paclitaxel metabolites correspond to peak labeled IV' to VIII'. (b) Chromatogram of a pretreatment bile sample.

141

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Time (min) Figure 5. Biliary metabolites of docetaxel in rat. Unextracted bile samples were injected directly onto the HPLC column. (a) Chromatogram of a bile sampled 1 h after docetaxel treatment. Peak VI corresponds to the metabolite hydroxylated on the t-butyl of the side chain at C-13 (2.1.9) while peaks V and VII correspond to the two isomeric hydroxyoxazolidinones of docetaxel (2.1.10). Other peaks labeled M correspond to minor docetaxel derivatives. (b) Chromatogram of a pre-treatment bile sample. (c) Chromatogram of a bile sample obtained 1 h after the i.v. administration of the docetaxel formulation alone. Similarly, a docetaxel metabolite t h a t is hydroxylated on the t-butyl of the C-13 side chain w a s 23-fold less cytotoxic t h a n docetaxel in a P-388 l e u k e m i a model (IC5o 0.34 pg/ml a n d 0.015 pg/ml, respectively). The two i s o m e r s of the hydroxyoxazolidinone of docetaxel (2.1.10, V and VII) a n d

the

oxazolidinedione

( 2 . 1 . 1 1 , XVI) w e r e e s s e n t i a l l y devoid of

cytotoxicity [14]. Thus, the hepatic m e t a b o l i s m of both paclitaxel a n d docetaxel l e a d s to a r e d u c t i o n in t h e p h a r m a c o l o g i c a l effect of t h e s e d r u g s . T h e s e h y d r o x y l a t i o n p r o c e s s e s s u g g e s t t h a t liver c y t o c h r o m e P-450 e n z y m e s are involved in the m e t a b o l i s m of the taxanes. D e s p i t e t h e s i m i l a r i t i e s of t h e i r c h e m i c a l s t r u c t u r e s , m e t a b o l i s m of paclitaxel a n d docetaxel are different.

the hepatic

In c o n t r a s t to paclitaxel,

142 which possesses an amido function at C-3', docetaxel has a t-butyl carbamate at this position, and lacks the acetyl group at the C-10 of the taxane ring (Figure 1). These differences lead to distinctly different cytochrome P450 hydroxylation sites. The m a i n metabolites of paclitaxel r e s u l t from hydroxylation at different sites of the taxane ring, the C-2 benzoate, and the side chain at C-13, depending on the species [3, 4, 7]. In contrast, in all species studied, including man, there is a single site of docetaxel hydroxylation, i.e. at the C-13 side chain, and this is f u r t h e r subject to species-specific modifications. The interspecies differences observed in the biliary metabolism of paclitaxel and interspecies variations in the proportion of the different docetaxel metabolites indicate t h a t it is desirable to investigate taxoid metabolism in humans. The use of microsomes constitutes a valuable tool, since liver h u m a n microsomes are more accessible to investigators t h a n h u m a n patients with biliary drainage catheters [3, 23]. 4.2.4. Glucuroconjugates and Sulfated Derivatives. The importance of the hepatic hydroxylation of taxoids suggests t h a t these metabolites could be sulfated or further glucuroconjugated. However, Monsarrat et al. failed to detect paclitaxel biliary metabolites with a molecular mass over 1,200, and detected no changes in the chromatographic properties of paclitaxel metabolites by incubating rat or h u m a n bile samples treated with paclitaxel in the presence of glucuronidase and sulfatase, although some endogenous compounds present in the bile both in control and treated animals were conjugated [3,4]. These o b s e r v a t i o n s s u g g e s t e d the absence of glucuroconjugated and sulfated derivatives of paclitaxel in bile. Similar observations have been made with docetaxel by Gaillard et al. [16]. Thus, taxoid metabolism seems to be limited to the phase of hydroxylation, and the hydroxylated metabolites of paclitaxel and docetaxel do not appear to be subjected to further conjugation. 4.2.5. Conclusions Among the three principal metabolic modifications of taxoids identified to date in the organism, hydroxylation appears to be the most important. The reversible epimerization of the hydroxyl group at C-7 occurs with both paclitaxel and docetaxel.

It seems to be more readily achieved in docetaxel,

143 where it introduces additional complexity in the characterization of the metabolites. This process, however, may be of minor pharmacological importance since it does not modify the cytotoxicity of the drug. Hydrolysis of paclitaxel and docetaxel has been observed at C-13, leading to the cleavage of the side chain and to pharmacologically inactive derivatives. However, baccatin III has been detected in rats only as a minor paclitaxel metabolite. Metabolites resulting from the hydrolysis at C-13 of paclitaxel and docetaxel were detected as very minor species in h u m a n s . Although paclitaxel, in contrast to docetaxel, possesses an acetyl group at C-10, the derivative resulting from C-10 hydrolysis was barely detected in the urine and remained undetectable in plasma and bile. Hydroxylation is pharmacologically i m p o r t a n t , since hydroxylated compounds are the major metabolites of paclitaxel and docetaxel, leading in all cases to less cytotoxic derivatives which are readily eliminated. Although paclitaxel and docetaxel have only minor chemical differences (Figure 1), their main sites of hydroxylation are distinct. In all species, docetaxel is mainly hydroxylated at the t-butyl of the side chain, whereas in paclitaxel the benzamide phenyl group remains unmodified. Moreover, the hydroxylated docetaxel is f u r t h e r modified, leading to the f o r m a t i o n of hydroxyoxazolidinone and oxazolidinedione derivatives. Depending on the species, paclitaxel is hydroxylated either at the phenyl group at C-3' or at the C-2 benzoate, as well as on the taxane ring at C-6 and C-19. In humans, 6hydroxypaclitaxel is the principal hydroxylated metabolite of paclitaxel. Besides the biliary metabolites which have been identified, at least 5 and 8 minor derivatives of paclitaxel and docetaxel, respectively, have not been characterized. However, it is known that none of these metabolites correspond to secondarily glucuroconjugated derivatives. Among the taxoids, only the metabolism of paclitaxel and docetaxel has been investigated so far. Therefore, it is not yet possible from the above conclusions to derive generalizations about taxoid metabolism. 4.3. C Y T O C H R O M E P 4 5 0 E N Z Y M E S I N V O L V E D IN T H E M E T A B O L I S M OF TAXOIDS.

Studies performed in rats and h u m a n s have d e m o n s t r a t e d t h a t the major metabolites of paclitaxel and docetaxel recovered in the bile correspond

144 to mono- and dihydroxy derivatives. Thus, hepatic metabolism of paclitaxel and docetaxel is mediated by a monooxygenase system supported by cytochrome P450. About 35 isozymes of cytochrome P450 have been identified, purified, and/or cloned from the human liver [30]. They display overlapping substrate specificity and are differently regulated by exogenous chemicals and at specific phases of ontogenesis [31, 32]. The identification of the cytochrome P450 isozymes (CYP) involved in paclitaxel and docetaxel metabolism can be achieved by studying the in vitro biotransformations of paclitaxel in the presence of human liver microsomes. Moreover, this experimental approach may allow us to understand the possible effects of other drugs on the clinical activity of paclitaxel. 4.3.1. Paclitaxel The in vitro metabolism of paclitaxel by human microsomes has been investigated by Cresteil et al. [24], Kumar et al. [25] and Harris et al. [26]. In vitro, paclitaxel was metabolized by human liver microsomes (Figure 6), but no metabolites could be detected using human kidney microsomes [24]. Besides paclitaxel, and in the presence of NADPH only, Cresteil et al. observed two metabolites which were identified by comparison with reference compounds. The major peak corresponded to 6-hydroxypaclitaxel (metabolite VIII', 2.1.7), the main paclitaxel metabolite generated in vivo by humans [3]. The minor peak corresponded to the monohydroxylated derivative 2.1.3 (metabolite VII'). Kumar et al., working with radiolabeled paclitaxel, also observed a major peak corresponding to 2.1.7 and two unidentified minor compounds [25]. Similar observations were made by Harris et al., who characterized 6hydroxypaclitaxel, metabolite VII', and the dihydroxypaclitaxel metabolite VI' (2.1.6) [26]. These metabolites were absent when NADPH was omitted from the reaction mixture, and their formation was strongly inhibited by carbon monoxide, which binds to the active site of cytochrome 450 [25, 26]. The amount of 6-hydroxypaclitaxel greatly exceeded the amount of metabolite VII', both in the bile of the human patient and in the in vivo situation. Kinetic measurements are difficult, due to the poor solubility of paclitaxel in aqueous solutions. However, in vitro the synthesis of 6hydroxypaclitaxel and metabolite VII' followed typical monophasic MichaelisMenten kinetics, indicating the involvement of a single enzyme system for each of these metabolites. Cresteil et al. reported an apparent Km of 15 ~M for

145 both metabolites [24] . This value was very similar to the Km (18 tiM and 16-21 tiM) measured by Kumar et al. [25] and Harris et al. [26], respectively, for 6hydroxypaclitaxel. Cresteil et al. determined t h a t the a p p a r e n t rate of synthesis of 6-hydroxypaclitaxel (Vm=120+ 30 pmol/mirdmg protein with 50 tiM paclitaxel) was roughly 22-fold higher than the apparent rate of synthesis of metabolite VII' (5.3 _+1.4 pmol/mirdmg protein) [24]. The a p p a r e n t rate of synthesis of 6-hydroxypaclitaxel appears to be highly variable. Harris et al. [26] reported a 7-fold higher rate (830-930 pmol]min/mg protein), but Kumar et al. [25] published a value which was 6-fold lower (19.3+14 pmol/min/mg protein). 0,04

PACUTAXEL

0,01 0,03 "

DOCETAXEL

B

0,00 0,00

viir

0,02 " t

0,004-IIII

o,oo J

0,01 t

o 0 | 10 20 Time (rain)

' 30

0

10

20

30

40

50

Time (rnin)

Figure 6. In vitro synthesis of taxoid metabolites. Paclitaxel (A) and docetaxel (B) were incubated with human liver microsomes. All minor peaks (*) were also present when NADPH was omitted from the reaction mixture. See Figures 2 and 3 for the chemical structures of paclitaxel and docetaxel metabolites, respectively. Attempts to characterize the cytochrome P450 isozymes (CYP) involved in these reactions have led to inconsistent conclusions. Cresteil et al. [24] and Harris et al. [26] both concluded t h a t paclitaxel was hydroxylated at the p a r a position of the phenyl at C-3' (metabolite VII') by CYP3A and t h a t 6hydroxypaclitaxel was synthesized by a different cytochrome. In contrast, K u m a r et al. [25] proposed t h a t the formation of 6-hydroxypaclitaxel was catalyzed by CYP3A.

146 It is clear from the results of Cresteil et al. [24] and Harris et al. [26] that CYP3A is involved in the synthesis of the minor metabolite VII' (Figure 7). PACLITAXEL

CYP3A/

~YF2C8

METABOLITE M4 METABOIATE M5 p-0H-phenyl-C3'-paclitaxel 6-0H-paclitaxel CYP2C8~x~

~/~YP3A

DIHYDROXYPACLITAXEL

I CYP3A HYDROXYDOCETAXEL (VI) CYP450? J HYDROXYOXAZOLIDINONES (V AND VII)

~

CYP450? OXAZOLIDINEDIONE (XVI)

Figure 7. Metabolic pathways for paclitaxel and docetaxel in the presence of human liver microsomes. Its synthesis was correlated (r=0.82) with CYP3A content [24]. Similarly, Harris et al. observed that the best correlation (r2=0.94) was found between the formation of this metabolite and testosterone 6-hydroxylation associated with CYP3A [26]. Drugs known to reduce CYP3A-dependent monooxygenase activities (orphenadrine, erythromycin, testosterone, and troleandomycin) were potent inhibitors (50-80%) of the formation of metabolite VII' [24, 26]. In contrast, quercetin, naringenin and kaempferol, potent inhibitors of the synthesis of 6hydroxypaclitaxel, had no inhibitory effect on the synthesis of metabolite VII'

147 [26]. Immunoglobulins directed against CYP3A (2 mg IgG/nmol P-450, i.e. 0.7 mg IgG/mg protein) inhibited its synthesis by 75-80% [24-26]. Finally, human CYP3A4, expressed in B lymphoblastoid cells in E. coli and in vaccinia-based systems, catalyzed the transformation of paclitaxel into metabolite VII' [26]. In agreement with the implication of CYP3A in the hydroxylation of the p a r a position of the phenyl ring at C-3', Harris et al. [26] observed that overexpressed human CYP3A metabolized 6-hydroxypaclitaxel to the dihydroxypaclitaxel derivative VI', 2.1.6 (Figure 2). The involvement of CYP3A in the synthesis of metabolite VII' is further supported by the observations of Walle et al. [22]. Using rat hepatocytes, they observed that metabolites VII' and VI were probably synthesized by CYP3A, since their formation was strongly inhibited by midazolam and verapamil. Both Cresteil et al. [24] and Harris et al. [26] performed a comparative study on the formation of several paclitaxel metabolites by h u m a n liver microsomes. Cresteil et al. [24] first studied the relationship between the rate of formation of metabolite VII' and 6-hydroxypaclitaxel. No correlation was observed between the formation of the two metabolites (r=0.64), suggesting that they were synthesized by different P450 isozymes. An identical conclusion was reached by Harris et al., who also observed a low correlation (r2=0.24-0.28) for the formation of these two species [26]. These authors agreed that metabolite VII' was synthesized by CYP3A, and that 6-hydroxypaclitaxel was synthesized by a different cytochrome isozyme. Kumar et al. reached the opposite conclusion [25]. Using microsomes prepared from 6 h u m a n livers, they observed that 17-ethynyl estradiol, a substrate for CYP3A, inhibited the formation of 6-hydroxypaclitaxel by 73%, while substrates of CYP1A2, 2B, 2C (tolbutamide), 2D6, and 2E were ineffective in this respect. In contrast, Cresteil et al. reported that tolbutamide inhibited the synthesis of 6-hydroxypaclitaxel by 30% [24]. Kumar et al. also observed that several substrates for CYP3A (midazolam, verapamil, testosterone, quercetin) led to 60-80% inhibition, but that erythromycin and troleandomycin were devoid of inhibitory effects [25], in agreement with the observations of Cresteil et al. [24]. Kumar et al. correlated the synthesis of 6-hydroxypaclitaxel with the metabolism of midazolam (r=0.95) and 17-ethynyl estradiol (r=0.87) [25]. Finally, they observed that antibodies against CYP3A1 inhibited the formation of 6-hydroxypaclitaxel.

148 However, Cresteil et al. [24] and Harris et al. [26] performed the same experiment using antibodies against CYP3A. They both observed inhibition of formation of metabolite VII' (75-80% in the presence of 2 mg IgG/nmol of P450) but failed to observe a significant inhibition in the synthesis of 6hydroxypaclitaxel, even using concentrations of IgG of 5 and 16 mg/nmol P450, respectively. In this respect, Kumar et al. reported a 70% inhibition of the synthesis of 6-hydroxypaclitaxel in the presence of 20 mg IgG per mg of microsomal protein (approximately 60 mg IgG/nmol of P450) [25], i.e. a concentration 4-10 times higher than the ones used by Cresteil et al. [24] and Harris et al. [26] in order to obtain inhibition of the formation of metabolite VII'. Thus, it is possible that the effect reported by Kumar et al. was nonspecific, i.e. the antibodies cross-reacted with other cytochromes. In agreement with this view, Kumar et al. failed to observe the formation of 6hydroxypaclitaxel using extracts prepared from a cell line overexpressing CYP3A4 and showing a 10-fold higher testosterone 6-hydroxylase activity than extracts from wild-type cells [25]. This result was confirmed by Harris et al. [26], who detected only metabolite VII' when paclitaxel was incubated with h u m a n CYP3A4. The conclusions reached by Kumar are difficult to reconcile with those of Cresteil and Harris. However, these discrepancies could partially reflect the relative amounts of active cytochrome isozymes in microsome preparations, as suggested by the lower rate of synthesis of 6-hydroxypaclitaxel observed in the preparations of Kumar et al. [25]. Cresteil et al. [24] and Harris et al. [26] agree that the synthesis of 6hydroxypaclitaxel was not under the control of CYP3A, but their experimental data were interpreted differently. Cresteil et al. [24] suggested that 6hydroxypaclitaxel was synthesized by CYP2C, while Harris et al. [26] favored the involvement of an unknown P450 isoform. Cresteil et al. [24] studied the correlation between paclitaxel m e t a b o l i s m and i m m u n o c h e m i c a l l y determined levels of individual P450 enzymes (CYP2C, 2El, 1A2, 3A, 4A1, and 2D6), as well as monooxygenase activities supported by CYP2C (diazepam demethylation), CYP2E1 (chlorzoxazone hydroxylation) and CYP1A2 (methoxy- and ethoxyresofurin dealkylations). They observed a positive correlation only between the synthesis of 6-hydroxypaclitaxel and CYP2C (r=0.72) as well as with the N-demethylation of diazepam (r=0.89) supported by CYP2C [33]. They also observed that diazepam inhibited the formation of 6-

149 hydroxypaclitaxel by 60%. Moreover, Cresteil et al. [24] observed that 6hydroxypaclitaxel was barely detectable using liver microsomes from newborns 1-7 days of age (<1 pmol/min/mg protein), while its rate of synthesis increased abruptly using liver microsomes from 3 to 6 month old infants (75 +50 pmol/min/mg protein). In contrast, the formation of metabolite VII' was detected using microsomes from newborns. This increase in the formation of 6-hydroxypaclitaxel occurred concomitantly with the onset of CYP2C, whereas the early increase in the formation of metabolite VII' was associated with the increase in CYP3A4 content and testosterone 6~ hydroxylase (Cresteil et al., manuscript in preparation). Because CYP2C is absent from human fetal liver and increases during the first months after birth, i.e. after CYP3A [24], it was tempting to correlate the rise of 6-hydroxypaclitaxel synthesis with the presence of CYP2C. In agreement with this view, Harris et al. [26] reported that the synthesis of 6hydroxypaclitaxel was not affected by antibodies against CYP3A4, but was inhibited by 85% in the presence of antibodies against CYP2Cmp (10 mg IgG/nmol P450). However, it was also inhibited by 60% using antibodies against CYP2A6 (10 mg IgG/nmol P450). Harris et al. [26] underlined that diazepam (100 gM) was not selective, because it inhibited the synthesis of 6-hydroxypaclitaxel by 70% and the rate of formation of metabolite VII' by 51%. They observed that high concentrations of mephenytoin (substrate of CYP2C19) and tolbutamide (substrate of CYP2C9-10) did not inhibit the synthesis of 6-hydroxypaclitaxel, and that correlations with mephenytoin 4'-hydroxylation and tolbutamide methylhydroxylation were low (r2=0.01 and 0.08, respectively). Therefore, further investigations are required to more clearly define the metabolic pathway in the formation of 6hydroxypaclitaxel, which is the principal metabolite of paclitaxel in h u m a n s . 4.3.2. Docetaxel The in vitro metabolism of docetaxel by liver microsomes has been investigated by Zhou-Pan et al. [34], Gaillard et al. [16], and Cresteil and Monsarrat [unpublished] (Figure 6 and 7). Using human microsomes, ZhouPan et al. detected four metabolites, but did not characterize them [34]. Using rat liver microsomes, Gaillard et al. [16], characterized, in addition to the 7-epi derivative, four docetaxel metabolites. The most prominent one corresponded to the docetaxel derivative hydroxylated on the t-butyl of the C-13 side chain

150 (metabolite VI, 2.1.9, Figure 3). Two of the minor metabolites (V and VII, 2.1.10) corresponded to the two isomeric hydroxyloxazolidinones. The apparent Michaelis-Menten constant (10 ~M) was similar to the value determined for paclitaxel, and the apparent maximum rate of synthesis varied from 18 to 160 pmol/min/mg protein [34]. Published experimental data suggested that the main metabolism of docetaxel by h u m a n liver microsomes was due to the isozymes of the CYP3 subfamily [15, 34, 35, Cresteil and Monsarrat, unpublished results]. Zhou-Pan et al. [34] reported significant correlation (r=0.76, 0.63, and 0.62) between the formation of these metabolites in the presence of 29 distinct preparations of h u m a n liver microsomes and erythromycin N-demethylase activity. No correlation was observed for the fourth docetaxel derivative (r=0.07), but it was not clear whether this derivative corresponded to a metabolite or simply resulted from a 7-epimerization, since its structure was not determined. In agreement with the central role of CYP3A in docetaxel metabolism, Marre et al. reported no correlation between the synthesis of docetaxel metabolites and aniline hydroxylase (r=0.35; CYP2E1) and debrisoquine 4-hydroxylase (r=0.34; CYP2D6) activities [35], while S a n d e r i n k et al. [15] failed to inhibit docetaxel metabolism with acetanilide (CYP1A2), a - n a p h t o f l a v o n e (CYP1A1/2), caffeine (CYP1A2), tolbutamide (CYP2C8-9), low concentrations of quinidine (CYP2D6) and aniline (CYP2E 1). In contrast, Marre et al. [35] and Sanderink et al. [15] observed that substrates and/or inhibitors of CYP3A were able to inhibit the metabolism of docetaxel by h u m a n hepatocytes in culture (95, 90 and 77% inhibition with ketoconazole, troleandomycin, and nifedipine, respectively). Moreover, Marre et al. [35] reported that the metabolism of docetaxel increased 1.44-fold when h u m a n liver hepatocytes were induced with rifampicin, and Sanderink et al. [15] indicated that, among five inducers of different cytochrome P450 enzymes subfamilies, docetaxel metabolism was increased 6-fold in liver microsomes obtained from rats treated with dexamethasone. Immunoinhibition with anti-CYP3A antibodies led to a reduction in the metabolism of docetaxel from 35% to 60% with 2 mg IgG/nmol P450 in microsome preparations obtained from two distinct individuals, whereas preimmune IgG had no effect [Cresteil and Monsarrat, unpublished results]. Using liver h u m a n microsomes, Zhou-Pan et al. [34] reported an a p p a r e n t Km of 15 ~M and an apparent maximum rate of synthesis of

151 docetaxel metabolites varying from 7 to 165 pmol/min/mg protein. Similarly, Sanderink et al. [15] published a mean rate of 25+6 pmol/min/mg protein with h u m a n liver microsomes. These values were similar to those reported from the s y n t h e s i s of 6-hydroxypaclitaxel. Using the s a m e microsomal preparations, Cresteil et al. observed a highly positive correlation between the metabolism of docetaxel and the production of metabolite VII' of paclitaxel (r=0.93) and no correlation with the rate of formation of 6-hydroxypaclitaxel (r=0.48). Furthermore, the extent of formation of docetaxel metabolites (2-3 pmol/min/mg p r o t e i n ) w a s comparable to the rate of formation of metabolite VII' and much lower t h a n t h a t of 6-hydroxypaclitaxel [Cresteil et al. u n p u b l i s h e d results]. Docetaxel m e t a b o l i s m by mouse and dog liver microsomes fits a one-enzyme model, but a two-enzyme model was necessary in order to account for the experimental data obtained with rats and h u m a n liver microsomes [15]. Although the results published so far indicate a prominent role for CYP3A, some additional cytochrome P450 isozymes could jointly act in docetaxel metabolism both in rats and humans (Figure 7). 4.3.3. Conclusions. It is clear from the experimental results published so far t h a t the h u m a n metabolism of paclitaxel and docetaxel involves hydroxylation by different cytochrome P450 enzymes (Figure 7). CYP3A has been demonstrated to play the major role in the metabolism of docetaxel. In contrast, an unknown cytochrome P450 enzyme, different from CYP3A, is involved in the synthesis of 6-hydroxypaclitaxel, the most prominent h u m a n metabolite of paclitaxel. H u m a n CYP3A is responsible for the synthesis of a minor hydroxylated metabolite. At the present time it is not possible to suggest general rules accounting for the hydroxylative hepatic metabolism of taxoids and it is clear that further investigations with other taxoid derivatives are necessary. It would be of great interest to characterize the cytochrome P450 enzyme responsible for the formation of 6-hydroxypaclitaxel and to determine whether CYP3A is involved in further modifications of the first hydroxylated metabolite of docetaxel. More importantly, in view of the differences among the cytochrome P450 enzymes involved in the hepatic metabolism of paclitaxel and docetaxel, in vitro metabolism studies of newly synthesized cytotoxic taxoids in the presence of h u m a n liver microsomes may facilitate preclinical screening of these agents.

152 4.4. PHARMACOLOGICAL DISPOSITION OF TAXOIDS

Knowledge of the dispositon of taxoids and their metabolites in the organism could be used in optimizing their clinical activities. Moreover, this information could provide some clues about some treatment failures and could suggest protocols to modulate the clearance of these drugs. 4.4.1. Pharmac0kinetics and Disposition. The pharmacokinetic behavior of paclitaxel and docetaxel in humans has recently been reviewed in great detail by Rowinsky et al. [1] and Bruno and Sanderink [2]. High dosages of both paclitaxel and docetaxel showed either biphasic or triphasic pharmacokinetic behavior in plasma, when sensitive analytical methods were used. For 3 h infusions, the disposition of paclitaxel in plasma was optimally modeled by two- or three-compartment models [29, 36]. However, nonlinear pharmacokinetic profiles have been observed when paclitaxel was administered at high doses over short periods, suggesting a saturation of the elimination and/or tissue distribution processes. Apparent distribution volumes of paclitaxel (mean VDSS=volume of distribution at steady state) were much larger (48 to 182 1/m2) than the volume of total body water suggesting that paclitaxel bound to plasma proteins and/or to other tissue elements and was readily excreted [28, 37-47]. Binding to plasma proteins and tissues" The peak plasma paclitaxel concentration in humans (Cmax) depends on the infusion duration. With 3 h infusion periods, the peak plasma concentration reached 2.5-12.5 pM (135-300 mg paclitaxel m 2) [36, 48-50]. Similarly, maximal plasma concentrations of docetaxel of 5 ~M have been reported [51-53]. The maximum concentration of paclitaxel in plasma decreased to 3.1-4.1 ~M, 0.7-0.9 ~M and 0.05-0.08 pM when 6, 24 and 96 h infusion periods were used, respectively [37, 38, 41-47, 54, 55]. Thus, infusions varying from 3 to 24 h achieved Cmax values that are several orders of magnitude higher than cytotoxic drug concentrations. More than 95% of paclitaxel may bind to plasma proteins, as demonstrated both by equilibrium dialysis and ultrafiltration methods [38, 41, 42, 55, 56]. This binding was reversible [10], allowing the rapid elimination of paclitaxel from the plasma compartment.

153 Taxoids are highly hydrophobic, and it is likely that they bind not only to cellular microtubules, but also to various hydrophobic cellular organelles. This view is supported by the observations of Manfredi and Horwitz [9]. They have demonstrated that intracellular paclitaxel was bound in a stoichiometric fashion to tubulin assembled in microtubules, but the drug also bound unspecifically to other undetermined cellular components. The distribution of radiolabeled paclitaxel and docetaxel has been recently investigated in animal tissues [57-60]. In rats, Gaver et al. observed that the amount of radioactivity in tissues was higher after i.v. t h a n i.p. administration [60]. Using i.v. injection, 86% of the initial dose was recovered in tissues after 3 h, while only 46% was recovered after i.p. injection. In both cases, the amount of radioactivity in tissues decreased with time: 50% and 46% remained after 24 h, 5% and 15% after 48 h, and less than 1% after 5 days from i.v. and i.p. injections, respectively. Thus, all the injected paclitaxel was slowly but completely eliminated. A similar observation was made by Marlard et al., who reported that less than 0.8% of the initial amount of radiolabeled docetaxel was present in mice after 4 days and was completely eliminated within 7 days in dogs [61] . Maximal concentrations in tissues sampled 2 h after a d m i n i s t r a t i o n of radiolabeled paclitaxel were observed in the portal triad, glomeruli and renal medulla (0.84, 0.67 and 0.68 ~M respectively). Tissue concentrations were lower in choroid plexus, liver, spleen, heart, lung and muscles (0.27, 0.23, 0.23, 0.21, 0.20 and 0.06 ~M). All these values corresponded to cytotoxic drug concentrations. In contrast, the concentration of paclitaxel in the cerebral fluid was low (0.7 nM) and radiolabeled paclitaxel was undetectable in testes and brain [58]. These observations are in a g r e e m e n t with the results of Klecker et al. [57], Eiseman et al. [62] and Gaver et al. [60]. Eiseman et al. reported blood/testes and blood/brain partition coefficients of 0.14 and 0.01, respectively, whereas the blood/liver partition coefficient was much greater (4.1-5.2). Similarly, very small quantities of radioactivity were observed by Gaver et al. [60] in testes and different nervous system tissues (cerebellum, cerebrum and spinal cord). A wide tissue distribution of radiolabeled docetaxel was reported by Gaillard et al. in mouse and dog [61]. As was the case with paclitaxel, radiolabeled docetaxel was not detected in the central nervous system.

154 One can conclude that taxoids are widely distributed to most tissues. At least in the case of paclitaxel, its concentration is particularly high in tissues that are possibly involved in organ barrier filtration, such as portal triad, glomeruli, renal medulla and choroid plexus [1]. Very low levels of paclitaxel were observed in the testes and central nervous system in rats, and a very low level of docetaxel was also observed in the central nervous system of rodents and dogs, i.e. tissues which are considered "tumor s a n c t u a r y " sites. Moreover, radiolabeled paclitaxel was not detected in the peripheral nervous system, although peripheral neurotoxicity is the major dose-limiting nonhematological effect of paclitaxel [58]. Clearance: Over a long period of time, renal clearance and biliary excretion together account for the overwhelming majority of taxoid disposition. However, over shorter periods (24 h) following treatment, only 25% of the a d m i n i s t e r e d dose of paclitaxel was excreted in the bile and urine, in agreement with the initial wide tissue distribution throughout the organism

[8]. Total urinary excretion of unmetabolized paclitaxel represented 10% of the administered dose in rats [4, 59] and from 1.4 to 8.2% (cumulative mean 5.5%) in h u m a n patients [1]. No metabolites were identified in these early studies, but Monsarrat et al. identified several paclitaxel derivatives in the urine of patients treated with paclitaxel [8]. Using reference compounds, Monsarrat et al. [3, 8, 20] identified by HPLC and mass spectrometry the major h u m a n paclitaxel metabolite, 6-hydroxypaclitaxel, and the 7-epimer derivative of paclitaxel. Both compounds were present in very low quantities, accounting for 1.5% and 0.8% of the total urinary excretion, respectively. In late periods of urine collection (16-24 h after the end of the infusion period), Monsarrat et al. also characterized the two hydrolyzed derivatives of paclitaxel, i.e. baccatin III and 10-deacetyl paclitaxel (1.2% of total u r i n a r y excretion) [8]. Thus, in humans, paclitaxel metabolites were quantitatively negligible in urine, and intact paclitaxel accounted for 97% of total renal clearance. Similarily, 2.5% -6.5% of administered radiolabeled docetaxel was recovered in the urine of rats, mice, rabbits, dogs and h u m a n s t r e a t e d with docetaxel. One can conclude t h a t renal excretion contributes little to the overall systemic clearance of taxoids. In rats, more than 98% of injected radiolabeled paclitaxel was excreted in the feces over a 6-day period [59].

Similarly, more t h a n 85% of injected

155

radiolabeled docetaxel was recovered in the feces of mice and dogs [61]. In humans, 80% of the total dose of radiolabeled docetaxel was excreted in the feces over a 7-day period, and the majority of the excretion occurred during the first 48 h [63]. These observations demonstrate that biliary excretion accounts for the majority of taxoid disposition in animals and humans, which is in agreement with the early studies of Monsarrat et al. [3, 4]. 3O

O "O

Total o

20"

w

U

VIII'

m,

r

f

lO-

PACLrrAXEL

/...~ VII' 0 m

0

I0

20

infusion ~ period

30

4o 50 Time (hours) ix~ infusion period

Figure 8. Biliary excretion of paclitaxel in a h u m a n patient.

The cumulative biliary excretion of paclitaxel and its metabolites was studied both in rats and in humans using a percutaneous drainage catheter, after complete biliary obstruction due to an extensive cholangiocarcinoma [3, 4, 21]. In the human patient (Figure 8) and in rats, paclitaxel and paclitaxel metabolites excreted via the biliary route accounted for 20% and 40% of the total administered dose of paclitaxel, respectively, during the 24 h period after the infusion. In the human, 3_+1% of the total dose was recovered in the bile as unmetabolized paclitaxel, and 12_+2% was recovered in the rat bile. In the human patient, the main hydroxylated metabolite (6-hydroxypaclitaxel) accounted for 60% of all biliary taxoid derivatives and for 12+_2% of total drug disposition, while the minor hydroxylated metabolites VIII' accounted for 10%

156 of all biliary taxoid derivatives in bile and for 2_+1% of the total administered dose. Biliary excretion of docetaxel has been examined by Vuilhorgne et al. [14] during the 48 h following infusion. Radioactive docetaxel and its metabolites were extracted from the feces of various animals (rats, mice, rabbits, dogs) and h u m a n s treated with docetaxel. In the feces of humans, rats and mice, docetaxel and its metabolites accounted for 48%, 46% and 66% of the total administered dose, respectively. In humans, 3% of the total dose was recovered in the feces as unmetabolized docetaxel, whereas the corresponding figure for animals was 9.5% (rat) and 1.5% (mouse). The hydroxylated metabolite of docetaxel (VI) accounted for 6.5% of the administered dose in humans, while the two hydroxyoxazolidinones (V and VII) of docetaxel accounted for 15.2% and the oxazolidinedione derivative (XVI) for 23.3%. The relative proportions of these metabolites varied depending on the species. For example, metabolite XVI represented 51% of all biliary docetaxel metabolites in humans, but only 14% and 18% for rats and mice, respectively. In contrast, hydroxylated docetaxel (VI)represented 14% of biliary metabolites in humans and 15% on mice, but as much as 47% in rats, and the two isomers V and VII represented 34% of all biliary metabolites in h u m a n s and 32% in rats, but as much as 59% in mice. Thus, quantitative aspects of both paclitaxel and docetaxel metabolism differ according to the species. Moreover, in humans, biliary excretion of paclitaxel and its metabolites over a 24 h period is lower (20% of the total administered dose) than the biliary disposition of paclitaxel and its metabolites in rats (48% of total administered dose). In both cases, however, metabolites were preferentially excreted via the biliary route, where they accounted for 17% and 45% of total drug disposition (Figure 8). To conclude, hepatic metabolism of taxoids seems to be the main route of disposition. 4.4.2 Clinical Interest. For both paclitaxel and docetaxel, metabolism results in inactivation and preferential excretion of metabolites by the biliary route. Hence, biotransformation of taxoids by liver P450 enzymes seems to constitute a critical factor in determining their pharmacological efficiency. A better u n d e r s t a n d i n g of taxoid biotransformation could potentially facilitate the development of more effective treatments.

Our knowledge of paclitaxel and

157 docetaxel metabolisms is recent, and the biosynthesis of 6-hydroxypaclitaxel, the main paclitaxel metabolite in humans, is not fully understood [26]. Despite these difficulties, some observations already show the potential applications of these pharmacological studies. For example, in a patient treated for an ovarian carcinoma with hepatic metastases, plasma concentrations of paclitaxel, 6-hydroxypaclitaxel and 7epi-paclitaxel were ten times higher than in other patients without hepatic dysfunction [Armand and Monsarrat, unpublished observations]. In agreement with this observation, the concentration of 7-epi-paclitaxel recovered in urine, from 16 to 24 h after the infusion, was increased 15-fold. The biotransformation of paclitaxel is also affected by prior treatment with associated drugs. Barbiturates were shown to accelerate the formation of metabolite VII' by h u m a n liver microsomes, whereas 6-hydroxypaclitaxel formation remained unchanged [24]. The same microsomes were also more active in the hydroxylation of docetaxel [Cresteil et al., manuscript in preparation]. These observations clearly emphasized the importance of biliary excretion and the role of concurrently administered medications on the biotransformation of taxoids by the human liver. Since 17% and 45% of paclitaxel and docetaxel are excreted via the biliary route as less active hydroxylated metabolites, it is tempting to try to reduce the excretion by preventing the metabolism mediated by cytochrome P450 enzymes. Due to differences between rats and humans in paclitaxel metabolites and in liver cytochrome P450 enzymes involved in the metabolism of paclitaxel, the observations in rats could not be directly applied to human patients. However, they may point to some tools for increasing clinical efficacy. It has been established that the paclitaxel derivative hydroxylated at the p a r a position of the phenyl group at C-3', corresponding to the major paclitaxel metabolite in rats (V) but to a minor metabolite in human (VIII'), is synthesized by cytochrome P4503A [22, 24, 26]. In agreement with this view, its synthesis was strongly inhibited (90%) by low concentrations (2 ~M) of ketoconazole, a commonly used antifungal medication [64]. Accordingly, coadministration of paclitaxel and ketoconazole to rats produced a moderate increase in plasma levels of paclitaxel (18%), but dramatically decreased liver metabolism and biliary excretion. In rats treated with ketoconazole, the amounts of paclitaxel and paclitaxel metabolites recovered in the bile during the 6 h infusion period were decreased by 86% and 90%, respectively. This

158 effect could not be attributed to a decrease in the biliary flow, which was unaffected by the treatment by ketoconazole, i.e. 8.97 ml and 7.39 ml for the 6 h infusion period in control and ketoconazole-treated rats, respectively. In a g r e e m e n t with the reduced biliary clearance of paclitaxel derivatives, the amount of paclitaxel metabolites in rat liver was reduced by 73% (1.1 nmol/g and 0.3 nmol/g in rats treated with and without ketoconazole, respectively), whereas the amount of paclitaxel remained unchanged (5.3 nmol/g and 5.5 nmol/g, respectively). Such an approach may be used to explore the possible reduction of elimination and inactivation of docetaxel, mediated both in animals and h u m a n s by cytochrome P4503A [15, 34, 35, Cresteil and Monsarrat, unpublished results]. A similar study has been performed with cimetidine, which is given as a premedication in order to reduce h y p e r s e n s i t i v i t y reactions associated w i t h paclitaxel therapy. Although cimetidine is a broad inhibitor of microsomal hydroxylation, Klecker et al. [64, 65] failed to observe a variation of biliary excretion of paclitaxel and its metabolites and their amounts in liver extracts. However, the clinical significance of this observation is unclear, since it is recognized that rats do not constitute a suitable model for studying paclitaxel metabolism in humans [3, 24, 26]. If it can be assumed that a decrease in the hepatic metabolism of taxoids could lead to an increase of drug activity, then induction of hepatic metabolism could potentially result in reduced activity and toxicity. Fetell et al. have hypothesized t h a t the induction of liver cytochrome P450 enzymes could account for the reduced plasma levels of paclitaxel in some patients [66]. It has been demonstrated that paclitaxel is active in vitro against glioma cells. Thus, patients with a newly diagnosed multiform glioblastoma, a brain cancer, were treated with paclitaxel as a 96 h infusion (140 mg/m 2) before radiotherapy. Pharmacological studies performed during the first cycle of chemotherapy failed to reveal any response and concomitantly showed minimal toxic side effects. The mean paclitaxel plasma level (16.8_+8.6 nM) was substantially lower than those reported (77_+6 nM) with identical dose and schedule in lymphoma or breast cancer. It is tempting to speculate that the absence of both activity and toxicity may be due to the induction of hepatic cytochrome P450 enzymes by anticonvulsants (barbiturates, phenylhydantoin) taken by the patients. This explanation suggests that dose escalation could result in increased activity.

159 4.4.3. Conclusions These examples demonstrate the potential usefulness of modulating paclitaxel and docetaxel activity in the clinic through a modification of their hepatic metabolism. The similarities between docetaxel metabolism in humans and laboratory animals allow the use of animal models in pharmacological studies. This is apparently not the case for paclitaxel. The experimental modulation of the activities of cytochrome P450 enzymes is going to be difficult to approach except with variously induced human hepatocytes cultured in vitro or with mammalian cells expressing the appropriate human cytochrome P450 enzyme [67]. These attempts will require close collaboration between basic research and clinical practice. The recent discovery of 6hydroxypaclitaxel in the plasma of cancer patients opens new possibilities of developing such coordinated approaches [8, 27, 28].

Acknowledgements Our work was supported by grants from "L' Association pour la Recherche sur le Cancer" (M.W., B.M.). We thank Dr. N. Johnson for his critical reading of the manuscript. The constant and kind interest of Dr. P. Potier is greatly appreciated. R~IERENCF~

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3.

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10. 11. 12. 13. 14.

15. 16.

17. 18. 19. 20.

21.

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162 38.

39.

40. 41. 42. 43.

4.

45. 46.

47.

48.

49. 50.

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163 51.

52. 53.

54.

55. 56. 57.

58. 59. 60. 61.

62.

63.

64.

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164 65.

66.

67.

Reed, E.; Sarosy, G.; Jamis-Dow, C.; Klecker, R.; Kohn, E.; Link, C.; Christian, M.; Davis, P.; Collins, J. Proc. Amer. Assoc. Cancer Res. 1993, 34, 395 (abstract). Fetell, M.R.; Grossman, S.A.; Balmacoda, C.; Leu, J.G.; Erlanger, B.F.; Rowinsky, E.; Khandji, G.; Yue, N.; Zeltman, M. Proc. Am. Soc. Oncol. 1994, 13, 179 (abstract). Note added in proof: Recently, using cDNA-expressed-human cytochrome P450 enzymes, Rahman et al. reported that cytochrome P450 2C8 efficiently catalyzes paclitaxel 6-hydroxylation in contrast with other cytochrome P450 enzymes (1A2, 3A3, 3A4, 3A5, 2B6, 2C9, and 2C9R144C). Moreover, paclitaxel 6-hydroxylation correlates with hepatic CYP2C8 content (r2=0.8), but not with CYP2C9 (r2=0.4). These observations confirm and extend the previous conclusions by Cresteil et al., who assigned the formation of 6-hydroxypaclitaxel to a CYP2C enzyme [24]. See: Rahman, A.; Korzekwa, K.R.; Grogan, J.; Gonzalez, F.J.; Harris, J.W. Cancer Res. 1994, 54, 5543.

The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B.V. All rights reserved

165

5 PACLITAXEL (TAXOL| CHEMISTRY AND STRUCTURE ACTIVITY RELATIONSHIPS

-

Shu-Hui Chen Bristol-Myers Squibb Pharmaceutical Research Institute, P.O.Box 5100, Wallingford CT 06492-7660, U.S.A.

Vittorio Farina Boehringer Ingelheim Pharmaceuticals, 900 Ridgebury Road, Ridgefield, CT 06877, U.S.A.

5.1. I N T R O D U C T I O N

Studies on the chemistry of paclitaxel I (1.1.1, Figure 1 ) a n d its derivatives began more than 30 years ago, when this compound was isolated from the bark of T a x u s b r e v i f o l i a by Wani and Wall [1]. In these early years, in spite of the a n t i t u m o r activity displayed by the substance, interest in its chemistry was modest, as was the supply of the drug. The field, therefore, advanced very slowly, mostly through the efforts of Kingston [2]. The lowyielding process that afforded paclitaxel from the bark of the yew tree appeared to be a limitation to the development of the compound as a therapeutic agent. Later, the isolation of 10-deacetylbaccatin III, 1.1.2, in r a t h e r high yields from 1Taxol| is a registered trademark of the Bristol-Myers Squibb corporation. The generic name "paclitaxer' is used throughout this chapter.

166 the leaves of Taxus baccata [3] opened the doors to the first semisynthesis of paclitaxel[4] and a more potent analog, Taxotere @ (docetaxel) [5] (1.1.3, Figure 1) utilizing a renewable source and thereby alleviating the supply problem substantially. As both paclitaxel and Taxotere | advanced through the clinical studies, the potential of these agents became obvious and generated considerable excitement [6]. Both paclitaxel and 10-deacetylbaccatin III, through the efforts of the Bristol-Myers Squibb company, the co-developer of the drug, became available in large amounts both to in-house chemists and outside collaborators for structure-activity relationship studies. These developments in the U.S., and also the continued efforts of the French groups at Gif and Rhone-Poulenc, led to a flurry of discoveries in the chemistry of these complex diterpenoids [7]. Paclitaxel has been approved by the Food and Drug Administration (FDA) for the t r e a t m e n t of refractory advanced ovarian cancer and breast cancer in 1992 and 1994, respectively. Due to these exciting developments, paclitaxel research has continued unabated. Years of efforts from many synthetic groups [8] finally culminated into two independent total syntheses of paclitaxel [9,10]. The syntheses are quite long and are unlikely to contribute much to the SAR in this area, at least in the near future, but represent an exciting academic achievement. O R~ II NH

O

18 R'O~,

O OH 1/19

....7~-i.~,~, 7 1,~ -,,16

o~3\

OH

7 ......6A..~ ,o

HO

6

O

'~O 1.1.1 Taxo] R = Ph, R ' = Ac 1.1.3 Taxotere R = t-Boc, R ' = H

Me

O

....

HO

:

t~ OAc

o

Bz() 1.1.2 1.1.4

R = H 10-deacety] baccatin II] R = COCH 3 baccatin I I I

Figure 1: Some medicinally important taxanes and the paclitaxe] | numbering system

At the same time, the search for n a t u r a l taxanes possessing core modifications compatible with bioactivity has begun to produce encouraging

167 results, most notably the discovery by Klein that 9-dihydrotaxanes, prepared by semisynthesis, have good biological activity [11], as well as recent reports on the activity of the 14-OH taxanes [12,13]. Research on the chemistry of paclitaxel continues for obvious reasons: the poor water solubility of paclitaxel, its rather poor in vivo potency, and its tendency to rapidly induce multi-drug resistance make it a sub-optimal drug [14]. It is hoped t h a t some of the new analogs may display improved properties in one or more of the above areas. For the pharmaceutical houses, due to the anticipated large sales of paclitaxel, the i n t e r e s t in devising unique, p r o p r i e t a r y and p a t e n t a b l e analogs of paclitaxel is obviously very strong. In addition, its high cost of production is enticing medicinal chemists to try to u n d e r s t a n d which portions of the molecule are actually interacting with its biological target and which have only minor functions or serve as scaffolding for the pharmacophore. As SAR studies become more complete, the first efforts to design a totally synthetic paclitaxel surrogate are appearing [15]. It is conceivable that more efforts in rational analog design will be reported in the near future. This chapter reviews all the relevant chemistry and structure-activity relationship studies up to the end of October 1994. Due to the existence of previous reviews on the subject [7], although comprehensive, this review will especially stress most recent key developments in the chemistry of taxanes and their implications for the SAR. We focus only on the chemistry of the core. Modifications of the C-13 side chain, which is crucial to the activity, form the topic of Chapter 6. 5.2. CHEMISTRY

5.2.1. Reactions at C-1/C-2 Functionalization of the tertiary alcohol function at C-1 without affecting the rest of the core is no trivial matter. Accordingly, few derivatives with C-1 substitution are known. Chen et al. initially reported a C-1 xanthate derivative [16a]. Subsequent reports by Klein et al. on derivatives of 9-dihydrotaxol [17] demonstrated the tendency of baccatin to undergo a C-2->C-1 benzoyl shift under basic conditions, leading, after t r e a t m e n t with carbon disulfide and methyl iodide, to 1-benzoyl-2-xanthyl derivatives. A re-examination of the earlier work [16b] verified that this is indeed the case in the baccatin series also (Scheme 1). Thus, 2.1.3 represents the first example of a C-1 acylated baccatin

168 III derivative. C-2 deoxygenation led to the deoxy analog 2.1.4. Similar results have been reported by the Kingston group as well [16c]. C-1 substitution does not imperil C-13 side chain introduction for biological evaluation. As outlined in Scheme 2, xanthate 2.1.3 was desilylated and then selectively resilylated at C-7 to give 2.1.7./~cylation according to the method of Holton [18], employing ~-lactam 2.1.10 as the side chain source, gave the desired C-1 functionalized paclitaxel derivative 2.1.9, after a final desilylation [19]. In search of a suitable C-1 protecting group, Chen and co-workers were led to utilize the novel dimethylsilane (DMS) group. Apparently, introduction of the bulkier trimethylsilyl group was difficult. As illustrated in Scheme 1, DMS was successfully introduced to the C-1 position of baccatin derivative 2.1.2 to give 2.1.5 in almost quantitative yield. Selective removal of DMS from C1 was accomplished with tetrabutylammonium fluoride at 0 ~ [20].

.o ....,,co 2o. TESO. . . . . . . . . . HO BzO

HO ~c

2.1.1 R=H -~ i 2.1.2 R=TES J

MeS 2.1.3

l iii AcO TESO....

O OTES

AcO

~0

TESO. . . . . . . . . .

20TES

Me2HSiO BzO 2.1.5

2.1.4

Conditions: (i) 5 equiv TESC1, imidazole, DMF, rt, 87%; (ii) Nail, THF+CS2, 70 ~ then MeI, 56%; (iii) 2 equiv Bu3SnH, AIBN, PhMe, 100 ~ 92%; (iv) 3 equiv Me2HSiC1, imidazole, DMF, 0 ~ 97%; (v) Bu4NF, THF, 0 ~ 84% of 2.1.2. [TES=triethylsilyl]

169 The significance of DMS protection of the C-1 hydroxyl group in the selective deacetylation at C-4 will be discussed in section 5.2.2. C-1-Benzoyl-2-deoxybaccatin 2.1.4 was converted to the corresponding C7 silylated analog 2.1.12 via a desilylation/mono-silylation sequence. The paclitaxel side chain was then attached onto 2.1.12 using Holton's protocol, to give 1-benzoyl-2-debenzoyloxytaxol, 2.1.14, after standard deprotection (Scheme 3) [19]. [ S c h e m e 21

AcO ~., RO ....\ \ BzO

Bz,

O ,~ OR'

P h ~ O

iii

=

AcO

MeS2CO 2.1.3 R=R'=TES ~ i 2.1.6 R=R'=H 2.1.7 R=H, R'=TES ~ i i

....

OR

O TESO,",,

-

O OR

NH O

Ph ],-" N. oJBz

O

BzO =MeS2CO

OAc

"

2.1.8 R=TES ) i 2.1.9 R=H

2.1.10

Conditions: (i) Py, 48%HF, CH3CN, 5 ~ 2.1.3 to 2.1.6, 76%; 2.1.8 to 2.1.9, 84%; (ii) TESC1, imidazole, DMF, 0 ~ 86%; (iii) LiHMDS, THF, -45 ~ then 2.1.10, 86%. [ Scheme 3 /

AcO RO ....

O OR'

Bz" NH

AcO ~'

O

-i i i -~ p h ~ . O

O

,,,, BzO

2.1.4 R=R'=TES ~ i 2.1.11 R=R'=H "h ii 2.1.12 R=H, R'=TES

2.1.13 R=TES "h Jr i 2.1.14 R=H

Conditions: (i) Py, 48%HF, CH3CN, 5 ~ 2.1.4 to 2.1.11, 79%; 2.1.13 to 2.1.14, 87%; (ii) TESC1, imidazole, DMF, 91%; (iii) LiHMDS, THF, 0 ~ then 2.1.10, 83%.

170 Under acidic conditions, a skeletal rearrangement of the A ring occurs, perhaps initiated by carbonium ion formation at C-1. A representative example is shown in Scheme 4. Kingston prepared ring-contracted paclitaxel analog 2.1.16 by reacting derivative 2.1.15 with methanesulfonyl chloride, followed by desilylation [21]. Later, similar rearrangements were observed by Potier [22] and Chen [23] under a variety of acidic conditions. In order to assess the contribution of the C-2 benzoate moiety to binding, a selective procedure for debenzoylation at C-2 was needed. Kingston has described a number of protocols for deacylation reactions of baccatin III. Under basic conditions, C-10 is deacylated first and, in certain cases, even the C-4 acetate is more labile than the C-2 benzoate [24]. Special conditions are therefore needed for selective C-2 deacylation.

AcO ph/~~_

0

Bz.. NH 0

Bz" NH 0 0 ....

phiaL-

i "

2.1.15

oH

O,

III

-<

o 2.1.16

Conditions: (i) MsC1, NEt3, -15 ~ to 0 ~ 20%; (ii) HF, Py, THF, 0 ~ to rt, 55%.

The first succesful attempt to selectively cleave the C-2 ester in a polyacylated baccatin derivative was reported by Farina et al., and made use of tin alkoxides or oxides. The reaction was postulated to occur by prior pentacoordination of the tin reagent to the C-1 hydroxy group, leading to rapid C-2 debenzoylation. Unfortunately, this was followed by backside attack of the C-2 alkoxide onto C-20, leading to oxetane opening, with formation of novel tetrahydrofuran derivative 2.1.18 in good yield (Scheme 5) [25]. The selectivity for the C-2 ester over the other four acetate groups is remarkable. Especially diagnostic here, in the 1H NMR spectrum, was the geminal coupling constant of the C-20 hydrogens, which increased from 8.3 Hz in 2.1.17 to 11.6 Hz in 2.1.18, and suggested a ring-expansion reaction.

171

Scheme 5~ AcO

AcO,,

O

AcO

OAc i

ii

~

AcO,

Sl I l l

Iii

o

OBz 2.1.17

O

.o, ~ .

O 2.1.18

r,,,oOc.

Conditions: (i) Bu3SnOMe, LiC1, NMP, rt, 67%.

Similar t e t r a h y d r o f u r a n - c o n t a i n i n g baccatin derivatives were later described by other r e s e a r c h groups, which used alkaline or reducing conditions to cleave the C-2 ester [11, 22, 24]. For example, the Gif group found t h a t reduction of 7-triethylsilyl-10-deacetyl baccatin 2.1.19 with lithium aluminum hydride at room temperature for a short period of time gave a 3:1 mixture of debenzoylated product 2.1.20 and the rearranged baccatin 2.1.21 in 80% overall yield (Scheme 6) [22].

.o

....

~

HO"

.o

....

o 2.1.20

2.1.19

HO .... H O 2.1.21 Conditions: (i) LiAIH4, THF, rt; 2.1.20 (60%), 2.1.21 (20%).

172 Selective C-2 debenzoylation in the presence of C-4 and C-10 acetates was finally achieved under a variety of conditions (Scheme 7).

AcO

O

AcO

OTES iorii

T E S O .... _

.

0

O

OTES

= T E S O ....

or iii

"

HO

2.1.2 Conditions: (i) NaOMe, MeOH, rt, 25%; (ii) Red-A1, THF, 0 ~

H

O

OH

2.1.22

87%; (iii) KOBu-t, THF, rt,

69%. Chen et al. reported that, whereas methoxide-promoted debenzoylation of 2.1.2 gave low yields of 2.1.22 [26], use of Red-A1, which presumably also precoordinates at C-l, led to the desired product in high yield [27]. Potassium tbutoxide was later reported by Datta et al. to perform similar function [28]. More recently, Kingston reported a simple procedure to debenzoylate paclitaxel derivatives. Thus, 2.1.15 was treated with NaOH under phase-transfer conditions to afford 2.1.23 in fair yields. Reacylation with a variety of substituted benzoates afforded a number of C-2 modified paclitaxel analogs for SAR studies (Scheme 8) [29]. Simultaneous, high-yielding C-2/C-10 deacylation in a 13-keto baccatin derivative could be carried out with catalytic bicarbonate in methanol/water [30]. The selective C-2 debenzoylation in baccatin derivatives has allowed chemists to prepare a number of paclitaxel analogs modified at C2. The simplest modification at C-2 is actually obtained by hydrogenation under forcing conditions, affording C-2 cyclohexanoate derivatives [24, 31]. In addition to the already mentioned approach directly from paclitaxel [29], baccatin derivatives modified at C-2 have been prepared by DCC-mediated coupling; introduction of the side chain afforded novel paclitaxel C-2 esters (Scheme 9) and carbamates (Scheme 10) [27]. As shown in Scheme 10, the carbamate linkage in 2.1.27 was prepared by reacting 2.1.22 with p-nitrophenyl isocyanate and pyridine in benzene at 70 ~ Standard desilylation and selective C-7 resilylation, followed by side chain acylation gave the desired

173 carbamate 2.1.29 in low yield, due to the unavoidable competitive formation of the C-1/C-2 cyclic carbonate 2.1.30 [27].

BZ'NH 2.1.15

O

p h ~ / [ L_"

AcO

20TES

0 ....

OTES

-

HO

OH

2.1.23

ii, i i i

Bz.. Ph

AcO

O

NH O . OH

0 ....

HO 2.1.24

OH

O,

R.~--[~ 0

Conditions" (i) 2 N NaOH, Bu4NHSO4, C6H6, 43%; (ii) DCC, DMAP, ArCOOH, PhMe, 50 ~ (iii) 5% HC1, MeOH, rt.

The formation of carbonate 2.1.30 underscores the instability of the C-2 carbamate linkage toward nucleophilic attack by the neighboring C-1 hydroxyl group. Similar cyclic derivatives have been reported in other publications also. For example, treatment of 2.1.22 with phosgene afforded 2.1.31 in high yield. Other attemps to protect the diol function as ketal or acetal led to complex mixtures of products, in which 2.1.32 (Scheme 11) predominated. A C2 benzyl ether could not be prepared either [19]. Another derivative that was easily prepared was the cyclic thiocarbonate 2.1.33, shown in Scheme 11 [16a]. One of the strategies for generating the C-2 benzoate in the total synthesis of paclitaxel is to utilize these C-1/C-2 cyclic carbonates. Treatment with phenyllithium in tetrahydrofuran at -78~ desired C-2 esters.

yielded regioselectively the

174

AcO

.O OTES 2.1.10

TESO ....

2.1.22

_

,,,

~Ac

0~==:0

2.1.25

R Bz,

AcO NH

0

O OH

\

....<

~~o

R a) p-MeOC6H4b) p-NO 2C6H4c) c-C6H lid) CH3~-

OH HO

-

'

O~=:=0

2.1.26

R Conditions: (i) 4 equiv DCC, xs DMAP, RCOOH, PhMe; (ii) Holton's protocol [18].

AcO

O

.h2

2.1.22

.o'~_ ~;No

2.1.27 2.1.28

R1 =R2 =TES ii R1 =TES, R2 =H J

~----~ ~ Bz. iii

TESO,,,,

AcO NH

0

0 OH

\

,,,Ph

o~'. 2.1.10

....< R10

Bz 2.1.29 2.1.30

!o OR 2

R1 =H, IRe =CONHC6H4-pNO2 R1,Ra =-C(O)-

Conditions: (i)p-NO2C6H4N=C=O, py, C6H6, 70 ~ 86%; (ii) Py, 48%HF, CH3CN, 5 ~ then TESC1, imidazole, DMF, 0 ~ 62%; (iii) LiHMDS, THF, -40 ~ 2.1.10, then (ii), 2.1.29 (15%), 2.1.30 (50%).

175 As illustrated in Scheme 12, Nicolaou et al. demonstrated t h a t treating carbonate 2.1.34 with five equiv of PhLi gave 2.1.35 in good yield [30]. Similarly, Holton showed t h a t t r e a t m e n t of carbonate 2.1.36 with 2.1 equiv of PhLi afforded the corresponding C-2 benzoyl derivative 2.1.37 in excellent yield [9]. These derivatives were subsequently transformed into paclitaxel. Complete defunctionalization of C-2 can also be achieved through the intermediacy of 2.1.22. As shown in Scheme 13, treatment of 2.1.22 w i t h sodium hydride in THF/carbon disulfide afforded xanthate 2.1.38, usually accompanied by a small amount of cyclic thiocarbonate 2.1.33. Barton deoxygenation smoothly afforded the desired C-2 deoxy derivative 2.1.39, which was acylated as usual to afford the paclitaxel analog 2.1.41. Attempted Barton deoxygenation on 2.1.33 and related analogs, on the other hand, brought about a number of interesting skeletal rearrangements, which are described fully in section 5.2.9. AcO

O OTES

T E S O .... :

.

O

OH

2.1.22

AcO

AcO

20TES

iii

T E S O ....

O OTES

T E S O .... O

O

OTES

S

2.1.33

2.1.31 T E S O .... _

HO

0

,,

OH

c

2.1.32 Conditions: (i) COC12,py, CH2C12, 0 ~ 87%; (ii) Nail, CS2+THF, MeI, then 1 equiv Nail, 83%;

(iii) acidic or basic conditions.

176

AcO

O OTES

HQ

OBOM

TBSO"' O -

",Ac

O

0 I ii

2.1.34 I

i

AcO

O OTES

2.1.36

HO%

OBOM

TBSO .... HO

2.1.37

\

2.1.35

Taxol 1.1.1

Conditions: (i) 4 equiv PhLi, THF, -78 ~

80%; (ii) 2.1 equiv PhLi, THF, -78 ~

Ac..~,(3 i

2.1.22

OBz

O OTES .

= TESO ....

85%.

,.....

L,._ ~u

.

.

.

1/ h

/ v

R 2.1.38

R=OCS2Me "~ ii t

2.1.39 R=H

HO .... AcO.

~O OTES

BZ'NH

O

ph~~['-O_

AcO~,

,60 OH

....

oH

HO

HO 2.1.40

OAc

2.1.41

Conditions" (i) Nail, THF+CS2(5:I), then MeI, 2.1.38 (61%), plus 2.1.33 (21%); (ii) Bu3SnH, AIBN, PhMe, 100 ~ 89%; (iii) Bu4NF, THF, rt, 85% (iv) TESC1, imidazole, DMF, 0 ~ 87%; (v) LiHMDS, THF, -40 ~ then 2.1.10, then Bu4NF, THF, rt, 63% overall.

177 5.2.2. Reactions at C-4: Deacetylation and Reacylation Early work on C-4 deacetylation was reported by Kingston [24] and Potier [22]. As outlined in Scheme 14, t r e a t m e n t of 1.1.4 under forcing hydrogenation conditions gave 2.2.1 in good yield. Methanolysis gave several products, including 2.2.2 and 2.2.3. Finally, extensive hydrolysis gave completely deacylated product 2.1.20 in fair yield.

AcO

~O OH

AcO i, ii

HO.... NO

Bz

=

,~O OTES iii

HO....

o

HO

O

1.1.4

O

~=2.2.1

O IIII

.o

HJ~"

....

HOA~~cO O

O

"

O

HO% ,~O OTES

He/. . HO

Hb

~

2.1.20

Conditions:

(i) Pt/H2, 90%; (ii) TESC1, Py, 80%; (iii) 0.5 N NaOMe, MeOH, rt, 69%.

Thus, it seems that deacetylation of the more hindered C-4 acetate is faster than deacylation at C-2, which is somewhat surprising. Kingston has

178 postulated that this occurs because of an initial base-catalyzed intramolecular acetyl transfer from C-4 to C-13, followed by rapid deacetylation. In agreement with this, if the C-13 hydroxyl is protected, the order of reactivity is reversed. In any case, the difference in reactivity is not satisfactory under these basic conditions, and completely selective C-4 deacetylation cannot be carried out [24]. Similarly, Potier's group reported attempted selective deacylation of C-4 under a number of basic and reductive condition, but in every case mixtures of products were obtained [22]. A solution to this problem was formulated by Datta et al., who treated 7triethylsilyl baccatin III 2.2.4 with potassium t-butoxide, yielding the desired 4deacetyl derivative 2.2.5 in fair yield (Scheme 15). This was explained once again by assuming acetyl transfer from C-4 to the free hydroxyl at C-13. The use of the bulkier base presumably retards direct deacylation at C-10 and C-2, which was highly competitive with methoxide [28].

AcO

O OTES

AcO i

HO ....

/

[

,," HO .... O

~O

BzO

BzO

2.2.4

2.2.5

Conditions: t-BuOK, THF,-20~

O OTES

58%.

Kingston et al. applied this chemistry to the preparation of 4-deacetyl-10acetyltaxotere, 2.2.8 (Scheme 16) and its related paclitaxel analog [32]. The starting material, 2.2.6, was prepared by the above t-butoxide-promoted hydrolysis of 2.2.1 which, in this case (temperature is not specified by the authors) apparently led to both C-4 and C-10 deacylation, contrary by the report of Datta and coworkers [28]. Coupling with synthon 2.2.7 was followed by C-10 reacylation and final deprotection.

179

h/h/t/

H OIIII

Boc~NZO

~

H

2.2.7

2.2.6

Boc,, NH O

\

AcO

O

OH

OH

HO BzO 2.2.8 Conditions: (i) DCC, DMAP, PhMe, 90%; (ii) Ac20, DCC, 4-(pyrrolidino)pyridine, THF, 65%; (iii) HCOOH, rt, 46%.

A more general solution to C-4 deacylation, which does not depend on the presence of a free hydroxyl group at C-13, was reported by Chen et a/.[20]. Since Red-A1 reduction of baccatin III seems to take place at the C-2 ester because of coordination with the hydroxyl group at C-l, it was felt that blocking this functionality would redirect the r e a g e n t elsewhere in the molecule, and hopefully to C-4, perhaps by prior coordination with the oxetane oxygen, the most basic oxygen in the molecule. A suitable C-1 blocking group was therefore developed. The a l r e a d y m e n t i o n e d dimethylsilyl (DMS) was found to be optimal. Both its introduction and its removal could be carried out under mild conditions. As illustrated in Scheme 17, reduction of 2.1.5 with excess Red-A1 afforded the desired C-4 deacylated baccatin 2.2.9, together with a smaller a m o u n t of the C-4/C-10 di-deacylated baccatin 2.2.10. The undesired baccatin 2.2.10 was easily reacetylated in situ to 2.2.9 by the method of Greene [4]. A more efficient approach to C-4 deacylated derivatives utilized C-13 trimethylsilyl

derivative

2.2.11 as the s u b s t r a t e (Scheme 18). Baccatin

derivative 2.2.12, bearing three different types of silyl groups, was treated with Red-A1 and then quenched with a s a t u r a t e d solution of t a r t r a t e , giving the

180 desired C-4/C-13 diol 2.2.13 in acceptable yield [20]. Loss of the TMS group occurred during the acidic work-up, as planned. Derivative 2.2.13 is obviously an ideal substrate for side chain attachment and the preparation of C-4 modified taxanes.

T

SO ....

Me2HSiOJ B ~

H OA~~cO

2.1.5

AcO

OTES

TESO ....

!0

Me2HSiO

+

~

O OTES,

TESO .... ~

Me2HSi

BzO 2.2.9

Conditions:

HO

BzO

2.2.10

(i) Red-A1,THF, 0 ~ (3:1) 2.2.9/2.2.10 (84%); (ii) Py, AcC1, 5 ~ 84%.

I Scheme18 /

AcO TMSO,,,

n

O

us se

~o,7~. ~ ~d~o 2.2.11 R=H "h i 2.2.12 R=DMS

AcO ii

O

OTES

HO .... Me2HSiO BzO

(

o

2.2.13

Conditions: (i) Me2HSiC1, imidazole, DMF, 0 ~ 92%; (ii) Red-A1, THF, 0 ~ then Na,K tartrate, 50-60%.

181 Preparation of 4-deacetyltaxol can be carried out quite efficiently beginning with paclitaxel, and using the selective 2,4-deacylation reaction reported by Kingston et al. A s shown in Scheme 19, 2.2.16 is obtained in high overall yield by phase-transfer promoted deacylation followed by rebenzoylation [32]. Synthesis of a 4-deacylated taxane for biological evaluation was also reported by the Gif group, following similar chemistry [33]. As we have seen, several solutions exist for selective C-4 deacylation and, not surprisingly, a number of workers have recently focused on the reacylation (or other derivatization) of C-4 for SAR studies at this position. This operation turned out to be non-trivial. During the course of an extensive deacylation/acylation study on baccatin derivatives [24], Kingston found that selective benzoylation of 2.2.17 led to derivative 2.2.18. The hindered C-4 hydroxyl group in 2.2.18 could not be acetylated even under forcing conditions and only 2.2.19 was obtained.

BZ'NH O ph~ L ~ . . , O .... -

~sss 9

~

HO"

iii

Ace,,.

BZ'NH 0 ph~~"O. OH

i ph/~,~ -

o

2.2.14

ii

AcO

BZ'NH O 0 ....

O

ss ss ~

HO'# 2.2.15

OH

OH

....

2.2.16 Conditions: (i) Benzyltrimethylammonium methoxide, CH2C12; (ii) PhCO2H, DCC, DMAP,

PhMe; (iii) 5% HC1, MeOH (42% overall). However, direct acetylation of 2.2.17 under standard conditions gave tetraacetyl derivative 2.2.20 in modest yield, where the C-4 carbinol was successfully acetylated (Scheme 20). Given these observations, Kingston speculated that the

182 inaccessibility of C-4 hydroxyl group in 2.2.18 towards acetylation is due to the steric hindrance imposed by the bulky C-2 benzoate moiety. In any case, the yields reported are too low to draw any definite conclusions. Chen et al. reported a general solution for derivatization of the C-4 alcohol moiety, and this has led to the preparation of several C-4 modified paclitaxel analogs [19, 20, 34]. As shown in Scheme 21, the fully protected baccatin derivative 2.2.9 (obtained as in Scheme 17) has only the C-4 carbinol open for derivatization. Treatment under highly basic conditions with a variety of acyl chlorides afforded the C-4 analogs in high yields. These were all converted to the respective paclitaxel derivatives via the usual Holton procedure.

Aco..... o" ,~. o~ ~'ii

2.2.17

AcO

.... . o "

o

i i~,,~

AcO . . . . . .

2.2.18

AcO

O OTES

O

,,

.o"-~~o~o~~ 2.2.20

AcO .... "

HO

H

OBz

2.2.19

Conditions: (i) DCC, DMAP, PhCOOH, 14%; (ii) Ac20, CH2C12, DCC, rt, 24%.

THF, DCC, 55 ~

89%; (iii) Ac20,

183 The authors stress that the success of this protocol depends highly on the following two facts: (i) protection of the C-1 hydroxyl group as a dimethylsilyl ether (DMS) allows clean C-4 deprotonation without any complication; (ii) the choice of a strong base, such as LiHMDS or BuLi (weaker bases are ineffective), is crucial.

AcO O ~ F - ~ .'~ . ~ ~ "// OTES t

AcO O , ~ , ~, P ~// " OTES

i

so ....

z x__#~

/

-

~

ii

so,,,

,%

~

OBz O , ~ R 2.2.9

AcO ~, HO ....

O

//

2.2.21

OR'

Bz. 2.1.!0

,.... O OBz O , ~ R

=

O O

AcO NH

-

O

II

= P h ~ O

....

OH

H

'~'

o" BzO

O,~R

0 2.2.22 2.2.23

2.2.24

R'=H

R'=TES

O

iii

R "a, CH3; b, CC]3; c, CH2CH3; d, CH2CH2CH3; e, CH(CH3)2; f, (CH2)3CH3; g, (CH2)4CH3; h, Ph; i: p-NO2-C6H4;

Conditions: (i) LiHMDS, THF, 0 ~

then acyl chloride, 70-85%; (ii) Py, 48% HF, CH3CN, 5 ~

(iii) TESC1, imidazole, DMF, 0 ~

Similar strategies led to the successful preparation of C-4 carbonates (Scheme 22). Once again, the corresponding paclitaxel analogs were prepared without complications by Holton acylation of 2.2.27a,d.

184

AcO ~, i

2.2.9

TESO ....

AcO

O // OTES HO

,,

DMSd ~

H d WOBz

2.2.25

AcO

isssss

i i i i

~ ..-.~ O OBz OCO2R

Bz.

O

O OH

OCO2R

2.2.26

R'=H

2.2.27

R'=TES

~ iii

2.1.10 y

Ph

: OH

R: a, Me;b, Et; c, n-Pr; d,p-NO2-C6H4

O ....\

HO

2.2.28

&Bz OCO2R

(i) LiHMDS, THF, ROCOC1; (ii) Py, 48%HF, MeCN, 5 ~ (iii) TESC1, imidazole,

Conditions:

DMF, 0 ~

AcO

O OTES

TESO ....

i

DMSO

OBz O

2.2.25d

AcO~,

AcO

O~

DMSd ~ ~ N 0 2

NH O P h ~ O OR

I"11-~,,,.0 OBz OCONHR

AcO

O OH

i ....

HO Bz(3

OCONHR

2.2.32

R" a, n-Pr; b, cyclopropyl; c, cyclobutyl Conditions:

ii

2.2.29

Bz. 2.1.10

OBz OCONHR 2.2.30 R'=H iii 2.2.31 R'=TES

._~ TESO .... ~ , ,% . /, / ~ OTES ~ "

O

,~O OR'

HO ....

O

(i) RNH2, THF, rt; (ii) Py, 48% HF, MeCN, rt; (iii) TESC1, imidazole, DMF, 0 ~

185 To prepare C-4 carbamates (Scheme 23), these authors resorted to a stepwise strategy. Reactive carbonate 2.2.25d was the key intermediate. Its reaction with amines led to displacement of the p-nitrophenol moiety and produced a number of carbamate derivatives. Conversion to the paclitaxel analogs by standard Holton acylation was complicated by side reactions (vide infra), and afforded only modest yields of the desired 2.2.32. Scheme 24 shows the synthesis of the C-4 aziridine carbamate. It was found more practical here, instead of using aziridine as a nucleophile, to construct the 3-membered ring stepwise.

AcO

2.2.25d

NO ....

0 iii

,,,,

HdW

.-%1o

OBz OC020BHn-p-N02 2.2.33 R=H ~ ii 2.2.34 R=TES

AcO

0

AcO

HO ....

iv

tt o

Hd_= H.-%10 OBz 0 H 2.2.35

"~ N 0

HO ....

OBz 0

BZ'NH~v ~ 0 Ph

_ OH

,,,,

2.2.36

OH AcO

2.1.10

0

N~

0~

0

~-r 9

OH

0 .... HO

2.2.37

: OBz

0 O~

N~

Conditions" (i) Py, 48% HF, MeCN, rt; (ii) TESC1, imidazole, DMF, 0 ~ (iii) NH2(CH2)2OH, THF, rt; (iv) PPh3, DEAD, THF, rt, 50%.

186 In the event, p-nitrophenyl carbonate 2.2.33 was silylated at C-7, then quantitatively converted to 2.2.35 by t r e a t m e n t with ethanolamine. Standard Mitsunobu conditions gave aziridine 2.2.36. In this case, the paclitaxel side chain was introduced onto 2.2.36 in high yield to give the desired C-4 aziridine carbamate 2.2.37, after standard desilylation. A clue as to the problems associated with the Holton acylation of carbamates 2.2.31 was provided when acylation with modified lactam 2.2.40 was attempted under standard conditions. Surprisingly, acylation at the C-4 c a r b a m a t e nitrogen was observed as the major pathway, as outlined in Scheme 25. In two cases, the unwanted 2.2.39 predominated, while the desired C-4 carbamate analogs 2.2.38 were only minor products. This problem is then due to the acidic nature of the carbamate proton. The high-yielding acylation of the aziridine analog 2.2.36, which bears no such acidic proton, suggests that protection of the C-4 carbamates to remove the N-H function should result in a generally high-yielding C-13 acylation.

AcO ~,

B~ ..NH 0 i, ii 2.2.31a,b

AcO

O ~ OH

" -~0,

~-

III

2.2.~a,b

OH

HO

OBz 0 ~/--- NHR

HO..../

Hd ~OBz~O 2.2.39a,b

0

TESO,%

R ,,..[ ~ / Boc-N H

Conditions: (i) LiHMDS, THF, 2.2.40; (ii) Py, 48%HF, MeCN, rt.

,,,,,

oJ-' N. Boc 2.2.40

187 Recently, Georg and co-workers have also reported the synthesis of a C-4 modified analog of paclitaxel (2.2.43, Scheme 26). The approach relies on the intermediacy of cyclic carbonate 2.2.41, which is used as a protecting group for the C-1/C-2 diol system while performing C-4 acylation with a large excess of acylating agent. Although no further examples are reported, the procedure may have general utility [35].

BZ'NH Ph

O

"

_

AcO ~

O -//- OTES

0 ....

~ H

2.2.14

Ph

.

o,

,, - p.A_Ao:_

_

iii

~.-O Bz.

NH -

AcO

O OH

~

Ph/~~__ O.... OH

2.2.43

OH

O~H

O

0

OH

....

OTBS 2.2.41

BZ'NH

HO

LO OTES ~

O

_O OTES

0 ....

2.2.15 AcO~

BZ'NH

AcO

BZ.NH

OBz

0

2.2.42

I

=

?=o

20TES

II

Ph/~flL'" O.......... OTBS _

~0 HO

AcO

O

i v, v

/

Conditions: (i) t-BuOK, H20 (2 equiv), THF, -20 ~

75%; (ii) (Im)2CO (20 equiv), THF, 55 ~ C ; (iii) (iBuCO)20 (20 equiv), DMAP (20 equiv), CH2C12, rt, 67% overall; (iv) PhLi (10 equiv), THF,-78 ~ (v) Py, HF, 0 ~ to rt, 48% overall.

Kingston and co-workers recently reported a study aimed at deoxygenation of the C-4 position [36]. Deacylation of suitably protected

188 paclitaxel and C-1/C-2 protection gave 2.2.41 as in Scheme 26. Conversion to the xanthate proceeded uneventfully, and Barton deoxygenation geve the desired 4deacetoxy derivative. Final deprotection turned out to be difficult because of the competitive acid-catalyzed oxetane ring opening, which was reported to be faster than in the 4-acetoxy-bearing analog, in spite of the lack of anchimeric assistance by the ester group (see section 5.2.3). Nevertheless, HF in pyridine gave the target 2.2.45 in fair yield (Scheme 27).

AcO ~'

Bz. NH 0 i

p h ~ . J [ , . O _ ....

2.2.41

OTBS

AcO

BZ-NH 0 Ph

-

0 ....

0 ~O

2.2.44 ii, i i i

00TES

(~CS2Me

O O

sss S

2.2.45

(i) Nail, CS2, MeI, 90%; (ii) Bu3SnH, AIBN, PhMe, 90 ~ 80%; (iii) PhLi, THF, -78 ~ 60%; then HF, py, rt, 66%. Conditions:

5.2.3. The chemistry of the oxetane moiety In a study dealing with the reaction of paclitaxel with electrophilic reagents [21], Kingston found t h a t reacting paclitaxel with Meerwein's reagent (triethyloxonium tetrafluoroborate) gave, in low yield, 2.3.1, a compound where the oxetane ring had been opened. The presence of the diol moiety was subsequently confirmed by formation of acetonide 2.3.2. The A ring contraction reaction, already briefly mentioned in section 5.2.1, apparently also took place, due to the acidic conditions employed. Kingston noted that reaction of paclitaxel with acetyl chloride directly yielded both A-ring contraction and oxetane opening, in addition to C-5 acetylation (Scheme 28). The oxetane-opening chemistry was independently investigated by Gu~ritte-Voegelein [37], using baccatins as substrates.

189

BZ'NH Paclitaxel 1.1.1

0

phil_

0 .... -

sts

OH

OH 2.3.1

BZ.NH

O

ph~~Jl"O

AcO~, ....

.

2.3.2

iii

OAc

L O OH

OH

Paclitaxel 1.1.1

OBz

~k

OBz OAc

\

Bz. NH

AcO ~

O

P h ~ O , ,

'"O

/

O ... // uAc

'

OH 2.3.3 -

'

\

OBz

O

"'OAc OcH

Conditions: (i) Et3OBF4, CH2C12, 35-51%; (ii) Me2C(OMe)2, p-TsOH, CH2C12, 95%; (iii) MeCOC1, then H20, 68%.

Reaction of baccatin derivative 2.3.4 with anhydrous zinc chloride in toluene was found to give 2.3.5, again with an opened oxetane ring and a contracted A ring. Treatment of 2.3.4 with aqueous acid, on the other hand, gave a mixture of three products, i.e. 2.3.6, featuring an intact oxetane ring, in addition to regioisomers 2.3.7 a n d 2.3.8 [22] (Scheme 29). T r e a t m e n t of paclitaxel and baccatin derivatives with Lewis acid has been discussed in a rather comprehensive study [23]. Use of strong acids, such as boron halides or TMSBr gave, in addition to A-ring contraction, oxetane opening with complementary regiochemistry (Scheme 30), but weaker Lewis acids smoothly led to oxetane opening w i t h o u t concomitant A-ring transformations (Scheme 31). Once again, two regiochemical modes are possible for the oxetane solvolysis, and the ratio of 2.3.12 to 2.3.13 depends on the Lewis acid used [23].

190

Scheme 29

_

HO

TrocO ,~

O //

TrocO

O.Troc

....

O

= HO ....

,,,

0 HO

: AcO OBz

2.3.4

~

' \

OBz

OAc

2.3.5

ii

_

TrocO ~i,

O //-

TrocO

_OTroc

HO ....

O

HO .... 0

HO'\

",

OBz

OBz

Hd \

2.3.6

2.3.7 2.3.8

Conditions: (i) ZnC12, PhMe, 80 ~

R2

OR 1

RI=AC , R2=H R I=H, R2=Ac

50%; (ii) 1 N HC1, AcOH, rt, 2.3.6 (20%), 2.3.7 (30%), 2.3.8

(17%).

BZ'NH Ph

O

AcO

2

OH

: O .... ()Cbz 2.3.9

HO

0

ii

" AcO OBz BZ.N H

O

AcO

ph-~.~_ 2.3.10

R 1 =Ac, R 2 =H

2.3.11 R 1 =H, R 2 =Ac

O

-

O .... '"O R2

OCbz

\

OBz

Conditions: TMSBr, CH2C12, rt, 2.3.10, 79%; (ii) BBr3, CH2C12, rt, 2.3.11, 75%.

OR1

191

Scheme 31 /

~,, II O.H

AcO

az.

2.3.9

, ewi.

NH

0

acia

0 OCbz

Lewis acid SnC14 TiCl4 BF3

2.3.12 (%) 17 57 43

2.3.13 (%) 69 15 43

0

....

. . . . . . OBz

,,OR 2 OR 1

2.3.12 R1 =Ac, R2 =H 2.3.13 R1 =H, Re =Ac

A variety of paclitaxel and baccatin substrates were submitted to Lewis acid treatment, and similar products were obtained. Some qualitative kinetic information could be gleaned from these studies: for example, paclitaxel itself reacts very slowly with tin tetrachloride. Once the C-2' hydroxyl group is blocked as a carbonate, however, high reactivity is restored. Also, 7-epi derivatives react very sluggishly in comparison with their 7(~) counterparts. From all these observations, a mechanism can be proposed for the oxetane opening reaction that requires the anchimeric participation of the C-4 acetate group, and is consistent with proposals by Kingston [21] and Chen [23] (Scheme 32). The reaction is initiated by complexation of the Lewis acid with the oxetane oxygen, which is probably the most basic atom in the molecule. The acetoxy group is positioned for participation by backside attack onto C-5, leading to acetoxonium ion 2.3.15. Indeed, when the C-4 acetoxy group is not as readily available for nucleophilic displacement (it is hydrogen bonded to the C7 epi-hydroxyl group, and it apparently interacts with the C-2' hydroxyl group in paclitaxel), the overall reaction is sluggish at 0 ~ and only proceeds at room temperature. Intermediate 2.3.15 can isomerize to acetoxonium ion 2.3.17, via strained orthoester 2.3.16. Alternatively, 2.3.15 can be hydrolytically quenched to yield the hemiorthoester 2.3.18, which can further unravel to afford 2.3.13, bearing a C-5 acetoxy group. Similar quench of 3.2.17 leads to 2.3.12, featuring a C-20 acetoxy group [23]. As a consequence, the regioselectivity of the reaction is a reflection of the ratio of the two acetoxonium ions, 2.3.15 a n d

192 2.3.17, and this ratio seems to be highly dependent on the Lewis acid used, either due to steric or electronic reasons.

OH

OH OSnCl4- H27

oH// I

OH ..

.[,~

Me

T|

2.3.15

~O H

2.3.13

2.3.12

)=0 2.3.14

2.3.18

Me

+_SnCi4_ Me

"0

0-~

It o. ~

o.l H20 OSnCI4-

Oto 2.3.17 Me

OH Oe~:

M

2.3.19

H

F u r t h e r evidence supporting the above mechanism came from the observation that, when 2.3.9 was treated with tin tetrachloride for 2 h at 0 ~ an unusual product was isolated in c a . 20% yield. It was assigned structure 2.3.20 (Scheme 33) aider extensive NMR characterization [23]. The formation of 2 . 3 . 2 0 can be explained by assuming t h a t the conformation of ring C in 2.3.17 can flip to a boat, in which the 7-OH group can attack the C-20 methylene carbon, to lock the C ring permanently in a boat-like conformation. Compound 2.3.21 was isolated in low yield when 7,13diacetylbaccatin III was solvolyzed under similar conditions. The product is the result of a 1,2 hydride suprafacial shift from C-5 to C-4 in an intermediate of the type 2.3.17, and leads to inversion at C-4, as shown by NMR data.

193

Bz" NH O

AcO

O

AcO

O

p h ' ~ ' J l " O _ .... OCbz

AcO ....

O //

OAc

.~, O

'"O HO

OBz

OBz

OAc

2.3.21

2.3.20

Attempts to trap acetoxonium ions 2.3.15 and 2.3.17 with nucleophilic reagents in the presence of Lewis acids proved fruitless, but stirring baccatin III in ice-cold trifluoroacetic acid in the presence of a large excess of phenyl dimethylsilane produced acetals 2.3.22 and 2.3.23 in fair overall yield, suggesting that the two acetoxonium ions were indeed both present, at least under these particular conditions [23] (Scheme 34). As shown in Scheme 35, in order to f u r t h e r confirm the diol functionality present in structures 2.3.12 and 2.3.13, both compounds were cleaved using lead tetraacetate in acetonitrile. While 2.3.13 gave the expected product 2.3.24 in high yield, 2.3.12 cleanly produced the unexpected 2.3.25, the result of C-20 to C-7 acetyl migration followed by exocyclic cleavage. Endocyclic cleavage was only a minor product [23].

AcQ

O

i Baccatin III, 1.1.4

OH,

+

=HO ....

N O IIII

..

"OH

"" 0 -

OBz

O.~H

2.3.22

Conditions:

CF3CO2H, PhMe2SiH, 0 ~ 2.3.22(42%), 2.3.23(9%).

HO BzO 2.3.23

\

H

H

194

IScheme 35 /

AcO Bz. NH O 2.3.13

~,

= ph.,.-'L,,~O. .... OCbz

BZ'NH O -

Ph

"

:

OBz

AC!~ 0'"

OCbz 2.3.25

'"OAc HO

2.3.24

2.3.12

O OH

0

20Ac

i

sis

OH

HO

"

OBz

O

Conditions: (i) Pb(OAc)4, MeCN, rt, 71-80%.

The chemistry of the novel 2.3.20, featuring a bridged C ring, was also studied. As shown in Scheme 36, t r e a t m e n t of 2 . 3 . 2 0 u n d e r s t a n d a r d acetylation conditions only led to a C-5 acetate derivative, 2.3.26, via a formally i n t r a m o l e c u l a r transesterification process. Oxidation of 2.3.20 gave the expected C-5 keto analog. When 2.3.20 was treated with DAST, an interesting rearrangement took place, leading to another bridged analog, 2.3.29, via a 1,2 alkyl migration through 2.3.28 [19]. 5.2.4. Reactions at C-7 Epimerization of the C-7 hydroxyl group to the 7((~) isomer 2.4.1, presumably via a retroaldol/aldol sequence, was first described by Kingston in his a t t e m p t s to promote radical reactions at t h a t position [38]. A more convenient way to effect epimerization at C-7 is to treat paclitaxel with base, as shown in Scheme 37 [19]. The 7((z) isomer is apparently more stable than paclitaxel. The most direct way to assess the importance of the binding of the C-7 hydroxyl group within its biological target is to replace it with a hydrogen

195 atom. Thus, it is not surprising that several groups have engaged in research aimed at such deoxygenation reaction.

BZ"NH O 2.3.20

phil'_

AcO

2

O

0 ....

OCbz

HO

-

2.3.26

OAc

O Bz

AcO

O

O

BZ"NH O phil_

0 , ~ ::SF2NEt2

0 ....

HO

2.3.27

2.3.28

t ss

, co 2 P h ~ O .

= OBz

o\

....

OCbz 2.3.29

HO

OBz

(i) Ac20, Py, CH2C12, 77%; (ii) Jones reagent, Me2CO, rt, 82%; (iii) DAST, CH2C12, 0 ~ 74%. Conditions:

Bz" NH 0 Taxol 1.1.1

AcO ~,

0 //

.OH

i, orii, o r i i i

p h / ~ ~ _ . 0 .... 8N

o OBz 2.4.1

Conditions:

rt, 65-80%.

(i) AIBN, PhMe, reflux; (ii) DBU (2.5 equiv), PhMe, reflux, 84%; (iii) Nail, THF,

196 Initial attempts involving Barton type reaction on C-7 derivatives such as thionobenzoates [39], selenocarbonates, oxalates [19] proved fruitless. After this quite extensive search, both Kingston et al. [40] and Chen et al. [41] found an identical solution to the problem of C-7 deoxygenation via xanthates. Chen and Farina prepared both 7-deoxytaxol and 7-deoxytaxotere via 7-deoxybaccatin, as shown in Scheme 38.

AcO

0

,.,.

9

AcO O -~_~.~.

OCS2Me

iii

R O ....

0

0 HO

: OBz

OAc

HO

1.1.4 AcO

OAc

2.4.2 R=H 2.4.3 R=TES O

AcO

R O ....

-~

/~

O HO

: OBz

" OBz

2.4.4 R=TES 2.4.5 R=H

ii O

,9-L,

Ph

-

OAc

O .... O

OH HO TESO,

) iv

%

,,"

o•1

Ph

OBz

OAc

2.4.7 R=Bz 2.4.8 R=Boc

N. R

2.1.10 R=Bz 2.4.6 R=Boc

Conditions: (i) Nail, THF+CS2, MeI, 57%; (ii) TESC1, imidazole, DMF, rt, 74%; (iii) Bu3SnH, AIBN, PhMe, 110 ~ 83%; (iv) TBAF, THF, rt, 74%; (v) LiHMDS, THF, -40 ~ then 2.1.10 or 2.4.6; then 1 N HC1, CH3CN,-5 ~ 53% of 2.4.7; 77% of 2.4.8.

Formation of the C-7 xanthate was complicated by some competitive C-7 epimerization. Deoxygenation proceeded smoothly to afford 2.4.4. Deprotection and attachment of the side chains gave 2.4.7 and 2.4.8 [41]. Kingston produced a 7-deoxy derivative directly from 2'-protected paclitaxel using the same chemistry (Scheme 39) [40].

197 Incorporation of fluorine atom into biologically active molecules has become an i m p o r t a n t facet of medicinal research. Consequently, C-7 fluorinated paclitaxel was an interesting synthetic target. T r e a t m e n t of 2'protected paclitaxel derivative 2.3.9 with two equivalents of DAST led to 7(cz) fluoro analog 2.4.11, together with side product 2.4.13, which features a 7,19cyclopropane ring. If a larger excess of DAST was used, the already discussed A ring contraction also took place. The structure of 2.4.11 seems secure on the basis of NMR studies, including appropriate values of JH,F. Also, the structure of 2.4.13 was confirmed by NMR studies and, after cleavage of the side chain, by a single crystal X-ray of the corresponding baccatin analog (Scheme 40) [42, 43].

,,,

gz..

AcO

S H

O

OH

_~I

~O' ~

SMe ii

ph: ~ ~ _ O, OTES

2.4.7 -

HO

OBz

OAc

o

iii

o

OBz

2.4.9

2.4.10

Conditions: (i) Nail, CS2, THF, then MeI, 60%; (ii) Bu3SnH, AIBN, PhMe, refl.; (iii) dil. HC1,

49%.

gz.

i

2.3.9

=

AcO~ ~0 F NH 0

ph-2- _

1111

OR Bz.

AcO

O

NH 0

_

HO

: OBz

0

OAc

2.4.11 R=Cbz 2.4.12 R=H

ii

0 2.4.13 R=Cbz 2.4.14 R=H

ii

O.... OR HO

-

AcO

OBz Conditions: (i) 2 equiv DAST, CH2C12, rt, 2.4.11 (55%), 2.4.13 (32%); (ii) H2, Pd/C, EtOAc, 2.4.12 (88%), 2.4.14 (90%).

198 Standard cleavage of the Cbz group gave the modified 2.4.12 and 2.4.14 for biological evaluation. A subsequent study examined solvent effects in the DAST fluorination reaction [44]. When the reaction was run in THF/ether instead of dichloromethane, 2.4.11 was the major product, accompanied by 10-12% of a new side product, the very interesting 6,7-dehydrotaxol analog 2.4.15 [44, 45] (Scheme 41). An analogous derivative was later prepared by Kelly and co-workers, using a C-7 triflate and effecting its base-promoted elimination in high yields [46].

AcO~, gz.

2.3.9

NH

,O

O

= phil'_

0 ....~

/

~

+ 2.4.11

OR

HO

: OAc OBz 2.4.15 R=Cbz ~ ii 2.4.16

R=H

Conditions: (i) 2 eq. DAST, THF/Et20, 2.4.15 (10%), 2.4.11 (45%); (ii) H2, Pd/C, EtOAc, 87%.

Note that saturation of the C-6/C-7 double bond was not possible even under forcing conditions [19]. A separate study deals with the fluorination of 7epi-taxol derivatives [47]. As shown in Scheme 42, with this substrate the DAST reaction produces only cyclopropane derivatives, no fluorination being detectable in this case. Further treatment of 2.4.13 with DAST gave A ringcontracted product 2.4.18. The lack of fluorinated products in this case suggests that attack of fluoride at C-7 from the ~ face is too hindered. Instead, an unusual participation of the trans-diaxially positioned angular methyl group ensues, leading to protonated cyclopropane 2.4.20 (Scheme 43). Similar intermediates probably occur in 1,2 Meerwein rearrangements. The deactivating effect of the C-9 carbonyl was postulated to prevent the completion of the 1,2 shii~, therefore leading to an isolable cyclopropane derivative. When the substrate was the 7(~)OH group, direct methyl participation is electronically unfavorable, and in this case (Scheme 44) the reaction proceeds probably through a carbonium ion, a

199 common i n t e r m e d i a t e t h a t easily explains the formation, in addition to 2.4.13, of the fluorinated compound 2.4.11 and the olefinic analog 2.4.15.

AcO BZ'NH Ph

O :

O OH ---~

~ O ....

=

OCbz

O HO

OAc OBz

2.4.17

AcO BZ'NH

0

\

Ph 'o. --

O

)

BZ'NH

/~%.

Ill

phil'_

I

--

oc z

AcO ~'

O

O /J

Ollll

ocbz

~ " " A: H OU c OBz

2.4.13

~

o

ii

2.4.18

Conditions: (i) 2 equiv DAST, CH2C12, rt, 2.4.13 (28%), 2.4.18 (31%),.; (ii) 4 equiv DAST,

CH2C12, rt, 2.4.18 (81%).

Bz. NH

AcO

O

phil__

0

~

.... 0 DAST=

O ....

OCbz

O

HO 2.4.17

BZ'NH

O

AcO ~

"

OBz

OAc

O

ph/~/[L'O,,, OCbz ' 2.4.13

H 2.4.19 X=OSF2NEt2

-X-I H H H+

0

0 = HO

: OAc OBz 2.4.20

H

200

O 2.3.9

X

DAS~

O O

2.4.21

-X ._-

+

H X=OSF2NEt2

$ methyl participation H H

~+ F

O

H

2.4.22

_H§

O 2.4.20

2.4.11

_H+

2.4.15

2.4.13

Formation of the 7(~)-fluoro derivative seems to be prevented in each case by the steric hindrance to approach of fluoride from the top face of the molecule. Some recent results by Klein have indirectly confirmed this mechanism: as shown in Scheme 45, when the C-9 carbonyl is absent in the substrate, and the C-7 hydroxyl group is activated as a transient triflate, a 1,2 methyl shift is observed, leading to skeletal r e a r r a n g e m e n t and eventually to B-ringcontracted product 2.4.25. Interestingly, a small amount of cyclopropanecontaining product 2.4.26 was found here also. The authors suggest that the peculiar conformation of the C ring may be at the origin of this very unusual cyclopropanation reaction [48]. Oxidation of taxanes with various agents was studied by Kingston [49]. Treatment of paclitaxel with chromic acid yielded first the C-7 keto derivative 2.4.27. Reacting this ketone with DBU in CH2C12 at room t e m p e r a t u r e or simply c h r o m a t o g r a p h y on S i 0 2 caused oxetane ring opening via ~elimination, leading to 2.4.28. Saturation of the 5,6-double bond and subsequent reaction with warm methanol led to lactone 2.4.29 (Scheme 46). Others have found that t r e a t m e n t of 2.4.27 with DBU leads not to 2.4.28, but to isomeric

201 enone 2.4.30, where the strong base has catalyzed 1,2 acetyl shift from C-4 to C20 [19, 33].

AcO ~,

-

OH

AcO

OH

fOH -I

AcO ....

i

= AcO ....

J, 0

0 HO

OBz 2.4.23

OAc 2.4.24

OAc

AcO .... ~ - ~ ~ , , , ,

~

AcO

Me

~]~

+

AcO ....

'~

OH

"" 0

HO

: OBz 2.4.25

OAc

HO

OBz 2.4.26

OAc

Conditions: Tf20, CH2C12, Py, rt; 2.4.25 (56%); 2.4.26 (8%).

Esterification at C-7 is a rather straightforward operation. Kingston reported that paclitaxel is rapidly acetylated at C-2', but C-7 acetylation requires more forcing conditions, involving DMAP and DCC. Selective 2'deacetylation under mildly basic conditions afforded C-7 acetyltaxol [50]. Baccatin III yields 7-acetyl baccatin under standard acetylating conditions (Ac20, pyridine), the C-13 position being somewhat more difficult to acetylate. A C-7 carbonate derivative of baccatin was described in the same paper [51]. Using similar methods, other workers have reported the synthesis of more complex esters, including water-solubilizing moieties, to be used as possible paclitaxel prodrugs [52-54]. Synthesis of sulfonate esters has also been reported under standard conditions [45, 55]. The same paper also reports the preparation of C-7 carbonates. While simple unhindered chloroformates react smoothly at C-7, more substituted ones react only with difficulty, if at all [19]. The same study [45] also describes the stepwise preparation of C-7 carbamates, which turned out to be rather challenging, since typical one-step procedures failed due to the

202 hindered nature of this secondary hydroxyl group. Two methods for the preparation of carbamates are shown in Scheme 47.

AcO i Paclitaxel 1.1.1

0 //

BZ'NH Ph

0

0

~

: OH

/j

0

0 ....

" 0 OBz

2.4.27

0

AcO BZ'NH

/,0

0

iii Ph

OH

-

0 ....

9

OH 0

2.4.?,8 2.4.29

AcO

O O

Bz" NH O

Ph~O OH 2.4.30

.... HO

Ac

Conditions: (i) Jones reag., acetone, 50%; (ii) Si02, 70%; (iii) Pt/C, MeOH, H2, 77%.

In the first case, the C-7 chloroformate is formed in s i t u and immediately quenched with the required amine. In the second case, pnitrophenyl carbonate 2.4.33 is readily prepared and isolated. Treatment with amines then yielded the desired derivatives [45, 55]. Silylation at C-7 (usually the sturdy triethylsilyl group is used) represents the preferred protection procedure for the C-7 hydroxyl. Standard fluoride deprotection regenerates the hydroxyl group [21]. Silylation of the 7(a) epimer of paclitaxel is difficult because of intramolecular hydrogen bonding between the 7-OH group and the C-4 acetate moiety, but it has been recently achieved by operating in highly polar solvents [45].

203

AcO ~,

Bz" NH O

O // OH hth/tt

p h ~ / [ l ' - _ O .... OCbz

O OBz

2.3.9

AcO ~

Bz" NH O

O // .OCONHBu

p h / ' ~ ~ _ O.... OH

O OBz

2.4.31

AcO

O

Bz" NH O

OH iv

ph/~~_ O .... OAIIoc

O HO

OAc OBz

2.4.32

AcO

O

O 10/NO2 O.~O

Bz. NH O p h ' / ~ ~ _ O.... OAIIoc AcO

BZ'NH P h i l _ : O O''' ,-

O HO

v, v i

O

~

//

OH

2.4.33

_OCONHR 0

-

OBz

" OAc OBz

2.4.34 2.4.35

R=(CH2)3CO2H R=(CH2)2NMe 2

Conditions: (i) COC12, Py; (ii) n-BuNH2, 72% overall; (iii) H2, PcYC, EtOAc, 90%; (iv) pNO2C6H4-OCOC1, Py, CH3CN, 78%; (v) H2N-(CH2)3-CO2All,THF, rt, 93%; or H2N-(CH2)2NMe2, THF, rt, 91%; (vi) Pd2dba3, CH2C12,PPh3, triethanolamine, rt, 68% for 2.4.34; 65% for 2.4.35 [Alloc= allyloxycarbonyl].

204 5.2.5.Reactions at C-9/C- 10 The ketone function at C-9 in paclitaxel is exceedingly resistant to many reagents that traditionally attack the carbonyl group, and consequently it has escaped t r a n s f o r m a t i o n until recently, when Commerqon reported its reduction by electrochemical means [56]. As s h o w n in S c h e m e 48, w h e n T a x o t e r e | was reduced eletrochemically, 9(a)-dihydrotaxotere 2.5.1 and 9([~)-dihydrotaxotere 2.5.2 were produced with very little stereoselectivity. Treating 7-epi-taxotere 2.5.3 under identical conditions led only to the 9(~)-dihydro derivative 2.5.4. On the other hand, when the electrolytic reduction was carried out at -1.95 to -1.90 V in the presence of CaC12, which is presumed to alter the electron density at the C-9/C-10 hydroxyketone moiety by forming a tight complex, C-10 deoxygenation was achieved in modest yield (vide infra).

HQ RI R2 OH B~ Taxotere 1.1.2

0

phil_

0 .... OH

0

HO

: OAc OBz

2.5.1 R 1 =OH, R2 =H 2.5.2 R1 =H, R2 =OH

H B~ 7-epi-Taxotere 2.5.3

OHoH

H O

ii

....

6H 2.5.4

o HO

OBz

OAc

Conditions" (i) E= -1.85V (SCE Hg cathode), MeOH, NH3, NH4C1; 2.5.1 (40%) and 2.5.2 (24%); (ii) same as i, 63%.

Klein et al. reported the synthesis of 9(a)-dihydrotaxol from the naturally occurring 13-acetyl-9(cx)-dihydrobaccatin III [11, 57]. As depicted in Scheme 49, the synthesis began with the protection of the C-7/C-9 diol moiety of 2.5.5

as an acetonide. The crucial C-13 deacetylation was achieved

205 chemoselectively in acceptable yield by the use of n-butyllithium under carefully optimized conditions. The paclitaxel side chain was then readily attached onto 2.5.7 according to Holton's protocol, affording 2.5.8 in fair yield. Final removal of the dimethylketal protecting group from 2.5.8 then furnished the desired 9(r dihydrotaxol 2.5.10 [11] in modest overall yield. The Abbott group has also reported the acylation of 2.5.7 with a large variety of side chains, bearing modified C-3' substituents, from which useful and extensive SAR information has been obtained (vide infra) [58].

Me O-A< 'Me i

AcO

OH

AcO

OH

-

0 i

AcO . . . . . . . . .

= RO .... O

HO

O

OAc OBz

2.5.5

HO

OAc OBz

2.5.6 R=Ac~ 2.5.7 R=H J i i

Aco

Me

Bz"~ NH 0 Ph

:

~

=iii'iv

,,,Ph

o/~N-Bz

0 .... 0

OH

2.5.8

EEO,%

HO

2.5.9

: OAc OBz AcO ~,

Bz. NH

OH OH

O

p h / ~ ' v ~ O _ .... OH 2.5.10

s HO

0 OBz OAc :

Conditions: (i) Me2C(OMe)2, CSA, 97%; (ii) n-BuLi, THF, -44~ 46%; (iii) n-BuLi, then 2.5.9; (iv) 0.5% HC1 in EtOH, 67%; (v) CSA, MeOH, 56% [EE=Ethoxyethyl].

206 Datta et al. recently reported the first example of enolization of the C-9 keto group in a fully functionalized taxane [59]. As shown in Scheme 50, t r e a t m e n t of 2.5.11 with potassium t-butoxide led to carbonate 2.5.12 in fair yield. This was t h e n converted into the corresponding paclitaxel and Taxotere | derivatives for biological evaluation. Many efforts have been devoted to the complete deletion of functionality at C-10 in order to examine its effect on biological activity. The first synthesis of 10-deoxytaxol was unexpectedly achieved by Chen et al. during their attemps to fluorinate such position. The synthesis began with 10-deacetyltaxol 2.5.15, which was obtained by Lewis acid-promoted methanolysis of paclitaxel [23] (Scheme 51). Treatment with trichloroethyl chloroformate then gave the 2',7diprotected derivative 2.5.16. Subjection of 2.5.16 to Yarovenko's reagent (C1FHCCF2NEt2) in CH2C12 at room temperature surprisingly yielded dienone 2.5.17, together with a small amount of a C-12 fluorinated enone [60].

os

TrocO O ~L /7 OTroc

\

i HO ....

e/SSso

O

HO ....

OBz

/

OTroc

9....

O OBz

2.5.12

2.5.11

/h//h/v,v

Ph~,,CO2H B o c - N ~ ,~. O 2.2.7

RH~____/~L

OH

Ph- ~' v0 .... OH

0

HO

: OAc OBz

2.5.13 R= PhCO 2.5.14 R= t-BuOCO

Conditions: (i)t-BuOK, THF, -30 ~ to 0 ~ (58%); (ii) 2.2.7, DCC, DMAP, PhMe, 70 ~ 62%; (iii) HCO2H, rt (71%); (iv) PhCOC1, NaHC03 or (Boc)20, NaHCO3 (62-72%); (v) Zn, AcOH, MeOH, 60 ~ 65% for 2.5.13, 69% for 2.5.14.

207 This reaction is remarkable since Yarovenko's reagent is a fluorinating reagent, and dehydration products are rarely obtained. Removal of the protecting group from 2.5.17 was readily accomplished with zinc in a mixture of acetic acid and methanol, to afford 2.5.18 in high yield. Finally, 2.5.18 was found to undergo smooth catalytic hydrogenation to afford 10-deoxytaxol 2.5.19. Owing to its exciting biological activity, 10-deoxytaxol has been the subject of several investigations. For example, Chen at al. reported that defunctionalization at C-10 can be obtained by radical methods using 10thionocarbonates [16a]. As shown in Scheme 52, 10-deacetyl baccatin is an appropriate s t a r t i n g material. When C-7 is protected, thionocarbonate formation proceeds at C-10 vs. C-13 with excellent selectivity. Barton deoxygenation affords 2.5.22, and acylation according to Holton then affords 10deoxytaxol in good overall yield [19]. An analogous C-10 deoxygenation, utilizing a xanthate, was described by Kingston [61].

HO gz..

NH

O

OR

0

phil_

ii

O ....

OR HO 2.5.15 R=H 2.5.16 R=Troc) i

l

O " OBz

OAc

0

BZ'NH

0

phil__

0 ....

OR

0 BZ'NH 0 phi_: 0 .... OH 2.5.19

OBz 2.5.17 R=Troc,~ 2.5.18 R=H 2 i ii

OH

0 HO

iv

-OAc OBz

Conditions: (i) TrocC1, Py, CH2C12, 0 ~ 46%; (ii) Et2NCF2CHFC1, CH2C12, 47%; (iii) Zn, MeOH, AcOH, 40 ~ 81%; (iv) H2, PcYC,EtOAc, 68%.

208 Efforts have been made at deleting the C-10 acetoxy group directly from paclitaxel. Since the C-10 acetoxy group is a doubly activated moiety (i.e. allylic and cz-keto) its removal may be achieved in principle by a direct Barton deoxygenation reaction. Indeed, Chen et al. reported t h a t t r e a t m e n t of 7-epitaxol 2.4.1 with 6-8 equivalents of tributyltin hydride and AIBN in toluene at 100~ afforded directly the corresponding 10-deoxy derivative 2.5.23 in excellent yield (Scheme 53) [62]. When paclitaxel was treated under the same conditions, however, only 2.5.23 was obtained in 39% yield, together with some 7-epi-taxol and unreacted starting material, thus suggesting that the function at C-7 plays a role in this radical deoxygenation [62].

H

,~O OTES

R

O

OTES

i

HO ....

=

HO ....

O

O HO

OAc OBz 2.5.20 R=OC(S)OC6F5 ii 2.5.21 R=H ,,/ 0 OR

OBz 2.1.19

iii

BZ-NH ph N - : ~ _ .

0 ....

TESO~I'Ph OR O Bz 2.1.10

0

HO 2.5.22 R=TES 2.5.19

R=H

OBz J]

OAc

iv

Conditions: (i) n-BuLi, THF, -40 ~ then C6F5C(S)C1, -20 ~ 74%; (ii) Bu3SnH, AIBN, PhMe, 90 ~ 99%; (iii) n-BuLi, THF, -40 ~ then 2.1.10, 0 ~ (iv) dil. HC1, CH3CN, 0 ~ 76% overall from 2.5.21 to 2.5.19.

The first one-step synthesis of 10-deoxytaxol directly from paclitaxel was reported by Holton using SmI2 as the reducing agent [63]. Similar chemistry was also reported by other authors [64, 65]. Electrochemical conditions were

209 also applied successfully to the reduction of C-10 acetoxy moiety, as mentioned above [56]. Scheme 53 / gz.

7-epi-taxol 2.4.1

z~ NH

O_H

0

ph/~~_

0 ,, O

OH HO

: OBz

OAc

2.5.23 Conditions: (i) Bu3SnH, PhMe, AIBN, 100 ~

88%.

When the SmI2 deoxygenation was conducted with an excess of the reagent and for prolonged periods of time, 10-deoxygenation and C-9 reduction were reportedly achieved simultaneously. Taxotere | gave instead a mixture of two products, since in this case the C-10 hydroxy is not as good a leaving group as the acetoxy and direct carbonyl reduction can compete with (~-reduction. Once the C-9 carbonyl is reduced, C-10 deoxygenation cannot obviously occur, and 2.5.25 is produced, along with the expected 2.5.26 (Scheme 54) [64]. Using 9-dihydrobaccatin III as a starting material, Klein and coworkers reported, in preliminary form, the selectively deoxygenation at C-9 using t r a d i t i o n a l Barton chemistry, as well as C-7 and C-9 double deoxygenation. Their chemistry is highlighted by the elegant synthesis of 7,9,10-trideoxytaxol, 2.5.31, the analog with the most defunctionalized northern half prepared to date (Scheme 55) [17]. Details of this chemistry have not yet appeared. Selective C-10 deacetylation in paclitaxel is not a facile operation. Kingston et al. reported that t r e a t m e n t of paclitaxel with zinc bromide in methanol yielded 10-deacetyltaxol in low yield together with its C-7 epimer [21]. Chen et al. reported a C-10 deacetylation study in which several other Lewis acids were examined, without substantial improvements [23]. However, with the ready availability of 10-deacetyl baccatin III as a convenient source of bioactive taxanes, this synthetic operation is no longer synthetically important. Although acylation at C-10 is well precedented [4, 52] the first general approach to C-10 modified taxanes was reported only recently by Kant [66].

210 Ethers, esters, carbonates, carbamates, and sulfonates were all prepared in good yields under mild conditions (Scheme 56).

BZ'NH 0

\

OHoH

mh Oo....

Paclitaxel 1.1.1

!o HO

OBz

2.5.24

Boc..

.o,, 2"o.

NH 0

phil"_

O,

OH

o

2.5.25

i

Taxotere, 1.1.2

.

B~

0

phil_

+

OHoH

0 .... oH

HO 2.5.26

Conditions:

-

OBz

(i) SmI2, 83% for 2.5.24; 40% for 2.5.25; 50% for 2.5.26.

As outlined in Scheme 56, 2.1.19 was treated with 1.05 equiv of n-BuLi at -40 ~ in THF, followed by the addition of 1.2 equiv of the electrophile. Derivatives 2.5.32 were then directly acylated with the paclitaxel side chain (~-lactam method) for biological evaluation. 5.2.6. Reactions at C-13 Modification of the C-13 position is a critical operation that may profoundly affect the biological activity, due to the important role of the phenylisoserine side chain. The C-13 hydroxyl group of baccatin III is often silylated in order to protect it from functionalization during complex synthetic operations. Use of the TMS and

211 TES blocking groups has been exemplified widely t h r o u g h o u t the chapter. Acetylation in pyridine was described as requiring h a r s h conditions by several workers [51, 67], but more recently such acetylation could be carried out at room temperature in CH2C12 with acetic anhydride [23]. Carbonates have also been used at C-13 as protecting groups, especially the convenient Troc group [67]. Oxidation of 7-TES Baccatin III at C-13 using MnO2 affords the corresponding enone, which can be reduced back to the baccatin derivative with borane [68]. The various methods that have been discovered to introduce the phenylisoserine side chain at C-13 are described in Chapter 6. [ Scheme 55 SMe S AcO ~, AcO ....

:

OH

AcO ---~0 ~, --"

OH i _

sSess

ii

AcO ....

2.5.5

2.5.27 HO ~,

H O ....

OH

iii

9.....

HO .... ~

iv

HO

2.5.28

gz..

v

OH

2.5.29 NH

0

\

.... < Cbz-N~

0

2.5.30

HO BzO 2.5.31

Conditions: (i) LiHMDS, CS2, then MeI; (ii) Bu3SnH; (iii) MeLi, then CS2, MeI; (iv) Bu3SnH;

(v) 2.5.31, then H2, Pd/C, then (PhCO)20, aq. MeOH (no solvents, temp. nor yields).

212

Scheme 56 / HO

HO ....

0

//

RO

.OTES

r

HO....

0

/I

OTES

r

0

0

OBz

OBz

2.1.19

2.5.32 Electrophile

R

Yield(%)

AcC1 BzC1 n-BuCOC1

COCH3 COC6H5 COBu-n

c-C3H5COC1 MeOCOC1

COC3H5-c 78 CO2Me 75

M e2SO4 Me2NCOC1 PhNCO MeSO2C1

Me CONMe2 CONHPh SO2Me

90 85 75

85 72 78 68

Conditions: (i) n-BuLi, THF,-40 ~ then electrophile,-40 ~ to 0 ~

5.2.7. Reactions at C-14 Recently, Appendino et al. reported the isolation of 14([~)-hydroxy-10deacetylbaccatin 2.7.1 from the needles of T a x u s w a l l i c h i a n a Zucc [69]. Due to the presence of an additional hydroxyl group at the C-14 position, the new taxanes derived from 2.7.1 upon C-13 acylation can be expected to possess substantially improved water solubility vs. paclitaxel and docetaxel, and perhaps also better in vivo antitumor activity. With this in mind, two groups set out to prepare 14(~)hydroxytaxol and 14(~)-hydroxytaxotere, as well as a number of related analogues [12, 13, 70]. The relative reactivity of the four hydroxyl groups in 2.7.1 was independently studied by Kant and Ojima [12, 13]. It was found that the reactivity of these groups toward acylation decreases in the order C-7 > C-10 > C-14 > C-13. Therefore, the a t t a c h m e n t of the phenylisoserine side chain to the C-13 position requires appropriate protection at C-7, C-10 and C-14. Toward this end, 2.7.1 was converted

213 into the 7,10-diprotected derivative 2.7.2. C-1 and C-14 were t h e n protected as carbonate 2.7.3, orthoformate 2.7.4, or acetonide 2.7.5, as shown in Scheme 57.

,,," H

HO ~

0 /f

O_H

H ,,

TrocO ~,

0 //

OTroc

i

O,

0

HO

OH

: OBz

_

O~

OAc

OBz

TrocO

2.7.1

O

2.7.2

OTroc

HO ....

i i , or iii, or i v

0

O RO

OR'

: OAc OBz

2.7.3 R,R'=C(O) 2.7.4 R,R'=CH(OEt) 2.7.5

Conditions: (i) 4 equiv TrocC1, py, 80 ~ 55%; (ii) 2 equiv TrocC1, py, 80 ~ TsO)20, 92%; (iv) 2,2-Dimethoxypropane, (p-TsO)20, 89%.

HO

0

R,R'=C(Me) 2

75%; (iii) CH(OEt)3, (p-

OTES

i

ii

2.7.1

~

HO .... 0 HO

HO

O

OTES

HO ....

OH

O

y

EtO

O

OAc OBz 2.7.7

OAc

2.7.6 iii

O

" OBz

RO

20TES

= HO ....

or i v

0

Oyo EtO

OBz

OAc

2.7.8 R=Ac 2.7.9 R=TES

Conditions: (i) TESC1, imidazole, DMF, 92%; (ii) CH(OEt)3, PPTS, THF, 90%; (iii) LiHMDS, AcC1, THF, 0 ~ 75%; (iv) LiHMDS, TESC1, THF, 0 ~ 78%.

214 Alternatively, the C-7 hydroxyl group was protected as a triethylsilyl ether, affording 2 . 7 . 6 in high yield. Compound 2.7.6 was next converted to 1,14orthoformate 2.7.7. C-10 acetylation yielded 2.7.8, and silylation afforded 2.7.9 (Scheme 58). Using protected baccatin derivatives 2.7.4 and 2.7.9, 14(~)-hydroxy-taxotere 2.7.11 was readily obtained in good overall yield using the protocols illustrated in Scheme 59. In addition, cyclic carbonate analog 2.7.12 and acetonide 2.7.13 were also prepared in a similar fashion. Similarly, the coupling between 10-acetoxybaccatin derivative 2.7.8 with the appropriate ~-lactams 2.1.10 and 2.4.6 afforded the desired 14(~)-hydroxytaxol 2.7.14, and 10-acetyl-14(~)-hydroxytaxotere 2.7.15 for biological evaluation (Scheme 6O).

2.7.9

Kant i, i ~ iii

RO,,, 9

iII

Ph

2.7.4

N "Boc 2.4.6 R=TES 2.7.10 R=CH(Me)OEt

I

OJ--'

~,~ 0 II t-BuO ''~k" NH

HO O

I /

O

OH

\

OH 2.7.11

Ojima iv, v, vi

0 HO

O" B z

OH

OAc

O t-BuO"JJ~NH p h i - i l l _"

O

HO

O

OH

0 .... 0

RO

OR'

OBz

2.7.12 2.7.13

R,R'=CO R,R'=CMe 2

OAc

Conditions: (i) LiHMDS, THF, 0 ~ then 2.4.6, 75%; (ii) 10 N HC1, CH3CN, -5 ~ (iii) NH4OH, THF, 0 ~ 67% overall. (iv) NaHMDS, THF, -40 ~ then 2.7.10, then 0.5% HC1, EtOH, rt, 96%; (v) HCOOH, dioxane, rt, then THF, MeOH, NaHCO3, rt, 73%; (vi) THF, 0.5 N HC1, Zn, 0 ~ 73%.

215 I n t e r e s t i n g l y , as shown in Scheme 61, direct acylation of 2.7.2 u n d e r s t a n d a r d conditions yielded two novel C-14 side chain-bearing t a x a n e derivatives (2.7.16 and 2.7.17) in fair yield. This observation is in a g r e e m e n t with the finding t h a t the hydroxyl group at C-14 can be acylated more readily t h a n the one at C-13 [13]. Scheme 60 /

O

AcO

O

.

(i, i i ) or [i i i, i i) 2.7.8 TE SO,,,,

,,, Ph

oJ-' N. R

ph~/[l"O. OH

.... 0 HO

2.1.10 R=Bz 2.4.6 P~Boe

OH

2.7.14 R=Ph 2.7.15 R=t-OBu

: OBz

OAc

Conditions: (i) LiHMDS, THF, -40 ~ then 2.1.10, 65%; (ii) 10 N HC1, CH3CN, -5 ~ then NH4OH, THF, 0 ~ 55% for 2.7.14, 62% for 2.7.15; (iii) LiHMDS, THF, -40 ~ then 2.4.6, 75%.

,,"

HO

~, //O O.H

i, or ii, then iii 2.7.2

OH

I

~

N. R

2.7.18 R=Bz 2.7.10 R=Boc

RC(O)HN

O

O=z

2.7.16 R=Ph 2.7.17 R=t-OBu

Conditions: (i) NaHMDS, THF, -40 ~ then 2.7.18; (ii) NaHMDS, THF, -40 ~ then 2.1.10; (iii) Zn, AcOH, MeOH, 52% overall for 2.7.16; 50% overall for 2.7.17.

5.2.8. Skeletal r e a r r a n g e m e n t s This section will discuss baccatin,

some interesting skeletal r e a r r a n g e m e n t s

of

m o s t l y concerning the A and B rings. These r e a r r a n g e m e n t s are

usually initiated by radicals or carbonium ions. In addition, a few recent reactions t h a t are more properly classified as degradation reactions will also be discussed. A n u m b e r of radical-based deoxygenation reactions were carried out on baccatin III derivatives [16a].

In this connection, it was discovered t h a t formation of a

216 radical at the C-7 position of the baccatin core results in a complex skeletal rearrangement (Scheme 62). Tributyltin hydride-mediated deoxygenation of 2.8.1 gave 52% of the desired 7,10-dideoxy baccatin III 2.8.2 and 25% of its tetracyclic isomer 2.8.3 [16a]. Other radical conditions were examined, and the distribution of products characterized. When Ph3SnH was used as the reducing agent, in addition to 2.8.2 and its isomer 2.8.3 (ratio, c a . 3:1), methyl ether 2.8.4 was also isolated in 30% yield; when (TMS)3SiH was employed as the reducing agent, in addition to 2.8.3, two new products, enol acetate 2.8.5 and C-12 exomethylene derivative 2.8.6, were also obtained (Scheme 63). The formation of 2.8.3, 2.8.5 and 2.8.6 can be rationalized by invoking a cascade of radical rearrangements, as shown in Scheme 64.

O

Scheme 62 t O IIII

O

\

0

)CS2Me HO

HO ....(

+

0

HO

: OAc OBz O

: OAc OBz 2.8.1

0 Me

HO ....~

Conditions:

2.8.2

O HO

A c = OBz

2.8.3

(i) Bu3SnH, AIBN, PhMe, 80 ~ 2.8.2 (52%); 2.8.3 (25%).

The initially formed radical 2.8.7, a ~-keto radical, can isomerize, via alkoxy radical 2.8.8, to 2.8.9. This places the radical-bearing C-8 at a close distance with respect to the C-11/C-12 double bond, and a 5-exo cyclization to 2.8.10 takes place. Surprisingly, this radical is not quenched by the tin hydride, perhaps due to the hindered nature of the radical-bearing C-12. The major pathway for radical 2.8.10 is the remote intramolecular hydrogen abstraction of H-3, to provide 2.8.11. Radical 2.8.10 also suffers a disproportionation reaction to give the minor product 2.8.6 only when (TMS)3SiH was employed as the reducing agent. Radical 2.8.11 is evidently also sterically hindered toward direct reduction, and undergoes an unusual oxetane fragmentation reaction to give 2.8.12. The resulting a-alkoxy

217 radical is then trapped by tributyltin hydride to yield 2.8.3. When the reducing agent used is tributyltin deuteride, the product is specifically labeled only at the C-5 methoxy group [16a]. However, in the case of the (TMS)3SiH reduction, in addition to 2.8.3, alkoxymethyl radical fragmentation with loss of formaldehyde gives allylic radical 2.8.13, presumably due to slow trapping of 2.8.12 by the rather unreactive silane reagent. After this cascade of six sequential intramolecular reactions, radical 2.8.13 is finally quenched by the silane to give 2.8.5 (Scheme 64).

O i or i 2.8.1

"

i= H O ....

I +

O IIII

jO %

or i i i HO

OAc OBz

HO

2.8.2

OBz OAc 2.8.4

0

O

H,,,~~~L~ +

HO

OMe ....~

HO

HO":

+

c

I

: OBz 2.8.3

HO O

H

HO ....

~

= OBz 2.8.5

OAc

O HO

= OBz

2.8.6

Conditions: (i) Bu3SnH, AIBN, PhMe, 80 ~ (ii) Ph3SnH, AIBN, PhMe, 80 ~ (iii) (TMS)3SiH, AIBN, PhMe, 90 ~ The proposed mechanism for the C-7 methyl ether formation is shown in Scheme 65. Addition of the triphenyltin radical onto xanthate 2.8.1 leads to the unstable intermediate 2.8.14. Usually, this intermediate undergoes ~-scission to afford a carbon radical. However, in this case the highly reactive hydrogen donor triphenyltin hydride was able to trap 2.8.14 to form 2.8.15.

218

0

2.8.1

"H

~HO'"

= HO'"

0 Ho BzO

OAc

BzO

2.8.7 0 HO"

H OAc

2.8.8

H

0

~

ai

H Ola0a

-

H"

~

HO BzO

OAc

HO BzO

2.8.9

~

OBz

H OAc

2.8.10

0

H ,,~ , ? - - %.,~, ~ H17-HO ....(, ~ ~

2.8.6

0

)~

.w,,, ._ H','" - HO"'

H

OAc

OBz

2.8.11

.

CH20 ] T

HO .... ~

2.8.13

c 2.8.12

0

/N;,,"~

HO

+H" OCH-------~2.8.3

: OBz

/) "

+H"

~ 2.8.5

OAc

Elimination of triphenyltin thiomethoxide gave C-7 thionoformate 2.8.16. Further reduction by excess triphenyltin hydride gave thioacetal 2.8.18, which was finally converted to methyl ether 2.8.4 through another C-S bond cleavage reaction. The same study [16a] describes a series of complex radical rearrangements that arise via C-1 and C-2 carbon radicals. As shown in Scheme 66, treatment of cyclic thiocarbonate 2.8.19 with tributyltin hydride and AIBN failed to give either a C-1 or a C-2 deoxy

derivative. Instead, after t r e a t m e n t of the crude product

mixture with trifluoroacetic acid, two new products, 2.8.20 (major) and 2.8.21 (minor), were obtained. Inspection of the 1H-NMR spectra dearly showed that both

219 compounds were the results of skeletal rearrangements. Their structures were confirmed by X-ray crystallography.

SSnPh3 O O+ SMe + Ph3Sn"

+ H"

~ HO,,,

2.8.1

0

BzO

SSnPh3

S

2.8.14

O O~J~H

O O+SMe HO,,,'~

. Ph3SnSMe ~HO....

H O

HO BzO

O H

OAc O O -''~

2.8.15

+ Ph3Sn"

2.8.16

SSnPh3

~ ~ HO,,,

OBz

+ H" O HO BzO

OAc 2.8.17

~

. / ~ ~ ~ ~~SSnPh3

, ~

~"~

H O ....

+ Ph3SnH=__

0 \{/. H O .... .

'---f.-.4/__"H HO

BzO

2.8.18

OAc

O HO

" BzO

OAc

2.8.4

A mechanistic rationale for the observed products is presented in Scheme 67. The initial adduct resulting from addition of the tributyltin radical to 2.8.19 apparently leads to both of the two conceivable fragmentation products, radicals 2.8.22 and 2.8.23. It is likely that intermediate radical 2.8.22 is hindered by the presence of its neighboring tributyltin thiocarbonate residue at C-l, and consequently it is not rapidly trapped by tin hydride to yield the corresponding 2-deoxybaccatin derivative

220 as seen with C-2 xanthates. Instead, intramolecular processes, as already seen for the C-7 radical, take over and 2 . 8 . 2 2 undergoes a t h e r m o d y n a m i c a l l y unfavorable and quite unusual 4-exo cyclization to the cyclobutylcarbinyl radical 2.8.24. Since the cyclobutylcarbinyl radical is highly unstable, opening with concomitant ~-elimination leads to 2.8.26 and, after desilylation, 2.8.20. 00TES

[ Scheme 66 TESO,,,,~

0 -

"~0

S

0

HO ....

I

Ac 2.8.19

~ii 0

OH +

OH

HO .... 0

/ 2.8.20 Conditions:

6H oAo 2.8.2

(i) Bu3SnH, AIBN, PhMe, 100 ~ 77%; (ii) CF3CO2H, THF, H20; 2.8.20, 42%; 2.8.21,

8% overall. Radical formation at C-1 is also apparently taking place in a competitive fashion: radical 2.8.23 undergoes a cyclopropylcarbinyl r e a r r a n g e m e n t to yield, following hydrolytic deprotection, A ring-contracted product 2.8.21. In addition to radical-initiated ones, cationic r e a r r a n g e m e n t s are very common in taxane chemistry. We have already discussed the A-ring contraction reaction apparently initiated by C-1 carbonium ion formation (Scheme 4) [21]. Rearrangements initiated by C-7 carbonium ion formation lead to the already discussed cyclopropane derivatives (Scheme 42) [47, 48] and, in the case of 9dihydrotaxanes, to B-ring contracted analogs [48], as shown in Scheme 45. These B-ring contracted taxanes could be deprotected at C-13 and acylated under standard conditions for biological evaluation [48].

221 Whereas Lewis or Bronsted acid treatment of taxanes leads to A-ring contraction, as already discussed, recently Khuong-Huu et al. reported an alternative ring enlargement reaction that proceeds under acidic conditions. ES

TESO ....(X

/~'.._ .~. ~J~ 0,~6

Bu3Sn'/

/

S

OAc Bu3Sn"

2.8.19

i~ OTES TESO ....

T E S O , , , ~

o_ 9 OAc ~--SSnBu 3 O [ 2.8.22

Bu3SnS~/F6

~

~T-SSnBu3 O 2.8.24

O 1

2.8.23

TESO,,

OAc

TESO ....(k

I~'.. 4 . . L

.... -H/_ ~ ._-~o

Bu3SnS.~ ~ 2.8.25

OAc

O I +H

+"1 O OR RO ....

RO"' O

2.8.26 R=TES ) 2.8.20 R=H

2.8.27 R=TES) 2.8.21 R=H

222

Scheme 68 I AcO

AcO

OAc OAc

~,

OAc

.--

OAc

ii

HO ....

AcO ....

O

O OH

2.8.28 -

HO~'

OAc

OBz

AcO

HO

OH

2.8.29 OAc

S

OAc

OAc

/OAc

-

......

OH

,

OAc

....._

H 0 -

2.8.30

"'-0--/OH 2.8.31

-

HO ....

0

&o Ph

2.8.32

Conditions: (i) KCN, MeOH, rt, 65%; (ii) Camphorsulfonic acid, DMF, C6H6,reflux, 90%.

As shown in Scheme 68 [71], t r e a t m e n t of peracetylated baccatin derivative 2.8.28 with KCN in methanol led to deacylation at C-13, C-2 and C-4, affording 2.8.29. Tetrol 2.8.29 was w a r m e d overnight in DMF-benzene solution in the presence of a catalytic amount of camphorsulfonic acid, leading to r e a r r a n g e d product 2.8.31, which contains a 10-membered ring and a tetrahydrofuran ring, presumably via intermediate 2.8.30. A similar r e a r r a n g e m e n t of the oxetane to the tetrahydrofuran system was described in Schemes 5 and 6 (Section 6.2.1). It is unclear why the usual A-ring contraction does not occur here, and the authors do not offer any explanation.

223 However, it seems quite reasonable to explain this result by postulating that the Aring contraction requires the presence of the C-2 benzoate group and proceeds not via a C-1 carbonium ion, but via acyloxonium ions like 2.8.32. In the absence of an ester group at C-2, carbonium ion formation at C-1 is not a facile process and the pathway described above predominates. Another very complicated r e a r r a n g e m e n t , involving migration of C-2 benzoate and opening of the oxetane ring, is observed in the 14(~)-hydroxy baccatin series, and is outlined in Scheme 69 [72].

HO

20TES

HO....

0

" HO HphOh ~:~,~

HO~'

0 =~

OAc : H+ 2.7.6

~/

20TES

-- HO....

HO....

",,S

0

o.~o+ ~ O,~o~ Ph 2.8.34

~zo

o. o[2~ 2.8.35

I

0%. /? OTES Ho

.o

%./ _/? OTES ....

....

Bz

o, ~ H0~"~?o....--+h o. 0 2.8.86

BzO/

~

~

d"") ~

2.8.37

When 2.7.6 was refluxed in benzene in the presence of a catalytic amount of pyridinium p - t o l u e n e s u l f o n a t e (PPTS), rapid d i s a p p e a r a n c e of the s t a r t i n g

224 material took place, and 2.8.37 was isolated in 40% yield. A plausible mechanism for this r e a r r a n g e m e n t is also depicted in Scheme 69. Migration of the benzoate to C-14, presumably via C-1/C-2 and C-1/C-14 oxonium ions 2.8.33 and 2.8.34, triggers the Wagner-Meerwein A-ring contraction. This is accompanied by the already discussed C-4 assisted opening of the oxetane ring, eventually yielding the stable orthoformate 2.8.37 by trapping of 2.8.36 by the unacylated C-2 hydroxyl. A different type of A ring contraction, this time accompanied by B ring expansion, was recently reported by Appendino and co-workers [73]. As shown in Scheme 70, when 7-triethylsilyl-10-deacetylbaccatin 2.1.19 was treated with excess MnO2, cyclopentenone derivative 2.8.40 was obtained in low yield, in addition to the desired 13-keto product 2.8.38. The authors postulate the intermediacy of adiketone 2.8.39, which undergoes an a-ketol rearrangement. The driving force for this ring contraction may be the release of the angular strain due to the presence of four adjacent sp 2 centers in 2.8.39.

HO

O

OTES

H O .... O HO

OBz

O

: OAc

O HO

2.1.19

: OBz

": OAc

2.8.38

HO

0

OTES

O O O HO 0

OBz

OAc

2.8.40 Conditions: (i) MnO2, EtOAc, CH2C12,rt; 2.8.38 (40%)+ 2.8.40 (24%).

OBz

2.8.39

-: OAc

225 In order to assess the contribution of an i n t a c t A ring of the baccatin framework to the a n t i t u m o r activity, Ojima and co-workers prepared a novel class of nor-seco paclitaxel analogues 2.8.44 and 2.8.45, as shown in Scheme 71. The synthesis began with the oxidative cleavage of the A ring of 2.7.1, a n a t u r a l product isolated from T a x u s w a l l i c h i a n a Zucc [69], with periodic acid. This gives 2.8.41, which i m m e d i a t e l y cyclizes in situ to provide the h e m i k e t a l 2.8.42. The C-7 hydroxyl group of 2.8.42 was selectively protected as the triethylsilyl ether, and the aldehyde moiety was reduced with sodium cyanoborohydride to afford 2.8.43 in fair yield. Final side chain a t t a c h m e n t onto 2.8.43 was performed using the ~-lactam approach to provide the desired nor-seco paclitaxel and docetaxel analogs 2.8.44 and 2.8.45 [74] (Scheme 71).

HO

H 0'"'

O

HO

-

OHC

i

~

,'

~

o

HO

OH

:

"

0

OBz OAc

2.8.41

2.7.1

,4~

OHC

BzO C)Ac

OH

OTES

rO

O

i~, iii

HO

,,,..-

,

O O

O H()

HO

BzO (gAc

2.8A3

2.8.4,2

R'NH

iv, v

~ /0,,,, OEt 0

,,,Ph

)_, N.

_O. ~

- pha-2-_ OH

COR

2.7.18 R=Ph 2.7.10 R=OBu-t

BzO C)Ac frO

.....J

0

O.H

~

~

-.:'H HO

~

BzO C)Ac

2.8.44 R=Bz 2.8.45 R=t-Boc

Conditions: (i) H5IO6, 92%; (ii) TESC1, NEt3, DMAP, 76%; (iii) NaBH3CN, pH 6. 80%; (iv) NaHMDS, THF, -40 ~ 2.7.18, or 2.7.10; (v) 0.5% HCI, rt; then Bu4NF, THF, -10 ~ 82% for 2.8.44; 58% for 2.8.45.

226 Recently, Khuong-Huu and co-workers reported on the chemistry of oxidative cleavage of the B ring of baccatin at the C-1/C-2 and C-9/C-10 segments [75]. Since the reverse operation is often the key step in total syntheses aimed at the paclitaxel skeleton [8-10], the value of this work is in providing valuable materials for the study of these reductive cyclizations. The debenzoylation of C-2 was described in section 5.2.1. A typical product of these studies, 2.1.20, was protected at C-13 as a triethylsilyl ether, giving 2.8.46, which was then subjected to sodium metaperiodate oxidation, affording 2.8.47 in good yield (Scheme 72).

ES

Ro ....k

7."-,,_ ,L Hal"

_= - O H

2.8.46

R=TES J

HO 2.1.20 R=H

Conditions:

00TES

b

T SO,,,

0

-

OH i

2.8.47

(i) TESC1, py, rt, 100%; (ii) NaIO4, EtOH, pH 5 buffer, 75%.

The C-9/C-10 bond cleavage reaction is shown in Scheme 73. 10-Deacetylbaccatin III, 1.1.2, was selectively protected to yield 2.8.48 in two steps. The removal of the carbonate groups and the subsequent C-7 epimerization were achieved on treatment with zinc dust and DBU, respectively, providing 2.8.49 in excellent yield. Reduction of the C-9 keto moiety of 2.8.49 was effected remarkably well with BH3SMe2 in toluene, producing 9(~)-dehydrobaccatin derivative 2.8.50. The stereochemistry at C-9 was confirmed by NOE studies. Oxidative cleavage of the C-9/C-10 bond of 2.8.50 led to lactol-aldehyde 2.8.51, as a single isomer, in almost quantitative yield [75]. 5.2.9. Photochemistry No information on the photochemistry of paclitaxel was available until recently. During the course of the development of paclitaxel as a commercial antitumor drug, paclitaxel was subjected to a series of stability tests, including exposure to sunlight. In this test, traces of a paclitaxel isomer were isolated by s e m i p r e p a r a t i v e HPLC. After extensive NMR studies, the compound was

227 identified as the pentacyclic paclitaxel isomer 2.9.4 (Scheme 74), containing a new bond between C-3 and C-11 [76].

H

TrocO

OH

O OTroc

\ HO .... ( ~

Iii, i v

Lii

,~,,.

AcO ....

= O

0 HO

BzO

HO

OAc

" BzO

OAc

2.8.48

1.1.2

HO

AcO'"

AcO"'

OHoH

)

v, O

O HO

OAc

2.8.,50

2.8.49

o%IHoH OH

Aco,,,o .

" BzO

: BzO

2.8.51

OAc

Conditions: (i) TrocC1, py, 80 ~ 93%; (ii) Ac20, py, DMAP, rt, 100%; (iii) Zn, MeOH, reflux; (iv) DBU, PhMe, 80 ~ 80%; (v) BH3-SMe2, PhMe, 0 ~ 84%; (vi) NaIO4, EtOH, pH 5 buffer, rt, 97%.

Since very limited amounts of 2.9.4 could be produced by sunlight exposure, a more efficient method for further studies was highly desirable. An approach to this problem utilized a photochemical reactor. A very good conversion (55%) of paclitaxel into 2 . 9 . 4 was achieved. M e c h a n i s t i c a l l y , this r e m a r k a b l e photochemical transformation can be considered to follow in part the well-known oxa-di-~-methane r e a r r a n g e m e n t (Scheme 74). Nakanishi [77] was the first to describe a similar bond formation between C3 and C-11 in taxinine. However, taxinine differs from paclitaxel in that an enone chromophore is present at C-11/C-12/C-13, and it is undoubtedly the excitation of

228 this f u n c t i o n t h a t i n i t i a t e s the r e a r r a n g e m e n t . In the p r e s e n t case, the photoexcited moiety m u s t be the ~,y-unsaturated ketone, and the m e c h a n i s m shown in Scheme 74 was proposed.

AcO BzHN

O"

0

OH

hv

Paclitaxel 1.1.1

=

p h ~ . - J l_' - 0 .... OH 2.9.1

AcO~ BzHN

O"

" OBz

OAc

OH

0

Ph~O OH

H3-shift

.... 0 OBz

2.9.2

H

BzHN/~.flL_ H Ph

/

0

HO

,~

A

/.,,.., O H

O\1)..

~

Radical

_ O.... OH

Recombination

O : OAc OBz

HO

2.9.3

AcO zHm

o

-

0 OH

k H'

--

2.9.4

O IIII

OH

O HO

: OAc OBz

The excited state initiating the r e a r r a n g e m e n t m u s t be the TI(~,~*) of the C9 carbonyl group, which is r e p r e s e n t e d as the diradicaloid species 2.9.1, as postulated in the first step of the oxa-di-~-methane r e a r r a n g e m e n t . Diradicaloid 2.9.1 then r e a r r a n g e s to cyclopropylcarbinyl radical 2.9.2, and at this point the intramolecular hydrogen t r a n s f e r from C-3 to C-12 occurs, in the same vein as

229 described in section 5.2.8 w h e n discussing radical r e a r r a n g e m e n t s .

Finally,

t r a n s a n n u l a r bond formation in 2.9.3 leads to 2.9.4. Although the above m e c h a n i s m is reasonable on the basis of the literature, a more comprehensive study addresses the issue of w h e t h e r the C-9 keto group is directly excited, or w h e t h e r some of the a r o m a t i c groups in the molecule are involved in the a b s o r p t i o n and s u b s e q u e n t i n t r a m o l e c u l a r

energy transfer.

Confirmation of this possibility was sought by indirect m e a n s , i.e. each of the aromatic groups in the molecule were in t u r n deleted, and the effect on the yield of the pentacyclic products was examined [78].

R2

O

R2 I

,',H'"' ,

R1

//

O

al

hv ,..._

R 3 .....

R3

, O

O HO

OBz

HO

OAc

" OBz

OAc

2.2.4

R1 = OTES R 2 = OAc, R 3 = OH

2.9.6 (23%)

2.1.19

R1 = OTES R2= H, R3 = OH

2.9.7 (20%)

2.9.5

R1 = R2 = H

2.9.8

(21%)

1~ = OTES AcO

O

OTES

H O .... O HO

"

OAc O

2.9.9

As s h o w n

in

Scheme

75, b a c c a t i n

derivatives

were

studied

first.

Interestingly, the three baccatin III derivatives 2.2.4, 2.1.19 and 2.9.5, bearing a

230 benzoate group at C-2, when subjected to photolysis under standard reaction conditions (254 nm, Pyrex, 0.05 M in CC14, 20 h) cleanly gave the expected rearranged products 2.9.6, 2.9.7 and 2.9.8. In striking contrast to the above observations, attempted photolysis of 2.9.9 failed to produce any of the expected rearranged pentacyclic derivative. The above results suggest t h a t an aromatic ester moiety at C-2 is necessary for the photoisomerization to occur. The contribution

of the aromatic groups of the side chain to the

photochemistry of paclitaxel was next examined. The key substrate is the paclitaxel analog 2.1.26c, in which the C-2 benzoyl moiety has been replaced by the cyclohexanoyl ester.

i, ii 2.9.9

--

TESO,

AcO Ii

H Bz.N 0 = II P h ~ O OH

//

0

O.H

.... 0 -

0

2.1.26c 2.1.10 O AcO ~

H

Bz.N

o

/

HO

2.9.10

/I

0

OH

\IJl.

(5

OAc

~/N~O

Conditions: (i) LiHMDS, THF,-40 ~ 2.1.10, 85%; (ii) Py, 48%HF, CH3CN, 5 ~ 99%; (iii) hv, 254 nm, 40% of 2.9.10, 21%

of2.1.26c.

As shown in Scheme 76, the

photolysis of 2 . 1 . 2 6 c was performed as usual,

affording a 40% clean conversion to 2.9.10. The authors conclude that initial excitation of the 3'-N-benzoyl amide or the 3'-phenyl group in the side chain also serves to excite the C-9 keto function in an intramolecular fashion, since the

231 presence of the side chain restores the normal reaction mode absent in 2.9.9 [78]. Finally, to confirm t h a t photoinduced isomerization specifically requires a side chain endowed with an aromatic amide at C-3', analog

2.9.12 was prepared as

shown in Scheme 77. Under standard photolytic conditions, compound 2.9.12 was found, as expected, to be completely inert. This experiment clearly supports the hypothesis of an "antenna effect" between the C-3' benzamide (and/or the C-2 benzoate) and the C-9 ketone in this photochemical isomerization [78].

0 [~}/IL

AcO NH O

i, ii

/,O OH

2.9.9 --

Ill

O

2.9.11

0

Ok,~=O ~~/

2.9.12

Conditions: (i) LiHMDS, THF,-40 ~ then 2.9.11, 87%; (ii) Py, 48%HF, CH3CN, 5 ~ 94%.

Scheme 78

AcO

OH

o Paclitaxel 1.1.1

i or ii

= 2.9.4

o ....

OH 2.9.13

~

HO BzO: AcO

0

Conditions: (i) hv, PhMe, 300nm, 2.9.4 (31%); 2.9.13 (30%); (ii) hv, CCI4, 300 nm, 2.9.4 (41%); 2.9.13

(21%). The same study also reports confirmation of the t r a n s a n n u l a r bond formation by single crystal X-ray analysis of a typical r e a r r a n g e d product. Photophysical

232 studies on paclitaxel and the analogs described above may shed further light or the initial stages of this interesting photoisomerization. Photolysis of paclitaxel at different wavelengths also produced interestin~ results: at 300 nm, the new compound 2.9.13 was isolated in 30-40% yield, ii addition to 2.9.4 (20-30%) [78].

The formation of compound 2 . 9 . 1 3 can b~

rationalized by invoking the occurrence of a Norrish type I process after the photoisomerization step. The epimerization at C-7 may not be a photochemical event. Interestingly, only one configuration out of four possible at C-8 and (the former) C-10 was obtained. This stereoselectivity may be due to preference for the formation of the less strained pentacyclic ring system. 5.3. STRUCTURE-ACTIVITY RELATIONSHIPS A large array of paclitaxel analogs containing modifications within the diterpenoid core were evaluated in microtubule assembly or disassembly assays, as well as in vitro cytotoxicity assays against a number of tumor cell lines. Some of the more active analogs emerging from above in vitro assays were further tested in in vivo, usually in mice, against m u r i n e or h u m a n tumor xenografts. From the body of data discussed in this section, it is obvious t h a t an imperfect correlation exists among the several in vitro tests, and even more so between in vitro potency and in vivo efficacy. We briefly discuss the most common biological assays and try to highlight their differences.

(i) In Vitro Microtubule Assays Initial Rate of Tubulin Polymerization (Swindell-Horwitz-Ringel Method) [791: This assay determines the initial rate of tubulin polymerization. The assembly of microtubule protein (MTP) in the presence of paclitaxel or analogs is performed as follows: MTP (1.5 mg/mL) is incubated at 35 ~ with 15 pM paclitaxel or analogs (added as DMSO solution; 1% final DMSO concentration) in the absence of GTP. The assembly reactions are followed by turbidity measurements at 350 nm. The value for paclitaxel in this assay is used as an internal standard, and experiments with analogs are usually performed in parallel.

This

assay

is

kinetic

in

nature,

and

does

not

measure

thermodynamic affinity for the binding site (nor is it clear t h a t a correlation

233 between the two exists). In addition, it yields no information on the types of microtubules formed (i.e. length, shape, bundles etc.). It is therefore expected to be a rough measurement of activity, useful perhaps to eliminate the inactive compounds, but oi~en it does not correlate well with cytotoxicity. Extent of Microtubule Assembly (Himes Method) [80]" The aim of this assay is to determine the extent of assembly at different concentrations of the analogs and then calculate an ED50. The assembly reaction is done at 37 ~ in PEM buffer (0.1M PIPES, pH 6.9, 1 mM EGTA, and 1 mM MgSO4) at a protein concentration of 1 mg/mL (10 ~M) in the presence of paclitaxel or analogs and 0.5 mM GTP. The reaction is again monitored by the increase in the apparent absorbance at 350 nm. This method is different from the above in t h a t the kinetics of the process are not considered, and only thermodynamic factors are. These data may correlate better with cytotoxicity, which is usually measured over many hours or even days. Microtubule Disassembly (Potier Method) [52]" This assay was developed by Potier's group on the basis of the unusual stability of microtubules formed in the presence of paclitaxel. In this experiment, a solution of MTP (2 mg/mL) is assembled at 37 ~ in the presence of paclitaxel or analogs, followed thereafter by disassembly via lowering the temperature to 4 ~

The initial rate of microtubule disassembly in the presence of the

compound is monitored by the drop in turbidity at 400nm. The initial rate of microtubule disassembly of paclitaxel is set as a standard. This assay also measures a kinetic parameter, and it is not obvious t hat it should correlate well with cytotoxicity. It is also not evident t hat the correlation with assembly data will be good. EC(O.Ol) Expression (Long Method) [81]" The potencies of the different analogs are expressed as an effective concentration (EC0.01), which is defined as the analog concentration capable of inducing the tubulin polymerization at an initial rate of 0.01 OD/min at 37 ~ as measured at 350 nm. The rates of polymerization are determined at several concentrations and EC0.01 values calculated for each analog by interpolating the appropriate region of the polymerization curves obtained. This method is of

kinetic nature, but it yields data that are concentration based. In view of the poor predictive value of tubulin polymerization data, this review focuses on cytotoxicity results r a t h e r t h a n microtubule assemby or disassembly.

234 (ii) I n Vitro Cytoto~dcity Assays A number of murine or h u m a n cancer cell lines, such as P388, B16, HCT-116 and KB, have been used for the determination of in vitro cytotoxicity of paclitaxel and its analogs [82]. The in vitro IC50 value measures the drug concentration required for the inhibition of 50% cell proliferation, usually after prolonged (2-3 days) incubation. One must note that the success of the analog in this assay will depend on its stability within the culture medium as well as intracellularly; the ability to penetrate the cell, most likely by passive diffusion, is also critical, and highly ionic derivatives may not be bioactive because of poor lipophilicity. In addition, cells that express the m d r (multidrug resistance) phenotype may have more or less reduced sensitivity vs. paclitaxel and analogs. Unfortunately, this issue has not been widely addressed in the literature for paclitaxel analogs, although it is obviously very important, and will not be discussed here (for further details, see chapter 7). It is not clear whether cell lines that are resistant to paclitaxel may also be resistant to all its analogs. The choice of the cell line may be dictated by several factors, such as availability, ease of culture, attempted correlation with in vivo data, clinical relevance etc. One must be cautioned that cytotoxicity of each analog may vary substantially from one cell line to another, and may depend on the exact cell culture protocol employed.

(iii) In Vivo Assays The ultimate test of the efficacy of a drug in a model system is an in vivo assay, usually in mice. Obviously, the success of each analog will depend, in addition to cytotoxicity, on a n u m b e r of other factors, such as proper administration, biodistribution, metabolism, and systemic toxicity (or lack thereof) to the animal. Several protocols have been designed for paclitaxel and docetaxel; usually the more s t r i n g e n t tests involve the use of h u m a n xenografts and i.v. drug a d m i n i s t r a t i o n (i.e. distal t u m o r model), but i n t r a p e r i t o n e a l models are also commonly used. The drugs are usually evaluated in terms of their ability to delay tumor growth or prolong life span, as measured at the maximum tolerated dose (MTD, i.e. dose of drug that is not appreciably toxic to the animal) vs. a control untreated group [83]. Obviously,

235 one does not expect (and usually does not find) complete correlation between in vitro and in vivo results, and therefore one should resist making exaggerated

claims of potent activity based only on cell culture data. 5.3.1. Paclitaxel Analogs Modified at C-1/C-2 As oulined in section 5.2.1, derivatization at C-1 is very hard to achieve. Compound 2.1.9 (Figure 2), where a benzoate has m i g r a t e d to C-l, is essentially inactive in a tubulin polymerization assay [19]. Ring contraction due to solvolysis at C-1 leads to a number of interesting analogs, e.g. 2.1.16, which have tubulin-polymerizing activity, but have very poor activity against tumor cells [21].

BZ'NH 0 Ph~O oH

Bz.

AcO

0 OH

,

"'

~o AcO

NH 0

OCS2Me 0 OH

p h / ~ ~ _ 0 ....

2.1.16

OH

-

"\ BzO

Bz.NH 0 mh4... o.

2.1.9

AcO.2o. ,~

....

2.1.41

~

HO

Figure 2: Simple C-1/C-2 modified paclitaxel analogs

236

R'CO. NH O Ph~O

AcO

O OH

....

6. HO

-

RC(O)O Table 1: Cytotoxicity of Paclitaxel Analogs Modified at C-2 Cpd.

R

R'

IC50/IC50

Cell Line

Ref.

(paclitaxel) a 2.1.29

p-NO2-C6H4-NH

Ph

>200

HCT-116

27

2.1.26a

p-MeO-C6H4-

Ph

>20

HCT-116

27

2.1.26b

p-NO2-C6H4-

Ph

>100

HCT-116

27

2.1.26c

c-Hex

Ph

11

HCT-116

27

2.1.26c

c-Hex

Ph

56

P-388

31

2.1.26d

Me

Ph

>20

HCT-116

27

3.1.1

o-C1-C6H4-

Ph

0.01

P-388

29

3.1.2

m-C1-C6H4-

Ph

0.0014

P-388

29

3.1.3

p-C1-C6H4-

Ph

150

P-388

29

3.1.4

m-CN-C6H4-

Ph

0.33

P-388

29

3.1.5

m-N3-C6H4-

Ph

0.002

P-388

29

3.1.6

m-NH2-C6H4-

Ph

1,500

P-388

29

3.1.7

m-CF3-C6H4-

Ph

15

P-388

29

3.1.8

m-F-C6H4-

Ph

0.35

P-388

29

3.1.9

2-Furyl

Ph

25

UCLA-P3

84

3.1.10

2-Thienyl

Ph

4.2

UCLA-P3

84

3.1.11

2-Naphthyl

Ph

>1,000

UCLA-P3

84

3.1.12

c-Hex

t-BuO

1.1

B-16

64

3.1.13

c- H ex

t-BuO b

11

P-388

31

(a) Concentration of analog that inhibits cell proliferation by 50% divided by concentration of paclitaxel that achieves same result. (b) This compound is a 10-deacetyl derivative. These observations suggest t h a t the i n t a c t A-ring s u b u n i t is an i m p o r t a n t structural element for cytotoxicity. The C-2 benzoate clearly plays a role in the cytotoxicity of paclitaxel, since 2-deoxytaxol, 2.1.41, is essentially inactive [26].

237 Due to the important role of the C-2 substituent for proper binding, it is clear t h a t small modifications at this site may lead to optimization of the activity, and it is not surprising t h a t several groups have reported efforts in this direction (Table 1). Several conclusions may be derived from the results in the table, even though cytotoxicities are generally reported for different cell lines. First of all, introduction of p a r a substituent at the benzoate invariably leads to loss of activity (see 2.1.26a,b and 3.1.3). Thus, the fit of the benzoate within the binding site seems to be r a t h e r tight, unless complex conformational changes are engendered by the p a r a substitution. It is unclear whether an aromatic ester at C-2 is needed for activity. Both Chen et al. [27] and Ojima et al. [31] have shown t h a t reduction of the C-2 benzoate to a cyclohexanoate leads to substantial loss of activity (see 2.1.26c and 3.1.13). In contrast, Georg et al. [64], working in the 10-acetyltaxotere series (see 3.1.12), report no loss of activity on hydrogenation of the C-2 benzoate. These apparent discrepancies are very difficult to interpret. Smaller aliphatic C-2 esters [27] (see 3.1.26d) and heteroaromatic ones [84] (see 3.1.9 and 3.1.10) are also poorly active. The most productive modifications to date have been carried out by Kingston et al., who reported large increases in cytotoxicity for some C-2 oand m - s u b s t i t u t e d benzoates [29]. In addition to i m p r o v e m e n t s in the cytotoxicity vs. paclitaxel, these analogs are clearly more effective in promoting microtubule assembly in vitro. These compounds, especially 3.1.2 and 3.1.5, are therefore promising, although neither data vs. resistant cell lines nor i n vivo evaluation have been reported. 5.3.2. Paclitaxel Analogs Modified at C-4 Chen and co-workers have extensively explored the SAR at C-4. Table 2 shows some of the highlights. As with C-2, deacylation or deoxygenation at C-4 leads to complete loss of activity (see 2.2.16 and 2.2.45). Introduction of large groups is also deleterious (2.2.24h), but aliphatic esters slightly larger than acetyl lead to improved activity (2.2.24j). Small carbonates and carbamates at C-4 are also quite active, especially in conjunction with improved side chains (for details on improved side chains, see chapter 6).

238

a'~. NH

O

\

AcO

O OH

Iii1< OH HO

BzO

X

Table 2: Cytotoxicity of Paclitaxel Analogs Modified at C-4 Cpd.

R

R'

X

IC50/IC50

Ref.

(paclitaxel) a 2.2.45

Ph Ph

Bz Bz

OH H

>25 n.d. b

19, 32 36

2.2.24h

Ph

Bz

OBz

100

20

2.2.28a

Ph

Bz

OCO2Me

0.9

19

2.2.24j

Ph

Bz

OCO-c-Pr

0.4

20

3.2.1

Ph

Bz

3.9

19

4.5 d

35

2.2.1{}

Ok\ 2 - - N-~J ~

l-o 3.2.2

t- oc

O N.

l-o / - - 3.2.3

Ph

Bz c

",,1

OCO-i-Pr

(a) Measured in HCT-116 cells. (b) Very poor tubulin polymerization activity. (c) This compound is a 10-deacetyl derivative. (d) Measured in B-16 melanoma cells. Georg et al. have also reported a derivative modified at C-4, 3.2.3, which is slightly less active t h a n paclitaxel. Upon further exploration, modification of the C-4 function is likely to afford very potent derivatives. 5.3.3. Paclitaxel Analogs Modified at the Oxetane Ring Many of the derivatives described in section 5.2.3, in which the oxetane ring has been opened, were evaluated for their ability to polymerize tubulin [23]. None of t h e m displayed any measurable activity and they are therefore useless in defining the SAR at this locus. Most of them are missing an acetoxy group at C-4 which, as we discussed above, is crucial for binding. This is in agreement with Kingston's early results [2]. Thus, the oxetane seems to play

239 an essential role in the binding of paclitaxel to microtubules. It is not known whether the oxetane acts to rigidify ring C and point the C-4 acetoxy group in the appropriate direction for binding, or whether the oxetane oxygen itself is a binding element. The derivatives that would answer this question (e.g. the one bearing a cyclobutane ring in place of the oxetane) have not yet been described. 5.3.4. Analogs Modified at the C-7 Position C-7 Xylosyltaxol, 3.4.1 was isolated from the bark and leaves of T a x u s b a c c a t a , and shown to be more potent t h a n paclitaxel in the tubulin polymerization assay [3, 85]. Other analogs were prepared either from baccatin derivatives or from paclitaxel itself. Most of the analogs tested (Table 3) have in vitro activity comparable to paclitaxel and docetaxel. With the exception of large lipophilic substituents such as the silyl ethers (see 3.4.8 and 3.4.10), any modification seems well tolerated, with some analogs being slightly more cytotoxic than paclitaxel. It seems likely that the C-7 substituent is not engaged in significant interactions at the binding site, and t h a t chemical modification at this position will only serve to m o d u l a t e the activity (perhaps via altered solubility, metabolism, biodistribution). Some of the C-7 derivatives were tested in vivo. With the exception of the carbonate derivative 3.4.6, none of the

compounds compared favorably with paclitaxel at their MTD. Interestingly, even cyclopropane derivative 2.4.14 retains in vitro activity, in spite of the slight conformational alteration imparted to the C ring vs. paclitaxel. 5.3.5. Analozs Modified at the C-9 Carbonyl This section examines primarily the effect of reducing the C-9 carbonyl on the cytotoxicity. Some analogs carry also C-10 and/or C-7 modifications and are discussed here for the sake of convenience. As with the C-7 position, even with the small database available, it is clear t h a t most modifications, including complete defunctionalization, are well tolerated at C-9. None of the modifications effected led to complete loss of activity. The derivative 2.5.31, f e a t u r i n g a completely defunctionalized northern half, is only 5-6 times less active than paclitaxel. Compounds with a partially hydroxylated northern segment have activities of the same level as paclitaxel and docetaxel (Table 4).

240

R "NH O

AcO

O X

p h - - " L ~ _ O .... OH HO

" BzO

Table 3: Cytotoxicity of Paclitaxel Analogs Modified at C-7 Cpd.

R

X

In vitro

Cell

In vivo

IC50/IC50

Line

activityb

Ref.

paclitaxel a 3.4.1

Bz

([~)Xylosyl

n.d.c

-

3

3.4.2

Bz

([~)OAc

1.3

J774.2

85

3.4.3

Boc

(~)L-Phenyl

0.44

P-388

52

3.4.4

Bz

([~) L-Alanyl

2.3

B-16

86

3.4.5

Bz

([~)N,N-dimethyl

2.3

S-16

86

3.4.6

Bz

(~)OC02Et

3.4.7

Bz

3.4.8

Bz

3.4.8 2.4.1

alanyl d

glutaryl 1.5

HCT-116

289 (40)

([~)OCONHBu

1.0

HCT-116

157 (50)

([3)OMs

0.9

HCT-116

Bz

([~)OSiEt3

>20

HCT-116

Bz

(a)OH

0.5

HCT-116 126-154 (30-

-

-

45 45 45 45 45

32) 3.4.10

Bz

(a)OSiMe3

>20

HCT-116

2.4.7

Bz

2.4.8

Boc

H

1.0

HCT-116

157 (50)

45

H

0.4

HCT-116

156 (64)

45

2.4.16

Bz

2.4.14

Bz

A6-dehydro

1.2

HCT-116

161 (60)

45

A7,19-cyclopropa

2.0

HCT-116

2.4.12

Bz

156 (80)

45

(a)F

2.9

HCT-116

185 (132)

45

3.4.11

Boc

(a)F

1.2

HCT-116

147 (40)

45

-

45

(a) See Table 1 for definition. (b) I n v i v o data in unstaged M109 model. Values indicate T/C at the MTD (mg/Kg/inj., in parentheses). Paclitaxel gave T/C values of 183-276 at MTDs of 50-75 mg/Kg/inj. (c) IC50 0.4 v s . paclitaxel (1.0) in tubulin polymerization assay. (d) A C-10 deacetyl derivative.

241

R "NH 0 ph-~,~_

X

Y Z

0 ....

OH

0 HO

" BzO

OAc

Table 4: Paclitaxel Analogs Modified at the C-9 Carbonyl Cpd.

R

X

Y

Z

IC5o/IC5o (paclitaxel)a

Cell Line

Ref.

2.5.1

Boc

(~)OH

(a)OH

2.5.2

Boc

( [ 3 ) O H (~)OH

(~)OH

1.9b

P-388

56

([~)OH

2.0b

P-388

56

2.5.4

Boc

(~)OH

(a)OH

(a)OH

3.2b

P-388

56

2.5.10

Bz

3.5.1

Bz

([~)OAc

(a)OH

(~)OH

8-10

P-388

17

([~)OAc

H

(~)OH

0.5

P-388

17

3.5.2

Bz

([~)OAc

H

H

1

P-388

17

2.5.31 2.5.24

Bz Bz

H H

H (~)OH

H (~)OH

5-6 14

P-388 B-16

17 64

2.5.25

Boc

(~)OH (~)OH

B-16

64

Boc

(~)OH (~)OH

1.1

2.5.26

(~)OH H

1.8

B-16

64

(a) See Table 1. (b) This value is referenced to docetaxel, not paclitaxel. Reduction of the C-9 carbonyl yields active a or 13carbinols (see 2.5.1 and 2.5.2). A 10-deoxy-9-dihydroderivative is much less bioactive t h a n paclitaxel, b u t simply switching the side chain to the one from docetaxel restores the activity (see 2.5.24

vs.

2.5.26). These observations reinforce the notion t h a t the northern

half of the molecule does not intimately interact with the microtubule binding site. 5.3.6. Analogs Modified at the C-10 Position Table 5 shows some of the analogs t h a t bear modified C-10 substituents. Some also bear modifications at C-7 and are discussed here for the sake of convenience. Although it is well known t h a t introduction of polar esters or other functions at C-10 leads to loss of activity [52], minor modification with

242 relatively small substituents at this position have been shown to lead to active analogs [66].

X RHN

0

P h i _ _-

Oy

\

0 ....<

OH

0 HO BzO

OAc

Table 5: Paclitaxel analogs modified at C-10 [19, 60, 66] Cpd.

R

X

Y

IC50/IC50 paclitaxel a

3.6.1 2.5.19 3.6.2 3.6.3 3.6.4 3.6.5 3.6.6 3.6.7 3.6.8 3.6.9 3.6.10 3.6.11 3.6.12 3.6.13 3.6.14

Bz Bz Boc Bz Bz Boc Bz Bz Bz Bz Boc Bz Boc Bz Boc

=O H H H H H OCO-n-Bu OCO-c-Pr OCONMe2 OMe OMe OCO2Me OCO 2Me OCOPh OCOPh

(~)OH (~)OH (~)OH (a)OH H H (~)OH (~)OH (~)OH (~)OH (~)OH (~)OH (~)O H (~)OH (~)OH

14 1.5 0.5 1.0 7.2 3.5 1.4 1.0 0.4 5.0 0.5 1.2 0.6 0.9 0.9

(a) See Table 1. Cell line: HCT-116 in all experiments. While a 10-keto group leads to a substantial loss of activity (see 3.6.1), the functionality at C-10 can be completely removed without loss of activity (see 2.5.19, 3.6.2 and 3.6.3) [60, 63, 64, 65]. On the other hand, deletion of both the C-7 and C-10 functions leads to some drop in cytotoxicity (see 3.6.4 and 3.6.5). In general, functions including esters, carbonates, carbamates and ethers are all

243 conducive to good activity, i.e. comparable with paclitaxel. These observations once again support the theory that the functionalities in the northern half of the core are not involved in binding to the microtubule. 5.3.7. Analogs Modified at C- ll/C- 12 Very little information is available on the role of the bridgehead C-11/C12 double bond vs. the bioactivity of the taxanes, since all natural taxoids are endowed with such double bond, and it is very difficult to modify it chemically. Chen and co-workers have reported some unusual chemistry at C-10 that results in the formation of dienone systems at C-10->C-18 [60]. C-12 fluorinated derivatives, where the double bond has moved into conjugation with the C-9 carbonyl, were also obtained as side products. Biological evaluation of some of these compounds (see Figure 3) shows that migration of the C-11/C-12 double bond leads to some loss in activity. Ten-fold drops in cytotoxicity (vs. paclitaxel) are seen with dienones 2.5.18, 3.7.1 and 3.7.2. The fluorinated derivatives are also ten-fold less active than paclitaxel, except for derivative 3.7.3, which bears a ~-methyl grouop at C-12, and is over 100-fold less active [19]. It is likely that, due to the importance of the C-13 side chain in the binding process, its exact spatial positioning is crucial to the activity of these analogs. Even slight conformational changes in the A ring might simply alter the spatial relationship of the side chain vs. the other binding elements in the molecule (the C-2 and C-4 esters). 5.3.8. Analogs Modified at the C-14 Position The biological activity of analogs bearing a functionalized C-14 has been explored in a preliminary fashion by Kant et al. [12] and Ojima et al. [13, 70]. The 14-(~)OH analog of paclitaxel, 2.7.14 (Table 6) shows slightly reduced cytotoxicity vs. the parent drug. I n v i v o evaluation showed that this derivative is essentially devoid of antitumor activity [12]. Switching the side chain to the one found in docetaxel, as predicted, results in a slightly improved performance (see 2.7.15 and 2.7.11) [12]. Even a cyclic carbonate at C-14/C-1 is compatible with good activity, but only in the presence of the docetaxel side chain, the paclitaxel analog being remarkably less active (2.7.12 vs. 3.8.1). A C-1/C-14 acetonide (see 2.7.13) is deleterious to activity.

244

#

BzHN O

O

R

O ....~ OH

2.5.18 3.7.1 3.7.2

R=(~)OH R=(~)OCOCHFC1 R=(cz)OH

ICso/ICso(pacl) 9.5 9.5 10 O

BzHN O RI,,,,~..~ ~ , ~ o ....

Z'.. i

BzO 3.7.3 3.7.4 3.7.5

J,

OAc

RI= F; R2 =Me; 1~3 = (a)OH RI= Me; R2 =F; R3 = (a)OH RI= Me; R2 =F; R3 = (~)OH

Figure 3: Paclitaxel analogs modified at C-10/C-12 and their

IC5o/IC5o(pacl) 230 13.5 9.5 in vitro

cytotoxicity (HCT-116)

Analogs where the side chain was introduced at C-14 instead of C-13 were much less active than docetaxel [13, 70]. Not enough is known about the SAR at C-14 to draw final conclusions as to the involvement of this position at the binding site. Since most of the analogs in Table 6 have similar activity to paclitaxel, it is likely that the C-14 functionality does not perform a binding function, and therefore only minor changes in cytotoxicity can be realized by fine-tuning such functionality. 5.3.9. Misce!!.aneous Analogs Klein reported on the synthesis of novel paclitaxel derivatives featuring a contracted seven-membered B-ring, 3.9.1 and 3.9.2 (Figure 4 ) [ 4 8 ] . Interestingly, these compounds were of comparable activity to paclitaxel in the

245 in vitro P-388 cytotoxicity assay. More work needs to be done to assess the

potential of these unusual analogs.

FI~HN

PhA

O

\

2

OH

_- Lo .... OH

_ R40 OR 3 0 i =

O

Table 6: Cytotoxicity of Derivatives Modified at C-14. Cpd.

R1

R2

R3,R4

IC5o/IC50 paclitaxel a

Cell Line

Ref.

2.7.14

Bz

Ac

H,H

4.0

HCT-116

12

2.7.15 2.7.11 2.7.12

Boc Boc Boc

Ac H H

3.8.1

Bz

H

2.7.13

Boc

H

H,H H,H C=O C=O C(Me)2

1.0 1.0 1.0 17 7.5

HCT-116 HCT-116 A121 A121 A121

12 12 13, 70 13, 70 13, 70

(a) See Table 1. The Ojima group described the synthesis and biological evaluation of two novel nor-seco analogs of paclitaxel and docetaxel, 2.8.44 and 2.8.45 [74]. These compounds are 20-40-fold less potent than paclitaxel in a number of tumor cell lines. These results thus clearly indicate the importance of the A-ring for the proper binding of paclitaxel and docetaxel to their biological target. A pentacyclic paclitaxel derivatives (2.9.4), prepared by photochemical irradiation of paclitaxel by Chen et al. [76], failed to show any activity in the tubulin polymerization assay as well as in cytotoxicity assays. The core of this molecule is grossly distorted with respect to the one in paclitaxel, and no activity would be expected. A recent report from Commerqon et al. provides the first example of a C19 modification [88]. The fact that a C-19-hydroxylated docetaxel analog (3.9.3) exhibits slightly better activity than the parent drug in the tubulin disassembly

246 assay suggests that chemical modifications at C-19 may lead to useful derivatives. AcO R.N o ,, X,o.c o. 3.9.1 R=Bz 3.9.2 R=Boc OBz OAc RHN

O

0 _. // OH

\

~ ~OHo

,

O .

HO

BzHN "

Ogz OAc

AcO I ~, / H,,,

0

O

/? OH \l~ 2.9.4

o ....

HO

dR HO

BocHN

O

P h ~ O

~

/

O/OH

~ 3.9.3

~

~-~o

HO BzHN

OAc

....

o.

0

OBz OAc

AcO

O

I~,,,,,II

jOH ~oH'" 3.9.4

p h ~ - ~ ' O _ ....

o.

2.8.44 R=Bz 2.8.45 R=Boc

~ HO

~~o (DBz OAc

Figure 4: Miscellaneous paclitaxel and docetaxel derivatives

247 Finally, among the many docetaxel metabolites isolated, one (3.9.4) features a novel core functionalization, i.e. a hydroxyl group at C-6. Such hydroxylation leads to a 30-fold drop in activity vs. docetaxel, i.e. detoxification of the drug [89]. 5.3.10. Conclusion Although much work remains to be done in this area, a qualitative picture of the SAR of paclitaxel is beginning to emerge. At least three functional elements, i.e. the C-13 side chain (see chapter 6) and the C-2 and C-4 esters, are intimately involved in interactions at the binding site. It appears that the northern half of the molecule and the tetracyclic skeleton (including an intact oxetane) function essentially as a molecular scaffolding to hold these binding elements in the proper orientation. Some uncertainty still exists about a possible binding role for the oxetane oxygen and the C-1 hydroxyl group. Modifications of the essential functions may therefore lead (and in some cases this has been achieved in cell culture) to more potent paclitaxel analogs, through further optimization of the fit with the microtubule site, whereas modifications at the non-essential positions may modulate the activity by changing the physico-chemical parameters of the molecule or via other secondary effects. R~'EgENCIi~

.

.

.

.

.

7.

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58. 59. 60. 61. 62. 63. 64.

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252 69. 70. 71. 72.

73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87.

Appendino, G.; Gariboldi, P.; Gabetta, B.; Pace, R.; Bombardelli, E.; Viterbo, D. J. Chem. Soc. Perkin Trans. 1 1992, 2925. Ojima, I.; Park, Y.H.; Sun, C.M.; Fenoglio, I.; Appendino, G.; Pera, P.; Bernacki, R.J.J. Med. Chem. 1994, 37, 1408. Py, S.; Khuong-Huu, F. Bull. Soc. Chim. Fr. 1993, 130, 189. (a) Appendino, G.; Varese, M.; Gariboldi, P.; Gabetta, B. Tetrahedron Lett. 1994, 35, 2217. (b) Appendino, G.; Cravotto, G.; Enrifi, R.; Jakupovic, J.; Gariboldi, P.; Gabetta, B.; Bombardelli, E. Phytochemistry 1994, 36, 407. Appendino, G.; Jakupovic, J.; Cravotto, G.; Varese, M. Tetrahedron Lett. 1994, 35, 6547. Ojima, I.; Fenoglio, I.; Park, Y.H.; Sun, C.M.; Appendino, G.; Pera, P.; Bernacki, R. J. Org. Chem. 1994, 59, 515. Py, S.; Pan, J.W.; Khuong-Huu, F. Tetrahedron 1994, 50, 6881. Chen, S.H.; Combs, C.M.; Hill, S.E.; Farina, V.; Doyle, T.W. Tetrahedron Lett. 1992, 33, 7679. Chiang, H.C.; Wood, M.C.; Nakanaira, Y.; Nakanishi, K. J. Chem. Soc., Chem. Commun. 1967, 1201. Chen, S.H.; Farina, V.; Huang, S.; Gao, Q.; Golik, J.; Doyle, T.W. Tetrahedron 1994, 50, 8633. Swindell, C.S.; Krauss, N.E.; Horwitz, S.B.; Ringel, I. J. Med. Chem. 1991, 34, 1176. Georg, G.I.; Cheruvallath, Z.S.; Himes, R.H.; Mejillano, M.R.; Burke, C.T.J. Med. Chem. 1992, 35, 4230. Long, B.H. unpublished results. Suffness, M. Ann. Rep. Med. Chem. 1993, 305. Rose, W.C. Anti-Cancer Drugs 1992, 3, 311. Nicolaou, K.C.; Couladouros, E.A.; Nantermet, P.G.; Renaud, J.; Guy, R.K.; Wrasidlo, W. Angew. Chem. Int. Ed. Engl. 1994, 33, 1581. Lataste, H.; S~nilh, V.; Wright, M.; Gu~nard, D.; Potier, P. Proc. Natl. Acad. Sci. USA 1984, 81, 4090. Mathew, A.E.; Mejillano, M.R.; Nath, J.P.; Himes, R.H.; Stella, V. J. Med. Chem. 1992, 35, 145. Klein, L.L.; Yeung, C.M.; Li, L.; Plattner, J.J. Tetrahedron Lett. 1994, 35, 4707.

253 88. 89.

Margraff, R.; B6zard, D.; Bourzat, J.D.; Commer~on, A. Bioorg. Med. Chem. Lett. 1994, 4, 233. Harris, J.W.; Katki, A.; Anderson, L.W.; Chmurny, G.N.; Paukstelis, J.V.; Collins, J.M.J. Med. Chem. 1994, 37, 706.

The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B. V. All rights reserved

255

6 THE C H E M I S T R Y OF THE TAXOL| S I D E CttAIN: S Y N T H E S I S , MODIFICATIONS AND CONFORNATIONAL STUDIES Joydeep Kant Johnson Matthey Inc., Biomedical Materials, 2003 Nolte Drive, West Deptford, New Jersey 08066, U.S.A.

6.1. INTRODUCTION

Taxol | (1.1.1), a complex antineoplastic diterpene isolated from Taxus brevifolia by Wani and Wall [1], has recently been approved for the treatment of cisplatin-refractory ovarian cancer and metastatic breast cancer [2,3]. The cytotoxicity of this drug is due to microtubule-mediated interruption of mitosis, which occurs through tubulin polymerization and formation of extremely stable and non-functional microtubules, which are abnormally resistant to depolymerization [4]. The limited availability of Taxol | from natural resources and its high clinical importance have stimulated considerable interest, within the synthetic community, toward the synthesis of this complex molecule and its analogs, with the aim of designing better antitumor drugs. Over the years, many approaches to the total synthesis of Taxol | have appeared in the l i t e r a t u r e [5], and recently Holton and Nicolaou, independently, have recorded successful approaches to this challenging target [6, 7]. Due to the length of these approaches, the total synthesis of Taxol| may not be feasible on an industrial scale. Taxol | was initially produced

256 exclusively by a tedious extraction procedure from the bark of the Pacific Yew; one Kg of the drug is isolated from the bark of 3,000 yew trees [8,9]. 0 18

AcO 0 ~ . _~19 OH

~3\

'2' ',,16,~ 4

'4

~

~'0

0

1.1.1 Taxol |

To circumvent this problem, Greene, Potier and coworkers [10] developed an efficient semi-synthetic approach. The chemistry involves an enantioselective synthesis of (2'R, 3'S)a-hydroxy-~-amino acid derivative 1.1.2 and its coupling to suitably protected 10-desacetylbaccatin III (10-DAB), 1.1.3, at the C-13 position. To date, the semi-synthetic approach appears to be the most practical way of producing Taxol| a large scale. Since 10-DAB is isolated from the needles of the widely distributed T a x u s b a c c a t a (yield: ca. lg/Kg dry leaves), a renewable source of 10-DAB is available in large quantities [11]. O

OH

Ph" ~ NH

O

- R oH 1.1.2

0

HO . . . . .

,,

n No 1.1.3

Thus, the success of the semi-synthetic approach relies on an efficient asymmetric synthesis of 1.1.2, followed by its attachment to 1.1.3. Therefore, in recent years, various academic and pharmaceutical research groups have

257 focused their efforts on the development of practical asymmetric syntheses of the side chain and its attachment to the baccatin core. Furthermore, these strategies have allowed the synthesis of a variety of side-chain analogs of Taxol| in this way helping to establish Structure-Activity Relationships (SAR) in search of more effective anti-tumor drugs. Since the discovery of Taxol| many reviews and accounts have appeared describing its chemistry and biology [5,12-19]. This chapter focuses only on the chemistry of the side chain. The first section describes a variety of chiral and non-chiral approaches to the synthesis of 1.1.2 and its mode of attachment to 1.1.3. The second section deals with the chemistry and biology of the side-chain modified analogs and their SAR. The final section summarizes what is known about the conformation of the Taxol | side chain in relation to the core and the biological mode of action. 6.2. SYNTHESES OF THE SIDE CHAIN 6.2.1. Asymmetric Epoxidation/Dihydroxylation Approaches Greene and co-workers employed the Sharpless asymmetric epoxidation followed by regioselective ring opening of the chiral epoxide 2.1.2 in their synthesis of the Taxol| side chain [20]. The key intermediate, 2.1.2 was prepared by Sharpless epoxidation from cis-cinnamyl alcohol 2.1.1, followed by oxidation of the epoxy alcohol and esterification of the resulting acid. The regiospecific ring opening of 2.1.2 using trimethylsilyl azide and zinc chloride afforded a - h y d r o x y - ~ - a z i d o amino acid 2.1.3 in high yield. Next, Obenzoylation followed by reduction produced the desired 2.1.5 via O->N-benzoyl migration. Although the epoxy alcohol was isolated with an enantiomeric excess (e.e.) of only 78%, a single recrystallization of 2.1.5 from chloroform afforded pure material in >95% e.e. (Scheme 1). The above chemistry was significantly improved by replacing the epoxidation step with the Sharpless dihydroxylation reaction. Treatment of i n e x p e n s i v e m e t h y l c i n n a m a t e , 2 . 1 . 6 , in aqueous acetone with dihydroquinidine-4-chlorobenzoate (DQCB) and N-methylmorpholine N-oxide (NMMO) in conjunction with a catalytic amount of OsO4, produced the diol 2.1.7 in fair yield with an e.e. of >98% [21]. Chemoselective tosylation at C-2 followed by mild base treatment afforded epoxide 2.1.2 which was converted to

258 2.1.5 in a one-pot procedure. Similarly, the side chain of Taxotere | 2.1.9, an

analog of Taxol |

was also synthesized (Scheme 2).

[Scheme 1]l Ph CH2OH \--/

i, i i, i i i

iv

=

2.1.2

oyp.

~

OMe OH 2.1.3

0

vi

O

phil_

Ph/JJxN H

N3 O II P h ~ O M e

=

~

O2Me

2.1.1

v

N3

0 ph , ~ ~ ~

0

phil"_

OMe

OH

0

2.1.5

2.1.4

Conditions: (i) t-BuO2H, Ti(OiPr)4, L-diethyl tartrate, CH2C12, -30 ~ 61%; (ii) RuC13, NaIO4, NaHCO3, CC14, CH3CN/H20; (iii) CH2N2, ether, 84% overall; (iv) TMSN3, ZnC12, H3O+, 90%; (v) PhCOC1, NEt3, DMAP, CH2C12, 94%; (vi) H2, PcYC,MeOH, then Ar, 89% (>95% e.e.).

Scheme 21 OH Ph@co2Me

i

0

~ Ph

0 OMe

=

p

O2Me

OR

2.1.6

/" 2.1.7 R = H ii ~ 2.1.8 R = OTs N3

iv

=

~ Ph

2.1.2 O R'~NH

O

O

v

OH

2.1.3

OMe

or v i

ph/~~_

OMe

OH

2.1.5 R=Ph 2.1.9 R=tBuO

(i) DQCB, NMMO, Os04 (cat.), acetone/H20, 51% (>98% e.e.); (ii)p-TsC1, NEt3, CH2C12, 88%; (iii) K2C03, H20, DMF, 91%; (iv) NAN3, AcOH, MeOH/H20, 95%; (v) PhCOC1, NEt3, DMAP; H2, Pd/C, EtOAc, 92%; (vi) Boc20, H2, Pd/C, EtOAc, 92%. Conditions:

259 Coupling of 2.1.5 to baccatin was initially reported to be a very difficult operation, probably due to the hindered nature of the C-13 hydroxyl group in baccatin [10]. The C-2 hydroxyl group in 2.1.5 was protected as an acid-labile ethoxyethyl ether and the ester was hydrolyzed to the free amino acid 2.1.10. Treatment of 7-triethylsilyl (TES) baccatin III (2.1.11) in toluene with 6 equiv of 2 . 1 . 1 0 , 6 equiv of di-2-pyridyl carbonate (DPC), and 2 equiv of 4(dimethylamino)pyridine (DMAP) at 80 ~ for 100 h produced the C-2', C-7protected 2.1.12 in 80% yield (yield based on only 50% conversion). The protecting groups were removed by using 0.5% HC1 in ethanol to give Taxol| in good yield (Scheme 3). This method suffers from two major drawbacks: esterification required excess amounts of the expensive chiral amino acid (6 equiv or more) and only 50% conversion was observed even under forcing conditions [10].

[Scheme 311 OAc

O

_

Ph

HO ....

NH --

O

i

ii

p hv- ~ _, , ~ O H

.o" BzO

~c

~

o.yo

2.1.11

2.1.10

O P h ' ~ NH

O

3 ii

P h ~ O

....

,

Taxol, 1.1.1

~ 0

2.1.12 Conditions: (i) DPC, DMAP, PhMe, 80 ~

80%; (ii) HC1, EtOH/H20, 0 ~

89%.

The esterification step was significantly improved by Commer~on and co-workers. The phenylisoserine side chain was introduced as an oxazolidine,

260 which

underwent

esterification

under

standard

DCC/DMAP

coupling

conditions in high yield [22]. The methodology avoided the use of an excess of enantiomerically pure amino acid 1.1.2, and the coupling yield was over 90%. The key intermediate was again a chiral epoxide (2.1.15, a homolog of 2.1.2), which was synthesized by condensation of the boron enolate of (4S, 5R)-3b r o m o a c e t y l - 4 - m e t h y l - 5 - p h e n y l - 2 - o x a z o l i d i n o n e (2.1.13) w i t h b e n z a l d e h y d e followed by t r e a t m e n t with lithium ethoxide, to produce chiral epoxide 2.1.15 in high optical purity.

Scheme 411

0 0 BrV~NA O \ / ,, ~ -~ Ph

OH 0 0 i = p h ~ ~ J ] ~ N~]O Br ~ ,~ -" Ph

2.1.13

2.1.14

N3 0 P h / ~ ~ - OEt

ii

0 r_ ph~--~CO2Et 2.1.15

Boc,.

NH2 .0. iv

= p h , / ~ ~_

OEt

iii

v

NH 0

P h Z ~ ~ - OEt

vi

_

OH

2.1.16

OH

OH

2.1.17

2.1.18

Ph~.~,- "c02H

Ph~}~.~,,-"C02Et vii

Boc~ N ~ O 2.1.19

Boc~N~ 0 2.1.20

Conditions: (i) NEt3, Bu2BOTf, CH2C12,-70 ~ to rt, then PhCHO,-78 ~ to 0 ~ 58%; (ii) EtOLi, THF, -75 ~ to 15 ~ 81%; (iii) NAN3, EtOH, NH4C1, 60 ~ 99%; (iv) H2, Pd/C, EtOAc, 86%; (v) Boc20, NaHCO3, CH2C12, 20 ~ 76%; (vi) CH2=C(Me)OMe, PTSP, PhMe, 80 ~ 99%; (vii) LiOH, EtOH/H20, 20 ~ 100%.

Epoxide 2.1.15 was converted to N-protected amino ester 2.1.18 via the azide method. Conversion to acetonide 2.1.19 was followed by saponification to afford

261 the free acid 2.1.20 in high yield (Scheme 4). T r e a t m e n t of 1.5 equiv of 2.1.20 with 2.1.22, using 1.6 equiv of DCC and 0.5 equiv of DMAP in toluene at 80 ~ afforded high yields of 2 . 1 . 2 4

without

any

detectable

epimerization.

Deprotection using formic acid followed by N-acylation and removal of the Troc group afforded Taxol | in high yield. Taxotere | (2.1.29) was similarly prepared (Scheme 5).

[ Scheme 5 OR~

0

O =

HO.... H d B ~

i:I(~A'~cO

Ph~ i

O"'

Boc-~_O

HO BzO 2.1.23 RI=CO2CH2CCI 3 2 . 1 . 2 4 R1 = Ac

Troc = C02CH2CCI 3

OR 1 O N_H2 0 ph/'Z',.,v~ O.... -

~

OR1 0 0 T r o c

.... J]~

I

2.1.21 RI= CO2CH2CCI 3 2.1.22 R 1 = Ac

ii

,

II

iii

~

~ S

o.

H

121

O

2.1.25 RI= CO2CH2CCI 3 2 . 1 . 2 6 R] = Ac FI10 R2HN Ph

O _ OH

R2HN

OH o

0 .... OH H d ~ ~BzO .:

2.1.27

O

RI=CO2CH2CC]3, 2.1.28 RI=AC , R2=Bz

I~I(~A~cO

R2=Boc

O

....

- ,

O BzO OAc 2.1.29 RI=H, R2=Boc (Taxotere| 1.1 1 RI=AC, R2=Bz (Taxo]|

(i) DCC, DMAP, PhMe, 2.1.20, 80 ~ 98%; (ii) HCO2H, 20 ~ 78% (2.1.25), 80% (2.1.26); (iii) Boc20, NaHCO3, THF, 20 ~ 87% or BzC1, NaHCO3, EtOAc, 87%; (iv) Zn, AcOH, MeOH, 60 ~ 89-90%. Conditions:

262 In a following paper, Greene has shown t h a t even the C-2 epimer of 2.1.20 can function as a suitable precursor to Taxol | and Taxotere | Indeed, C-2 epimerization during the acylation reaction is virtually complete, and this opens the possibility of using, as esterifying agents, side chain precursors that are not stereochemically homogeneous at C-2 [23]. In a similar approach, Didier and co-workers prepared diastereomeric mixtures of oxazolidinecarboxylic acids 2.1.30 by treating amino ester 2.1.9 with chloral [24]. Coupling of 2.1.30 with 2.1.21 by the DCC/DMAP procedure produced 2.1.31. Treatment with zinc in acetic acid removed the two carbonate groups and hydrolyzed the oxazolidine to provide 2.1.32, which was converted to Taxotere | by the usual procedure (Scheme 6).

[Scheme 6~ CCl 3

I

HN~"O

Ph"

i =

CO2H

2.1.30 NH 2 .O

I

0130~ ~ , , , _ . O ' I~ Hn H

OH

TrocO~,

O i1|

HO Bz()

2.1.31

O

\

/O OTroc

OH iii

2.1.29

OH HO

-

BzO

"

~Ac

2.1.32 Conditions: (i) 2.1.21, DCC, DMAP, PhMe, 99%; (ii) Zn, AcOH, EtOAc, rt; (iii) Boc20, MeOH, rt, 70% overall.

Jacobsen's approach to the Taxol | side chain employed (salen)Mn(III) complex 2 . 1 . 3 3 to effect the asymmetric epoxidation step [25]. Partial h y d r o g e n a t i o n of commercial ethyl phenylpropiolate (2.1.34) to Z - e t h y l c i n n a m a t e (2.1.35) using Lindlar's catalyst followed by epoxidation with commercial bleach in conjunction with 2.1.33 afforded the Z - ( R , R ) e p o x i d e 2.1.15 in >95% e.e. The trans isomer of 2.1.15, however, was a significant byproduct in the epoxidation reaction. Nonetheless, in a highly regioselective

263 ring-opening process, the mixture of cis and trans epoxides gave 2.1.36 in high yield upon t r e a t m e n t with ammonia. Intermediate 2.1.36 was subsequently transformed into the desired product 1.1.2 under standard conditions. The low cost of the reagents makes this an attractive and practical approach to the side chain (Scheme 7).

Scheme 7 / O'McI%

tBu

tBu

U

2.1.33 0 Ph

~

CO2Et

i =

2.1.34

Ph\

/CO2Et

ii

~

h~--~C P O2Et

2.1.35

2.1.15 O

NH 2 Q iii=

Ph

: OH _

2.1.36

NH 20. NH 2 ~

,v

Ph

: OH

2.1.37

P h / [ L NH OH

v

=

O

ph/~.~_

OH

OH

1.1.2

Conditions: (i) H2, Lindlar cat., 84%; (ii) NaOC1, (R,R)-2.1.33 (6 tool %), 4-phenylpyridine-Noxide (0.25 equiv), CH2C12, 56%, >95% e.e.; (iii) NH3, EtOH, 100 ~ 65%; (iv) Ba(OH)2, then H2SO4, 92%; (v) PhCOC1, NaHCO3, then HC1, 74%.

Sharpless reported a six-step asymmetric synthesis of the Taxol| side chain using the novel chiral catalyst (DHQ)2-PHAL [26]. Asymmetric dihydroxylation of 2.1.6 afforded diol 2.1.38 in 99% e.e. In a one-pot procedure, 2.1.38 was protected as a cyclic orthoester and regioselectively opened by acetyl bromide to produce a 6:1 mixture of bromo acetates, favoring the desired stereoisomer 2.1.39. T r e a t m e n t of 2.1.39 with sodium azide followed by hydrogenation gave the N-acetyl derivative 2.1.40 which was converted to 1.1.2 in fair overall yield (Scheme 8).

264 A very similar approach was also reported by Koskinen, although the enantioselectivity of the dihydroxylation step was not reported [27]. Utilization of Z-cinnamates in this approach would eliminate the need for the double inversion operation at C-3, but these substrates are dihydroxylated in very poor e.e. [27].

[ Scheme 8/ 0 ph

v

OH OCH3

0

Br

OH

2.1.6

CH 3

2.1.39 0

Me'~NH

iii

OCH 3 _ OAc

~

2.1.38 0

0

0

p h ~ l J L.. O C H OH

2.1.40

Ph'~NH

3

iv, v

0

ph..~.jl ~ OH

(~H 1.1.2

Conditions: (DHQ)2-PHAL (0.5 tool %), K2OsO2(OH)4, NMMO, t-BuOH, rt, 72% (99% e.e.); (ii) MeC(OMe)3,p-TsOH, CH2C12, rt, then AcBr, CH2C12,-15 ~ 60%; (iii) NAN3, DMF, 50 ~ H2, Pd/C, MeOH, rt, 74%; (iv) 10% aq. HC1, heat; (v) PhCOC1, 2 N aq. NaOH, rt, 72% overall. Potier and his group attempted an oxyamination reaction directly on 13cinnamoyl baccatin 2.1.41 in order to develop a semi-synthesis of Taxol | and Taxotere (ii) [28]. Negligible stereoselectivity was observed when employing the classical Sharpless catalytic procedure; two pairs of threo and erythro isomers 2 . 1 . 4 2 - 2 . 1 . 4 5 were obtained in poor yields along with recovered starting material (ca. 50%). The isomers were separated and converted to Taxol | and Taxotere | derivatives using standard conditions. This procedure allowed the synthesis of all possible diastereomers and regioisomers for SAR studies (vide infra) (Scheme 9). 6.2.2. Chiral Pool Approach Greene and collaborators employed (S)-phenylglycine (2.2.1), available from the natural chiral pool, as a starting material in the synthesis of Taxol | and Taxotere (i!) side chains [29]. Aminoacid 2.2.1 was reduced to the alcohol,

265 followed by in s i t u protection of the amino group to afford 2.2.2 or 2.2.3. An a t t e m p t at a t a n d e m S w e r n oxidation/addition of v i n y l m a g n e s i u m bromide to the crude aldehyde g e n e r a t e d racemic 2.2.5.

Scheme 9/ AcO ~r~

O Ph

o,k_

O ~~

r~

s .....~ ~ . ~ k

AcO

R2 .O

;

__~ Ph

O

OTroc

o ....

"' Ho" o~z~ ~ ~ o HO OBz 2.1.41 2.1.42 (2'R,3'S)R]=OH, I~2=NHCO2tBu 2.1.43 (2'S,3'R)RI=OH, I~2=NHCO2tBu 2.1.44 (2'R,3'B)RI=NHCO2tBu,R2=OH 2.1.45 (2'S,3'R)RI=NHCO2tBu, R2=OH =

Conditions: (i) t-BuOCONCINa, AgNO3, OsO4, H20/CH3CN.

[Scheme 1011

0

N_H2

RJJ~NH i =- ~ O H

=

~ ~ O

0

OH

RflL'NH ii =

H

'L2

2.2.1 O

2.2.2 R = Ph 2.2.3 R = Ot-Bu

R~ N H iii

R~ N H iv, %

2.2.5 R - P h 2 . 2 . 6 R = Ot-Bu

2.2.4 O

-

0 -

OH

2.1.10 R = Ph ~

2.2.7' R = Ot-Bu

Conditions: (i) LiA1H4, PhCOC1 or Boc20, 74-79%; (ii) (COC1)2, DMSO, i-Pr2NEt, CH2C12,-78

~ to-35 ~ (iii) CH2=CHMgBr, -78 ~ RuC13, NaIO4, NaHCO3, 62-82%.

62% overall; (iv) CH2=CHOEt, H3 O+, 90%; (v) cat.

266 However, by adding the Swern oxidation product to the vinylmagnesium bromide in a m i x t u r e of T H F - C H 2 C 1 2 gave 2 . 2 . 5 with good s y n diastereoselectivity (9:1) and no racemization. Intermediate 2.2.5 was further elaborated to 2.1.10. Side chain 2.2.7 was similarly synthesized (Scheme 10). 6.2.3. Lithiobenzylamine Method A conceptually novel approach to the synthesis of the Taxotere | side chain employed the dianion of N-Boc-benzylamine 2.3.1 [30]. Treatment of such dianion with acrolein produced a 6:1 mixture of amino alcohols with a bias for the desired s y n alcohol 2.3.2 . The diastereomers were separated and the s y n alcohol was protected as a (trichloroethoxy)methyl ether, 2.3.3. Oxidative cleavage of 2.3.3 gave a racemic acid which, upon resolution using (+) ephedrine, gave the optically pure enantiomer 2.3.4 (Scheme 11). Scheme 11/

NHBoc

Boc.

Boc. NH -

i =

-

ii

~

NH_

. ~ O w C C i

2.3.1

2.3.2

3

2.3.3

Boc.NH 0 II1,1v ....

_

-

~

OH 0 ~ CCI3

2.3.4

(i) sec-BuLi, acrolein, THFfrMEDA,-78 ~ 49%; (ii) BrCH2OCH2CC13, proton sponge, CH3CN, 70 ~ (iii) RuC13, NaIO4, NaHCO3, CC14/ CH3CN/H20, rt, 80%; (iv) (+)ephedrine, 83%. Conditions:

6.2.4. Diastereoselective Michael Approach Davies reported a synthesis of the Taxol|

side chain

based upon

asymmetric tandem conjugate addition-electrophilic hydroxylation of

tert

butyl

cinnamate 2.4.1 [31]. The chemistry involved conjugate addition of the homochiral lithium (S)-(a-methylbenzyl)-benzylamide 2.4.2 to 2.4.1, followed by hydroxylation

of the i n t e r m e d i a t e

enolate

with

(+)-(camphorsulfonyl)

267 oxaziridine to produce the a n t i product 2.4.3 in high diastereoselectivity. In order to adjust the stereochemistry at C-2, the intermediate 2.4.3 was then subjected to hydrogenolysis followed by methanolysis and benzoylation, to afford the a n t i hydroxy amide 2.4.4 in high yield. Multi-step inversion at C-2 via oxazoline 2.4.5 gave the well-known methyl ester 2.1.5 in high yield and optical purity (Scheme 12). [Scheme 1211

-"

0

Ph

Ph

"

OBut

//~" N J

+ ph/~N/~Ph Li

2.4.1

/

i ~ Ph

OBut 2.4.3

Ph

Ph'~NH

O

Ph

O

N,~O OMe

OH

ii, iii, iv

OH

2.4.2

O

0

Ph'~NH

=-

OMe Ph

-

Ph

O : OH

OMe

O 2.4.4

2.4.5

2.1.5

Conditions: (i) (+)-(Camphorsulfonyl)oxaziridine, THF,-78 ~ 89%, >98% d.e.; (ii) H2, Pd/C, AcOH; (iii) HC1, MeOH; (iv) PhCOC1, NEt3, 96% overall; (v) DEAD, PPh3, THF, 0 ~ 77%; (vi) HC1, MeOH; (vii) NaHCO3, 91% overall.

6.2.5. The ~-Lactam Method The racemic synthesis of the Taxol |

side chain v i a ~-lactams was

reported by Palomo and co-workers; their chemistry utilized as a key intermediate azetidine-2,3-dione 2.5.1 [32], which in turn can be prepared by a number of [2+2] protocols [32, 33]. Diastereoselective reduction of 2.5.1 using sodium borohydride afforded exclusively cis lactam 2.5.2. Protection of the C-3 hydroxyl group followed by oxidative removal of the N-aryl group furnished 2.5.3. Ring opening of the ~-lactam with chlorotrimethylsilane in methanol afforded the key intermediate a-hydroxy-~-amino ester 2.5.4, which was then converted to 2.1.5 in the usual way (Scheme 13) [32]. The first direct application of ~-lactams in the semi-synthesis of Taxol| was demonstrated by Holton [34]. Treatment of 2.1.11 with n-BuLi at-40 ~ in

268 THF chemoselectively generates the alkoxy anion at the C-13 position, and this readily attacks r a c e m i c [~-lactam 2 . 5 . 5 to afford a m i x t u r e of chromatographically separable diastereomers 2.5.6 and 2.5.7 in high yield.

[Scheme 13 ]1 O

O

Ph

H O,,,, i

,,,Ph I N

,,

oJ

C I"'~O,,,, II, II1=

.....

OMe 2.5.2

.~

NH 2 .C) =

2.5.3

OMe

2.5.1

iv

Ph

~

oJ

: OH

Ph

v

O Me

,,,Ph I N.

=

NH Ph

U

OH

2.5.4

O Me

2.1.5

Conditions:(i) NaBH4, THF/MeOH; (ii) CICH2COC1, Py, CH2C12, 80% overall; (iii)

(NH4)2Ce(NO3)6, CH3CN/H20, 0-5 ~ -70 ~ to 20 ~ no yield.

70%; (iv) TMSC1, MeOH; (v) PhCOC1, NEt3, CH2C12,

Scheme 14/ OAc Et3SiO,"

tS

.:.

,Ph

i

0

Ph

HO BzO 2.1.11

(+)2.5.5

(~SiEt3

OSiEt3

+ HO ....

oJ-'

BzNH O .-= II phJ"-.,v.--'~-_ O ....

O

OAct 2 0 S i E t 3

OAc BzNH

,%

+ O

2.5.6 Conditions: n-BuLi or LiHMDS, -40 ~ to 0 ~

O

O

ph-~~O

t3 ....

OS,E, 2.5.7 80-90% of 2.5.6+2.5.7.

H

~

0

269 Furthermore, when excess racemic 2.5.5 (ca. 5-7 equiv) was used, an 8:2 mixture of diastereomers, with a bias for the desired 2'R, 3'S diastereomer, was isolated (Scheme 14). However, the use of excess ~-lactam and the c h r o m a t o g r a p h i c separation of the diastereomers can be avoided if optically pure cis (3R, 4S)~lactam is employed. Thus, t r e a t m e n t of 2.1.11 with 1.6-1.8 equiv of resolved, enantiomerically pure 2.5.5 in the presence of n-BuLi afforded only 2.5.6 in high yield [34]. Holton's semi-synthetic approach to Taxol| therefore, required a practical synthesis of enantiomerically pure ~-lactams. This was reported by Ojima, Georg and co-workers [35-37]. Their c h e m i s t r y relied on an enantioselective enolate-imine cyclocondensation to synthesize the required 3hydroxy-4-aryl-~-lactams. T r e a t m e n t of the enolate of ester 2.5.8, bearing Whitesell's chiral auxiliary [(-)trans-2-phenyl-l-cyclohexanol)] with silylimine 2.5.9 gave exclusively 2 . 5 . 1 0 in high enantiomeric purity (96-98% e.e.). Conversion to 1.1.2 was accomplished under standard conditions (Scheme 15).

[Scheme 1511

Ph

0 /~OTIPS

Ph +

I(

TIPSO, "

i

Me3si/ 2.5.8

2.5.9

0 iv

Ih111 =

NH2~

ph~ V OH .

OH

2.1.37

-HCI

o,7

"H

2.5.10

ph ) 1 " NH

=

Ph "

0

ph/~~]-OH OH

1.1.2

Conditions: LDA (2 equiv), THF, -78 ~ to rt, 85% (96% e.e.); (ii) Bu4NF, THF, rt, 98%; (iii) 6 N HC1, reflux, 100 ~ (iv) PhCOC1, aq. NaHCO3, CH2C12, 70%. (TIPS=triisopropylsilyl). This versatile approach was also used to synthesize a number of taxane analogs modified at the side chain (vide infra). In a modification of the above protocol, Oppolzer's (-)-10-dicyclohexyl sulfamoyl-D-isoborneol was used by

270 Georg and collaborators to prepare enantiomerically pure 2.5.13, the Holton intermediate [38].

High enantioselectivity was observed in this case also

(Scheme 16). Georg and co-workers reported the first attempt at using an asymmetric Staudinger reaction to prepare the required optically pure 13lactam.

Using

a peracylated galactopyranosyl template, these workers

initially reported that a single diastereomer, 2.5.16, was obtained [39]. A later study, however, demonstrated that this chiral template affords little or no diastereoselectivity in the Staudinger reaction (Scheme 17) [40].

[Scheme 1611

Ph O SO2N(C6H11)2

2.5.11

TBSO,

Ph

NTMS

TBSQ

N- H O

2.5.9

2.5.12

,Ph N Ph "~ O

2.5.13

Conditions: (i) LDA, THF, 94% (97% e.e.); (ii) PhCOC1, NEt3, DMAP, CH2C12, 96% (TBS= t-

Butyl dime thylsilyl).

[Scheme 1711 OAc ,...-OAc Ph

AcO

OAc

H

ArO~ 0

2.5.14

CI

2.5.15

OAc ,~OAc O AcO

OAc ~.~OAc O Ar

Ph

2.5.16

i

AcO

_ Ph

2.5.17

Conditions: NEt3, CH2C12, rt, 75% (2.5.16:2.5.17= 2:3) [Ar=p-methoxyphenyl].

Ar

271 A highly diastereoselective approach to the Taxol | side chain via the Staudinger reaction was reported by Farina and co-workers [41]. The reaction between L-threonine-derived imine 2.5.18 and the acid chloride 2.5.19 under typical Staudinger conditions afforded the desired cis ~-lactam 2.4.20 in high yield and with good diastereoselection ( 3 R , 4 S / 3 S , 4 R >10:1). The chiral template was removed by t r e a t m e n t of 2.5.20 with t e t r a b u t y l a m m o n i u m fluoride, followed by mesylation/elimination to give 2.5.22 quantitatively; this was then ozonized to 2.5.23, and finally base-promoted hydrolysis of the acetate and the oxalamide groups afforded the target compound 2.5.24 in high optical purity (99.5%). Lactam 2.5.24 was eventually used in the preparation of Taxol | using Holton's protocol (Scheme 18).

{Scheme 18 ]J

o= =

,co

N..~~

+

i

=

,COoj"" -

O-" -Cl

CO2Me 2.5.18

,COoj"" -

OR ii N~~,,,. "I

"

CO2Me 2.5.19

2.5.20

"

I

OH :

N~-.~ CO2Me 2.5.21

R=SiPh2OBu t

m

,...

AcO.,,,

,,,Ph

iii

AcO,%

,,,Ph

N.O COeMe

2.5.22

CO2Me

2.5.23

HO,,,,

,,,Ph N. H

2.5.24

Conditions: (i) NEt3, CH2C12,-40 ~ to rt, 74%; (ii) Bu4NF, THF, AcOH, 82.5%; (iii) MsC1, NEt3, CH2C12, -78 ~ to rt; (iv) 03, CH2C12, -78 ~ the Me2S; (v) aq. NaHCO3, MeOH, 80% overall.

Bourzat and Commerqon reported a moderately diasteoselective Staudinger reaction using (S)-a-methylbenzylamine as chiral template [42]. The chiral imine 2.5.25 was treated with acetoxyacetyl chloride 2.5.19 in the presence of triethylamine, to give a 3:1 mixture of 2.5.26 and 2.5.27. This

272 mixture

was

subjected

to base-catalyzed hydrolysis

and

the

desired

diastereomer 2.5.28 was isolated by crystallization. Hydrolytic cleavage followed by removal of the chiral auxiliary by hydrogenation afforded the ester 2.1.9 (Scheme 19). This approach could also be used to prepare a number of analogs to be used for SAR purposes (vide infra).

(,Scheme 19]l AcQ i

Ph..11 Nv

""

,Ph " Nv Ph :

0~.._ I

Ph OH 3

0

+

Ph

~

Ph

Ph _ OH 3

2.5.27

H3C~NH

OH3

2.5.28

ii

0,/~- Nv

2.5.26

,Ph Nv

Ph

CH 3

2.5.25 HO,,,,

AcO.

0 : OH

B~ 0 Me

-

2.5.29

Ph

0 : OH

0 Me

2.1.9

Conditions: (i) AcOCH2COC1, NEt3, CHC13, 0 ~ to rt, 74% of 2.5.26 + 2.5.2"/(3:1); (ii) 1 M aq. KOH, THF, 0 ~ 67%; (iii) 6 N HC1, MeOH, reflux, 88%; (iv) H2, Pd/C, MeOH/AcOH then Boc20, CH2C12,aq. NaHCO3, 20 ~ 70%. Holton's asymmetric Staudinger approach to the synthesis of chiral cis3-hydroxy-4-arylazetidinones utilized the Evans strategy [43]. Under the Staudinger conditions, chiral acid chloride 2.5.30 and imine 2.5.31 afforded 2.5.32 in high yield and complete diastereoselectivity. Treatment of 2.5.32 with LiHMDS in dichloromethane followed by the addition of N-chlorosuccinimide gave 2.5.33 as a mixture of diastereomers, which were eventually converted into 2.5.1. Diastereoselective reduction using sodium borohydride as previously described produced the chiral alcohol 2.5.2. Hydroxyl group protection, Ndearylation and benzoylation afforded 2.5.36 (Scheme 20). A short synthesis, amenable to large scale production, of racemic 4-aryl and 4-heteroarylazetidin-2-ones was reported by Rey and coworkers at BristolMyers Squibb. Treatment of commercially available hydrobenzamide 2.5.37

273 with acetoxyacetyl chloride in the presence of triethylamine afforded cis ~lactam 2.5.38 as a diastereomeric mixture (at the aminal carbon) in good yield. Removal of the N-benzyl group by hydrogenation afforded racemic 2.5.39 (Scheme 21) [44].

[Scheme 20]1

pPh

0

c,

I~ Ph

/'~ O. N,,

+

Ph ,,,

O oJ_,

i.__

MeO

2~~. "

2.5.311

2.5.31

pPh

2.5.3

OMe

pPh

.

o

O.~N,,,

i___Li... " ~

2.5.33

OMe

HO% ,,,Ph N

vi, vii

2.5.2 ~

I

iv

OMe

2.5.34

EEO,,,, 0# 1

0 0~"

,,, N.

2.5.35

Ph

,Ph v

2.5.1 EEO,%

r

viii

H

OMe

0

,,"

Ph

N.

COPh

2.5.36

OMe Conditions: (i) NEt3, CH2C12, -78 ~ 93%; (ii) LiHMDS, CH2C12, -78 ~ NCS, 95%; (iii) aq. AgNO3, CH3CN, 0 ~ (iv) SiO2, >95% overall; (v) NaBH4, MeOH, 0 ~ 100%; (vi) Ethyl vinyl ether, MsOH, 0 ~ 100%; (vii) (NH4)2Ce(N03)6, CH3CN, 87%; (viii) PhCOBr, Py, CH2C12, 0 ~ 98%. [EE=l-Ethoxyethyl].

6.2.6. Chiral Sulfinimine Approach Davis and co-workers described the use of chiral sulfinimines in their enantioselective approach to the Taxol | side chain [45]. Addition of the lithium enolate of methyl acetate to readily available, enantiomerically pure

274 2.6.1, followed by desulfinylation/benzoylation, afforded N-benzoyl-~-amino ester 2.6.3 in good yield.

[Scheme 21 ]~ Ph"71

Ir

/ Ph

NT~N Ph

A c (3, , Ph "" " 0,/~,._ NI ~ Ph y

i

A cQ ii

r

2.5.37

""

""

Ph

0/I/_ NI

=

H

2.5.38 N 7

2.5.39

Ph Conditions: (i) AcOCH2COC1, NEt3, EtOAc, 5 ~ >95%; (ii) H2, Pd/C, EtOAc, 78%.

S u b s e q u e n t a s y m m e t r i c h y d r o x y l a t i o n u s i n g (+)-(camphorylsulfonyl) oxaziridine gave a 86:14 s y n : a n t i mixture of 2.1.5 and 2.4.4 in fair yield. Chromatographic separation of the diastereomers afforded 2.1.5 in good enantiomeric purity (>93%) (Scheme 22).

[Scheme 2211

O

O

I

O

H

Ph~ "N

ph~S'NH Ph

Ph

2.6.1

0

OMe

2.6.2

A N__~~

i v _ Ph "-

Ph

ph.fl

O

. OMe OH 2.1.5

ph - - ~ ~ O M e 2.6.3

0

AN

+ Ph

~

Ph

OMe OH 2.4.4

Conditions: (i)CH2=C(OLi)OMe, -78 ~ 76%; (ii) CF3CO2H, MeOH; (iii) NEt3, DMAP, PhCOC1, 76% overall; (iv) LDA, LiC1, (+)-(camphorylsulfonyl)oxaziridine, -100 ~ to -78 ~ 58%.

6.2.7. Aldol Reaction Approaches A n u m b e r of asymmetric aldol-type approaches have been used to prepare taxane side chains. Three basic strategies have been utilized: a) Use of

275 a chiral aldehyde or imine (or equivalent) in conjunction with an achiral enolate; b) Use of a chiral enolate equivalent in conjunction with an achiral aldehyde or imine; c) Use of a chiral catalyst to promote reaction between achiral partners.

Combinations of the above three strategies are also

conceivable. Hanaoka and his group employed optically pure (+)-tricarbonyl(q6-2trimethylsilylbenzaldehyde)chromium (0) complex 2.7.1 in conjunction with the titanium enolate of thioester 2.7.2

to afford, after desilylation and

decomplexation, the a n t i - a l d o l product 2.7.4 in a highly diastereoselective manner. Mitsunobu reaction with hydrazoic acid gave the desired s y n azide 2.7.5 in high yield. Reduction with triphenylphosphine and water furnished the amino derivative which was benzoylated to afford 2.7.6 in high optical purity (> 98%). Deprotection with thallium nitrate in methanol followed by hydrogenolysis gave the target compound 2.1.5 (Scheme 23) [46,47]. [Scheme 2311

OH

0

o + Cr

TMS

oc'lco

_

OBn

2.7.1

CO

;

SBu t

Cr

Bu t

..... //,i//

TMS

oc'l'co

2.7.2

s

2.7.3

CO

O

.JL OH

O

N3

/~~,._

P h ~ S B u OBn 2.7.4

iv

t

=

Ph~

Ph

:

Ph--NH SB

OBn 2.7.5

0 Ph'~NH

vii

~

O

_

OBn

2.7.7

v, vi~

Ph~

Ph/[L'NH OMe

viii

=

Ph

:

SBu t

OBn 2.7.6

0

O

~v ~

Ut

O

O _

OMe

OH

2.1.5

Conditions: (i) TiC14,NEt3, CH2C12, -78~ 93%; (ii) Bu4NF, HF, CH3CN/THF, -78~ to 0~

(iii) hv, Et20,0~ 63% overall; (iv) HN3, PPh3, DEAD, C6H6, rt; (v) PPh3, H2OfrHF, 60~ (vi) PhCOC1, DMAP, CH2C12, 0~ 53% overall; (vii) Tl(NO3)3, MeOH, rt, 90%; (viii) Pd/C, H2, EtOH, 60~ 89% (>98% e.e.)

276 Yamamoto's approach to the synthesis of 2.1.5 employed a double diastereoselection strategy utilizing chiral Lewis acid 2.7.9 [48]. Reaction of chiral imine 2.5.25 with a-silyloxy (Z)-ketene acetal 2.7.8 mediated by chiral boron reagent 2.7.9 produced enantiomerically pure syn adduct 2.7.10 (syn/anti = 99/1, diastereofacial ratio = 99/1). Hydrogenolysis followed by benzoylation under Schotten-Baumann conditions gave N-benzoyl-(2R,3S)-phenylisoserine methyl ester (2.1.5) in good yield (Scheme 24).

[Scheme 24]l Ph ~

N~

Ph OMe .--/~-~0 + (Et)3SiO Si(Et) 3

Ph

Me

~NH i -_ (S)-2.7.9

2.7.8

2.5.25

0

phi2-< k_ OMe OH 2.7.10

0 il, i l i

P h / ~ NH

0

w -

.

ph~~l"OMe OH

2.1.5 Conditions: (i) CH2C12,-78 ~

~~]"-0

(S)-2.7.9

91%; (ii) H2, Pd/C, MeOH; (iii) PhCOC1, aq. NaOH, THF, 68%

overall.

Approaches utilizing chiral enolates were reported by Swindell and Greene. Swindell and coworkers attempted an asymmetric hetero-Diels-Alder reaction between N-benzoylbenzaldimine 2.7.11 and chirally modified ketene acetal 2.7.12 [49]. Aqueous work-up furnished, however, aldol products as a mixture of diastereoisomers, with 2.7.13 as the major product. The isomers were subjected to debenzylation followed by transesterification to provide a mixture containing mainly 2.1.5 (93:7 syn/anti, only one syn isomer) (Scheme 25). One should note that several chiral alcohols were examined as templates for the above operation and (1S,2R)-(+)-trans-2-(1-methyl-l-phenylethyl)-lcyclohexanol was the best one. Greene's approach utilized the Oppolzer template. Camphorsultam 2.7.14 was condensed with benzaldehyde N-(t-

277 butoxycarbonyl)imine 2.7.15, to provide exclusively 2.7.17 with complete (>99%) enantio- and diastereoselection. The chiral auxiliary was then oxidatively cleaved, to afford 2.7.17 in fair overall yield [50] (Scheme 26).

[Scheme 25]l

O

Me Ph.,,~ Me (Me)3SiO~L_ / ~ + BnO/_~ 0

O Ph~N//'-.ph 2.7.11

Ph/~NH i

Me O Ph.J/Me OBn

2.7.13

2.7.12

0 Ihtll

Ph'~ NH 0 Ph~OMe 2.1.5 OH

Conditions:

(i) C6H6, rt, 75%; (ii) Pd(OH)2, H2; (iii) MeONa/MeOH, 82% overall.

{scheme 26]l

O

BocHN O N

+

=

OBn

NBoc

02 2.7.14 BocHN

2.7.15

Ph

_ N OBn 02 2.7.16

O

Phi_

OH OBn

2.7.17 Conditions:

(i) LiHMDS, THF,-78 ~ 66%; (ii) LiOH, H202, rt, then aq. Na2SO3, 0 ~ 70%.

6.2.8. Enzymatic Approaches The s y n t h e s i s of all d i a s t e r e o m e r s of 3 - p h e n y l s e r i n e s and 3phenylisoserines in enantiomerically pure form using enzymatic resolution was first reported by HSnig [51]. Racemic butyryl ester 2.8.3, synthesized as

278 shown below in Scheme 27, was resolved by hydrolysis with P s e u d o m o n a s fluorescens. Both the alcohol 2.8.4 and the unreacted isomer 2.8.5 were isolated in high enantiomeric purity (>98% e.e.). Sih and co-workers reported the enzymatic resolution of 3-acetoxy-4phenyl ~-lactams 2.8.6-2.8.8 using bacterial lipases [52]. The most suitable lipase for the various transformations was the P s e u d o m o n a s lipase P-30. Under these conditions, the undesired enantiomer was selectively hydrolyzed.

[Scheme 27]~

0

N3 i, ii

ph~~-'~OE t

0

N3

= Ph"'Z"'~OEt==

+

0

Ph~OEt

2.8.1

N3

0

OH

2.8.4

N3

0

O\ ~

2.8.5

0

Conditions: (i) NAN3, aq. EtOH, NH4C1, 60%; (ii) (n-PrCO)20, py, H20, 92%; (iii) Pseudomonas fluorescens, 2.8.4 (26%, >98% e.e.), 2.8.5 (35%, >98% e.e.). These intermediates were converted to the C-13 Taxol | side chain 1.1.2 by standard protocols (Scheme 28). Patel and co-workers at Bristol-Myers Squibb have optimized this resolution on a large scale and have applied it to the commercial production of Taxol| [53]. Chen has enzymatically resolved racemic trans-phenylglycidic ester 2.8.9 by transesterification with lipases in organic media. Thus, incubation of racemic 2.8.9 with Lipase MAP-10 in hexane-isobutyl alcohol (1:1) afforded 2.8.10 and 2.8.11 in high enantiomeric excess. The product and the substrate were separated by fractional distillation. The individual enantiomers were subsequently converted to the Taxol| side chain 2.1.5 in a number of steps. It is noteworthy that both 2.8.10 and 2.8.11 were converted to optically pure 2.1.5, as shown in Scheme 29 [54].

279 Chen's chiral oxazoline 2.4.5 was utilised by Kingston in a new semisynthesis of Taxol| saponification of 2.4.5 with aqueous NaOH provided 2.8.14, which upon treatment with 7-TES baccatin 2.1.11 underwent a smooth coupling reaction to afford 2.8.15. Acid-catalyzed hydrolysis afforded Taxol| in good yield [55] (Scheme 30).

[ Scheme

2811

II

Ac O,,,, ,,,Ph

AcO,,,,

N

.,"Ph

oJ-' N OCH3 Ph

,,"Ph N

(+)-2.8.7

""~"~

ACE),,,,

O

Ph (+)-2.8.8

N ~ Ph

OH OH .

1.1.2 A Bristol-Myers Squibb group reported the synthesis of chiral phenylisoserine ethyl ester 2.8.17 via diastereoselective microbial reduction of the prochiral ketone 2.8.16 [56]. Microorganisms from H. polymorpha SC 13865 and H. fabianii SC13894 effectively reduced the ketone in high yield (>80%) and optical purity (>95%) (Scheme 31). Similarly, Kayser and Kearns used a yeast-mediated stereospecific reduction of chiral (~-keto ester 2.8.20, obtained in three steps from natural (S)phenylglycine 2.2.1, as shown in Scheme 32. Under these conditions, the desired 2.5.4 was obtained diastereomerically pure, although no information on the optical purity of this intermediate was reported [57]. Standard transformations were used to convert 2.5.4 into side-chain synthon 2.1.5, although no details of the experimental procedures used are given.

280

6.3. S I D E

CHAIN

MODIFICATIONS

FOR

STRUCTURE-ACTIVITY

RELATIONSHIP STUDIES 6.3.1. Simplified Side Chain Analogs Studies

of n a t u r a l

and

semisynthetic

congeners

of Taxol |

have

d e m o n s t r a t e d t h a t a taxane ring and an ester C-13 side chain are required for a n t i t u m o r activity, since b a c c a t i n

III 1.1.3 and N - b e n z o y l

(2'R, 3'S)-3'-

phenylisoserine 1.1.2 are devoid of significant activity [58]. [ S c h e m e 29~

ph\O \

i \

0 =

CO2Me

0

P h " ' L ' ~ c O2Me +

2.8.9

PhJ/---&"CO2Bu i

2.8.10

N3 ii, iii 2.8.10

2.8.11

O

BzHN

~

=-- Ph

:

OMe

=

Ph

_

OH N3

v//i,/x

ph./~CO2Bu, OH

O

2.8.12

./~ Ph NH

N3 vii__

ph/~~CO2Bui OCOPh

"l"P'h .,1. OMe

OH

O

2.8.13

0

Ph 2.4.4

2.1.5

O

vi

OMe

OH

2.1.3

2.8.11

O

i v, v

=

xi

~~-,H CO2Me

Ph, H

2.1.5

2.4.5

Conditions: (i)Mucor miehei lipase MAP-10, i-BuOH, 2.8.10 (42%, 95% e.e.), 2.8.11 (43%, 95%

e.e.); (ii) Et2NH2Br, Et2A1C1, CH2C12,-15 ~ 90%; (iii) NAN3, DMF, 65-70 ~ 80%; (iv) PhCOC1, DMAP, CH2C12, rt, 93%; (v) H2, Pd/C, 50 psi, 80%; (vi) NAN3, aq. acetone, NH4C1, refl., 95%; (vii) PhCOC1, DMAP, CH2C12, rt, 91%; (viii) H2, Pd/C, 50 psi, 93%; (ix) Na2CO3, aq. MeOH, rt, then CH2N2, 81%; (x) SOC12, CHC13, reflux, 73%; (xi) 1 N HC1, MeOH, 80%.

281

[Scheme 30 ~1

Ph 2.1.11

H-_,,,~~_-CO2Me Ph H

H_,,,~z~-~-CO2H Ph H 2.8.14

2.4.5

\ AcO

Ph 0

O OSiEt3 iii

Taxol,

O....<

5--"0 Ph

1.1.1 0

HO

OAc OBz

2.8.15 Conditions: 0.1 N NaOH, 96%; (ii) DCC, 4-pyrrolidinopyridine (cat.), PhMe, rt, 95%; (iii) 0.1 N HC1, 95 ~ 75%.

[Scheme 31 ]m O

O

Ph~NH

O

Ph@Oet O

i

PhANH =

2.8.16

O

Ph'~~-__ OEt OH 2.8.17

Conditions: (i) H. polymorpha SC 13865, or H. fabianii, 80% (>95% e.e.).

In order to better define the SAR of Taxol | for the design of more effective drugs and to understand the features of the Taxol | binding site on the microtubule, a variety of side chain modifications have been reported. Recently, a comprehensive list of C-13 side chain analogs with their biological data has been reported in a review article [19]. This section briefly discusses the types of modifications that have been described and summarizes the SAR within each group.

282

[Scheme 32~ _NH2

NH 2 9 HCI

- O H ;

"o,

0 2.2.1

0 2.8.18

NH 2 .O

NH 2 O

Ph

OMe

"

iv

=

O

ph..~,.~ : OH

2.8.20 9

;;

ON;;; 0 2.8.19 NH 2 .O

O

Me

v

=

P h ~ O M OTMS

2.5.4 BzHN

vi, v / /

NH2

*" Ph

2.8.21

O -

OMe

OH

2.1.5 Conditions: (i) SOC12, Et20,-10 ~ 100%; (ii) NaCN, Li2CO3, THF, refl., 58%; (iii) HC1, MeOH, 73%; (iv) Baker's Yeast, 72%; (v) TMSC1, Py, 73%; (vi) PhCOC1, K2CO3, MeOH/H20, 70%; (vii) KF, H20, 91%.

The biological activity of each analog is usually determined in vitro. Two assays are n o r m a l l y carried out, as already described in C h a p t e r 5, i.e. a tubulin polymerization assay, and a cytotoxicity determination against one or more cancer cell lines. I n v i v o testing is not described in this chapter. The reader is cautioned t h a t different laboratories carry out these tests in different manners

(i.e.

t u b u l i n a s s e m b l y o r d i s a s s e m b l y m a y be m e a s u r e d ,

and

different cell lines m a y be used). It should be made clear t h a t the data presented here serve more as a preliminary screen to discern the usefulness of certain modifications t h a n as a definitive ranking of all the compounds. The first a t t e m p t s at understanding the SAR at the C-13 side chain were reported by Swindell's [59] and Potier's [60] groups.

The analogs 3.1.1-3.1.6

with deleted side chain substituents were synthesized by the esterification of the corresponding acids with 2.1.22, using dipyridyl carbonate according to the Greene protocol. These new analogs were found to be much less active t h a n Taxol|

in a microtubule binding assay (Figure 1) [59].

Similarly, simple

283 cinnamoyl derivatives (3.1.7 and 3.1.8) as well as the dihydroxy compounds (3.1.9 and 3.1.10) displayed poor activity (Figure 2) [60]. In view results, it was concluded t h a t the phenylisoserine moiety plays important role in the binding of Taxol | to microtubules, and that the functionalities present on the side chain serve to preorganize it for [59]. Figure 1: Simplified Side Chain Analogs [59]

of these a very various binding

AcO~, 2 OH allnl

is

0 O

3.1.1 a

Me--CO2=

-

OH

3.1.2 a =

(31%)

Me\/CO 2- (38%) | OH

3.1.,5 R =

A Ph NH (60%) L~CO2_ OH

3.1.3 R = p h i l / C O 2 - (69~176 /~ OH 3.1.6 R = Ph NH 3.1.4 R = ph-/"-,~_. - _CO2 (68%) L~ICO2_ OH

OH

(NA)

Activity defined as % of microtubule assembly in the presence of analog (Taxol = 100%) [59]. NA=not available

6.3.2 Analogs with 2', 3' Isomeric Structures The French group synthesized a variety of side chain analogs with epimerized or transposed s u b s t i t u e n t s [28, 60]. The data showed t h a t inhibition of microtubule disassembly is sensitive to the absolute configuration at C-2' and C-3'; in all cases, the natural 2'R, 3'S configuration was found to lead to more potent analogs (tubulin binding assay and cell culture) (Table 1). Interestingly, analogs 3.2.5-3.2.8, with 2',3' transposed substituents, were significantly less active t h a n the parent compounds 1.1.1, 3.2.1 and 3.2.2. Furthermore, analogs with u n n a t u r a l 2'S,3'R configuration in the transposed

284 series were less active compared to the analogs with 2'R,3'S configuration. Taxotere | (2.1.29) was the most active compound in this series. It was proposed t h a t the changes in bioactivity are the result of changes in the side chain conformation imposed by the s t r u c t u r a l or configurational modifications [60]. Figure 2: Simplified Side Chain Analogs [60]

R10,

O OH

R2-'~'O .... (DBz ~Ac .ul

Cpd.

R1

R2

ID50/ID50 (Taxol) b

3.1.7

H

(E)-CH=CHC6H5

23

3.1.8

COCH3

(E)-CH=CHC6H5

100

H

CH(OH)CH(OH)C6H5

3

COCH3

CH(OH)CH(OH)C6H5

60

3.1.9 a 3.1.1{} a

(a) Threo compounds (2'R, 3'R+2'S, 3'S). (b) Concentration of the drug leading to a 50% inhibition of the rate of microtubule disassembly. The ratio ID50/ID50 (Taxol) is defined as 1 for Taxol [60]. 6.3.3 Analogs with Modifications at the C-3' Nitrogen A variety of analogs with modifications at the C-3' nitrogen have been reported by various groups. In general, modifications at this position are tolerated very well (Tables 2, 3, and 4). The most promising analog, Taxotere | (2.1.29, bearing a t-Boc on the C-3' nitrogen instead of a benzoyl group, and not bearing an acetyl group at the C-10 hydroxyl) is presently undergoing clinical trials in France and United States. Taxotere | more active t h a n Taxol |

has been found to be slightly

in the microtubule disassembly assay; furthermore,

it has shown a better cytotoxicity profile t h a n Taxol | against J774.2 and P388 cell lines [61,62] (see Table 1).

285 Table 1: Analogs with 2', 3' Isomeric Structures [28, 60]

R3

.O

Ph~O

........ HO O BzI-IdA~ O

R2

Cpd.

R1

R2

R3

ID50/ID5o

IC5o

(Taxol) a

P388b

Taxol (2'R, 3'S)

COCH3

OH

NHCOPh

1.0

0.27

3.2.1 (2'S, 3'R)

COCH3

OH

NHCOPh

4.5

7

Taxotere(2'R,3'S)

H

OH

NHCO2Bu t

0.5

0.13

3.2.2 (2'S, 3'R)

H

OH

NHCO2Bu t

30

-

3.2.3 (2'R, 3'S)

COCH3

OH

NHCO2Bu t

0.5

0.17

3.2.4 (2'S, 3'R)

COCH3

OH

NHCO2Bu t

30

7

3.2.5 (2'R, 3'S)

COCH3

NHCOPh

OH

10

6

3.2.6 (2'S, 3'R)

COCH3

NHCOPh

OH

110

-

3.2.7 (2'R, 3'S)

H

NHCO2Bu t

OH

10

-

3.2.8 (2'S, 3'R)

H

NHCO2Bu t

OH

160

-

(a) Drug concentration leading to a 50% inhibition of the rate of microtubule disassembly divided by the concentration of Taxol needed to achieve such inhibition [60]. (b) Concentration needed to inhibit cell proliferation by 50%(l~g/mL) in murine P388 leukemia cells. Except for Taxotere |

and N - g l u t a r y l analog 3.3.3, none of the other

analogs reported by the French group exhibited i n t e r e s t i n g levels of activity. Analog 3.3.2, having a free amino group at C-3', was also found to be less potent t h a n the N-amido analogs [60] (Table 2). Georg's group used the ~ - l a c t a m s y n t h o n a p p r o a c h to s y n t h e s i z e a series of N-benzoyl modified analogs (3.3.5-3.3.17) (Table 3).

Most of the

analogs displayed good cytotoxicity and were found to be active in a tubulin assay. In general, none of the analogs were clearly superior to Taxol |

or

286 Taxotere | based on these preliminary data. Compounds 3.3.8 and 3.3.11 were found to be very active in the microtubule assembly assay but were less active t h a n Taxol | in the in vitro cytotoxicity assay against B-16 melanoma cells. What is most notable from the data in Table 3 is that, in general, many small structural modifications have a deleterious effect on the bioactivity [63-66]. Swindell and coworkers also synthesized a small number of analogs modified at the N-benzoyl group and evaluated their microtubule assembly activity at a fixed concentration, as well as their cytotoxicity toward J774.2 cells. None of the analogs were found better than Taxol | [67] (Table 4).

Table 2. Analogs Modified at the N-Benzoyl Group [60]

R2

2 o.

O

phi-J]'-_

0,

iii

SSsso

OH

0

Cpd.

R1

R2

ID50/ID50 (Taxol)a

3.3.1

H

NHTs

5

3.3.2

COCH3

NH2

44

3.3.3

H

NH CO(C H 2)3CO O H

1

3.3.4

H

NHCO(C6H4)SO3H

5.5

(a) For definition, see Table 1. 6.3.4. Analogs with Modifications at the C-3' Phenyl Group The C-3' phenyl group was replaced by substituted phenyl groups and by hetero-aryl groups by Georg et al. and the analogs were evaluated in a microtubule assembly assay and against B-16 melanoma cells [63, 66, 68-70]. Some analogs with substitutions at the para-position of the C-3' phenyl group (especially methoxy or fluoro) displayed good activity. Introduction of a heteroaryl group at C-3' also led to very potent compounds, especially the 2-

287 pyridyl- (3.4.11) and the 2-furyl (3.4.14) derivatives. The latter was the most active compound in cell culture, and was at least 3 times more potent t h a n Taxol| (Table 5). Table 3. Analogs with Modifications at the N-Benzoyl Group [63-66] O

.~ NH O R1 Ph~O OH

AcO

O

OH

.... O

R1

ID50/ID50 (Taxol) a

B-16 IC50/IC50 (Taxol)b

Reference

3.3.5

p-Me-Ph

1.6

1.4

63

3.3.6

p-CF3-Ph

6.0

17.7

63

3.3.7

(Me)2CHCH2

1.18

2.65

64

3.3.8

(Me)3CCH2

0.74

3.13

64

3.3.9

(Me)3C

2.61

22.4

64

3.3.10

p-C1-Ph

2.4

1.5

65, 66

3.3.11

p-MeO-Ph

0.6

1.3

65

3.3.12

3,4-(C1)2-Ph

2.1

10.6

65

3.3.13

m-C1-Ph

2.0

1.8

65

3.3.14

m-N(Me)2-Ph

1.4

1.6

65

3.3.15

o-CH3-Ph

1.9

7.5

65

3.3.16

p-NO2-Ph

2.0

20.7

65

3.3.17

p-F-Ph

1.2

4.3

65

Cpd. i

i

(a) Microtubule disassembly assay, for definition see Table 1. (b) Drug concentration needed to reduce B 16 melanoma cell proliferation by 50%, divided by concentration of Taxol needed to achieve the same effect.

288 Table 4: Analogs with Modifications at the C-3' N-Benzoyl Group [67] O

.~ NH O R1

AcO

O

p h - ' ~ ~ _ _ O.... OH

OH

O

i|l|l

Cpd.

R1

microtubule assembly a (%1

IC50 J774.2 b (~M)

3.3.18

p-N3-Ph

90

0.28

3.3.6

p-CF3-Ph

80

0.90

3.3.19

PhCH20

94

0.19

3.3.20

p-CH3CO-Ph

84

1.6

3.3.21

p-PhCO-Ph

39

1.5

3.3.22

o-OH-Ph

93

0.56

(a) Percent of microtubules assembled v s . Taxol at a fLxed drug concentration (Taxol= 100). (b) Concentration needed to inhibit J774.2 cell proliferation by 50% (Taxol = 0.09gM) [67] I n t e r e s t i n g l y , the cyclohexyl analog 3 . 4 . 1 6 assembly properties t h a n Taxol |

exhibited b e t t e r m i c r o t u b u l e

and was found to be equipotent with Taxol |

in the cytotoxicity assay. This result shows t h a t an aromatic s u b s t i t u e n t at C3' is not necessary for biological activity. Ojima and co-workers used the [~-lactam chemistry discussed in section 6.2.5 to introduce side chain modifications in the Taxotere | series (Table 6). Specifically, the introduction of alkyl and alkenyl groups at C-3' was reported for the first time. The analogs were tested against various cancer cell lines as well as in the microtubule disassembly assay [71-73]. Analogs 3.4.17 and 3.4.24 were found to be equipotent to

Taxotere | in the m i c r o t u b u l e d i s a s s e m b l y

assay, b u t the same analogs were found less cytotoxic a g a i n s t P388 and ovarian A121 cancer cell lines.

289 Table 5: Analogs with Modifications at the C-3' Phenyl Group [63, 66, 68-70]

0 Ph~J" NH 0 R I ~ O OH -

OH ....

~oot ~

HO

- H-O OBz 6 iC

ii

Cpd.

R1

ID50/ID50 (Taxol) a

B-16 IC50/IC50 (Taxol) b

reference

3.4.1

p-Me-Ph

2.4

3.0

63

3.4.2

p-C1-Ph

1.9

2.2

66

3.4.3

p-MeO-Ph

0.5

1.0

68

3.4.4

m-C1-Ph

4.4

6.7

68

3.4.5

o-Me-Ph

5.1

>33

68

3.4.6

p-C1-Ph

1.9

2.2

68

3.4.7

3,4-(C1)2-Ph

7.1

40

68

3.4.8

3-N(Me)2-Ph

4.6

5.8

68

3.4.9

p-NO2-Ph

1.1

35

68

3.4.10

pF-Ph

1.1

1.2

68

3.4.11

2-pyridyl

0.7

0.8

69

3.4.12

3-pyridyl

0.5

27

69

3.4.13

4-pyridyl

0.4

1.3

69

3.4.14

2-furyl

0.9

0.3

69

3.4.15

3-furyl

0.9

3.3

69

3.4.16

cyclohexyl

0.29

0.91

70

i

(a) For definition, see Table 1. These data are referred to microtubule assembly experiments. (b) For definition, see Table 3.

290 Interestingly, 3-cyclohexyl derivative 3.4.24 appears much less active Taxotere|

t h a n the c o r r e s p o n d i n g Taxol |

vs.

analog 3.4.1{}, at least in cell

culture. (Table 5). This discrepancy, either due to assay dependence, cell line differences, or to a subtle interplay between the C-3' amide function and the C3' carbon substituent, underlines some of the difficulties encountered in trying to reach firm conclusions on the SAR of the taxanes. Table 6: Taxotere @ Analogs Modified at the C-3' Phenyl Group [71-73]

IIO

R~O\

ButOT,,. N H

\

O ~

OH

o ....

OH

_ OBz

HO

O

i

Cpd.

R1

R2

ID50/ID50 (Taxol) a

P388 IC50 (nM) b

A121 IC50 (nM) b

Taxotere

OH

Ph

0.70

9.9

1.2

3.4.17

OH

(Me)2CH

0.78

12.2

1.9

3.4.18

OAc

(Me)2CH

3.4.19

OH

(Me)3C

1.45

125

3.4.20

OH

Ph-CH=CH

1.45

264

3.4.21

OH

(Me)2C=CH

0.64

12.8

3.4.22

OAc

(Me)2C=CH

3.4.23

OAc

MeCH=CH

3.8

0.46 -

-

0.90

(a) For definition, see Table 1. (b) Concentration needed to inhibit cell proliferation by 50%. A121 is an ovarian cancer cell line. 6.3.5. Analogs with Modifications at C-2' v

A v a r i e t y of pro-drugs have been synthesized by derivatizing the C-2' position of Taxol|

These analogs typically have no activity in the tubulin

291 assay. The chemistry and biology of these derivatives are the subject of a separate chapter in this book (see Chapter 3). Besides prodrugs, not too much is known about the role of the C-2' hydroxyl group in microtubule binding. Kant et al. found that replacing the hydrogen bond donor C-2' hydroxyl group with the hydrogen bond acceptor methoxy group resulted in considerable loss of cytotoxicity. The C-2' deoxytaxol derivative, synthesized using Barton's chemistry, was 75 fold less cytotoxic than Taxol| vs. the HCTll6 human colon carcinoma cell line. Replacing the hydroxyl group with another hydrogen bond acceptor, fluorine, also resulted in loss of cytotoxicity (Figure 3). The observed loss was attributed either to the failure of the phenylisoserine side chain to adopt a "preferred conformation" required for effective binding or to the need for participation of the 2'-hydroxyl group in intermolectflar hydrogen bonding at the receptor site [74].

0 Ph~NH

2~

0

phil_

0 ,,

cpd

HCT116 ICso (~tM)

Taxol 3.5. I 3.5.2

0.004 0.297 0.866

3.5.3

0.475

ii

R

0

3.5.1. R = H 3.5.2. R = O M e 3.5.3. R = F

Figure 3" C-2' modified analogs [74] 6.3.6. Side Chain Analogs of 9(R)-Dihydrotaxol An Abbott group reported a variety of side chain analogs of the potent 9(R)-dihydrotaxol 3.6.1 [75,76]. These side chains were synthesized by the ~lactam method and coupled to the 9(R)-dihydrobaccatin using Holton's chemistry. Analogs were evaluated by a microtubule assembly assay and in vitro cytotoxicity assays against several human and murine tumor cell lines (Table 7). Many of the compounds prepared have outstanding in vitro antitumor activity.

292 Table 7: Side Chain Analogs of 9(R)-Dihydrotaxol [75,76]

H R1CO-N

AcO

O

R2-~~_ OH

OHO

H

O .... NO

. (3Bz

O

R1

R2

ID50/ID50 (Taxol) a

3.6.1

Ph

Ph

0.76

0.053

3.6.2

3.6.3

t-BuO 2-furyl

Ph Ph

0.87 0.73

0.0025 0.034

3.6.4

methyl

Ph

3.17

2.02

3.6.5

3.6.6

t-butyl t-butyl-CH2

Ph Ph

2.36 1.03

0.083 0.052

3.6. 7

t-butyl-N H

Ph

1.06

0.017

3.6.8 3.6.9

t-amyl-O isopropyl-O

Ph Ph

0.73 0.75

0.0046 0.0092

3.6.10

neopentyl-O

Ph

0.82

0.019

3.6.11

adamantyl- O

Ph

1.54

0.01

3.6.12

isobutyl-O

Ph

1.03

0.028

3.6.13

ethyl-O

Ph

0.82

0.043

3.6.14 3.6.15

benzyl-O Bz

Ph MeOCH2

3.14

0.057 >0.1

3.6.16

Bz

PhOCH2

5.81

>0.1

3.6.17

t-BuO

4-thiazolyl

1.36

0.0014

3.6.18

t-BuO

methyl

1.08

0.043

3.6.19

t-BuO

3.6.20

t-BuO

vinyl ethyl

0.92 0.61

0.016 0.011

3.6.21

t-BuO

butyl

2.15

0.018

3.6.22

t-BuO

cyclohexyl

0.57

0.014

3.6.23

t-BuO

isobutyl

0.95

0.00036

3.6.24

t-BuO

benzyl

>17

>0.1

Cpd

IC50 (~g/mL) b P388

3.6.25 t-BuO pentyl 2.0 0.016 (a) ID50=Drug concentration that reduces tubulin concentration by 50%. (b) see Table 6.

293 The activity of the 3'-alkyl derivatives of Taxotere @ (in particular isobutyl analog 3.6.23 and cyclohexyl derivative 3.6.22) is especially remarkable. The authors briefly discuss the superior performance of these compounds in vivo [76] and comment that they are less toxic than Taxol| The authors also conclude that the C-3' substituent must fit within a hydrophobic pocket, and that this pocket is obviously limited in size (for example, the 3' benzyl derivative 3.6.24 is totally inactive). 6.3.7. Extended Side Chain Analogs I n t e r e s t i n g analogs of Taxol| and Taxotere (!i) with one carbonhomologated side chains were synthesized by Georg and co-workers. The compounds were p r e p a r e d using 10-DAB and Commerqon's modified oxazolidineacetic acid protocol. Unfortunately, neither 3.7.1 nor 3.7.2 (Figure 4) displayed any significant activity in a tubulin assembly assay. The lack of activity was attributed to unfavorable conformations of these analogs which might prevent binding to the receptor site on microtubules [77]. In addition, as discussed in section 6.3.6 (see 3.6.24, Table 7) there seems to be a well-defined limitation to the length of the side chain that can achieve proper binding, and addition of an extra methylene unit to such chain is totally deleterious here as well. O

AcO

O OH

R-~NH -

O .... OH

O

O

3.7.1. R = Ph 3.7.2. R = t-BuO Figure 4: Extended Side Chain Analogs [77] 6.4. CONFORMATIONAL STUDIRS Because the topology of the binding site on the microtubules and the bound state conformation of Taxol |

are unknown, potentially important

294 features of the binding site have been derived from the reported crystal structure of Taxotere | 2.1.29 [78]. The solution conformation of Taxol | was studied by NMR spectroscopy in non-aqueous solvents and it was found to be very similar to the one observed in the crystal structure of Taxotere | NOESY and ROESY experiments suggested that the side chain is folded under the diterpene core; this was further supported by NOEs observed between H-2' and the C-4-acetyl group. The observed small JH2'-H3' coupling constant of 2.7 Hz is indicative of hindered rotation within the side chain, which is attributed to the intramolecular hydrogen bonding [79-82]. Swindell and collaborators reported conformational analysis of methyl N-benzoylisoserinate (2.1.5)and methyl phenyllactate using MM2 calculations. These modeling studies suggested that the hydrogen bonding or electrostatic interactions involving the proximate C=O, OH, and NH moieties in the side chains are the major determinants of side chain conformation [59]. Gu~ritteVoegelein studied the conformation of Taxotere | and some side chain analogs and proposed a similar intramolecular hydrogen bonding [60]. Swindell proposed that these interactions "preorganize" the side chain for binding to microtubules [59]. However, recent NMR studies of Taxol| and Taxotere | in polar solvents (DMSO/H20 and MeOH) suggest a different side chain arrangement. It was proposed t h a t Taxotere | and all 2'R, 3'S active analogs display conformations in which the C-2 benzoate group holds the side chain in a particular spatial arrangement. This new conformation is stabilized by a hydrophobic interaction between the C-2 benzoate and the C-3' N-benzoyl or NBoc groups. Thus, hydrophobic interactions together with the network of intramolecular hydrogen bonding discussed above would be responsible for a specific orientation of the phenylisoserine side chain. This study also led the authors to suggest a binding process of active taxanes to tubulin: the first step is the recognition of the taxane core by the binding site; the second step is hydrophobic interactions between the C-2 benzoate and the C-3' amino substituents of the side chain. These interactions orient the side chain in such a position that the C-2' hydroxyl and the C-3' phenyl groups can interact with appropriate tubulin residues, leading to stabilization of the drug-receptor complex [83]. Scott and Swindell studied conformations of Taxol|

and its side chain

methyl ester 2.1.5 by NMR spectroscopy and molecular modeling in an

295 aqueous environment. Interestingly, for the side chain methyl ester 2.1.5, the coupling constant JH2'-H3' changed from 2.1 Hz in chloroform to 4.7 Hz in a 1:1 w ater/DMSO-d6. The conformational equilibrium of the methyl ester thus shifted from a gauche to an anti conformer. Similarly, JH2'-H3' for Taxol| also changed from 2.7 (CDC13) to 8 Hz in a mixture of water and DMSO; this points to the dominant contribution of conformers with large torsional angle and indicative of a different side chain conformational ensemble.

Molecular modeling studies found four low-energy conformations

for Taxol| in three of them, the 2' and 3' protons were in g a u c h e arrangement, similar to the crystal structure; the fourth conformer favored the anti arrangement and was the dominant Taxol| conformation in aqueous solution [84]. The Kansas group reported new NMR studies on Taxol| and Taxotere | in support of their "hydrophobic collapse" theory. These studies suggest that water strongly induces the same type of conformation in both Taxol| and Taxotere| the key interaction is the hydrophobic clustering of the 2-benzoyl, 3'-phenyl, and 4-acetyl groups; the N-benzoyl or Boc groups are not participating in organizing the side chain. The NOESY and ROESY experiments on Taxotere | in a 1:1 mixture of methanol/water and Taxol | in DMSO/H20 displayed cross peaks between the aromatic signals on the different rings. Based on the theory that the chemical shift changes of the signals arise from mutual ring-current effects, these new data demonstrate the closeness of these aromatic rings. The strongest interactions were observed between the meta and para protons of the 3'-phenyl and the ortho and meta protons of the C-2 benzoyl group; and furthermore, no cross peaks were observed between t-butyl of the N-Boc and C-2 benzoyl groups

[85]. Since no active Taxol|

analogs are known with deleted 2-benzoyl and 3'-

phenyl groups, it is quite likely that these groups are providing the putative "preorganization" of the side chain conformation most relevant to binding, via hydrophobic clustering rather than intramolecular hydrogen bonding. Structural information on the microtubule binding site and its complex with the drug is needed to understand more about the active conformation of Taxol |

and its tubulin polymerization mechanism.

296 RE~'EtCENCES ~

~

3. 4. 5. ~

o

o

9. 10. 1 1 ~

12. 13. 14. 15. 16.

.

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85.

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The Chemistry and Pharmacology of Taxol and its Derivatives V. Farina, editor 9 1995 Elsevier Science B.V. All rights reserved

301

7 THE BIOCHEMICAL P H A R M A C O L O G Y OF T A X O L | A N D M E C H A N I S M S OF RESISTANCE Lisa M. Landino and Timothy L. Macdonald Department of Chemistry, University of Virginia, Charlottesville, VA 22901, U.S.A.

7.1. INTRODUCTION Microtubules, like DNA and RNA, are conserved structures in all eukaryotic cells with numerous and diverse functions. By forming a scaffolding network within cells, microtubules maintain cell shape. In neurons, microtubules form a track through the axons along which organelles and proteins are transported. In addition, microtubules are the primary components of the mitotic spindle which forms during cell division. All eukaryotic cells utilize a spindle, composed of microtubules, to segregate chromosomes during mitosis [1]. The principal component of microtubules is the protein tubulin, a heterodimer composed of two similar subunits, (~ and ~-tubulin. The molecular weight of each subunit is approximately 50 kDa. Sequence analysis of the major a - a n d ~-tubulin gene products expressed in mammalian cells shows about 40% homology between the two tubulins [2]. The ability to reversibly assemble into microtubules (polymer) and

302 disassemble to the tubulin heterodimer (monomer) is an intrinsic property of t h i s d i m e r i c protein. A l t e r n a t i n g a - a n d ~-tubulins assemble longitudinally into protofilaments which t h e n join l a t e r a l l y w i t h other protofilaments to form the cylindrical microtubule s t r u c t u r e . A typical microtubule is composed of 13 protofilaments with an outer d i a m e t e r of 28 nm and an inner diameter of 14 n m [1] (Figure 1).

Figure 1. Assembly of tubulin monomers into a microtubule structure.

303 In addition, a number of aberrant structures can be formed in vitro including rings and sheets. Purified tubulin can be assembled in vitro at 37~ in the presence of GTP, Mg 2+, and a calcium chelator [3]. Two molecules of GTP bind to each a,~ heterodimer and, upon incorporation into the microtubule polymer, one molecule of GTP is hydrolyzed to GDP and phosphate. In vivo, r e g u l a t i o n of microtubule a s s e m b l y and disassembly is mediated by microtubule-associated proteins (MAPs) and fluctuations in intracellular calcium concentrations [4]. The numerous and diverse functions of microtubules require facile, yet regulated, assembly and disassembly in response to intracellular stimuli. MAPs are thought to be principally associated with tubulin through charge interactions at specific sites along the microtubule polymer and these interactions can be modulated by intracellular signaling events such as MAP phosphorylation [5]. MAPs are a large and diverse family of proteins which co-purify with tubulin through in vitro cycles of assembly and disassembly. Ion exchange chromatography separates the negatively charged tubulin from the more basic MAPs [6]. Only a handful of the most abundant MAPs, such as MAP1, MAP2, and tau, have been characterized in any detail, although many other minor proteins associate with tubulin in vivo. The precise roles of these associated proteins is of considerable interest since the regulation of microtubule function is critical to cell growth and viability. The process of tubulin polymerization to microtubule structures can be dissected into discrete steps. Tubulin polymerization requires a slow nucleation phase, followed by rapid elongation of the microtubule structure, and ultimately an equilibrium p l a t e a u is reached. The f i l a m e n t o u s microtubule structures are in equilibrium with a pool of unpolymerized (z,~ heterodimers both in vivo and in vitro. The concentration of unpolymerized tubulin monomer at this steady state (in equilibrium with polymer) is referred to as the critical concentration (Cc). At tubulin concentrations below the critical concentration, microtubules will not form. A number of factors influence the Cc in vitro, including t e m p e r a t u r e , pH, and the presence of MAPs or drugs such as colchicine or Taxol| [7,8]. In 1981, Carlier and Pantaloni had observed that GTP hydrolysis was not coupled to tubulin polymerization, but that it occurred after monomer incorporation [9]. Their investigations suggested that GTP hydrolysis was

304 not a driving force in the assembly of microtubules, but r a t h e r hydrolysis of GTP-bound t u b u l i n m o n o m e r s in the assembled microtubule induces a conformational

change

in

the

microtubule

structure.

Such

a

conformational role of nucleotide t r i p h o s p h a t e hydrolysis (both ATP and GTP) has emerged as a fairly common p h e n o m e n o n in biological systems [10]. Investigation of the interactions of any tubulin-specific compound with t u b u l i n a n d m i c r o t u b u l e s r e q u i r e s an u n d e r s t a n d i n g of the dynamic properties of microtubules [11,12].

As a dynamic s t r u c t u r e , there is a

constant flux between the monomeric and polymeric forms.

If the rates of

m o n o m e r addition and loss at the microtubule ends are identical, no change in microtubule length at equilibrium is detected. While GTP-bound (z,~ heterodimers are incorporated into the plus (+) or assembly end of the microtubule, GDP-bound tubulin heterodimers are lost from the minus (-) or disassembly end of the microtubule.

This dynamic property of flux or

t r e a d m i l l i n g is observed in vitro using purified tubulin; however, the presence of MAPs can suppress this dynamic behavior [13] (Figure 2).

(+) ASSEMBLY

(-) DISASSEMBLY MICROTUBULE AT EQUILIBRIUM % (~0

Figure 2. Microtubules have distinct assembly (+) and disassembly (-) ends. At equilibrium, the rates of monomer addition and loss are constant and no net change in microtubule length is observed. This dynamic behavior, known as treadmilling or flux, can be suppressed by antimitotic drugs such as vinblastine or Taxol| Another dynamic property of microtubules is dynamic instability [14]. An individual microtubule can undergo t r a n s i t i o n s of l e n g t h e n i n g and s h o r t e n i n g which are observable both in vitro and in vivo.

The growth

305 phase is stabilized by a GTP-cap which exists at high t u b u l i n concentrations, since the rate of monomer addition will transiently exceed the rate of GTP hydrolysis. However, if GTP-bound tubulin concentrations are low, the rate of GTP hydrolysis will exceed the rate of monomer addition. The stabilizing GTP-cap at the assembly end (+) will be lost and this will induce rapid depolymerization. This dynamic property can also be suppressed by the presence of MAPs [15]. Suppression of the dynamic properties of microtubules by antimitotic agents has been suggested as a plausible mechanism of cytotoxicity. A number of in vitro and in vivo studies by J o r d a n and Wilson have d e m o n s t r a t e d t h a t microtubule dynamics can be s u p p r e s s e d by substoichiometric concentrations of vinblastine and Taxol| [16-21]. The results of their studies and the mechanistic implications of their findings are presented in section 7.2. 7.2. THE T U B U L I N / M I C R O T t ~ U I ~

SYSTEM AS A D R U G TARGET

The tubulin/microtubule system has captured the a t t e n t i o n of medicinal chemists since it is the target of a number of synthetic molecules and natural products including colchicine, vinblastine, and Taxol | [1, 7, 8]. The structures of these natural products, as well as the semi-synthetic Taxol| derivative, Taxotere | are shown in Figure 3. The therapeutic utility of compounds which interact with tubulin and microtubules results initially from their ability to disrupt normal spindle function. Such compounds are often called spindle poisons or antimitotic agents. The compounds t h a t bind to tubulin are chemically divergent and there is evidence that they bind to the ~-subunit of the tubulin heterodimer [22, 23]. Colchicine and vinblastine bind to the unpolymerized tubulin monomer at two distinct sites, as demonstrated by competitive binding assays [7, 8]. Monomer binding shifts the monomer-polymer equilibrium toward the depolymerized state. The majority of compounds which bind to tubulin inhibit its polymerization in vitro. The exception is the diterpenoid Taxol| isolated from the bark of the Pacific yew tree, Taxus brevifolia. Taxol| has received much attention due to its antitumor activity, its unique s t r u c t u r e and m e c h a n i s m of action. Unlike colchicine and vinblastine, which inhibit tubulin polymerization, Taxol | stabilizes

306 microtubules and shifts the equilibrium toward the polymerized state. This novel mode of action sets Taxol |

apart and has sparked a renewed

interest in the tubulin/microtubule system as a chemotherapeutic target.

H•O

eee##J

Me0 . ~ ~.~...~ 'NHAc

Ac

MeO

HMeO~N~OH ! Me

~

0 OMe

Vinblastine (E=COOMe)

Colchicine

O

R1"~NH

0

Phil_

R20~, 20H o .... _

OH

0

.o

Taxol, RI=Ph; R2=Ac Taxotere, Rl=t-BuO; R2=H Figure 3. Structures of the most important antimitotic agents The molecular pharmacology of Taxol | has been examined in detail by Horwitz and coworkers [24-28]. These researchers demonstrated that this drug preferentially binds to the microtubule polymer rather than to the tubulin monomer. Taxol | binds reversibly to microtubules with a binding constant of -1 ~M. In addition, the Taxol | binding site is distinct from the sites occupied by GTP, colchicine, and vinblastine.

The microtubules

formed in the presence of Taxol | in vitro are extremely stable and have properties distinct from normal microtubules. Taxol | induces tubulin polymerization in the absence of GTP or MAPs and the microtubules are resistant to depolymerization induced by calcium ion. Taxol |

will induce

307 tubulin polymerization at low temperatures (4 ~ but, in this case, GTP or MAPs are required for maximal polymer formation [24]. A number of in vitro studies have been undertaken to investigate the interaction of Taxol| with tubulin and microtubules at both the kinetic and thermodynamic level. Since tubulin has not been crystallized, the specific molecular i n t e r a c t i o n s of antimitotic agents with the protein have remained elusive. However, a variety of biophysical methods have been employed which have provided much insight into tubulin-drug interactions [29]. In addition, since colchicine, vinblastine, and Taxol| bind at distinct sites, the thermodynamic relationships between these ligand binding sites and their effect on microtubule assembly can be addressed. Timasheff and coworkers have examined the combined effects of Taxol| and colchicine on the thermodynamics of tubulin polymerization in vitro [30]. The stabilizing effect of Taxol | is capable of overriding the destabilizing effect of agents such as colchicine and vinblastine. However, the stabilizing free energy for addition of each a,~ heterodimer gained upon binding of Taxol | is diminished for the tubulin-colchicine complex vs. pure tubulin. The binding of Taxol | provides more t h a n - 3 . 0 kcal/mol of free energy in the polymerization of pure tubulin into microtubules, whereas only -0.5 kcal/mol is gained for Taxol| polymerization of the tubulin-colchicine complex. This reduction in stabilizing free energy implies that energy is expended to overcome unfavorable factors such as sterics or geometric strain in the tubulin-colchicine complex. In addition, Taxol| polymerization of the tubulin-colchicine complex required both h e a t and GTP, whereas pure t u b u l i n would assemble at 10 ~ in the absence of GTP. The processes of tubulin polymerization and GTP binding and hydrolysis are linked under normal polymerization conditions. Essentially, the energy imparted by Taxol | binding to the microtubule decreases the cooperativity of these processes and, as a result, Taxol| tubulin polymerization does not require GTP binding or hydrolysis. In addition, Carlier and Pantaloni showed that Taxol| does not interfere with GTP binding or hydrolysis at the exchangeable GTP binding site [31]. Although Taxol| can induce polymerization of GDP-bound tubulin monomers, it is important to note that, if GTP is present, hydrolysis does occur.

308 Based on this thermodynamic analysis, the action of Taxol |

may be

exerted on the lateral interactions between protofilaments which are normally weak. Differences in the conformation of the tubulin-colchicine complex vs. pure tubulin at these lateral junctions would translate into different l i g a n d b i n d i n g s t r e n g t h s . Careful c o n s i d e r a t i o n of the thermodynamics of tubulin-ligand binding may provide insight into the in v i v o mechanism of these antimitotic agents. The selectivity of Taxol (ii) against particular tumor cell lines may stem from its ability to contribute maximal binding energy under specific physiological conditions. The presence of specific MAPs, for example, may dictate the thermodynamic consequences of Taxol | binding. Some of the most convincing evidence to explain the mechanism of antimitotic agents comes from the work of Jordan and Wilson [15-21, 32-34]. Early and simplistic models to explain the cellular m e c h a n i s m of colchicine and vinblastine suggested t h a t these compounds disrupted cellular function by preventing microtubule formation. Subsequent in vivo studies revealed that, at therapeutically useful concentrations of these depolymerizing agents, microtubules were still formed. In the case of t h e r a p e u t i c a l l y useful levels of Taxol (ii), all cellular tubulin was not polymerized, suggesting that antimitotic agents interact with their cellular target in a more subtle, but intriguing manner. The continuing efforts of these researchers have examined the interactions of antimitotic agents with microtubule ends and surfaces and have revealed how alterations in microtubule dynamics can modulate cellular function and viability. The o b s e r v a t i o n t h a t T a x o l | m i c r o t u b u l e s do not depolymerize significantly when diluted or cooled suggests t h a t the dissociation rate constant of tubulin monomer is decreased by Taxol | binding to the microtubule. Investigation of the dynamics of Taxol | stabilized microtubules has d e m o n s t r a t e d such a reduction in the dissociation rate constant of tubulin from microtubules [32]. The two dynamic properties of microtubules which have been investigated most thoroughly are dynamic instability and treadmilling. The dynamic behavior known as dynamic instability, in which an individual microtubule undergoes periods of growing and shortening, is not as well understood as treadmilling. However, on a simplistic level, the importance of such a behavior during mitosis and chromosomal segregation can be appreciated.

309 Individual

microtubules

must

grow

and

shorten

in

response

to

intracellular stimuli in order for cell division to occur. The dynamic instability of microtubules is influenced by the presence of microtubuleassociated proteins (MAPs). Although MAPs will suppress this behavior in vitro, it can still occur to a detectable extent in bovine brain microtubules, which contain approximately 70% tubulin and 30% MAPs [20].

Like

dynamic instability, treadmilling is also suppressed by MAPs [21]. The net addition of GTP-bound monomers and loss of GDP-bound monomers at the microtubule ends occurs at the polymer steady state. This equilibrium phenomenon has been demonstrated in vitro; in addition, mitotic spindle microtubules, which are very dynamic st ruct ures, display the same behavior. Agents such as v i n b l a s t i n e

and colchicine i n h i b i t microtubule

assembly at substoichometric concentrations in vitro. vinblastine

to d i s r u p t

microtubule

formation

at

The ability of

a ratio

of 300:1

(tubulin:vinblastine) supports a mechanism whereby the drug binds to the ends of microtubules, not to soluble tubulin monomer [33]. Wilson et al. have also d e m o n s t r a t e d t h a t tubulin exchange at microtubule ends is inhibited by 50% when only -1.2 molecules of vinblastine are bound per microtubule [34]. Wilson and coworkers propose t h a t s u p p r e s s i o n of microtubule dynamics at low concentrations of vinblastine correlates with mitotic block at the m e t a p h a s e / a n a p h a s e boundary.

In addition to suppression of

microtubule dynamics, vinblastine also enhances the length of time that a microtubule spends in an a t t e n u a t e d state in which no growing or shortening of the microtubule is detected. Notably, Wilson's laboratory has demonstrated t h a t low vinblastine concentrations did not prevent mitotic spindle formation, but still blocked HeLa cells at metaphase.

The mitotic

spindle was fully formed in the proper bipolar configuration; however, the alterations induced by low vinblastine concentrations were much more subtle

as

determined

by

immunofluorescence

microscopy.

These

alterations were sufficient to prevent the transition from metaphase to anaphase.

The overall structure of the microtubules and the kinetochore

was not affected by the drug, but the number of microtubule-kinetochore attachments was decreased. Microtubule-kinetochore attachment is vital to chromosomal segregation during the later stages of mitosis.

310 The concentrations

of vinblastine

which induced

these

subtle

alterations in the mitotic spindle, without inducing depolymerization of the microtubules, were in the low nanomolar range (0.1 - 6 nM). Following incubation with 2 nM vinblastine for 18-20 hours, no HeLa cells were detected in anaphase for 2 - 48 hours after treatment with the drug. This indicates that mitotic block at the metaphase/anaphase boundary is not transient. In addition, a substantial percentage (31-38%) of cells in interphase treated with low nanomolar Taxol| concentrations (3-10 nM) had multiple nuclei. Jordan et al. examined the mechanism of mitotic block and inhibition of cell proliferation by low concentrations of Taxol| [16]. Despite their seemingly opposite mechanisms of action, Taxol| and vinblastine demonstrate nearly identical effects on the mitotic spindle in HeLa cells. Like vinblastine, Taxol|

induced mitotic block at the metaphase/anaphase

boundary. The concentration of Taxol| which induced 50% mitotic block in HeLa cells was only 8 nM. This concentration of Taxol | which resulted in significant mitotic arrest did not increase microtubule mass in vivo as had been anticipated for this microtubule-stabilizing agent. The effect of low Taxol | concentrations on mitotic spindle organization was also determined. Immunofluorescence microscopy revealed t h a t spindles which had been treated with nanomolar levels of Taxol| were nearly identical to spindles formed in the presence of low concentrations of other antimitotic agents including vinblastine.

Astral microtubules were more

obvious in Taxol| cells when compared to the control cells. This is in agreement with previous reports of Taxol| aster formation [1719]. In addition, the distance between the spindle poles at opposite ends of the cell was significantly shorter for the Taxol| cells relative to the control cells (4.0 +/- 0.4 ~m vs. 7.4 +/- 0.2 ~m). As the concentration of Taxol| increased, alterations in spindle formation became more pronounced. The chromosomes, which are normally aligned at the metaphase plate, were attached to microtubules which extended from the centriole like the spokes of a wheel. The normal bipolar configuration, with the two centrioles at opposite ends of the cell, was not observed in HeLa cells treated with micromolar concentrations of Taxol|

Some chromosomes were attached to astral microtubules which

also extended from the centriole.

311 At Taxol |

concentrations above 10 nM, an increase in the mass of

microtubule polymer was detected.

Maximal enhancement of microtubule

polymer levels was achieved at 330 nM Taxol | polymer mass.

In addition, at high Taxol |

with a 500% increase in

concentrations, microtubule

bundles were observed. Some loose bundles were detected in a few cells incubated with 33 nM Taxol| however, 1 pM Taxol| induced formation of massive microtubule bundles.

The effects of Taxol|

on HeLa cells at

concentrations above 10 nM, such as induction of microtubule bundles and increased polymer mass, are not surprising and have been previously reported for a number of cell lines. The most notable effects of Taxol | reported by Jordan et al. are those that occur at low drug concentrations [16]. In addition to investigating the effects of nanomolar levels of Taxol | on HeLa cells, the effects on microtubule dynamics in vitro were examined. Video-enhanced differential interference c o n t r a s t microscopy was employed to assess the dynamic instability of Taxol| microtubules assembled from pure bovine brain tubulin. In the presence of 0.1 and 0.5 ~M Taxol| both the microtubule growing and shortening rates were affected. These rates were inhibited by approximately 50% with 0.1 ~M Taxol (ii). In addition, the exchange of tubulin subunits at microtubule ends was inhibited substantially at 0.1 llM. This exchange process is defined as microtubule dynamicity and is defined as the number of tubulin dimers exchanged per second. Microtubule dynamicity was inhibited by 70 % with 0.5 ~tM Taxol| Taxol (i!) also influenced the length of time that a microtubule spent in an attenuated state.

The attenuated state defines that period in which a

microtubule is neither growing nor shortening, i.e. a static state. It is important to note that these concentrations of Taxol | 0.1 and 0.5 ~M, did not induce any detectable increases in microtubule polymer mass. Suppression of microtubule dynamics by antimitotic agents is an attractive mechanism for the biochemist/medicinal chemist. Although the primary effect of these compounds in vitro is thermodynamic (shifting of the monomer-polymer equilibrium), the in vivo effect which ultimately may prevent mitosis appears kinetic in nature.

Through subtle alterations in

the rates of monomer gain and loss at the microtubule ends, chromosomal segregation is disrupted.

312 7.3. THE BASIS F O R THE THERAPEUTIC EFFECTS OF TAXOL | One of the p r i m a r y goals of this review is to explore how the biochemical effects of Taxol | eventual cellular consequences. Taxol |

on tubulin and microtubules relate to its Although the primary cellular target of

appears to be t u b u l i n and microtubules, the in vivo steps

subsequent to Taxol |

binding are ill-defined.

In addition, Bhalla and co-

workers have d e m o n s t r a t e d t h a t the ul t i m at e consequence of Taxol | t r e a t m e n t in a leukemic cell line is apoptosis [35]. directed, energy-dependent process of cell death.

Apoptosis is a geneOne h a l l m a r k of this

process is fragmentation of cellular DNA into multiples of 200-base pair units by a Ca 2+ and M g 2 + - d e p e n d e n t endonuclease. characteristic

morphological

changes

are

also

A n u m b e r of

observed,

including

chromatin condensation, nuclear disintegration, and the formation of apoptotic cell bodies [36-39]. A number of potent and very effective chemotherapeutic drugs such as cytosine arabinoside, etoposide, cisplatin and doxorubicin have been shown to induce leukemic cell death by this apoptotic pathway.

In addition,

microtubule-selective agents such as colchicine and vinblastine have also been shown to induce apoptotic cell death in leukemic cells. Notably, the primary cellular targets of these drugs can vary, yet the same cellular markers of apoptosis are observed in leukemic cells t reat ed with these drugs.

This suggests t h a t interaction with the pri m ary target perturbs

cellular function in such a m a n n e r t hat a common signaling mechanism initiates cell death. The influence of antimitotic agents on microtubule dynamics described above attempts to rationalize how cells are sensitive to a diverse group of molecules. and Taxol |

Despite their seemingly opposite modes of action, vinblastine share a n u m b e r of common effects on microtubules, as

demonstrated by the work of Jordan and Wilson [15-21, 32-34]. However, their findings provide no clues about the selectivity of Taxol |

against a

select group of tumor types. Why does Taxol |

exhibit selective cytotoxicity toward some cancer

cells? The available data show that Taxol | has a narrow therapeutic index with significant efficacy against ovarian and breast tumors. which will ultimately define the selectivity of Taxol |

The factors

may be numerous.

313 Since all eukaryotic cells have intact tubulin]microtubule systems, the sensitivity of specific cell types to Taxol| must be influenced by factors which may be distinct from the tubulin/microtubule system. Table 1. Observed Effects of Taxol |

Biochemical Effects of Taxol|

References

Shifts monomer-polymer equilibrium toward polymeric state (affects thermodynamics of tubulin polymerization)

24-26

Affects the kinetics of monomer addition and loss at microtubule ends (microtubule dynamics)

15-21

Alters cofactor requirements (GTP, Mg 2+, etc. ) for tubulin polymerization

9, 31

Stabilizes microtubule against depolymerization induced by Ca 2+, dilution, or low temperatures

24-28

Cellular Effects of Taxol| Induction of microtubule bundling in leukemic cell line

70-75

Induction of mitotic arrest at the G2/M interface

24, 70

Induction of apoptosis in Taxol|

leukemic cells

Expression of multidrug transporter in resistant cells T axol| mimics the effects of LPS in murine macrophages

35 80-84 103-108

The tubulin-microtubule system is carefully regulated by a variety of physiological modulators, and it is likely that these endogenous regulators

314 will exert a significant influence on the effects of intracellular Taxol | Moreover, because the presence and nature of many modulating agents varies according to cell type and stage of development, these factors may contribute to the selectivity of Taxol (!i) in cellular targeting. The following sections will describe the effects of Taxol (ii) on the tubulin/microtubule system that have been identified at the biochemical and cellular levels. 7.4. C E L L U L A R

TUBULIN

CONCENTRATIONS

AND

ISOTYPE

COMPOSITION INFLUENCE TAXOL | SENSITIVITY The microtubules which form the mitotic spindle and the cytoskeleton within cells vary in their stability, ease of formation, and dynamic properties. The constant flux between soluble tubulin monomers and polymer is a fundamental property of microtubules which dictates the functional activities and diversity of microtubules. Cabral et al. have demonstrated that the levels of polymerized tubulin within cells correlates with drug resistance to antimitotic agents [40]. The antimitotic agents discussed thus far have all been shown to interact with microtubule ends and influence microtubule dynamics. Subtle changes in microtubule stability ( a d j u s t m e n t s in the monomer/polymer equilibrium) can influence both sensitivity and resistance to antimitotic agents. In an attempt to elucidate a novel mechanism of drug resistance (see section 7.7) the levels of polymerized tubulin in different drug-sensitive and drug-resistant cells were measured. In these experiments, Cabral et al. found t h a t CHO cells which were resistant to Taxol | but not colchicine or vinblastine, contained lower levels of polymerized tubulin ( - 2 0 - 3 0 % ) w h e n compared to the wild-type cells (-40%). These researchers suggested that the addition of Taxol | to the Taxol(ii)-resistant cells would enhance the cellular level of polymerized tubulin to near the wild-type level of polymer, but the increase in polymer mass would not be sufficient to affect viability. Conversely, CHO cells which were r e s i s t a n t to microtubule-depolymerizing agents, such as colchicine or vinblastine,

contained higher levels of polymerized tubulin

(-50%). In addition to explaining a novel mechanism of drug resistance, this hypothesis presented by Cabral and coworkers may provide a simple

315 indicator of cell sensitivity to Taxol | [40]. A cell with normally high levels of polymerized tubulin may be more sensitive to Taxol | than a cell with a lower level of polymer.

Investigation of cellular polymer levels in a wide

variety of cell types could be very useful in determining sensitivity to Taxol| In most eukaryotes, but particularly higher vertebrates, both r and [3tubulin exhibit considerable polymorphism. Both subunits are encoded by small multigene families and both are subject to post-translational modifications, including the addition and subsequent removal of the Cterminal tyrosine of a-tubulin [41,42] the acetylation of r [43-45], the polyglutamylation of a- and ~-tubulin [46,47], and the phosphorylation of ~-tubulin [48-51]. Additionally, the expression of different tubulin genes is differentially regulated during development and is in part either tissueor cell specific [52-54]. The various gene products are referred to as isotypes; the v a r i a n t s arising by post-translational modification are referred to as isoforms. A fundamental, unresolved issue is the extent to which the structural and functional characteristics of different microtubule populations are determined by differences in the primary sequence of tubulin isotypes. The evolutionary conservation across species of distinct isotypes, as well as the differential expression of different members of tubulin gene families, indicates that at least some isotypes may be functionally specialized with distinct biochemical functions in vivo. This proposal is known as "the multitubulin hypothesis" [55-57]. A closely related issue is the role of posttranslational modification in the function of tubulin variants. The size of the tubulin gene families can range from one to two a- and ~- tubulin genes for fungi to five to seven a- and ~-tubulin genes for higher vertebrates [58]. Mammalian brain is the most frequently used source of tubulin for structural and kinetic studies. At least five r and five ~-tubulin isotypes are expressed in the adult mammalian brain, which suggests that as few as five and as many as 25 structurally distinct a,~dimers may exist in the brain [56]. The post-translational modifications of these isotypes increase this number further. The precise functional roles of isotype/isoform species is not presently dear. Recent studies by Luduena and coworkers suggest that ~-tubulin isotype composition has an effect on the in vitro assembly of brain tubulin

316 [59]. Both the maximal rate and the extent of polymerization increases when bovine brain tubulin depleted of class III ~-tubulin by immunoaffinity chromatography is stimulated to assemble by either MAP2 or tau. Similarly, whereas very little is known about the role of tubulin posttranslational modification, recent studies by Frankfurter and co-workers indicate that the charge heterogeneity of rat brain tubulin affects in vitro MAP-stimulated assembly [60]. They have d e m o n s t r a t e d t h a t polyglutamylation of rat brain class III ~-tubulin increases with development [61] and, more recently, that the critical concentrations of tubulin required for assembly for postnatal day 10 and newborn MAP-free tubulin were 2-fold and 3-fold higher, respectively, than for adult tubulin. Moreover, the MAPstimulated assembly of adult tubulin exhibited biphasic kinetics (indicative of at least two tubulin populations), whereas postnatal day 10 and newborn tubulin assembled with monophasic kinetics [61]. With the sole exception of the acetylation of a-tubulin at lysine 40, the remaining known post-translational modifications of tubulin occur within the C-terminal isotype-defining domain, approximately the last 15 residues. What is compelling about this observation is that the isotype-defining Cterminal domain is believed to contain the binding sites for MAP2 and Ca 2+ [62]. Moreover, both polyglutamylation and phosphorylation of ~-tubulin occur in this region and are developmentally regulated [47]. Consequently, these p o s t - t r a n s l a t i o n a l modifications may significantly alter the interaction of tubulin with a number of ligands during neural development. For example, the ionic interactions which control the association of MAPs with tubulin are localized to this C-terminal domain [63]. The further acidification of this already negatively charged region might strengthen MAP binding and serve, in part, to increase microtubule stability. Present evidence is consistent with isotypes and isoforms as playing important roles in the kinetics and thermodynamics of tubulin assembly. The underlying chemotherapeutic utility of tubulin-specific agents may result from their ability to selectively bind to and stabilize a discrete subpopulation of tubulins. Class III ~-tubulin has been shown to be responsible for the slow phase of colchicine binding. Selective removal of class III ~-tubulin and its associated r from MAP-free tubulin alters the kinetics of colchicine binding from biphasic to monophasic [64]. In addition, the slow phase of colchicine binding was not observed for class

317 III-depleted tubulin. Microtubule depolymerization induced by colchicine selectively depolymerizes class III ~-tubulin first. Recently, Lu and Luduena reported that the removal of class III ~tubulin and its associated r enhanced Taxol| microtubule assembly [65]. The Class III-depleted tubulin assembled more rapidly and to a greater extent than unfractionated tubulin in the presence of Taxol | The critical concentration (Cc) of tubulin required for assembly in the presence of 10 ~M Taxol| was only 0.16 mg/ml for the Class IIIdepleted tubulin as compared to 0.4 mg/ml for the unfractionated tubulin. Their studies provide further evidence that tubulin isotype composition can greatly influence the kinetics of tubulin assembly. It is intriguing to hypothesize that a cell could modulate its sensitivity to antimitotic agents through alterations in its isotype composition. In our laboratory, we investigated the isotype-selectivity of Taxol| induced tubulin polymerization. Our goal was to assess the contribution of ~-tubulin isotype composition and charge heterogeneity on the kinetics of Taxol| microtubule assembly. Analysis of tubulin charge heterogeneity was achieved by high resolution isoelectric focusing (IEF). Microtubules, formed from MAP-free tubulin with varying concentrations of Taxol| were subjected to isoelectric focusing and immunostaining with isotype-specific antibodies to assess any variations in isotypes polymerized. These studies showed that Taxol| did not exhibit any selectivity under the experimental conditions employed (unpublished results). Consistent with this observation, photoaffinity labeled analogs of Taxol | have been employed to identify the binding site on tubulin. Using a 3H-labeled 3'-(p-azidobenzamido)taxol, Horwitz and co-workers recently localized the Taxol| binding site to the N-terminal 31 amino acids of ~tubulin [66]. The N-terminal region of the tubulins, unlike the C-terminal region, is much more conserved across isotypes. Consequently, Taxol | binding to this N-terminal domain would not be expected to show any isotype selectivity. 7.5. TAXOL| :INDUCES MICROTUBUI~ BUNDLING The cylindrical structure of microtubules results in a fairly rigid structure with limited conformational motility. In vivo, individual

318 microtubules, composed of tubulin, are decorated with microtubuleassociated proteins (MAPs). Much evidence suggests that MAPs extend from the surface of the microtubule and influence interactions between adjacent microtubules. Longitudinally-associated microtubule bundles have been observed in a number of cell types. The formation of any unusual or aberrant microtubule structure which may suggest a plausible mechanism of action of antimitotic agents is of interest. The interplay of cellular proteins and the mitotic machinery may provide the most compelling clues about the cellular mechanisms of these agents. Taxol| has been shown to influence the number of protofilaments within a microtubule structure in vivo [67]. Whereas the majority of animal and p l a n t cells construct m i c r o t u b u l e s c o n t a i n i n g 13 protofilaments, developing wings of Drosophilia construct 15 protofilament microtubules [68]. Taxol| treatment of cultured wings induced formation of microtubules containing 12 protofilaments at microtubule-nucleating sites. Dye et al. have d e m o n s t r a t e d by video-enhanced differential interference contrast microscopy t h a t Taxol| microtubules, assembled in vitro from pure tubulin, became flexible and appeared wavy [69]. Notably, the addition of MAP2 or tau, two major MAPs, reversed this Taxol| flexibility. The addition of these MAPs did not cause Taxol| to dissociate from the microtubule, since flexibility was again observed when the MAPs were released from the microtubules by high salt concentrations. This alteration in the flexibility of microtubules is likely to result from slippage between adjacent protofilaments in the cylindrical microtubule structure. A decrease in the strength of the lateral or circumferential interactions induced by Taxol| would allow bending of the microtubule structure. Reversal of this phenomenon by MAPs may result from the ability of these proteins to bridge adjacent protofilaments and restore the strength of the lateral protofilament interactions. This observation of Taxol| microtubule flexibility and its reversal by MAP2 and tau may have relevant consequences in vivo. Although MAPs are always associated with microtubules in vivo, their distribution and levels of expression vary in different cell types. Thus, the microtubules in Taxol| cells may have enhanced flexibility, which

319 would affect their ability to perform crucial cellular functions requiring rigid microtubule structures. In fact, alterations in microtubule flexibility may explain the ability of Taxol| to induce microtubule bundling i n vivo. The formation of microtubule bundles in Taxol| cells has been observed. Microtubule bundles can be described as abnormal microtubule arrays composed of multiple microtubules which associate with each other longitudinally. Rowinsky and coworkers have d e m o n s t r a t e d t h a t the stability of these Taxol| microtubule bundles correlates well with cell sensitivity to Taxol| [70-75]. Taxol | t r e a t m e n t of cultured leukemic cells has been shown to induce mitotic arrest at the G2/M interface of the cell cycle. However, the formation of microtubule bundles in these cell lines was observed t h r o u g h o u t the cell cycle. Taxol| t r e a t m e n t induced microtubule bundling in sensitive leukemic cell lines such as HL-60 and LC8A, and also in the relatively resistant cell lines K562 and Daudi [70]. Microtubule bundling in the resistant cell lines was reversible both in the presence and absence of Taxol| and these cells accumulated in G2/M. Conversely, the microtubule bundles formed in the sensitive cell lines appeared to be irreversible and persisted in the presence and absence of Taxol| Thus, the stability, not simply the formation, of the microtubule bundles correlates with the cytotoxicity of Taxol | Another observed microtubule change in these T a x o l | leukemic cell lines was the formation of multiple aster-like aggregates of short microtubules [71-75]. Asters, also composed of microtubules, radiate from the centrioles in a star-like conformation during mitosis. The role of astral microtubules in mitosis is not well understood. The formation of multiple asters and microtubule bundles were independent events and did not occur simultaneously. In addition, the formation of these multiple "asters" did not correlate well with sensitivity to Taxol| Subsequently, these researchers demonstrated the cell cycle phase of the Taxol| formation of multiple asters and microtubule bundles [71]. The formation of Taxol| multiple asters occurred during mitosis, whereas cells containing microtubule bundles were in G0/G1, S, and G2 phases of the cell cycle. Both a Taxol| and a Taxol| resistant cell line displayed Taxol| asters only during mitosis. In addition, there were no notable variations in the cell cycle positions for microtubule bundle formation in the Taxol|

vs. the Taxol |

320 resistant cell lines. Hence, the sensitivity of a cell line to Taxol | does not correlate with microtubule bundle formation during a particular phase of the cell cycle or with the ability of the Taxol| cell to synthesize DNA (S-phase specific). The majority of studies have demonstrated that the cytotoxicity of the V i n c a alkaloids (such as vinblastine) and other tubulin-specific agents is due to their effect on microtubules, specifically the mitotic spindle. However, recent evidence suggests that the cytotoxicity of these compounds may result from effects on cells during interphase [72, 73]. Roberts et al. compared the effect of Taxol | on S-phase activity (DNA synthesis) in both a Taxol| (HL60) and a T a x o l | leukemic cell line (K562) [74]. Although the labeling of DNA with [3H]thymidine was scarcely affected in these two cell lines (a measure of Sphase activity), the formation of a high percentage of polyploid cells in the Taxol| cell line, K562, was observed. Following 24 hour t r e a t m e n t with Taxol | 66% of the Taxol| K562 cells contained multiple numbers of chromosomes (polyploidization), whereas only 8% of the Taxol| HL60 cells were polyploid. The induction of polyploidism in Taxol| cell lines relative to Taxol| cell lines suggests t h a t this may be a useful indicator of drug sensitivity. Resistance of a cell line to Taxol | may be predicted based on its ability to induce such a distinct alteration in chromosome numbers. Thus, Rowinsky and coworkers have demonstrated that the formation of persistent microtubule bundles correlates with Taxol | sensitivity in leukemic cell lines, although the formation of these persistent structures is not cell-cycle specific. In addition to direct effects on microtubules, Taxol | treatment induced polyploidism to a great extent only in Taxol| leukemic cell lines. These observations provide researchers with clear cellular markers to assess sensitivity to Taxol | [70-75]. The formation of stable microtubule bundles in Taxol| cells suggests that these structures play a crucial role in mediating the cytotoxic activity of Taxol | Therefore, the biochemical mechanisms of bundle formation and the consequences of this event have been investigated. Turner and Margolis have examined the bundling of Taxol| microtubules in vitro [76]. Their study examined the effect of ATP on bundle formation and also identified a protein factor which induced bundle

321 formation in vitro. The protein factor which was required for in vitro bundling did not copurify with tubulin through cycles of assembly and disassembly, which indicates that it is not a microtubule-associated protein (MAP). However, Cowan and coworkers have reported t h a t microtubule bundling into parallel arrays is induced by MAP2 and tau, both of which are MAPs [77]. Crosslinking of adjacent microtubules is achieved by a short C-terminal sequence present in both MAP2 and tau. This a-helical domain is distinct from the microtubule-binding site which serves to nucleate microtubule assembly. Construction of a n u m b e r of m u t a n t MAP2 proteins has allowed the domain which influences bundle formation to be identifed [78]. The microtubule bundles formed in the presence of Taxol| which were observed by T u r n e r and Margolis were sensitive to high c o n c e n t r a t i o n s of ATP, which s u g g e s t s a p h o s p h o r y l a t i o n - l i n k e d mechanism. In addition, elevated Ca 2+ concentrations (millimolar) relaxed Taxol| microtubule bundles and released s u b s t a n t i a l amounts of MAPs from these structures. Relaxation of Taxol| bundles by high concentrations of Ca 2+ and ATP implicates signal transduction pathways in the regulation of this drug-induced event. U m e y a m a and coworkers examined the dynamics of microtubules which were induced to bundle by MAP2C in vivo [79]. Both the incorporation of biotin-labeled exogenous tubulin into the microtubule ends and the turnover rate of microtubule bundles were investigated in this study. Incorporation of the biotin-labeled tubulin into preexisting microtubule ends was fairly rapid. However, the microtubule turnover rate, determined by photoactivation of caged fluorescein-labeled tubulin, was dramatically reduced. Normally there is a constant flux between the monomer and polymer pools of tubulin within cells. The abnormal microtubule bundling array induced by MAP2C in this study prevented this normally facile flux. The observation t h a t microtubule bundles have altered dynamic properties is not surprising. Although, in this case, the bundles were induced by MAP2C, cellular microtubule bundles formed in the presence of Taxol|

would be anticipated to exhibit similar losses in dynamics.

These

322 data lend further support to the hypothesis that suppression of microtubule dynamics by antimitotic agents is the mechanism of cytotoxicity.

7.6. MECHANISMS OF RESISTANCE TO TAXOL| R e s i s t a n c e to Taxol| h a s s u b s t a n t i a l l y l i m i t e d its clinical development. Due to its hydrophobic nature, Taxol| induces overexpression of the m u l t i d r u g t r a n s p o r t e r , P-glycoprotein [80, 81]. Pglycoprotein is a membrane-spanning glycoprotein which is found in most normal cells in low amounts, but its gene (mdr) is amplified in drugresistant cells. Enhanced levels of this protein increase the cell's ability to remove a c c u m u l a t e d hydrophobic drugs. The compounds which are extruded by this ATP-driven pump are diverse and affect a n u m b e r of different i n t r a c e l l u l a r targets in addition to the tubulin/microtubule system, such as DNA topoisomerase II. Roy and Horwitz first identified a phosphoglycoprotein associated with Taxol | resistance in J774.2 cells, a murine macrophage-like cell line [82]. Drug resistance was induced by growing the cells in the presence of increasing concentrations of Taxol| Cross-resistance to microtubule-specific agents, such as colchicine and vinblastine as well as compounds such as doxorubicin and actinomycin D, was observed. Analysis of membrane proteins in the Taxol| and wild-type cells by SDS-PAGE revealed a prominent protein band at 135 kDa in the Taxol| cells which was scarcely detected in the wild-type cells. The presence of this protein band correlated well with drug resistance to Taxol| In addition, colchicine- and vinblastine-resistant J774.2 cells displayed prominent phosphoglycoprotein bands at 145 and 150 kDa, respectively. Thus, the phosphoglycoprotein of the Taxol| cell lines is similar, but not identical, to the classic P-glycoprotein expressed in vinblastine- and colchicine-resistant cell lines. The full-length cDNA of the m d r l gene encodes a 170 kDa P-glycoprotein. The detection of these smaller membrane phosphoproteins in drug-resistant cell lines suggests that a number of heterogeneous drug transporters exist. Greenberger and co-workers have examined the biosynthesis of the heterogeneous forms of glycoproteins which are expressed in drug resistant J774.2 cells [83, 84].

The heterogeneity in vinblastine- and

323 colchicine-resistant cells lines was attributed to variations in glycosylation which resulted in altered electrophoretic mobility. In these vinblastineand colchicine- resistant cells, a single precursor (125-kDa) was rapidly processed to two forms of 135- and 140kDa. However, in Taxol| J774.2 cells, two distinct precursor proteins were expressed [83]. This observation demonstrated that different glycoproteins which mediate drug resistance could be expressed in response to different drugs. In order to f u r t h e r investigate this heterogeneity, the forms of P-glycoprotein expressed in independent cell lines were examined [84]. The results of this study indicated t h a t expression of the precursor forms of the d r u g - r e s i s t a n c e - a s s o c i a t e d proteins was not drug specific when examined over a range of cell types. A number of approaches have been employed in order to overcome Pglycoprotein-mediated Taxol| resistance. The sensitivity of resistant cells to Taxol| was restored when Taxol | was administered in conjunction with agents such as the cyclosporine derivative SDZ PSC 833 and the cyclopeptolide SDZ 280-446 [85]. Combination drug t h e r a p y may be necessary to overcome r e s i s t a n c e to Taxol | C u r r e n t l y Taxol | is administered with cremophor-EL, which enhances solubility and also drug accumulation intracellularly [86]. Recently, Taxol| has been administered in drug carriers such as liposomes. The interactions of Taxol | with lipids have been studied as well. The investigation of the interactions of Taxol | with dipalmitoyl phosphatidylcholine (DPPC) liposomes by a number of physical methods demonstrated t h a t the drug can partition into the membrane bilayer and perturb the m e m b r a n e [87]. Membrane fluidity and the lipid order p a r a m e t e r were affected. These findings are clinically relevant, since cellular uptake is a prerequisite for effective drug/target interactions. Bhalla and coworkers had previously demonstrated that t r e a t m e n t of leukemic cell lines with clinically relevant levels of Taxol| induced the c h a r a c t e r i s t i c morphological changes associated with apoptosis or programmed cell death [35]. In that earlier study, Bhalla and coworkers had also d e m o n s t r a t e d t h a t Taxol| t r e a t m e n t altered the levels of expression of two oncogene products, bcl-2 and c-myc. Bcl-2 has been d e m o n s t r a t e d to block apoptotic cell d e a t h t h r o u g h an ill-defined a n t i o x i d a n t p a t h w a y [88] and cellular levels of this mitochondrial

324 membrane protein appear to correlate with resistance to antitumor agents. Recently, Miyashita and Reed demonstrated that elevated cellular levels of the bcl-2 oncoprotein could block apoptosis induced by chemotherapeutic agents in a h u m a n leukemia cell line [89]. In addition, leukemic cells which are arrested at the G2/M transition of the cell cycle, such as those treated with Taxol| typically undergo apoptosis. In an attempt to further elucidate cellular mechanisms of resistance to Taxol| as well as other antitumor agents, Bhalla and coworkers have characterized a Taxol| h u m a n myeloid leukemia cell line, HL-60 [90]. Given previous results, these researchers examined the levels of expression of the multidrug transporter, P-glycoprotein, as well as the levels of bcl-2. These Taxol| cells overexpressed P-glycoprotein and, as a result, exhibited cross-resistance to other drugs including vincristine and doxorubicin. In addition, Taxol | t r e a t m e n t did not induce microtubule bundle formation in these resistant cells and the cells did not undergo apoptosis. Examination of the cellular levels of bcl-2 by Western blot analysis revealed t h a t the Taxol| cells did not have significantly elevated levels of bcl-2 relative to the Taxol| cells. This suggests t h a t Taxol | resistance in this cell line is not due to overexpression of bcl-2. The absence of microtubule bundles in the Taxol| resistant HL-60 cells is consistent with the observations of Rowinsky and coworkers, who have d e m o n s t r a t e d t h a t the presence of p e r s i s t e n t microtubule bundles correlates with sensitivity to Taxol| [70-75]. Although the levels of bcl-2 in the Taxol| leukemic cell line were not elevated, the role of bcl-2 in cellular resistance to antitumor agents is of considerable interest. Most recently, Willingham and Bhalla used an anti-bcl-2 monoclonal antibody and fluorescence immunocytochemistry to localize the bcl-2 protein during mitosis in h u m a n carcinoma cell lines, KB and OVCAR-3 [91]. In both these cell lines, interphase cells showed no detectable expression of bcl-2. However, cells undergoing mitosis displayed localized bcl-2 protein around condensed chromosomes. This pattern of bcl2 localization was observed in prophase, metaphase and anaphase in the two cell lines studied. Cells in telophase no longer contained detectable bcl2 protein. Taxol | treatment of these cell lines did not alter the distribution of bcl2 around the condensed chromosomes during mitosis, but mitotic arrest

325 was observed, and with continued Taxol (!i) treatment the cells showed the characteristic morphological changes associated with apoptosis. In additon, bcl-2 localization was lost. Previously, Taxol | had been demonstrated to induce apoptosis in leukemic cell lines [35]. These results of apoptosis induced by Taxol | in human carcinoma cell lines support a common mechanistic outcome of Taxol |

treatment in mammalian cells.

The relevance of these bcl-2 localization studies during mitosis is considerable. The observation that bcl-2 is expressed and associates with chromosomes at the initiation of mitosis, and that the protein disappears in telophase, suggests a protective role during the time the chromosomes are accessible to the cytoplasm.

Bcl-2 may protect the chromosomes from

degradation by the "apoptotic" nuclease at internucleosomal sites. If this were the case, overepression of bcl-2 could prevent apoptosis by preventing DNA damage. Since treatment with Taxol | induces mitotic arrest in these cells, the length of time the chromosomes are exposed in the cytoplasm is altered and the protective action of bcl-2 may be affected.

The continued

investigation of the role of bcl-2 in mitosis and apoptosis may provide important insights into the mechanism of antimitotic agents such as Taxol (ii). The levels of bcl-2 during mitosis in different cell lines could help address the issue of Taxol | selectivity against a particular tumor cell line. A novel mechanism of resistance to microtubule-specific agents has been observed by Cabral and co-workers [40, 92-98]. Random mutagenesis by ultraviolet irradiation was employed to generate drug-resistant CHO cell lines. Early investigation of these drug-resistant mutants produced by random mutagenesis demonstrated that resistance was not the result of amplification of P-glycoprotein or defects in drug accumulation. These drug-resistant CHO cell mutants were found to have altered a - a n d 13tubulins, as determined by two-dimensional electrophoresis [98]. However, the mutant tubulin subunits did not have altered drug-binding properties, which would be the simple hypothesis to explain the observed drug resistance.

Nonetheless, a direct effect on microtubules is supported by

evidence that cells resistant to a destabilizing drug such as colcemid are cross-resistant to other destabilizing agents even though their binding sites on tubulin are distinct.

In addition, cells r e s i s t a n t to microtubule-

destabilizing agents are hypersensitive to Taxol | agent.

a microtubule stabilizing

326 Random mutagenesis of CHO cells by ultraviolet radiation produced a n u m b e r of m u t a n t s which were r e s i s t a n t to Taxol | a microtubulestabilizing agent, and colcemid, a microtubule-destabilizing agent. In addition, a m u t a n t which required Taxol| for survival was also identified [96, 97]. In the absence of Taxol| the CHO cell m u t a n t which required Taxol| for survival initially accumulated in G 2 ~ , and did not undergo cell division (cytokinesis), but f u r t h e r increased its DNA content (became polyploid). This indicates t h a t the mutation does not halt progression through the cell cycle, but it prevents cytokinesis, the ultimate splitting of the parent cell into two daughter cells. The Taxol| m u t a n t did not form a functional mitotic spindle apparatus in the absence of Taxol | as d e t e r m i n e d by i m m u n o f l u o r e s c e n c e studies, a l t h o u g h non-spindle cytoplasmic microtubules were formed. The spindle microtubules which did form in the absence of Taxol| were shorter, fewer in number, and were kinetochore microtubules only. The mitotic spindle contains two forms of microtubules: the kinetochore microtubules, which radiate from the centrioles and form the contact site of microtubule a t t a c h m e n t to the chromosome at the centromere, and interpolar microtubules, which interdigitate between the chromosomes which are aligned at the metaphase plate [99]. In a d d i t i o n , i n t e r p o l a r microtubules known as asters extend from the centrioles in a star-like conformation. The kinetochore microtubules are more stable t h a n the interpolar microtubules and they are believed to be responsible for the chromosomal segregation which begins during anaphase [100, 101]. Initial studies of the Taxol| CHO cell m u t a n t suggested that the cells contained lower intracellular concentrations of tubulin, which allowed cytoplasmic microtubules to form, b u t not spindle microtubules. Addition of Taxol| would shift the monomer-polymer equilibrium t o w a r d the polymerized state and reduce the critical concentration of tubulin required for microtubule assembly. As a result, spindle microtubules would be able to form. However, quantitation of intracellular tubulin did not reveal significant alterations in the levels of total tubulin in these Taxol| cells. In order to further probe the nature of this resistance to antimitotic drugs, Minotti et al. have examined the levels of polymerized tubulin in drug-resistant CHO cells [40].

Since subtle changes in microtubule

327 stability (adjustments in the monomer-polymer equilibrium) are believed to be responsible for resistance to antimitotic agents in these mutants, the levels of polymerized tubulin in these cells may correlate with drug resistance. As anticipated, Taxol| mutants contained lower amounts of polymerized tubulin (-20-30%) relative to wild-type CHO cells (-40%). Thus, addition of Taxol| to these resistant cell lines would enhance the amount of polymerized tubulin, but the increase in polymer would not be sufficient to affect cell viability. Conversely, m u t a n t s r e s i s t a n t to microtubuledestabilizing agents, such as colcemid and vinblastine, contained elevated levels of polymer (-50%) relative to the wild-type cells. These effects on the levels of polymerized tubulin confirm that subtle alterations in the microtubule assembly properties of CHO m u t a n t s can confer drug resistance. This r e p r e s e n t s a common m e c h a n i s m of resistance to both microtubule-stabilizing and destabilizing agents. This mechanism is distinct from drug resistance mediated by overexpression of P-glycoprotein, a multidrug transporter. In addition, the sensitivity of a particular cell line to Taxol| or other antimitotic agents may be related to the level of polymerized tubulin in those cells. The levels of polymerized tubulin in different cell lines or tissues determined in a number of studies are highly variable. The selectivity of Taxol| against specific tumor types may be influenced by these variations in levels of polymerized tubulin. The need for more soluble Taxol| analogs has resulted in the development of Taxotere | a semi-synthetic analog with enhanced solubility and equal or better activity. Diaz and Andreu have recently compared the assembly of purified tubulin to microtubules induced by Taxol| T a x o t e r e | [102]. Their data confirm t h a t Taxol| and Taxotere | compete for the same binding site on microtubules. Notably, the critical concentration of tubulin required for microtubule formation was 2.1 +/- 0.1 times smaller with Taxotere | than with Taxol| 7.7. ALTERNATIVE AND SECONDARY E F F E C T S OF TAXOL| Although Taxol| interacts primarily with the tubulin]microtubule system, a number of in vivo Taxol| effects that appear independent of the tubulin]microtubule system have been reported. While investigating the

328 effects of microtubule targeting agents on tumor necrosis factor (TNF-a) receptors, Ding and co-workers observed that Taxol | profoundly affected murine macrophages [103]. Taxol | was found to induce TNF-(z release and decreased expression of the TNF receptor.

These effects mediated by

Taxol |

were identical to those elicited by bacterial lipopolysaccharide

(LPS).

These data suggest that the actions of LPS on macrophages are

mediated by an i n t r a c e l l u l a r t a r g e t also affected by Taxol |

The

intracellular target is suggested to be a microtubule-associated protein, rather than tubulin, and the presence of Taxol | function of an unidentified MAP.

may disrupt the normal

-In a subsequent study by Manthey and coworkers, the ability of Taxol | to induce gene expression and to generate a LPS-like signal in murine macrophages was examined [104]. Taxol | was found to increase the steady-state levels of LPS-inducible genes and to induce the tyrosine phosphorylation of several proteins with molecular weights of 41-45 kDa. Further elucidation of a common signaling pathway mediated by LPS and Taxol | may provide crucial insight into the cellular mechanisms by which Taxol | influences tumor growth. Continued efforts of Manthey and coworkers have demonstrated that the LPS-like activities of Taxol | in murine macrophages are distinct from the effects of Taxol | on microtubule structure and stability [105]. Two lipopolysaccharide antagonists, RsDPLA and SDZ 880.431, inhibited Taxol| TNF-r release, gene a c t i v a t i o n and tyrosine phosphorylation. However, these drugs were unable to inhibit Taxol | induced microtubule bundling in RAW 264.7 macrophages. Taxotere | a semisynthetic analog of Taxol | (see Figure 3), did not induce TNF-a release or gene expression but was more effective at inducing microtubule bundle formation. Microtubule bundling was observed in RAW 264.7 macrophages in the presence of as little as 75 nM Taxotere | whereas 300 nM Taxol | was required to achieve the same level of bundle formation. It is the view of Manthey and coworkers t h a t Taxol |

activates

macrophages through an as yet unidentified protein which also mediates LPS-induced signaling. Identification of the components of such a pathway will provide insight into the cellular mechanism of Taxol|

antitumor

activity. The antitumor activity of LPS has been documented and this activity is directly linked to LPS's ability to induce the release of TNF-a and

329 other cytokines [106].

In addition, LPS serves as a signal to activate

macrophage nitric oxide synthase. However, the observation t h a t the Taxol| analog, Taxotere | is unable to activate gene expression or TNF-a release suggests t h a t these secondary cellular effects of Taxol| on murine macrophages are not required for the therapeutic activity of Taxol| derivatives. Taxotere | with its e n h a n c e d w a t e r solubility and microtubule-stabilization activity, is currently in Phase II clinical trials and is highly effective against breast cancer and other solid tumors [102]. Continued efforts by Manthey and co-workers to elucidate the role of Taxol| LPS-like signaling have demonstrated that this antimitotic drug provides a second signal for murine macrophage tumoricidal activity [107]. The combination of LPS or Taxol| and IFN-y induced macrophages to lyse tumor cells. This synergism required a functional LPS signaling pathway, as d e m o n s t r a t e d by using both LPS-responsive and LPShyporesponsive macrophage cell lines. In addition, the combination of LPS or Taxol| and IFN-y induced expression of nitric oxide synthase and, consequently, increased nitric oxide secretion. Cellular levels of nitric oxide correlated with tumor cell killing and addition of the nitric oxide synthase inhibitor, NG-monomethyl-L-arginine, inhibited t u m o r cell killing. This work demonstrates that Taxol | has the potential to activate host a n t i t u m o r mechanisms which appear distinct from the effects of Taxol| on tubulin and microtubules. Ding and co-workers, who initially reported the shared activities of LPS and Taxol| have more recently examined the ability of Taxol| and LPS to induce the tyrosine phosphorylation of a microtubule-associated protein kinase, MAPK [108]. Phosphorylation is a common intracellular signaling event which has been shown to modulate the interaction of tubulin with MAPs. Tyrosine phosphorylation of MAPK, induced by Taxol | or LPS in murine macrophages, also triggered phosphorylation of an unidentified protein of approximately 86 kDa. MAPK, known as mitogenactivated protein kinase as well as a number of different names, is a family of s e r i n e / t h r e o n i n e k i n a s e s r e g u l a t e d by tyrosine a n d t h r e o n i n e phosphorylation. Induction of MAPK phosphorylation induced by LPS or Taxol|

was extremely rapid with near maximal levels of phosphate

incorporation within 1 minute.

Despite these recent findings, it is still

unknown whether these Taxol|

effects are all secondary to the interaction

330 of Taxol |

with microtubules. LPS has also been shown to bind to

microtubules. The possible links between microtubule binding and the LPS-like activities of Taxol |

seem to implicate microtubule-associated proteins in

the cellular mechanism of this antimitotic agent.

MAPK is a member of a

complex kinase cascade and the identification of u p s t r e a m activators of this cascade may provide scientists with clues as to the possible links between these two Taxol | activities.

It is important to note that a number

of proto-oncogene products including mos, fyn, and lyn are microtubuleassociated and some are tyrosine kinases [109, 110] which could act as upstream activators of MAPK.

Tyrosine phosphorylation of MAPK by a

microtubule-associated proto-oncogene product could be induced by a common Taxol|

pathway.

7.8. C O N C L U S I O N S

This review seeks to provide i n s i g h t s into the f u n d a m e n t a l mechanisms underlying the therapeutic utility of Taxol | by describing several critical biochemical and cellular effects of this antimitotic agent. U n d e r s t a n d i n g the interactions of Taxol |

with its p r i m a r y target, the

tubulin/microtubule system, at the biochemical level is vital to our ultimate understanding of the cellular consequences of Taxol | The ability of Taxol |

treatment.

and other antimitotic agents to suppress

microtubule dynamics both in vitro and in vivo provides the medicinal chemist with mechanistic insights as to how mitosis may be blocked. Ultimately, the roles of the numerous microtubule-associated proteins (MAPs) may provide some of the most compelling mechanistic clues as they may link the biochemical effects of Taxol |

on microtubules with the

cellular outcome of Taxol | treatment, which is apoptosis. Through continued efforts, f u n d a m e n t a l knowledge r e g a r d i n g the molecular mechanisms and the target sites of Taxol | cellular components will be acquired. fostering more effective Taxol |

interaction with

This knowledge can be applied to

use in cancer t r e a t m e n t , to predicting

additional sites for chemotherapeutic application, and to the development of additional therapeutic protocols.

331

o

2. 3. 4. ,

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7. .

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10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26.

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337

SUBJECT INDEX Abeotaxanes Acylations, of taxoids Aldol reaction Amentotaxus

Anhydrotaxinol Artifacts, isolation Austrotaxus

Baccatins NMR X-ray I,II III IV,V VI VII 10-deacetyl, III 1-hydroxy, I 14-hydroxy- 10-deacetyl bcl-2

Biliary excretion, of taxoids Brevifoliol Biogenesis, of taxoids Cardiotoxicity, of taxoids Clemeolide

20,22,23,45,57-62,86 23,25,174,179,182-187,203,212 275-277 8 34,36 38 8 15 93 94 16 16,95,97,165-255 16 16,61,62,77 16 16 73 75 323-325 154 15,85 25

Colchicine Conformation, of side chain Cremophor EL 3,11-Cyclotaxanes Cytochrome 450

43 27 323 306-314 293 104-110 20,22 143-51

Deacylations, of taxanes Dehydration, of taxoids Deoxygenations, of taxoids Disposition, of taxoids

171-173,177-181 198-201,207 176,188,189,196,208-211 152

Enzymes, in side chain synthesis 7-Epimers, metabolism

277-281 133,195

Fluorination, of taxoids Formulation, of taxoids

197-201 103-30

Glucuroconjugates, of Taxol Glycosidation, of taxoids

142 25

c-myc

338 Hydrolysis, metabolic Hypersensitivity, to Taxol

136 109

g-Lactams Linkers, self-immolating Liposomes LPS

267-274 117-24 125 328

MAPs Mass Spectra, of taxoids Michael reaction Microtubules bundles treadmilling Multi-drug Resistance

303,318,321,329 55 266 301-332 311,317-319 304 313,322-325

NMR Spectra, of taxoids Numbering, of taxoids

58-91 13

Oxazolidines Oxazolines Oxetane, reactions Oxidation, metabolic of Taxol

260-262 280 171,175,189-193

P-glycoprotein Paleotaxus grandis Paleotaxus jurassica Paleotaxus rediviva Plasma binding, of taxoids Prodrugs, of Taxol Pro-prodrugs, of Taxol Pseudotaxus

325 7 7 7 152 110-24 117-24 8

Reduction, of taxoids Representations, of taxoids Ring contraction, A, of taxoids

204,205,210 13 42,170

134-7 202

Skeletal rearrangements, of taxoids 32,222-4 Solubility, of Taxol 106 Spicaledonine 22 Sulfation, metabolic 142 Sulfinimines 273-274 Taiwanxan Tau Taxacultins Taxagifine

15,93 303 17 15,22,27,65,93,96

339 Taxchins Taxchinins Taxicins Taxines biology NMR spectra structures Taxinines A B, E, H, J, K, L M Taxomyces andreanae Taxus Taxus baccata Taxus brevifolia Taxus celibica Taxus cuspidata Taxus globosa Taxus wallichiana Taxus yunnanensis Taxusin Taxuyunnanines

43,44 84,89 10,11,16 12,16,20,34,72,78,80-83 17,94 17 17,87 31 8,45 7,8,9,39 9,31,45,105 8,9 9 9 9 8 93 16,27

Torreya Tubulin isoforms isotypes

8,45 301-332 315 315

Urinary excretion, of Taxol UV Spectra, of taxoids

154 56-57

Verticilloids Vinblastine

57 306,314

Winterstein esters

11,24,31,41

16,27 16,31,94 79