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THIRD-GENERATION PHOTOSENSITIZERS:
Synthesis, Characterization, and Liposome Interaction
of Promising New Benzoporphyrin Derivatives
BY
ANDREW NORMAN TOVEY
B.Sc, University of British Columbia, 1991
A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF
THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
in
THE FACULTY OF GRADUATE STUDIES
DEPARTMENT OF CHEMISTRY
We accept this as confonning
to the required standard
THE UNIVERSITY OF BRITISH COLUMBIA
September 1994
© Andrew Norman Tovey, 1994
In presenting this thesis in partial fulfilment of the requirements for an advanced
degree at the University of British Columbia, I agree that the Library shall make it
freely available for reference and study. I further agree that permission for extensive
copying of this thesis for scholarly purposes may be granted by the head of my
department or by his or her representatives. It is understood that copying or
publication of this thesis for financial gain shall not be allowed without my written
permission.
Department of C^BMiST^j The University of British Columbia Vancouver, Canada
DE-6 (2/88)
ABSTRACT 11
The goal of this project was to synthesize new compounds via the peripheral
modification of ring B BenzOporphyrin Derivative (BPD) dimethyl ester so that their
viability as photosensitizers could be assessed Various moieties were attached to the BPD
chromophore via ester, amide, amine and phosphorus-carbon linkages, with the rationale
behind each modification provided. Compounds (3)-(ll), (13)-(20), (23), (26), (27),
(29)-(33) were characterized by NMR, high resolution mass spectroscopy, and, in
some cases, elemental analysis and all are presently undergoing in vitro and in vivo
biological testing.
The interactions of (2), (4), (6), and (8) as well as the successful photosensitizer
Benzoporphyrin Derivative monoacid ring A (BPDMA) with unilamellar vesicles was also
investigated using fluorescence quenching of liposome-bound diphenylhexatriene hexanoyl
phosphatidylcholine (DPH-hPC). Stern-Volmer plots were derived for each compound
and the different results were discussed.
T A B L E OF CONTENTS 111
ABSTRACT .
TABLE OF CONTENTS
LIST OF TABLES .
LIST OF FIGURES .
LIST OF SCHEMES .
LIST OF ABBREVIATIONS
ACKNOWLEDGEMENTS .
CHAPTER 1 INTRODUCTION 1
1.1 Overview. . . . . . . . 2
1.2 Structural Features . . . . . . 6
1.3 Nomenclature . . . . . . . 7
1.4 Electronic Spectra and Absorption Properties . . . 12
1.5 Photodynamic Therapy . . . . . . 16
1.5.1 Introduction and History . . . . 16
1.5.2 Benzoporphyrin Derivative Monoacid Ring A (BPDMA) 20
1.5.3 Major Second Generation Photosensitizers . . 22
1.5.4 Third Generation Photosensitizers . . . 23
1.5.5 The Photodynamic Effect—Singlet Oxygen Production 24
1.5.5.1 Introduction . . . . . 24
1.5.5.2 Excited States of Photosensitizers . . 25
1.5.5.2.1 Type I Photoprocesses . . 27
1.5.5.2.2 Type II Photoprocesses . . 29
1.5.5.3 Determination of Singlet Oxygen Production 31
1.6 Fluorescence Quenching and Liposomes . . . 32
iii
vi
vii
xi
xiii
xv
1 V
1.6.1 Experimental Challenge
1.6.2 Liposomes
1.6.3 Liposomes as Drug Delivery Agents
1.6.4 Fluorescence Quenching
32
33
35
36
CHAPTER 2 SYNTHESIS OF BPD DERIVATIVES: RESULTS
AND DISCUSSION 40
2.1 Research Objective . . . . . 41
2.2 Synthesis of Ring B Benzoporphyrin Derivative
Dimethyl Ester (BPD) 43
2.3 Synthesis of Variable Alkyl Chain Ester Derivatives of BPD . 50
2.3.1 Rationale . . . . . 50
2.3.2 Via the BPD Vinyl Group . . . . 52
2.3.3 Via Transesterification . . . . . 54
2.3.4 Via Amide Formation . . . . . 57
2.3.4.1 Direct Amidation . . . . 57
2.3.4.2 Nucleophilic Displacement . . . 58
2.4 Synthesis of BPD Derivatives with Free Amine Functionalities . 63
2.4.1 Rationale . . . . . . 63
2.4.2 Via Amide Linkage . . . . . 66
2.4.3 Via Coupling to the BPD Vinyl Group . . 69
2.5 Synthesis of Analogous BPD Amines and Amides . . 72
2.5.1 Rationale . . . . . . 72
2.5.2 BPD Amides via Acyl Chloride Displacement . . 73
2.5.3 BPD Amines via Iodide Displacement . . . 73
2.6 Synthesis of BPD Derivatives with Hydroxyl Functionalities . 79
2.6.1 Rationale . . . . . . 79
2.6.2 Via Reduction of BPD .
2.6.3 Via Transesterification .
2.6.4 ViaAmidation .
2.6.5 Via the BPD Vinyl Group
2.7 Synthesis of Phosphonate Ester BPD Derivatives
2.7.1 Rationale .
2.7.2 Via Iodide Displacement
2.8 Summary . . . . .
82
86
86
87
88
88
89
91
CHAPTER 3 FLUORESCENCE QUENCHING: RESULTS
AND DISCUSSION 92
3.1 Fluorescent Probe Selection . . . . . 93
3.2 Composition and Creation of Liposomes . . . 97
3.3 Fluorescence Spectra of Liposome-Bound
Diphenylhexatriene (DPH) and BPD . . . . 98
3.4 Photobleaching of Liposome-Bound DPH . . . 99
3.5 Fluorescence Quenching and Stern-Volmer Plots . . 100
3.6 Summary . . . . . . 106
3.7 Future Work 107
CHAPTER 4 EXPERIMENTAL 108
4.1 General Methods for BPD Derivatives . . . . 109
4.2 General Methods for Fluorescence Quenching Experimentation . I l l
4.3 Synthesis of BPD Derivatives . . . . . 115
REFERENCES 150
LIST OF TABLES
Table 3.1: Percent Fluorescence Photobleaching of Liposome-Bound
DPH-hPC at Various Excitation and Emission Bandpasses
(EX 362nm, EM433, 200 flashes) 99
LIST OF FIGURES
Figure 1.1: Important Tetrapyrrolic Backbones . . . . . 2
Figure 1.2: Biosynthesis of Uroporphyrinogen III . . . . 3
Figure 1.3: Various Biologically Important Tetrapyrroles . . . 4
Figure 1.4: Delocalized Electron Pathways of Porphyrins and
Reduced Porphyrins . . . . . . . 6
Figure 1.5: Structure of [34] Annulene . . . . . . 7
Figure 1.6: Fischer Numbering System for Porphyrins . . . . 8
Figure 1.7: The Four Isomers of Uroporphyrin . . . . . 9
Figure 1.8: The Structure of Protoporphyrin LX . . . . . 9
Figure 1.9: Systematic Numbering System for Porphyrins . . . 10
Figure 1.10: The Structure and Systematic Name of Ring B BPD
Dimethyl Ester . . . . . . . 11
Figure 1.11: The Structure of BPD Diol . . . . . . 12
Figure 1.12: The Four Types of Porphyrin Spectra . . . . 13
Figure 1.13: The Typical Spectrum of a Protonated Porphyrin . . . 14
Figure 1.14: Typical Spectra of Chlorins and Metallochlorins . . . 15
Figure 1.15: Typical Spectra of Bacteriochlorins and
MetaUobacteriochlorins . . . . . . 15
Figure 1.16: The Various Components of Hematoporphyrin Derivative . . 18
Figure 1.17: The Ring A and Ring B Dimethyl Esters, Monoacid/
Monoesters and Diacids of BPD (Only One Enantiomer
of Each is Shown) . . . . . . . 21
Figure 1.18: Major Second-Generation Photosensitizers:
a) Mono-L-Aspartyl Chlorin t?6; b) Tin Etiopiirpurin;
c) Meso-tetra(m-hydroxyphenyl)chlorin . . . . 23
Figure 1.19: Modified Jablonski Diagram for a Typical Photosensitizer
Figure 1.20: The Type I Reaction of 2-Methyl-1,4-Naphthoquinone
(MQ) with Cholesterol in the Presence of
Light and Oxygen . . . . .
Figure 1.21: Examples of the Three Major Classes of Type II Reactions
Figure 1.22: General Liposome Structure . . . .
Figure 1.23: Schematic Representation of n-(9-Anthoyloxy) Fatty Acids
Showing the Transverse Positions of the 9-Anthoyloxy
Fluorescent Probe . . . . .
Figure 2.1: The Conversion of Hematoporphyrin LX to
Protoporphyrin LX Dimethyl Ester
Figure 2.2: Diels-Alder Reaction of Protoporphyrin LX Dimethyl Ester
(PPLX) with Dimethylacetylene Dicarboxylate (DMAD)
Figure 2.3: UV-Vis Spectra (CH2CI2) of Protoporphyrin LX
Dimethyl Ester and BPD 1,4-Diene Dimethyl Ester .
Figure 2.4: The Conversion of BPD 1,4-Diene Dimethyl Ester
to BPD 1,3-Diene Dimethyl Ester and Their Respective
UV-Vis Spectra in CH2CI2 . . . .
Figure 2.5: A H NMR (CDCI3), UV-Vis Spectrum (CH2CI2), and
Structure of BPD 1,3-Diene Dimethyl Ester .
Figure 2.6: Alkyl Ether Derivatives of Hematoporphyrin .
Figure 2.7: Alkyl Ether Derivatives of Methyl Pheophorbide a
and Chlorin erj Trimethyl Ester
Figure 2.8: The Chemical Shifts of the Vinyl Group of BPD
Dimethyl Ester . . . . .
Figure 2.9: *H NMR (CDCI3), UV-Vis Spectrum (CH2CI2) and
Structure of BPD Dipentyl Ester
IX
Figure 2.10: ! H NMR (CDCI3), UV-Vis Spectrum (CH2CI2) and
Structure of BPD Dibutyl Amide (14) . . . . 62
Figure 2.11: Amide-Linked Protoporphyrin Derivatives . . . . 64
Figure 2.12: Amide-Linked Morpholine Derivatives of
Hematoporphyrin LX . . . . . . . 65
Figure 2.13: lH NMR (DMSO-d6) and Structure of BPD
Di(N,N-dimethylethylenediamine)amide (16) . . . 68
Figure 2.14: Reaction of Deuteroporphyrin LX Dimethyl Ester and
Protoporphyrin LX Dimethyl Ester with N,N-
dimethylmethylenearranonium Iodide . ; . . . 6 9
Figure 2.15: *H NMR (DMSO-d6) and Structure of BPD
Eschenmoser Derivative (18) . . . . . . 71
Figure 2.16: Two Pairs of Amide- and Amine-Linked BPD Derivatives . . 72
Figure 2.17: The UV-Vis Spectra of BPD Dimethyl Ester (2) and Zn BPD
Dimethyl Ester (21) in CH2CI2 . . . . . 74
Figure 2.18: The UV-Vis Spectra of BPD Dimethyl Ester (2) and
BPD Diol (23) in CH2CI2 76
Figure 2.19: lH NMR (DMSO-d6) and Structure of Zn BPD diol (22) . . 77
Figure 2.20: Porphyrins and Chlorins with Hydroxy Groups:
a) Tetra(para-hyckoxyphenyl)porphyrin and Tetra(meta-
hydroxyphenyl)porphyrin; b) Tetra(para-hydroxyphenyl)chlorin
andTetra(meta-hydroxyphenyl)chlorin; c) Hydroxy
Octaethylchlorin Derivatives . . . . . . 80
Figure 2.21: Zinc Phthalocyanine Derivatives with Hydroxy Groups . . 81
Figure 2.22: UV-Vis Spectra (CH2Cl2/MeOH) of Zn BPD Triol (28)
and Zn BPD (21), and lH NMR (DMSO-d6) and Structure
ofZnBPDTriol 83
Figure 2.23: UV-Vis Spectrum of BPD Triol (29) in CH2Cl2/MeOH . . 85
Figure 2.24: Phosphonate Ester Substituted Phthalocyanines . . . 88
Figure 2.25: lH, 3 AP NMR (DMSO-d6) and Structure of BPD
Diphosphate Ester (33) . . . . . . 90
Figure 3.1: Various Diphenylhexatriene-Containing Fluorescent Probes . . 94
Figure 3.2: Idealized Representation of the Orientation of
Diphenylhexatriene Hexanoyl Phosphatidylcholine
(DPH-hPC) Fluorescent Probe in a Dimyristoyl
Phosphatidylcholine (DMPC) Bilayer . . . . 95
Figure 3.3: Structure of BPD Dimethyl, Dipropyl, Dipentyl,
and Diheptyl Esters . . . . . . . 96
Figure 3.4: Dimyristoyl Phosphatidylcholine (DMPC) and Egg
Phosphatidylglycerol (EPG) . . . . . . 97
Figure 3.5: Fluorescence Excitation and Emission Spectra of Liposome-
Bound DPH-hPC in Aqueous Solution and BPD Dimethyl
Ester in n-Methylpyrrohdinone . . . . . 99
Figure 3.6: Stern-Volmer Plots of Fo/F versus Concentration BPD
Derivative For BPDMA, and the BPD Dimethyl, Dipropyl,
Dipentyl, Diheptyl Esters . . . . . . 102
Figure 3.7: Probable Orientation of Hematoporphyrin LX Dipropionic
Acid in Lipid Bilayers . . . . . . 103
Figure 3.8(a): Proposed Orientation of BPDMA Within a Lipid Bilayer . . 105
Figure 3.8(b): Proposed Orientation of BPD Dimethyl Ester Within
a Lipid Bilayer . . . . . . . 105
Figure 3.8(c): Proposed Orientation of Longer Alkyl Chain BPD Diesters
(only BPD Diheptyl Ester is Shown) Within a Lipid Bilayer . . 105
xi
LIST OF SCHEMES
Scheme 2.1: The Conversion of BPD Dimethyl Ester (2) to BPD
Amine (3) and the Resulting Stereoisomers . . . . 53
Scheme 2.2: General Transesterification of BPD Dimethyl Ester (2) . . 55
Scheme 2.3: Direct Amidation of BPD Dimethyl Ester (2) . . . . 57
Scheme 2.4: Amidation of BPD Dimethyl Ester (2) Via Displacement
of BPD Diacid Chloride (35) 60
Scheme 2.5: Synthesis of BPD Derivatives with Free
Amine Functionalities . . . . . . . 66
Scheme 2.6: Quaternization of BPD Di(N,N-dimemylemylenedianiine)airiide
(16) Using Methyl Iodide . . . . . . 67
Scheme 2.7: Reaction of BPD Dimethyl Ester with N,N-dimethyl-
methyleneammonium Iodide to Form BPD
Eschenmoser Derivative (18) . . . . . . 70
Scheme 2.8: Synthesis of BPD Amides Using Acid Chloride Displacement . 73
Scheme 2.9: Synthesis of Two BPD Amine Derivatives Via
Reduction of BPD Dimethyl Ester (2) . . . . 75
Scheme 2.10: Synthesis of BPD Triol (29) from Zn BPD
Dimethyl Ester (21) . 8 4
Scheme 2.11: Synthesis of BPD Di(ethyleneglycol) Ester (30) from
BPD Dimethyl Ester (2) Via Transesterification . . . 86
Scheme 2.12: Synthesis of BPD Di(ethanol)amide (32) By
Direct Displacement . . . . . . . 87
Scheme 2.13: Synthesis of BPD Vinyl Hydrate (31) Via
Hydrobromination and Displacement . . . . . 87
Scheme 2.14: Synthesis of BPD Diphosphonate Ester (33)
from BPD Diol (23) .
LIST OF ABBREVIATIONS x i i i
ATP adenosine triphosphate
BPD Benzoporphyrin Derivative
BPDMA Benzoporphyrin Derivative Monoacid Ring A
br. broad
DMAD dimethyacetylene dicarboxylate
DMF N,N-dimethylformarnide
DBU 1,8-diazabicyclo[5.4.0]undec-7-ene
DMSO dimethylsulfoxide
DD3AL-H diisobutylaluminum hydride
DPH diphenylhexatriene
DPH-hPC diphenylhexatriene hexanoyl phosphatidylcholine
DMPC dimyristoyl phosphatidylchohne
d doublet
EI electron impact
EPG egg phosphatidylglycerol
HpD hematoporphyrin derivative
HDL high-density lipoprotein
Hz hertz
IUPAC International Union of Pure and Applied Chemistry
ISC intersystem crossing
J coupling constant
LDL low-density lipoprotein
LUV large unilamellar vesicle
LRMS low-resolution mass spectroscopy
Me methyl
m multiplet
MLV rnultilarnellar vesicle
NMR nuclear magnetic resonance
NMP n-methylpyrrolidinone
PPIX protoporphyrin LX dimethyl ester
PDT photodynamic therapy
RES reticulo-endothelial system
SUV small unilamellar vesicle
s singlet
t triplet
tic thin-layer chromatography
TEA m^mylamine
THF tetrahydrofuran
UV-Vis ultra-violet and visible
A C K N O W L E D G E M E N T S
First and foremost I would like to thank my supervisor, Dr. David Dolphin, for his support
and guidance throughout the course of this work. I would also like to acknowledge all the
members of my research group, past and present, in particular Dr. Ethan Sternberg, who kindly
provided constructive criticism and helpful suggestions at all stages of this work. I must also
thank Dr. Ross Boyle and Dr. Veranja Karunaratne for their fruitful discourse and for proof
reading portions of this thesis and Dr. Lawrence Mayer who was integral in the design of the
fluorescence experiments. Special thanks go to Mr. Mike Wong who was extremely supportive
over the course of my graduate studies. The assistance of Mr. Dan Debeyer and the employees of
QuadraLogic Technologies is also gratefully acknowledged.
I would like to extend my utmost thanks to the Science Council of British Columbia who
awarded me a G.R.E.A.T. award for two consecutive years.
Finally, I thank my family for their support and most of all my wife, Ann, to whom this
thesis is dedicated.
Chapter 1 Introduction
1.1 Overview
Porphyrins, chlorins, bacteriochlorins, isobacteriochlorins and corrins are the
macrocyclic backbones of many important natural pigments of life. These molecules each
consist of four pyrrolic moieties joined together directly or more commonly through
methine carbon bridges (Figure 1.1).
Porphyrin Chlorin Bacteriochlorin
IsobacteriocWorin Corrin
Figure 1.1 Important Tetrapyrrolic Backbones
The structural similarities between these compounds led to the belief that they have
a common ancestor. This ancestor is now known to be uroporphyrinogen III*, a
compound which is biosynthesized from four molecules of porphobilinogen, each of which
is formed from the condensation of two molecules of 5-ammolevulinic acid (Figure 1.2).
Q m i . i . - •>. <• •— ^OjH
5-aminolevulinic acid NH2
Porphobilinogen
C02H C02H Uroporphyrinogen III
Figure 1.2 Biosynthesis of Uroporphyrinogen HI
Porphyrins, and their reduced analogs, are responsible for myriad biological
functions. Some of these compounds are shown in Figure 1.3. For example, the iron(Il)
metallated complex of protoporphyrin IX, called protoheme or heme, is the prosthetic
group of hemoglobin and myoglobin as well as cytochromes, catalases and peroxidases .
Chlorophyll a, a chlorin pigment with a chelated magnesium ion, is responsible for the
harvesting of light energy in plants3. Another magnesium containing compound,
bacteriochlorophyll a, and its demetallated counterpart bacteriopheophytin a, both having a
bacteriochlorin structure, are the main components of the photosynthetic apparatus of
purple and green bacteria . Vitamin B12 has a cobalt metallated corrin structure and is the
prosthetic group for a number of important enzymes .
Almost sixty years ago the first porphyrins were isolated from oil shale which lent
credence to the belief that petroleum was derived from plant and animal remains**. This
discovery has given rise to the scientific field known as organic geochemistry, which
involves the isolation, characterization and total synthesis of these compounds,
appropriately named 'petroporphyrins'.
4
Bacteriochlorophyll a Bacteriopheophytin a
Figure 1.3 Various Biologically Important Tetrapyrroles
Recent experimentation on metallated and non-metallated porphyrins has determined
that many of these tetrapyrrolic macrocycles have features which make them particularly
well-suited for a new anti-cancer treatment called photodynamic therapy (PDT). This
promising treatment utilizes the fact that porphyrins preferentially biodistribute in
tumourous versus healthy tissue and that they are capable of producing singlet oxygen via
photosensitization when illuminated.
The continued interest in biosynthetic and synthetic aspects of tetrapyrroles coupled
with their utility in several other scientific disciplines, guarantees a healthy future for
research into porphyrins and their analogs.
1.2 Structural Features
Porphyrin Chlorin Bacteriochlorin
Figure 1.4 Delocalized Electron Pathways of Porphyrins and Reduced Porphyrins
Porphyrins are tetrapyrrolic macrocycles with the pyrrole moieties joined by
methine carbon bridges. They possess 227t-electrons, 18 of which, in a given resonance
structure, participate in a cyclic delocalized conjugation pathway (Figure 1.4). These
compounds are generally planar (although exceptions have been synthesized ) and they
adhere to Huckels' 4n+2 rule (where n=4 or 5) and thus porphyrins have been referred to
as 'aromatic'. The closed, conjugated electron pathway generates a diamagnetic ring
current in a way analogous to benzene. This is manifest in the L H NMR of porphyrins,
where the inner pyrrolic protons are shielded from the externally applied magnetic field and
typically appear between -2 and -5ppm. The protons located on the bridging carbons (the
'meso' protons labeled H m ) are deshielded by this ring current and this effect places their
resonances downfield at about lOppm^. The most profound ring-current effect for these
types of compounds has been seen with expanded porphyrins which have added ethylene
moieties between each pyrrole subunit. One such porphyrin, [34] annulene, with 34 7t-
electrons, gave a chemical shift difference of 31.5ppm between the inner and outer
protons^ (Figure 1.5).
Figure 1.5 Structure of [34] Annulene
Chlorins are porphyrins which have undergone a two hydrogen reduction in one of
the pyrrolic rings (Figure 1.4). They retain the carbon skeleton of porphyrins but with two
fewer Jt-electrons. There remains a closed aromatic system of electrons, so the system is
still predominantly planar and ring-current effects still occur. Bacteriochlorins are
tetrahydroporphyrins with opposite exocyclic 'double-bonds' having been reduced (Figure
1.4).
All these compounds can act as tetradentate ligands and can bind literally dozens of
metals in various oxidation states^ with the concomitant loss of the pyrrolic hydrogens.
Such metallations alter the electronic distribution within the macrocycle and hence change
its reactivity.
1.3 Nomenclature of Porphyrins and Structural Analogs
The nomenclature of porphyrin-type compounds is difficult to understand due to the
fact that in practice the trivial naming schemes of the past have not given way gracefully to
the new systematic naming schemes. The initial naming system was developed by Fischer
and others1*"13 in the 1920's and 1930's and employs a large amount of trivial names in
conjunction with a basic numbering system,
porphyrin nomenclature are discussed below,
of all porphyrins, porphin (Figure 1.6).
2 a 3
Figure 1.6 Fischer Numbering System for Porphyrins
The P-positions of the pyrrole moieties are numbered 1-8 and the methine carbons joining
the pyrroles (called 'meso' carbons) are labeled a-8. For the dihydroporphyrins, chlorins,
the skeletal structural is the same as porphin but the site of saturation is between the
carbons located at P-positions 7 and 8. For tetrahydroporphyrins, such as
bacteriochlorins, porphin is again used with the sites of saturation being between 0-
positions 7 and 8 and between 3 and 4.
Fischer used a 'type' system to characterize porphyrins with the same substituents
attached at the P-positions on the pyrrole rings and to distinguish between the various
isomeric possibilities. For example, the uroporphyrins are porphyrins with acetic (A) or
propionic (P) acid groups at the pyrrolic P-positions with one of each on each pyrrole
moiety. The four isomeric possibilities are shown in Figure 1.7 using lines denoting the
two p-positions of each pyrrole moiety 14.
Some of the more widely used aspects of
The basic scheme is shown for the simplest
P A P P P A P P
A / \ P k/ \ A A / \ P A / \ A
P \ ^ y A A \ ^ yK / A p \ / p
A P P P P P A A Type I TypeU Type III Type IV
Figure 1.7 The Four Isomers of Uroporphyrin
For systems with three different groups, A, B, and C, with one of A on each
pyrrole group, one of B on each of two pyrroles, and one of C on the remaining two (3-
positions, there are 15 different orientations. For example, the series of porphyrins given
the name protoporphyrin, where the three different substituents are vinyl (V), propionic
acid (P) and methyl (M), has 15 different isomers. The compound which happened to be
the ninth isomer in this series, protoporphyrin IX, is the only naturally occurring
protoporphyrin, the dimethyl ester of which is the crucial starting material in the making of
BPD dimethyl ester (Figure 1.8). Other important trivially named porphyrins are
etioporphyrin, rhodoporphyrin, and phylloporphyrin, on the spectra of which the general
classification of optical spectra of porphyrins is based (see section 1.4).
Figure 1.8 The Structure of Protoporphyrin LX
10
A revised form of porphyrin nomenclature was proposed in 196015 and adopted in
198816 by a IUPAC-IUB joint commission. It takes into account all the substituents
including the pyrrolic nitrogens in a systematic way. The simplest porphyrin, appropriately
called porphine, shows the new numbering scheme (Figure 1.9).
17 1 6 1 5 1 4 13
Figure 1.9 Systematic Numbering System for Porphyrins
The pyrrole rings are lettered clockwise, starting from the top left, A through D. The
carbons are numbered 1 through 20 starting from the leftmost ce-carbon of ring A and
proceeding in a clockwise fashion around the tetrapyrrole. The pyrrolic nitrogens are
numbered 21 through 24 with the numbering again starting at the ring A substituent. The
old type nomenclature has been retained in some cases for the sake of brevity i.e.
protoporphyrin IX is now protoporphyrin. Any compound that has the basic structure of
protoporphyrin would keep protoporphyrin as the root and the modifications would be
systematically added to the name. For example, the compound resulting from the methyl
esterification of the propionic acid chains of protoporphyrin is called protoporphyrin
dimethyl ester.
For systems with fused rings, the numbering of the ring is derived from the lowest
numbered carbon in the porphyrin skeleton. A letter or letters denoting to which pyrrole(s)
the fused ring(s) is(are) attached appear in brackets directly preceding the word porphyrin.
The letters are italicized b, g, 1, q corresponding to rings A through D. For example, the
structure and systematic name for BPD dimethyl ester is shown in Figure 1.10.
The systematic nomenclature, while much less ambiguous and more widely
applicable, is not exclusively used today presumably due to tradition and brevity.
Throughout this work, the trivial names 'BPD' and 'BPD dimethyl ester' will be used to
refer to the compound shown in Figure 1.10 for the sake of brevity and any modification to
BPD will be reflected in added text. For example, the BPD derivative where the chain
methyl esters have been reduced to the corresponding alcohols will be called 'BPD diol'
(Figure 1.11).
C H 3 0 2 C C 0 2 C H 3
Ring B BPD Dimethyl Ester
(71,72-bis(memoxycarrjonyl)-13,17-bis(2-(methoxycarbonyl)ethyl)-
2,7,12,18-tetramethyl-3-vinyl-7,71 -dihydrobenzo[g]porphyrin)
Figure 1.10 The Structure and Systematic Name of Ring B BPD Dimethyl Ester
Figure 1.11 The Structure of BPD Diol
1.4 Electronic Spectra/Light Absorption Properties
The electronic spectra of porphyrins and related compounds have been used in their
characterization since the late 1800's. In 1883, the intense absorption of hemoglobin
centered at 400nm was observed by Soret1^. This intense absorption is characteristic of
conjugated tetrapyrrolic molecules and is commonly referred to as the Soret or (3 band.
This band has by far the most intense absorption and generally has an extinction coefficient
in the range 100,000-400,000M"1cm"i. For porphyrins, there are also four accompanying
less intense lower energy absorptions commonly referred to as 'Q-bands' which appear in
the 450-650nm region.
The relative intensities of these four bands, denoted IV, IU, II, and I in decreasing
energy, place a particular porphyrin into one of four categories: etio-type, rhodo-type,
oxorhodo-type, and phyllo-type. The four spectral "types" are named from the porphyrins
which were first found to exhibit these characteristic spectra (Figure 1.12).
13
500 650 600 650 600 650 600 650
Wavelength (nm)
Figure 1.12 The Four Types of Porphyrin Spectra
The etio-type spectra are representative of all naturally occurring porphyrins and are
distinguished by methyl, ethyl, acetic or propionic acid groups on the P-positions of the
macrocycle. The rhodo-type spectra occur in porphyrins with a single conjugation-
extending group directly attached to a p-position such as a ketone, aldehyde, carboxylic
acid, ester or unsaturated moiety like a vinyl group. When there are conjugation-extending
groups on diagonally-opposite pyrrole rings the oxorhodo-type spectrum results. The
phyllo-type spectra result when the P-positions on the porphyrin are substituted with alkyl
or propionic acid groups but at least one P-position remains unsubstituted. The extinction
coefficients of these Q-bands are roughly one-fiftieth that of the Soret bands in most
porphyrin spectra. The type of substituent attached to the porphyrin not only changes the
relative intensities of the Q-bands but results in hypsochromic (blue-shifted) or
bathochromic (red-shifted) shifts of these peaks. For example, a vinyl group in
conjugation with the porphyrin 7t-electron system provides a bathochromic shift of the Q-
bands1^.
Much work has been done to interpret the electronic spectra of porphyrins. It is
well known that the colours associated with these compounds are due to the planar,
14
conjugated 7c-electron system. Piatt19 has proposed a simplified four-orbital model to
explain porphyrin spectra. This model attributes the four long wavelength absorptions to a
low energy 'Q' state (the so-named Q-bands) in which the transition dipoles nearly cancel
out. The Soret band is the result of a strongly allowed higher-energy excited state where
the transition dipoles add.
The electronic spectra of porphyrins change dramatically in acidic media. When all
four pyrrolic nitrogens are protonated a dication is formed.
<—» c Si & u > <L>
-1 ' i 5O0 600
wavelength (nm)
Figure 1.13 The Typical Spectrum of a Protonated Porphyrin
The four Q-bands collapse to a two band system, a result which has been attributed to the
approach of the porphyrin towards square planar symmetry^ (Figure 1.13). The change
in the spectra of metalloporphyrins is similar, metallated systems have a Soret band and
only two peaks in the visible region, normally called the a and P bands in decreasing
energy. The intensities and locations of these peaks are determined in part by the type and
oxidation state of the metal ion.
The most relevant changes in the electronic spectra of porphyrins with respect to
this work are those which occur upon hydrogenation. Chlorin-type compounds
(dihydroporphyrins) result from the reduction of one exocyclic double bond in the
porphyrin macrocycle. The spectrum shows the lowest energy band is bathochromically
15
shifted by 20-30nm and becomes much more intense in relation to the Soret band (the ratio
of Soret to band I is now closer to 5:1) (Figure 1.14)21. T n e Soret is generally red-shifted
as well but the intensity is approximately the same as in the corresponding porphyrin. The
remaining Q-bands are in evidence but are less distinct and still roughly one-fiftieth the
intensity of the Soret band. Metallation of chlorins produces a simplified spectrum in
comparison to the free-base chlorin, again due to the increase in symmetry, with a shift in
*
— chlorin
— metallochlorin
wavelength (nm)
Figure 1.14 Typical Spectra of Chlorins and Metallochlorins
Figure 1.15 Typical Spectra of Bacteriochlorins and Metallobacteriochlorins
the band I absorption (usually hypsochromic but dependent on the valence and type of
metal) (Figure 1.14). Some of the other Q-bands can be shifted or may disappear
altogether. Protonation of chlorins in acidic media causes many of the same changes as
metallation.
Further reduction of a chlorin produces a bacteriochlorin. The highest wavelength
absorption is red-shifted in relation to the corresponding chlorin and becomes much more
intense (Figure 1.15)22. This band I absorption appears at roughly 750nm with an
extinction coefficient of ca. 80,000cm" M " 1 . Metallation produces only a small change in
the spectra of bacteriochlorins.
The differences in electronic spectra described above are extremely important
diagnostic tools in the chemistry of tetrapyrroles. Not only can the identity of a compound
be determined or confirmed but the spectra are invaluable in monitoring the progress of
reactions and assessing the completion of work-up.
1.5 Photodynamic Therapy (PDT)
1.5.1 Introduction to PDT and the History of PDT drugs
Photodynamic therapy is a medical treatment which employs a combination of light
and drug to create cytotoxic ('cell-lethal') forms of oxygen (singlet-oxygen and superoxide
radical), as well as other reactive species, to bring about the destruction of cancerous or
unwanted tissue. While the term photodynamic therapy is relatively new (ca. 20 years
old), this form of treatment can be traced back to the ancient Egyptians, who used the
combination of orally ingested plants (containing light-activated psoralens) and sunlight to
treat vitilago^3.
Contemporary PDT began near the turn of the last century. Raab^4 used the
combination of acridine dye and light to destroy paramecia in 1900. This was followed by
the work of Jesionek and Tappeiner^S, who, in 1903, treated a skin cancer with topically
applied eosin and light. It was a few years later (1913) when the first porphyrin-type
compound, hematoporphyrin, was shown to have the 'photosensitizing' property, i.e. the
ability to bring about a photodynamic effect. In an unusual experiment, Meyer-J3etz26
administered 200mg of hematoporphyrin to himself and remained extremely sensitive to
light for weeks.
Relatively little work was done on these types of compounds in the next 50 years
until a mixture called 'hematoporphyrin derivative' (HpD) was prepared by Lipson and
Baldes^ i n 1960. This multiple component, first-generation photosensitizer was, and
currently is, prepared by the treatment of hematoporphyrin with 5% sulfuric acid in acetic
acid followed by alkaline hydrolysis. A number of monomelic porphyrins are produced in
this two step reaction (Figure 1.16). Also produced are many porphyrin oligomers 2^!,
with the linkages identified as ether32-34? ester30,35 carbon-carbon34,36 j-ipD was
found by these experimenters ^ to preferentially accumulate in certain cancerous tissues in
animals and attempts were made to utilize HpD for tumour imaging.
However, this potential anti-cancer modality was not investigated further until the
early 1970's, when the first sustained series of tests on animals and human cancers were
begun using HpD by Dougherty and others38-40> m a t m e field of clinical PDT truly began.
Dougherty is regarded by many to be the principle reason for the advancement of modem
PDT.
The bulk of work for the next decade was performed on HpD and scientists were
interested in whether there was a particular component (or components) of the HpD mixture
that were responsible for the observed photodynamic effects. It was found that the lower
molecular weight monomelic fraction showed no in vivo photodynamic activity but the
oligomeric, higher molecular weight fraction did show activity^1. A purified, synthetically
reproducible, portion of this latter fraction of HpD was patented in 1981 under the trade
name Photofrin™.
PhotofrinTM changed hands a number of times in the next few years until
QuadraLogic Technologies (QLT) obtained the patent from Johnson & Johnson in 1987.
In collaboration with American Cyanamid/Lederle
Hematoporphyrin 1) 5% sulfuric acid/acetic acid 2) IN aqueous NaOH
and Oligomers
Figure 1.16 The Various Components of Hematoporphyrin Derivative
19
Laboratories, QLT performed the Phase III (human) clinical trials. In addition, in the past
decade it is estimated^ that nearly 10,000 publications relating to PDT have appeared
detailing biological testing of almost all types of solid human cancers, most of which using
Photofrin™.
In April, 1993, Photofrin^M received approval for the treatment of papillary
bladder cancer from the Health Protection Bureau of Canada. Approval was also received
in the Netherlands for the treatment of esophageal cancer. Similar approvals are expected
for the treatment of other forms of cancer in Canada and in several other countries such as
Japan and the United States. It is interesting to note that this process has taken over 20
years since the initial biological testing of the drug.
In spite of the fact that PhotofrinTM n a s \)Qen approved for clinical use, it exhibits
several drawbacks. Firstly, as stated before, the purified portion of HpD that has been
patented as PhotofrinTM i s a complex mixture of porphyrin oligomers. This complexity
makes characterization and basic physical studies very complicated. Secondly, the
wavelength of maximum absorption is 630nm, a wavelength at which effective light
penetration through skin is low (ca. 4mm)43. Large tumours or tumours which are more
deeply-seated within the body, are difficult to treat with this drug. Thirdly, the clearance
rate of PhotofrinTM f r o m m e body requires 4-6 weeks post-injection^^ for systemic
concentrations to fall to acceptable levels, during which time the patient must remain in
subdued light to prevent skin phototoxicity.
The above detrimental characteristics of Photofrin have led scientists in this multi-
disciplinary field to look for more suitable PDT agents. Among the most important
requirements of a drug for use in PDT are the following: 1. The potential drug must be an
effective photosensitizer i.e the drug, upon activation by light, must be able to bring about
destruction of cancer cells. 2. The drug must have an appreciable absorption in the visible
region at wavelengths greater than 630nm where the depth of light penetration through the
20
skin is good. 3. The drug must be structurally and chemically defined and consist of,
ideally, a single isomer. 4. The drug should preferentially localize in the tissues for which
this therapy is being sought. 5. A successful drug must clear from the body in a relatively
short time post-irradiation to reduce the patients' skin sensitivity to light. 6. The drug
must have no 'dark' toxicity i.e administration and uptake of the drug should have no
adverse effect on the body other than the desired photodynamic effect. Along with these
important chemical, photochemical and biological features are some practical considerations
such as ease, yield, and cost of production.
1.5.2 Benzoporphyrin Derivative Monoacid Ring A (BPDMA)
In an effort to improve on the first-generation drug, new second-generation
photosensitizers were sought out. The aforementioned concerns of Photofrin had to be
addressed. In 1984, Morgan et. al.45 synthesized a new class of dihydroporphyrins
(chlorins) which were named benzoporphyrin derivatives (BPD) due to their structural
similarity with benzoporphyrins. The maximum absorption band appears at 690nm, a
wavelength at which the effective depth of penetration of light is twice as high as that at
630nm46. When benzoporphyrin derivatives (Figure 1.17) were initially tested against
various healthy and malignant cell lines (in vitro testing)47? it Was determined that the
dimethyl esters were virtually inactive and the diacids (BPDDA and BPDDB) showed ten to
seventy times the cytotoxicity of hematoporphyrin on a number of healthy and malignant
cell lines. Surprisingly, the cytotoxicity data for the two monoacid/monomethyl ester
derivatives, showed that the ring A and B monoacids (BPDMA and BPDMB in Figure
1.17, only one regioisomer shown) were five times more cytotoxic than the corresponding
diacids. Given these results, BPDMA was chosen as the drug for more extensive testing.
21
Me02C C0 2Me
Me02C
Me02C »»••:
Figure 1.17 The Ring A and Ring B Monoacid/Monoesters and Diacids of BPD (Only One Enantiomer of Each is Shown)
22
BPDMA, while not consisting of a single component, is a mixture of separable
regioisomers each consisting of a pair of enantiomers. As a result, characterization and
study of the BPD's is much easier than with Photofrin. Of particular importance clinically
is the fact that skin photosensitivity drops to near normal 72 hours post-injection^ using
BPDMA. The quantum yield of singlet oxygen has been determined to be 0.78^9 in
homogeneous solution and 0.46^0 m in vivo systems.
Several other groups in academia and industry are actively developing other second-
generation compounds to be used as photosensitizers in PDT. The following describes the
major candidates presently being evaluated in clinical trials.
1.5.3 Major Second-Generation Photosensitizers
Mono-L-aspartyl chlorin eft; This water-soluble dihydroporphyrin is currently
undergoing clinical trials with the support of Nippon Petroleum. The chlorin skeleton is
derived from chlorophyll a and the absorbance maximum is 664nm. The quantum yield of
singlet oxygen production is 0.70^ . (Figure 1.18 (a))
Tin Etiopurpurin: This lipophilic drug is the latest candidate to enter into clinical trials.
Supported by PDT Pharmaceutical, this chlorin-type molecule has a maximum absorbance
at 650nm and a singlet oxygen quantum yield of 0.65^2. (Figure 1.18 (b))
Meso-tetra(m-hydroxyphenyl)chlorin: Clinical testing is presently being performed
on this drug with the support of Scotia Pharmaceuticals. The absorbance maximum is
650nm. The quantum yield of singlet oxygen has not been accurately determined. (Figure
1.18 (c))
23
Figure 1.18 Major Second-Generation Photosensitizers: a) Mono-L-Aspartyl Chlorin e6; b) Tin Etiopurpurin; c) Meso-tetra(m-hydroxyphenyl)chlorin
1.5.4 Third Generation Photosensitizers
With three of the major problems associated with Photofrin having been addressed
with BPDMA i.e improved wavelength of light activation, improved clearance from the
body, and ease of characterization, there is interest in developing third-generation drugs.
While these proposed drugs will retain the aforementioned qualities of BPDMA, the desire
is to develop compounds that will have a higher affinity for cancerous versus healthy
tissue. Biodistribution data shows that BPDMA, while not accumulating specifically in
cancerous tissue, is preferentially retained in cancerous versus healthy tissue^ j We are
24
interested in compounds that will dramatically increase this preferential binding so that; 1.
The amount of healthy tissue destroyed during PDT will be minimized; 2. The drug
dosage can be decreased; 3. The light dosage level can be decreased.
There has been some research into the determination of which substituents of
photosensitizers give better biodistribution results. Furthermore, the work that has been
performed has not been standardized to any great extent and comparisons between parent
photosensitizers is inconclusive. Also, because the present second-generation drugs have
quite varied structures, a component of one that provides better biological results may not
provide the same for another class of photosensitizer. As a result, it is hard to predict a
priori which peripheral modifications to BPD dimethyl ester will yield improved
localization in cancerous tissue. Nevertheless, incorporation of the moieties which have
shown good results in other drugs is a logical place from which to start. Chapter 2
describes the components of other drugs which have been shown to provide good
biodistribution results and their attachment onto the periphery of BPD.
1.5.5 The Photodynamic Effect—Singlet Oxygen Production
1.5.5.1 Introduction
Although photodynamic therapy (PDT) involves the production, and subsequent
reaction, of singlet oxygen and other reactive species, it is not known to what extent these
species are responsible for the photodynamic effect. Tumour cytotoxicity has been
ascribed to direct reaction of reactive oxygen species and also to indirect effects related to
vascular collapse54-56 However, van Lier^?, and many others, believe that singlet
oxygen is the principle cytotoxic species responsible for the tumour response during PDT
of experimental animal tumours. In recent experimentation with hematoporphyrin
derivative (HpD), in molecular oxygen-poor systems, cells were found to be resistant to
PDj58 Clearly, the presence of oxygen and the subsequent production of singlet oxygen
25
is extremely crucial to the success of PDT. The following sections describe the various
photoprocesses that a photosensitizer can participate in and the methodology used to
qualitatively determine the production of singlet oxygen by a number of new derivatives of
ring B BPD 1,3-diene dimethyl ester.
1.5.5.2 Excited States of Photosensitizers
Photodynamic therapy requires a light activated photosensitizer (defined as a
compound capable of substrate modification after activation by light) to create a cytotoxic
form of oxygen called singlet oxygen. A modified Jablonski diagram (Figure 1.19)
describes the possible photoprocesses that a sensitizer can undergo. The sensitizing
compound in its singlet ground state (So) initially absorbs a photon to place it in an excited
state. The resulting first excited singlet state (Si) has a short lifetime (c.a. l-100ns) and
very few photosensitized reactions are mediated by this stated A compound in this state
can lose its energy radiatively through fluorescence (Si >Srj + hv) or through internal
conversion (Si >So + heat) and return to its ground state.
Of particular importance with regard to PDT is that the excited species can also
undergo a non-radiative process called intersystem crossing (ISC) (Si >Ti + heat)
which requires fast spin inversion resulting in the creation of the first excited triplet state
(Ti) of the sensitizer. The effectiveness of a particular photosensitizer is dictated by the
efficiency of this process as given by the ISC quantum yield i.e. the ratio of the triplet
sensitizer formed to the singlet sensitizer initially formed. For example, the ISC quantum
yield for the successful photosensitizer BPDMA has been determined to be 0.7960
Once the sensitizer is in the desired triplet state, it can return to the singlet ground
state radiatively via phosphorescence (Ti >So + hv) or through internal conversion
(Ti——>So + heat). However, the lifetime of the triplet state is typically much longer
than the singlet state (in the microsecond-millisecond range)**1 and these species can
26
1. Absorption of Light 2. Fluorescence 3. Internal Conversion 4. Intersystem Crossing (ISC) 5. Phosphorescence 6. Singlet Oxygen Production via Interaction of
Triplet Sensitizer and Triplet Oxygen
Figure 1.19 Modified Jablonski Diagram for a Typical Photosensitizer
27
interact with large numbers of other molecules in this time. For example, in the case of
BPDMA, the triplet state lifetime is ca. 25p:s62. These interactions can bring about two
principal types of reactions: 1. Electron or hydrogen transfer reactions such as with a
reducing substrate (Type I photoprocesses) and; 2. Energy transfer reactions with ground
state oxygen (Type H photoprocesses). The prevalence of these processes is dictated by
many factors such as substrate and oxygen concentration and both may compete in a given
system.
1.5.5.3 Type I Photoprocesses
In many cases, a sensitizer (SENS) in its excited triplet state (3SENS) can abstract
electrons or hydrogen atoms from a particular substrate (SUB) more readily than in its
singlet ground state.
3SENS + SUB > (SENS-)" + (SUB-)+
3SENS + SUB > SENSH+SUB-
These free-radical products (both the substrate and sensitizer) are very reactive and can
react in a number of ways. The semi-reduced forms of the sensitizer can react with ground
state oxygen to produce the reactive superoxide radical anion, ( O 2 O " (or its protonated
form, H O 2 O and in turn return to its ground state**3.
(SENS-)" + 3 02 > SENS + (02-)"
SENSH- + 3 02 > SENS + H02-
In addition, two molecules of the semi-reduced sensitizer can react to generate one molecule
of ground state sensitizer and one fully reduced. This latter species can react with triplet
28
oxygen to create hydrogen peroxide and the sensitizer. A multitude of other reactions can
also occur some of which involve the consumption of the sensitizer and some that
regenerate the sensitizer in its ground state. An example of a Type I process is the reaction
of 2-methyl-l,4-naphthoquinone (MQ) with 3(}-hydroxy-cholest-5-ene (cholesterol) in the
presence of oxygen (Figure 1.20). The epimeric 7a- and 7(3-hydroperoxycholesterols are
the main products.
7-oc-hydroperoxycholesterol 7-(5-hydroperoxycholesterol
Figure 1.20 The Type I Reaction of 2-Methyl-l,4-Naphthoquinone (MQ) with Cholesterol in the Presence of Light and Oxygen
29
The Type I Photoprocess has been exploited recently in the treatment of psoriasis
using 8-methoxypsoralen and UV-A radiation. It is thought that the psoralin undergoes a
Type I photoaddition to thymidine bases of DNA*^.
1.5.5.4 Type II Photoprocesses
Type II reactions are reactions whereby the triplet sensitizer imparts its energy to
ground state triplet oxygen (one of the few stable ground state triplet molecules) to
regenerate the sensitizer in its ground state and to produce singlet oxygen. Because there is
no net change in spin in this reaction, the reaction is very efficient (i.e. this is a 'spin-
allowed' reaction). Oxygen can exist in two singlet states; the first is an extremely short
lived (<10"H sec), higher energy (37.5kcal/mol) species and the second is a longer-lived
(ca. 4|is in water), lower energy (22.5kcal/mol) species*>5. Due to the lifetime of the
former species, the latter form of singlet oxygen is believed to be the only species involved
in the photosensitization reactions.
3SENS + 3 02 > SENS + ^ 2
Singlet oxygen can react with biological substrates in several ways. Figure 1.21
shows the three major classifications*'*' and examples of these reactions.
Hydrogen Abstraction and Addition (Ene Reaction)
HO OOH
5a-Hydroperoxycholest-6-en-3pVol
Cycloaddition Reactions
N H 3+
COO'
N H 3+
[2+2]
xo2
COO"
N
N H 3 *
[4 + 2] / \ \ COO"
'o2
Tryptophan Endoperoxide
N H 3+
/ \ COO"
o—o Histidine Endoperoxide
Oxygenation
C H 3 J COO"
Jo2
o*
C H f + T COO'
Methionine Sulfoxide
Figure L21 Examples of the Three Major Classes of Type II Reactions
31
1.5.5.5 Determination of Singlet Oxygen Production
One of the assumptions made when planning the derivatization of BPD was that
modification to the periphery would not appreciably alter its photophysical properties, in
particular its ability as photosensitizer. While this assumption is more than likely valid,
scientific experimentation was necessary to be certain. Recent experiments have shown
that the quantum yield of singlet oxygen production from the triplet state for BPDMA is
nearly unity in single solvent systems0^ and 0.46 in biological systems****. The easiest
way to determine if this quality has been retained is, then, to check that the new derivatives
do indeed produce singlet oxygen.
Quantitative determination of singlet oxygen production and quantum yield can be
performed in a number of ways, all of which are, unfortunately, beyond the scope of this
thesis. Upon perusal of the literature, an experimental technique was uncovered that could,
with a good degree of certainty, provide a qualitative assessment of singlet oxygen
generation. Recently developed by van Lier69
this method allows singlet oxygen
determination via the derivatives of cholesterol formed when sensitizer, oxygen, cholesterol
and light are combined. Because Type I and Type II photoprocesses give different
cholesterol products, this methodology can be used to determine which photoprocess is
being utilized by a new photosensitizer. Also interesting to note is that because cholesterol
is a major constituent of human cell membranes, the reactions that occur give a first hand
look at the possible modes of cellular attack in more complicated biological systems. The
methodology is described in the experimental chapter.
It has to be stated that the results of this experimentation can only provide insight
into the mode of photosensitization in a homogeneous solvent: the actual photodynamic
effect in biological systems is almost certainly due to several mechanisms.
3 2
1.6 Fluorescence Quenching and Liposomes
1.6.1 Experimental Challenge
An important property of BPDMA is its exceedingly poor solubility in water. This
property is also shared by almost all of the ring B BPD derivatives synthesized in this
work. This poses a problem with respect to biological testing because the testing done on
living animals and humans requires that the photosensitizer be administered by injection in
aqueous media. In order to solubilize BPDMA, liposomes are used to carry the
hydrophobic drug into the blood stream.
While there has been a great deal of research into which tissue BPDMA distributes
in living systems, little is known about the interaction of BPDMA with liposomes. An
important question then becomes "Is the localization of BPDMA due to the properties of the
liposome in which it is 'encapsulated' or due to BPDMA itself?". A related question is
"Does the BPDMA immediately diffuse out of the liposome upon contact with various
fractions of blood serum or does it remain encapsulated as it circulates throughout the
body?". Recent studies have shown that liposome-encapsulated BPDMA when mixed into
human plasma associates predominantly with various lipoproteins, in particular high- and
low-density lipoproteins (HDL and LDL)70. Also, preassociation of liposomal BPDMA
with LDL prior to intravenous injection led to better biodistribution results^. These
results support the findings that lipophilic photosensitizers are delivered to tumours in part
via an LDL receptor-mediated pathway^. Other possible routes of delivery include
passive diffusion of the photosensitizer across the cell membrane and uptake by scavenger
receptors^. The delivery of liposomal BPDMA, or of any drug, to various regions of the
body is extremely complex.
In an effort to shed some light on the questions posed above, it was felt that the
determination of how BPDMA and other derivatives interact with liposomes might be
useful. If BPD derivatives interact with liposomes in different ways, this might lead to
33
differential interaction with lipoproteins and point to why some derivatives biodistribute
better than others.
There has been some research into determining the behaviour of various
compounds, primarily biologically important molecules or drugs, within liposomes^^-78
One method of experimentation employs the use of fluorescent 'markers' imbedded within
liposomes. These markers, or "probes" as they are commonly referred to, are chosen so
that they fluoresce at a wavelength that corresponds to an absorption peak of the
compound. As the compound diffuses into the liposome, the intrinsic fluorescence of the
probe is reduced or "quenched" by the compound if it is in close proximity to the probe.
Changes in the degree and mode of this fluorescence quenching can provide information
about how different compounds interact differendy with the liposome species.
1.6.2 Liposomes
Liposomes, first described by Bangham et. a l 7 9 in 1965, are spherical structures
comprised of single or multiple bilayers (lamella) of phospholipids (or similar compounds).
A phospholipid consists of a polar head group (charged or neutral) and two non-polar acyl
chains of varying length and degree of unsaturation. These molecules are amphiphilic,
with the head group preferring polar solvents and the acyl chains preferring non-polar
solvents. When phospholipids are placed in an aqueous solution and left to equilibrate, the
phospholipids are oriented such that the contact of the acyl side chains and the bulk
aqueous phase is minimized. The only orientation which provides this feature is a
liposome (Figure 1.22).
33
differential interaction with lipoproteins and point to why some derivatives biodistribute
better than others.
There has been some research into determining the behaviour of various
compounds, primarily biologically important molecules or drugs, within Hposomes74-78
One method of experimentation employs the use of fluorescent 'markers' imbedded within
liposomes. These markers, or "probes" as they are commonly referred to, are chosen so
that they fluoresce at a wavelength that corresponds to an absorption peak of the
compound. As the compound diffuses into the liposome, the intrinsic fluorescence of the
probe is reduced or "quenched" by the compound if it is in close proximity to the probe.
Changes in the degree and mode of this fluorescence quenching can provide information
about how different compounds interact differendy with the liposome species.
1.6.2 Liposomes
Liposomes, first described by Bangham et al.79 in 1965, are spherical structures
consisting of single or multiple bilayers (lamella) of phospholipids (or similar compounds).
A phospholipid consists of a polar head group (charged or neutral) and two non-polar acyl
chains of varying length and degree of unsaturation. These molecules are amphiphilic,
with the head group preferring polar solvents and the acyl chains preferring non-polar
solvents. When phospholipids are placed in an aqueous solution and left to equilibrate, the
phospholipids are oriented such that the contact of the acyl side chains and the bulk
aqueous phase is minimized. The only orientation which provides this feature is a
liposome (Figure 1.22).
34
Cross-Section of Unilamellar Liposome
w h e r e o phospholipid polar headgroup
= hydrophobic acyl chains
Figure 1.22 General Liposome Structure
When phospholipids are dispersed in an aqueous phase, a heterogeneous mixture of
vesicles (liposomes) is generally formed. The components of this mixture can be
differentiated by diameter and the number of bilayers (the 'lamellarity'). All liposomes
having more than one bilayer are called multilamellar vesicles (MLV). Liposomes with one
bilayer are classed as either small unilamellar vesicles (SUV) or large unilamellar vesicles
(LUV), with a diameter of lOOnm being the general cut-off, although this is somewhat
ambiguous.
Liposomes can be simply formed in the laboratory by first dissolving the
phospholipid in an organic solvent and depositing the lipid on the sides of a flask by
removal of the solvent under vacuum. The resulting thin film is then hydrated by
introducing an aqueous solution followed by agitation of the flask. The liposomes thus
formed tend to be MLV with diameters in the range of several hundred to several thousand
3 5
nanometers* . Several factors such as hydration time, temperature and lipid composition
dictate the actual size distribution in a given system.
SUV can be prepared by sonication of a MLV suspension to generate liposomes in
the 25-50nm range. These liposomes can also be prepared using a French press which
forces a MLV suspension through a small orifice at 20,000psi. Repeated extrusion yields a
reproducible population of SUV again with diameters in the 30-50nm range**1.
LUV can be prepared in a number of ways including dialysis, column
chromatography, and centrifugation but these are all relatively time-consuming and/or
unreproducible. A new method has been recently introduced, however, that allows the
production of a relatively homogeneous population of liposomes in a reproducible and
facile way**2, A MLV suspension is placed in an extrusion apparatus and filtered under
pressure (up to 800psi) through a defined pore size polycarbonate filter (100, 200 or
400nm pore size). Repeated passes through these filters yields vesicles with diameters that
agree well with the pore size filter used.
1.6.3 Liposomes as Drug Delivery Agents
Liposomes have been researched as potential drug delivery agents in the past 20
years. Initially, it was thought that the encapsulation of a drug into liposomes followed by
intravenous injection would "disguise" the drug and thus prevent it being targeted by the
reticulo-endothelial system (RES). The RES is a critical part of the bodies defense system
which removes foreign matter as well as particulates, dead cells, and microorganisms.
Unfortunately, much of an administered dose of liposomes is removed by the RES** 3.
There are new liposome compositions called 'Stealth' liposomes which have been designed
to bypass the RES and thus circulate more effectively throughout the body84
Liposomes, are chosen as drug delivery agents for a number of reasons. One of
these is the fact that certain liposome compositions can entrap large amounts of water
36
soluble drug (up to 30 litres per mole lipid)85. The hydrophilic drug is dissolved in the
aqueous solution used to hydrate the lipid film and incorporated upon liposome formation.
Once injected into the body, these liposomes can deliver the drug over a sustained period of
time by slow diffusion across the liposome membrane, through perturbation by proteins or
via enzymatic degradation^.
Hydrophobic drugs can also be formulated within liposomes. The drug is initially
dissolved with the phospholipid in an organic solvent and the film is created. Hydration
with an appropriate aqueous solution yields liposomes with the compound imbedded within
the hydrophobic acyl chains. Compounds not normally soluble in aqueous systems are
now rendered water-soluble.
1.6.4 Fluorescence Quenching
The interaction of hydrophobic drugs with liposomes can be investigated in a
number of ways. One such methodology is fluorescence energy transfer or fluorescence
quenching. A fluorescent molecule (probe) is embedded within a population of liposomes
and aliquots of the fluorescence quencher (drug) dissolved in a small amount of solvent are
added. The use of this method requires that the drug absorb light (with high extinction
coefficient) at or near the wavelength of fluorescence emission of the probe^. As the
aliquots of drug are added, the energy of the probe is imparted to the drug (primarily via
dipole-dipole interaction^) only if the drug is within roughly lOnm of the probe^. The
fluorescence of the probe is quenched and the observed fluorescence is reduced.
There are two main types of fluorescence quenching, dynamic (collisional) and
static (complex forming)90. Dynamic quenching results when the quencher diffuses to the
probe during the lifetime of the excited state. A number of equations relate the
concentration of added quencher and the resulting collisional quenching of probe
fluorescence. The most simple of these is the classical Stern-Volmer equation:
37
F Q / F = 1 + kqXotQ] = 1 + K D [ Q ]
where F 0 and F are the fluorescence intensities in the absence and presence of quencher,
respectively, kq is the bimolecular quenching constant, x 0 is the fluorescence lifetime of the
probe, [Q] is the concentration of quencher and K D = k q X 0 is the Stern-Volmer quenching
constant.
In the case of static quenching, the quencher and probe form a non-fluorescent
ground state complex. Upon absorption of light, the complex returns to the ground state
without photon emission. This type of quenching is related to concentration of quencher
using the following equation:
F Q / F = 1 + K S [ Q ]
Here the fluorescence intensity and concentration terms are the same as for dynamic
quenching but the quenching constant is replaced by the association constant K S . This term
is defined as follows:
K S = [P-Q]/[P][Q]
where [P-Q] is the concentration of the probe-quencher complex and [P] and [Q] are the
free probe and quencher concentrations.
In homogeneous systems that contain a single class of fluorescent probes all equally
accessible to the quencher, fluorescence quenching can generally be interpreted by one of
these mechanisms. A plot of F 0 / F versus [Q] that yields a straight line, with y-intercept
equaling one, is indicative of this.
38
In more complex, non-ideal systems, a mixture of these quenching mechanisms can
take place. In these cases, the F 0 /F versus [OJ plots curve upward towards the y-axis^l.
It is quite difficult to separate the dynamic and static components in such systems but
modified Stern-Volmer equations have been constructed to account for this.
Regardless of the exact contributions of each type of quenching, the plots of Fo/F
versus [Q] can provide information about the location of a fluorescent probe or quencher
within a liposome. For example, long chain fatty acids have been modified so that the
same fluorophore is bound at various positions on the chain^2 (Figure 1.23). These fatty
acids were incorporated separately into dimyristoyl phosphatidylcholine liposomes and the
fluorescence of the 9-anthroyloxy moiety was quenched by addition of Cu(U). The results
showed strong quenching of the fluorophores bound near the liposome/water interface and
poorer quenching for the probes more deeply imbedded within the liposome. This
predominantly dynamic quenching enabled the assessment of the transverse location of the
variously bound fluorophores.
po 2 pcy p o 2 po 2- po 2-
Approximate Bilayer Center
Figure 1.23 Schematic Representation of n-(9-Anthoyloxy) Fatty Acids Showing
the Transverse Positions of the 9-Anthoyloxy Fluorescent Probe
39
The amount of fluorescence quenching is related to the proximity of quencher to the
probe and differential quenching of a lipid-bound probe with defined transverse location by
various hydrophobic drugs might provide insight into the way these drugs localize within
liposomes. However, a number of criticisms of this methodology have been expressed in
the recent literature. The first is that the location of fluorescent probe within the liposome is
ill-defined for many probes^3. How can one infer the localization of a particular drug
within the liposome if the location of the probe changes or is varied? Secondly, diffusion
of the probe between liposomes occurs for many probes. Thirdly, the incorporation of
fluorescent probe within a liposome may alter the local environment of the probe within the
liposome^, if these liposome/probe systems are to be models of pure liposomes, the
fluorophore must not markedly change the liposome properties.
Chapter 3 describes how these concerns were addressed in the selection of the
fluorescent probe used in this work as well as the incorporation of this probe into
liposomes. Further, it details the methods used to create and characterize the unilamellar
vesicles and the fluorescence quenching experiments performed with them using BPDMA
and four newly synthesized derivatives of BPD. The chapter is completed with discussion
of the results of these experiments followed by a summary and future research.
Chapter 2 Synthesis of BPD Derivatives:
Results and Discussion
41
2.1 Research Objective
Over the past three decades, there has been much interest in the chemical properties
and biological behaviour of reduced porphyrins, especially the chlorins. One of the
principal reasons for this is their potential use as drugs in the relatively new field of
photodynamic therapy (PDT). The first clinically tested drug, hematoporphyrin derivative
(HpD) and its purified form, PhotofrinTM^ w e r e described in some detail in the preceding
chapter. Given the recent inception of clinical PDT there has been a large amount of
biological testing, most of it conducted using HpD and PhotofrinTM. As a result, there has
been less research into the testing of second-generation drugs. This lack of structure-
activity relationships is one of the principal stumbling blocks hindering further
advancement of the field. With this said, the research that has been done has provided
some insight into the compounds which give the best biological activity.
The brief history of the benzoporphyrin derivatives developed in the introductory
chapter described the cytotoxicities of the ring A and ring B BPD compounds. It was
shown that the monoacid derivatives of ring A and ring B BPD had the best initial
biological results. Because the amount of biological testing required for approval of a new
drug is staggering, further testing can only be performed on one compound. The ring A
isomer of the benzoporphyrin derivative monoacid (BPDMA) was somewhat arbitrarily
chosen as the lone compound for the more comprehensive testing. However, in the
industrial production of BPDMA, an equal amount of the ring B material is produced as an
'unwanted' by-product and stored without further modification. Because the biological
results of ring B BPD are similar to ring A BPD and the ring B material can be acquired in
relatively large amounts, it was chosen as the compound for derivatization.
The object of this portion of research is two-fold. The first is to incorporate the
successful or promising chemical features of many recently published photosensitizing
agents into BPD in order to improve its biodistribution in malignant, versus healthy, tissue
42
or to provide selective intracellular binding properties. The success of a given derivative
will be expressed in results of in vitro and in vivo testing currently underway in this
laboratory.
Because the absorption maximum of BPD appears in a desirable region of the
visible spectrum, these peripheral modifications must add new functionality to the chlorin
backbone without substantially changing these desirable electronic properties. There are a
number of places on BPD where a desired functionality could be added and a number of
synthetic protocols were developed. This leads to the second object. There is a large
amount of in vitro and in vivo testing presently being conducted in this field and new data
on structure-activity relationships are being published constantly; therefore the synthetic
routes must be flexible enough to allow the addition of a wide range of substituents to
BPD.
The following sections describe the rationale behind the choices of the peripheral
modifications of BPD and detail the synthetic routes that were required to attach the
requisite groups.
43
2.2 Synthesis of Ring B BPD Dimethyl Ester
The crucial step in the synthesis of the benzoporphyrin derivatives is the Diels-
Alder reaction of protoporphyrin IX dimethyl ester (PPIX) and dimethylacetylene
dicarboxylate (DMAD). PPIX can be purchased from a number of sources but can also be
made in good yield starting from hematoporphyrin IX dipropionic acid. The latter method
of synthesizing PPIX was used in this laboratory.
Hematoporphyrin is initially heated in DMF to facilitate dehydration of the two
secondary alcohol groups to vinyl groups at positions 3 and 8, followed by esterification of
the propionic acid groups at positions 13 and 17 in methanol and acetic acid (Figure 2.1).
The two steps provide overall yields in the 60-80% ranged
Figure 2.1 The Conversion of Hernatoporphyrin K to Protoporphyrin IX Dimethyl Ester
The crude batch of PPIX was recrystallized from CH2Cl2/MeOH to give 60% of
the desired porphyrin. The identity of the pure compound was confirmed by *H NMR and
uv-vis spectroscopy prior to use.
Figure 22 Diels-Alder Reaction of Protoporphyrin DC Dimethyl Ester (PPDC) with Dimethylacetylene Dicarboxylate (DMAD)
u - • • — i — • — • — • — » _ i « » « — F i • < — • — • — i — • — • — . — . — • 400 500 600 700 800
WAVELENGTH (nm)
Figure 2.3 UV-Vis Spectra (CH2CI2) of Protoporphyrin DC Dimethyl Ester and BPD 1,4-Diene Dimethyl Ester
45
The PPIX formed above was reacted with an excess of dimethylacetylene
dicarboxylate (DMAD) in refluxing toluene (Figure 2.2) and the reaction progress was
monitored by tic and uv-vis spectroscopy. The uv-vis spectra of both the reactant and the
product are shown in Figure 2.3. The reaction was stopped after 27 hours when there
appeared to be no further change in the new absorption at 666nm. Tic showed two new
compounds running slower than the starting protoporphyrin. A tic cospot of the products
with known standards showed the slightly more polar compound to be the ring A isomer.
The yield of the above reaction is normally 20% for each of the regioisomers^^ and
isolation of enough material to carry out the desired derivatizations would be difficult.
Fortunately, the industrial preparation of BPDMA by Raylo Chemical involves separation
of the ring A and ring B BPD 1,4-diene dimethyl ester isomers directly after the DMAD
reaction. The ring B isomer is removed and stored without further modification. We were
thus able to procure gram quantities of this compound.
The Raylo ring B 1,4-diene dimethyl ester was found by tic to have substantial
impurities which were determined to be primarily the slightly slower moving ring A isomer
and smaller amounts of the initial protoporphyrin IX dimethyl ester. Purification by
chromatography on a large scale was unsuccessful due to the similarities between the two
regioisomers. After much experimentation, recrystallization from boiling ethyl acetate (ca.
30mL EtOAc per gram BPD) was found to give pure ring B isomer in 82% yield.
The pure ring B 1,4-diene derivative was treated with 1,8-
diazabicyclo[5.4.0]undec-7-ene (DBU) in dichloromethane to provide the BPD 1,3-diene
dimethyl ester in a 90% yield after chromatography (Figure 2.4). The reaction progress
was monitored by uv-vis spectroscopy which showed a new absorbance at 690nm
(Figure2.4). In the original paper^, when this rearrangement was carried out using
M e O , C C0 2Me
46
M e O z C C0 2Me Me02C C0 2 Me
WAVELENGTH (nm)
triethylamine (TEA) or DBU, two different products were formed that differed only in the
geometric arrangement of the carbomethoxy substituent attached at C-7* in relation to the
angular methyl group located at C-7 (Figure 2.4). After rearrangement of the BPD 1,4-
diene with TEA the methyl ester at C-71 and the methyl group at C-7 were in a cisoid
orientation. Treatment of the product derived from TEA rearrangement or the original 1,4-
diene with DBU yielded the conformer with the methyl ester and the methyl group in a
transoid orientation. This transoid orientation was confirmed by a positive nOe effect
observed for the C-7* proton and the methyl group at C-7. This data, coupled with other
findings, led the researchers to the conclusion that TEA rearrangement gave the kinetic
product and the DBU rearrangement gave the thermodynamic product. It is this
thermodynamically more stable, less strained conformer which is the desired starting ring B
BPD 1,3-diene dimethyl ester compound.
Before the peripheral modifications are described, the properties of BPD dimethyl
ester will be listed as a reference point for the syntheses that follow. The NMR is
shown in Figure 2.5. There are several important features to describe. The pyrrolic
nitrogens are shielded substantially from the external magnetic field and appear at roughly
-2.4ppm. Conversely, the protons located on the meso carbons (positions 5, 10, 15, and
20 using IUPAC) are deshielded by the aromatic ring current and appear between 9 and 10
ppm. These two sets of resonances bracket the signals of the remaining 38 protons. The
axial methyl group located at C-7 (1.78ppm, singlet), methyl ester-7* (2.95ppm, singlet),
methylene protons at 132 and 172 (3.16 and 3.20ppm, two overlapping triplets), methyl
groups-2, -12, and -18 and methyl esters at 132 and 172 (3.42, 3.47, 3.63, 3.65 and
3.67ppm, 5 singlets), methyl ester-72 (3.99ppm, singlet), methylene protons at 13* and
17* (4.17 and 4.29ppm, two triplets), proton at C-7* (5.07ppm, singlet), the vinyl group
at C-3 (6.17 (d), 6.37 (d), 8.11 (dd)), and finally protons 7 3 and 7 4 (7.45 and 7.83, two
doublets).
48
HDNva osav
49
The absorption spectrum of BPD dimethyl ester is shown in Figure 2.5. It is
characteristic of a chlorin but it has an unusually broad Soret band and the lowest energy
absorption at 690nm is bathochromically shifted some 30nm in comparison with typical
chlorins. This is due to the extended conjugation provided by the benzene-like moiety
attached to the chlorin periphery at carbons 7 and 8. Not only does electronic delocalization
occur through the 1,3-diene but it extends through to the carbonyl of the ester functionality
at C-7 2. Also interesting is the ratio of intensities of the Soret to the absorption at 690nm
which is roughly two to one as opposed to the five to one ratio typically observed for
chlorins.
BPD dimethyl ester is best solubilized in chlorinated organic solvents (chloroform
and methylene chloride) and dimethyl sulfoxide and is somewhat less soluble in ethyl
acetate, tetrahydrofuran, acetonitrile, and acetone. It is poorly soluble in methanol,
ethanol, benzene and toluene and virtually insoluble in water and hexanes. Clearly there is
solvation competition between the relatively non-polar aromatic chlorin backbone and the
more polar carboxylic ester functionalities.
50
2.3 Synthesis of Variable Alkyl Chain Ester Derivatives of BPD
2.3.1 Rationale
Although it is not established why certain porphyrins localize preferentially in
tumourous versus healthy tissue, many researchers believe that the polarity of peripheral
substituents plays an important role. To verify this, there have been recent syntheses of
photosensitizers with peripheral substituents of various alkyl chain lengths. Evensen and
others^ synthesized derivatives of hematoporphyrin which have ether linkages of variable
length. Figure 2.6 shows the various compounds. It is important to note that each of these
compounds is a mixture of diastereomers. These researchers performed testing of these
derivatives on a mouse mammary tumour and showed that inhibition of tumour cell growth
increased with increasing alkyl
Where R= H C H 3
C 2 H 5
C 3 H 7
C4H9
Figure 2.6 Alkyl Ether Derivatives of Hematoporphyrin
chain length. The methyl and ethyl alkyl chain ether compounds showed little or no activity
which supports previous testing of derivatives of hematoporphyrin99.
51
To follow up on this previous work, similar peripheral modifications were made to
methyl pheophorbide a and chlorin e(j trimethyl ester (two chlorophyll derivatives isolated
from Spirulina alga) by Pandey et. al .* u u (Figure 2.7). These compounds, again mixtures
of diastereomers, were injected into mice bearing transplanted tumours and irradiated with
665nm light. The amount of tumour damage was subjectively determined at various times
post-injection for a range of dosages and compared with Photofrin. Both unsubstituted
chlorin e(j and methyl pheophorbide a showed poor tumour response but the corresponding
hexyl ether derivatives showed responses comparable to Photofrin at reduced dosage
levels.
H H
0 2Me
R- C 3 H 7 , C6 H 13 R- C 6 H 1 3
Figure 2.7 Alkyl Ether Derivatives of Methyl Pheophorbide a and Chlorin e6 Trimethyl Ester
5 2
2.3.2 Via the BPD Vinyl Group
Alkyl ether derivatives of ring B BPD have recently been synthesized by Meunier
and others are currently undergoing biological testing as potential PDT agents and as
such were not resynthesized. Other means of attaching these alkyl chains were utilized.
Hydrobromination of the vinyl group of tetrapyrroles followed by nucleophilic
attack is well known in porphyrin chemistry. It was envisioned that if ammonia was used
as the nucleophile, the resulting amine could be used to add alkyl groups via amine or
amide linkages. However, the methyl ester at C-71 and the axial methyl group C-7 are in a
transoid configuration, and there are two such orientations which allow for this. The
starting material is thus an inseparable mixture of enantiomers and the introduction of a
chiral center will lead to a diastereomeric mixture which should be separable (Scheme 2.1).
The relevant features of the BPD X H NMR spectrum are the resonances resulting
from the vinyl group (Figure 2.8). The resonances are typical of an ABX system where
the doublet centered at 6.17 ppm (J=12Hz) is assigned to the H R (cis-coupled to Hx), the
doublet centered at 6.37 ppm (J=18Hz) assigned to H A (trans-coupled to Hx) and the
doublet of doublets centered at 8.11 ppm (Jcis=12Hz and Jtrans=18Hz) is assigned to Hx-
The geminal coupling between H A and H R is not seen in the case of BPD dimethyl ester
although the doublets are somewhat broadened. In the spectra of some derivatives of BPD
this coupling is seen and is roughly 1 Hz.
H B 6.17 (d, J=12Hz)
, H X 8.11 (dd, J=12Hz, 18Hz) z) 6.37 (d, J=18Hz) Hjf ^
Figure 2.8 The Chemical Shifts of the Vinyl Group of BPD Dimethyl Ester
Scheme 2.1 The Conversion of BPD Dimethyl Ester (2) to BPD Aniine (3) and the Resulting Stereoisomers
54
Treatment of a methylene chloride solution of BPD dimethyl ester (2) with
anhydrous HBr under a nitrogen atmosphere at 0°C yielded a bright green solution which
was immediately transferred to a saturated anhydrous NH3/THF solution via cannula.
Attempts to isolate the BPD hydrobromide prior to displacement led to the formation of
several products. The two-step reaction provided the desired BPD amine (3) (as a
diastereomeric mixture) with amination occurring at the C-3* (Markovnikov) position in
75% yield with a minor product being recyclable starting material. Low-resolution mass
spectroscopy gave the parent ion. Uv-vis spectroscopy showed a slight hypsochromic
shift of the maximum absorption band from 690nm to 686nm (due to the loss of
conjugation with the vinyl group) and the disappearance of the characteristic vinyl
resonances in the NMR. Because of the presence of diastereomers, full characterization
by lH NMR was difficult. Separation of the two isomers was extremely difficult by
chromatography, but a solvent system was developed that allowed qualitative separation.
Preparative thin-layer chromatography was attempted and was largely unsuccessful.
Given the unforeseen difficulty in separating the diastereomeric mixture, this
synthetic route to variable chain length BPD derivatives was abandoned. Certainly the
compounds could be reacted further but with the difficulty in separation, and the resulting
low yields of the separated diastereomers, other routes were explored.
2.3.3 Via Transesterification
Because the propionate esters of BPD have shown good reactivity towards acid-
catalyzed hydrolysis (the chemistry which enables synthesis of BPDMA), it was thought
that these methyl esters could be selectively transesterified with alcohols of increasing chain
length. This would introduce successively longer alkyl chain lengths at two peripheral
positions without generating any new stereoisomers.
where R= C 3 H7 (4) C 4 H9 (5) C 5 H 1 t (6) C 6 H 1 3 (7) C7H 1 5 (8) C 8 H 1 7 (9)
Scheme 2.2 General Transesterification of BPD Dimethyl Ester (2)
Toward this end, a general transesterification methodology was developed (Scheme
2.2). Experimentation led to the use of concentrated sulfuric acid as the best catalyst and
several hew ester-linked compounds were synthesized. In each case, BPD dimethyl ester
(2) was stirred with a large excess of the straight chain alcohols and a small amount of
dichloromethane (to solubilize the reagents) and a few drops of concentrated sulfuric acid
were added. The reaction time was on average three days at room temperature in the dark.
BPD dimethyl ester (2) was transesterified (with yields in parentheses) to BPD dipropyl
ester (4) (98%), BPD dibutyl ester (5) (98%), BPD dipentyl ester (6) (98%), BPD
dihexyl ester (7) (97%), BPD diheptyl ester (8) (92%) and BPD dioctyl ester (9) (93%).
Al l compounds were characterized by elemental analysis, A H NMR, and high resolution
spectroscopy.
The lU NMR and uv-vis of (6) are shown in Figure 2.9. One of the interesting
features of the NMR spectrum is the chemical shift of the terminal methyl groups of the
pentyl chains. Centered at 0.61ppm, this multiplet is shifted upfield roughly 0.25ppm
5 7
from where typical terminal methyl groups appear. This may point to the interaction of
these chains with the ring current generated by the chlorin. As the alkyl chains extend over
the chlorin, the diamagnetic ring current shields these protons from the external magnetic
field resulting in an upfield shift of the resonance. The uv-vis spectrum shows little or no
change from the starting dimethyl ester, which supports the assumption that modification of
the propionate ester chains of BPD will not appreciably change the BPD chromophore.
2.3.4 Via Amide Formation
The synthesis of amide-linked alkyl derivatives arose out of some concerns that
have been expressed about ester-linkages in the recent literature. Pandey et. al . l u2 and
Moan et. a l . * u 3 have theorized that hydrolysis or cleavage of ester-linkages may occur in
vivo. While this has not been quantitatively determined for photosensitizers, it is certainly
feasible that one or both of these reactions can occur in biological systems. Amide-linkages
are much less susceptible to hydrolysis than ester-linkages m us several variable
chain length diamide derivatives of BPD were synthesized to incorporate the desired
functionalities. Comparison of the biological results of these amides with the ester-linked
compounds should provide some insight into this question.
2.3.4.1 Direct Amidation
CH3-<N
CH302( 02CH3
CH3
02Me
RNH2, THF, 70°C
CH3
02Me
where R= C6H13 (10) C8H17 (11)
Scheme 2.3 Direct Amidation of BPD Dimethyl Ester (2)
58
A look at the literature showed that direct nucleophilic displacement of methyl esters
to the corresponding amides is quite successful using primary amines 1^. Thus, straight
chain primary amines were used. In the reactions a large excess of dry amine was heated
with BPD dimethyl ester (2) in freshly distilled THF (Scheme 2.3). The reaction time was
generally two days with the temperature roughly at 70°C. In each case, a large amount of
baseline material was noted on tic which was initially hard to rationalize due to the fact that
the reaction conditions were relatively mild. However, inspection of this poorly soluble
brown material by uv-vis spectroscopy showed a rhodo-type spectrum indicative of a true
benzoporphyrin. The generation of a product with similar spectroscopic characteristics has
been discussed in the literature 106 and was seen when BPD was treated with base for
prolonged periods. The compound is believed to be the benzoporphyrin derived from loss
of the axial methyl group at position 7 with subsequent rearomatization to the porphyrin.
The BPD diamides were chromatographed on silica gel and this baseline material was
retained at the top of the column. Small amounts of slightly faster moving material were
collected (the two possible monoester-monoamides). The slowest fractions were the
desired diamides. In this way, the following derivatives were synthesized (with
accompanying yields): BPD dihexyl amide (10) (49%) and BPD dioctyl amide (11)
(49%). Characterization of these compounds was by high resolution spectroscopy and 1 H
NMR.
2.3.4.2 V ia Nucleophilic Displacement
Given the mixed success of the previous method of direct displacement, a better
methodology was sought. Scheme 2.4 shows the reaction sequence. It has been shown
that the methyl esters of BPD can be selectively hydrolyzed in aqueous acid to the two
possible monoester/monoacid compounds (BPDMA and BPDMB) and if the hydrolysis is
59
performed at higher temperature, the corresponding diacid compound is formed1^. This
was indeed the case when the hydrolysis was allowed to proceed at room temperature in
contrast to the hydrolysis to BPDMA/BPDMB which proceeds at 0°C. The hydrolysis
was monitored by tic and because the chromophore of BPD does not change appreciably
when modifying the propionate chain esters, semi-quantitative assessment of the reaction
progress was possible by Uc. The reaction was quenched when all the starting material was
consumed, leaving a small amount of the two possible monoacids and primarily the desired
BPD diacid (12). The reaction was stopped at this stage because allowing the hydrolysis
to proceed further generates a slighdy slower moving compound which appears to be the
BPD triacid with the ester at C-7 2 being the third site of hydrolysis. This compound was
difficult to remove at later stages of synthesis and was therefore avoided. Attempts to
purify the BPD diacid after workup were largely unsuccessful but a small sample was
purified for characterization (AH NMR and high resolution mass spectroscopy) using
preparative Uc. In general, the crude diacid was carried over to the next reaction.
Once the diacid was synthesized it was reasoned that if the conversion could be
made to the BPD diacid chloride (35), we could not only generate the desired straight
chain diamide derivatives but a very powerful methodology by which a large number of
derivatives could be made. Assessment of the various reagents available led to the use of
oxalyl chloride for a number of reasons. The oxalic acid by-product decomposes to CO
and CO2 which drives the reaction to completion. Also, any excess reagent can be
removed at fairly low temperature (63°C) 1 U 8. Treatment of the crude diacid (12) with
oxalyl chloride in refluxing methylene chloride led to a bright green solution due to the
protonation of the pyrrolic nitrogens by the acid generated in the reaction. Removal of the
excess oxalyl chloride and solvent under nitrogen left a crude residue that was taken up in
methylene chloride and reacted directly with several dry amines to form the corresponding
BPD diamides. For this displacement reaction to be successful, four equivalents of the
60
(2) (12)
(COCI)2, CH 2CI 2, reflux
where R= CH 3 (13) C 4 H 9 (14)
Scheme 2.4 Amidation of BPD Dimethyl Ester (2) Via Displacement of BPD Diacid Chloride (35)
61
particular amine were required; 2 equivalents to deprotonate the pyrrolic nitrogens (to
return the chlorin to its dull green coloured, free-base form) and two equivalents for the
acid chlorides. In practice, enough amine was titrated in to deprotonate the nitrogens and
then an amount slightly exceeding this was added to finish the reaction.
Column chromatography allowed separation of the diamide from the faster moving
monoester-monoamides generated in the reaction. In this way, the following derivatives
were synthesized (with accompanying yields based on the starting BPD dimethyl ester
(2)): BPD dimethyl amide (13) (67%) and BPD dibutyl amide (14) (55%). The yields
of these three step reactions were comparible to the direct displacement methodology.
The A H NMR and uv-vis spectra of BPD dibutyl amide (14) appear in Figure
2.10. The main differences in the NMR spectrum of the dibutyl amide from the dimethyl
ester is the loss of the two methyl ester peaks located at roughly 3.5ppm and the addition of
the amide NH protons that appear as triplets at 6.27 and 6.71ppm. An interesting feature is
the upfield shift of the alkyl chain resonances, in particular the terminal methyl groups
which appear at 0.32 and 0.51ppm. This is again presumably the result of interaction of
these methyl groups with the diamagnetic ring current of the chlorin backbone as the alkyl
chains are long enough to fold over the planar macrocycle. The uv-vis spectrum is
essentially the same as the starting dimethyl ester in keeping with the initial assumption that
the chromophore of BPD will not be changed appreciably by these modifications.
62
63
2.4 Synthesis of BPD Derivatives with Free Amine Functionalities
2.4.1 Rationale
Very little of the published research on peripheral modification of photosensitizers
is comprehensive in scope. Typically, a few modifications are made to an existing
photosensitizer and these compounds are tested in vitro on one or a few cell lines and, more
rarely, tested in vivo on one type of tumour. However, a series of publications by
Woodbum and others 109-111 h a v e described the synthesis and biological testing of several
new derivatives of protoporphyrin IX dimethyl ester in a comprehensive way. These
papers detailed the in vitro cytotoxicity of the new compounds, their subcellular localization
as well as how these derivatives target DNA in an in vivo model. The work appears to give
rise to some general correlation between the structural feature of the photosensitizer and
biological activity. The following discussion describes the promising features of these new
drugs that were incorporated into BPD.
Amine functionalities were attached to the periphery of protoporphyrin IX via its
propionate ester chains (Figure 2.11). These amines, which at physiological pH are
cationic, were found to target DNA in their in vivo model. This is a particularly important
result It is known that the production and subsequent reaction of singlet oxygen is a major
cause of cell-kill during photodynamic therapy. However, studies have shown 112 m a t due
to the lifetime (ca. lu.s in tissue) and diffusion properties of singlet oxygen, any cellular
damage is restricted to a 0.1 \im radius. If a photosensitizer is activated and it brings about
the destruction of a crucial component of the tumour cell, such as DNA, eradication of the
cell will occur more effectively and at much smaller doses.
64
where R= NHfCH^NfCH^
NH(CH2)3N(CH3)2
Figure 2.11 Amide-Linked Protoporphyrin Derivatives
These same derivatives were also found to target mitochondria in an in vitro model.
This result is supported by earlier research113-114 which found cationic molecules to be
highly selective for mitochondria. The mitochondria contain the enzymes responsible for
electron transport and oxidative phosphorylation within cells. If a photosensitizer binds to
mitochondria within a tumour cell, photoactivation of the drug will significantly reduce the
amount of cellular ATP and lead to the death of the cell115.
Two derivatives of hematoporphyrin IX that were synthesized by this group are
shown in Figure 2.12. Biodistribution data showed that these two derivatives showed
excellent localization in tumourous tissue versus other tissues in the mouse model. Tumour
to skin biodistribution ratios for compound A were 7.2 to 1 and 2.5 to 1 for compound B.
These findings, coupled with other data, led this group to believe that the morpholine
functionality may have an important role in defining the biodistribution behaviour of the
photosensitizers. In order to capitalize on this, the morpholine moiety was incorporated
into BPD.
65
Figure 2.12 Arrude-Linked Morpholine Derivatives of Hematoporphyrin DC
As was mentioned in the introduction, a desirable quality of a photosensitizer is
water-solubility. Pandey and others11*' have recently shown that cationic, water-soluble
porphyrin and chlorin photosensitizers can be synthesized via quaternization of amines.
Similarly, Oleinick and others11^ have recently synthesized silicon metallated
phthalocyanines with a quaternary ammonium iodide functionality bound as an axial
substituent of the metal in their attempts to solubilize their photosensitizers in water. Water
solubility is important because most of the compounds presently being tested have poor or
no solubility in water and cannot be administered to cells or animals in aqueous solution.
This creates the necessity of drug solubilization media such as cremophore emulsion,
dimethyl sulfoxide (DMSO), or liposomes. Each of these media have problems associated
with their use. For example, cremophore emulsions are difficult to reproduce, some
hydrophobic compounds form aggregates in, or precipitate out of, DMSO118 and many
liposomes are susceptible to fusion with lipoproteins119 and are themselves
thermodynamically unstable. In addition, the contribution that each of these media make to
the resultant biological activity is poorly understood.
66
While it may be the hydrophobicity of the sensitizer that is the main reason for its
biodistribution properties, the advantages of water solubility are too many to overlook. In
addition, it is feasible that compounds that have both hydrophobic and hydrophilic regions
will be good candidates as photosensitizers. PDT drugs with this quality have been
referred to as 'amphiphilic' by Bonnett and Berenbaum who believe this feature is related to
the selectivity of a photosensitizer for tumourous versus healthy tissue^O f n e
hydrophilic moiety may facilitate water solubility in serum and thus lead to better systemic
distribution while the hydrophobic moiety dictates the tumour and eventual intracellular
targeting.
2.4.2 Via Amide Linkage
Scheme 2.5 Synthesis of BPD Derivatives with Free Amine Functionalities
The hydrolysis : acid-chloride formation : displacement methodology developed for
the straight chain alkyl amide derivatives was employed to make the compounds shown in
Scheme 2.5. BPD di(propylmorpholine) amide (15) and BPD di(N,N-
dimethylethylenediamine) amide (16) were synthesized in 70% and 52% yields,
respectively. Both compounds were characterized by high resolution spectroscopy and
NMR. The A H NMR spectrum of the former compound is shown in Figure 2.13.
6 7
Initially, the peak centered at 7.78ppm was difficult to assign. It appears to be a singlet and
integrates to two protons but upon further inspection it was found to be a AB system
resulting from the protons at 7 and 7 . In this solvent (d6-DMSO) the chemical shift
difference between these protons is much smaller than in CDCI3 and the protons are
essentially equivalent. All other new peaks were confirmed by decoupling experiments.
To determine if quatemization of the dimethylamino moieties of compound (16)
would render it water-soluble, it was dissolved in dry acetone and treated with a large
excess of methyl iodide (Scheme 2.6). After a 10 minutes stirring, the solvent and
unreacted methyl iodide were removed. The resulting solid was found to dissolve readily
in distilled water. Initial testing showed that roughly 5mg of this product were soluble in
lmL water. The BPD di(trimethylarnmoniumiodide) (17) was characterized by NMR.
Scheme 2.6 Quatemization of BPD Di(N,N-chmemylemylenediarrune)amide (16) Using Methyl Iodide
68
69
2.4.3 Via Coupling to the Vinyl Group
Recent publications by Pandey and others* 16,121 n a v e shown that Eschenmosers'
reagent, N,N-dimethylmethylenearnrnoniurn iodide, reacts with various porphyrin-type
compounds in a specific fashion. For example, deuteroporphyrin IX dimethyl ester, a
close relative of protoporphyrin IX dimethyl ester differing only in that there are protons
instead of vinyl groups at positions C-3 and C-8, reacts in excellent yield to produce 3,8-
bis[(dimethylamino)methyl]deuteroporphyrin IX dimethyl ester. However, for porphyrins
containing vinyl groups, the reagent was shown to react regioselectively at the vinyl
positions. In this way, protoporphyrin IX dimethyl ester gave the corresponding
bis[(dimethylamino)methyl] adduct (Figure 2.14). The initial electrophilic attack is
followed by loss of a proton to regenerate the double bond.
Deuteroporphyrin DC dimethyl ester
Protoporphyrin K dimethyl ester
Figure 2.14 Reaction of Deuteroporphyrin DC Dimethyl Ester and Protoporphyrin DC Dimethyl Ester with N -dUrnemylrnemyleneainmonium Iodide
70
BPD dimethyl ester (2) was treated with a large excess of Eschenmoser's reagent
in dichloromethane and gave primarily a much slower moving compound in 88% yield after
stirring for two days in the dark at room temperature (Scheme 2.7).
Scheme 2.7 Reaction of BPD Dimethyl Ester with N,N-dlmemylmethyleneammonium Iodide to form BPD Eschenmoser Derivative (18)
The l H NMR spectrum (Figure 2.15) showed the loss of the doublet at 6.37ppm denoting
the cis-coupled proton at C-3 2 , the loss of the doublet of doublets centered at 8.1 lppm due
to the proton at C-3*, and the appearance of two new signals at 7.00ppm (doublet of
triplets, Jcis=7Hz and Jtrans=17Hz) and 8.20ppm (doublet, Jtrans=l7Hz) denoting the
trans-coupled proton at C-3 2 and the trans-coupled proton at C-3*, respectively. After
final characterization by high resolution mass spectroscopy, the product was determined to
be the BPD Eschenmoser derivative (18). In this way, an amine functionality was added
to BPD via the vinyl group without generating any new stereoisomers.
Quatemization of the dimethylamino moiety of this compound led to the formation
of several compounds. The fact that the trimethylamino moiety is a good leaving group
coupled with the stability of the resulting carbocation (the positive charge can be delocalized
throughout the chlorin) is presumably the source of this problem.
71
7 2
2.5 Synthesis of Analogous BPD Amines and Amides
2.5.1 Rationale
Some of the free amines that have been incorporated into photosensitizers have been
added via amide linkages (see previous section). But what effect does the amide
functionality or the carbonyl of that amide have on the biodistribution and targeting
properties of these derivatives? In an effort to determine if this moiety is important, the
synthesis of two pairs of compounds was envisioned (Figure 2.16). The two pairs would
differ only in the 133 and 173 positions which would be either carbonyls or methylene
groups. Distinct differences in the in vitro testing might provide insight into this question.
Figure 2.16 Two Pairs of Amide- and Amine-Linked BPD Derivatives
2.5.2 BPD Amides via Acyl Chloride Displacement
The BPD dimorpholine (19) and dipiperidine (20) amides were synthesized using
the hydrolysis and acid chloride methodology in 79% and 70% yields, respectively
(Scheme 2.8).
Scheme 2.8 Synthesis of BPD Amides Using Acid Chloride Displacement
2.5.3 BPD Amines via Iodide Displacement
A new methodology was developed for the synthesis of the corresponding BPD
diamines. It was felt that if the propionate chain esters could be selectively reduced to the
corresponding diol, there would be a synthetic handle for the desired transformation.
Diborane reagents were avoided due to their reactivity with the vinyl group at
position 3. In fact, a small scale reaction performed on the BPD dimethyl ester using
borane-tetrahydrofuran complex yielded a product which was found (by *H NMR) to be
BPD dimethyl ester but with the loss of the typical resonances due to the vinyl group.
Hydride reducing reagents appeared to be the best candidates for the desired
transformation. Because we were interested in the selective reduction of the propionate
chain esters, it was felt that a bulky hydride reagent might be appropriate.
74
When the free-base chlorin was treated with dusobutylaluminum hydride (DIBAL)
the solution immediately turned from dark green to bright red. This was an unusual result
given the assumption that modification of the propionate esters should not noticeably affect
the BPD chromophore. The uv-vis spectrum of this compound showed the loss of the
normal BPD spectrum and the presence of a porphyrin-type spectrum. This product was
not characterized but the compound may be the benzoporphyrin described earlier (section
Metallation of tetrapyrroles via the pyrrolic nitrogens is commonly used by
porphyrin chemists to inhibit unwanted reactions. The metal of choice in the literature is
zinc(II). A methylene chloride solution of BPD dimethyl ester (2) was treated with a four
fold excess of zinc(II) acetate in methanol to yield Zn BPD (21) in 99% yield (Scheme
2.9). The reaction progress was monitored by tic and uv-vis spectroscopy. The free-base
BPD spectrum was changed to one typical of metallochlorins where the Q-bands are
simplified due to the increase in symmetry obtained by metal insertion and the absorption
maximum is hypsochromically shifted to 672nm. The spectra of free-base and metallated
BPD dimethyl ester appear in Figure 2.17.
2.3.4.1).
• l.OOO-i
BPD
0.0000 400 SOO 600 700 •00
MAVELFN6TM
Figure 2.17 The UV-Vis Spectra of BPD Dimethyl Ester (2) and Zn BPD Dimethyl Ester (21) in CH2CI2
7 5
1. D IBAL-H, THF 2 aqueous ammonium chloride
I .TsCI , pyridine, C H 2 C I 2
2. Na l , acetonitri le 3. R H , acetonitr i le
Scheme 2.9 Synthesis of Two BPD Amine Derivatives Via Reduction of BPD Dimethyl Ester (2)
76
The best conditions for the desired reduction were found to be treatment of Zn BPD
(21) in tetrahydrofuran with just over 4 equivalents of 1.0M DEBAL-H in hexanes at 0°C
under nitrogen. The yield of Zn BPD diol (22) after quenching with saturated ammonium
chloride and chromatography on silica (5%MeOH/CH2Cl2 eluent) was a moderate 52%.
The A H NMR is shown in Figure 2.19. The spectrum revealed the loss of two of the
singlets between 3.5 and 4ppm which correspond to the propionate methyl esters. The six
methylene groups, two of which are new, were somewhat difficult to assign so a series of
decoupling experiments were performed. The multiplet at 2.20-2.3 lppm was irradiated
and the multiplets between 3.68-3.76 and 3.82-3.88ppm were both simplified. Irradiation
of the lowest field of these peaks gave simplification of the 2.20-2.3 lppm resonance.
Irradiation of the 3.68-3.76ppm peak not only simplified the 2.20-2.3lppm peak but also
converted the overlapping triplets centered at 4.68 and 4.69ppm (which are due to the OH
protons) to singlets. Based on this data the following assignments were made: 2.20-
2.31ppm=2xRCH2CH2CH20H; 3.68-3.76ppm=2xRCH2CH2CH20H; and 3.82-
3.88ppm=2xRCH2CH2CH20H, where R is the BPD backbone.
Demetallation to the free-base BPD diol (23) was facilitated by treatment of the Zn
BPD diol (22) with trifluoroacetic acid in dichloromethane in a 99% yield. The uv-vis
spectra of BPD dimethyl ester and BPD diol are shown in Figure 2.18.
0 ' • • ~ 400 500 600 700 800
Figure 2.18 The UV-Vis Spectra of BPD Dimethyl Ester (2) and BPD Diol (23) in CH2CT2
77
To incorporate the desired amines, standard tosylation was used. A
dichloromethane solution of the BPD diol (23) was treated with a ten-fold excess of para-
toluenesulfonyl chloride and a small amount of pyridine. A smaller amount of the tosylate
led to incomplete conversion to the ditosylate. The reaction was allowed to proceed at 0°C
overnight at which time dc showed good conversion to a faster moving compound than the
diol. The compound was worked up but chromatography led to the formation of new
compounds. Low resolution mass spectroscopy (electron impact) gave the parent (M +)
ion. The crude BPD ditosylate (24) was carried over to the iodination step.
Iodination was necessary because direct displacement of the tosylate with piperidine
was incomplete. The crude BPD ditosylate (24) was dissolved in acetonitrile and a
thirteen fold excess of sodium iodide was added. The rriixture was heated for 10 minutes at
which time a tic cospot with the crude ditosylate showed conversion to a slighdy faster
moving compound and a small amount of baseline material. This BPD diiodide (25) was
also sensitive to chromatography but less so than the ditosylate and a small amount was
purified and immediately characterized by *H NMR. In general, this crude residue was
carried over for the final displacement
The crude BPD diiodide (25) was reacted individually with piperidine and
morpholine by stirring at room temperature in the dark. The diamines were
chromatographed to yield the BPD dimorpholine (26) and dipiperidine (27) amines in
64% and 51% yields from the BPD diol, respectively. Both compounds were characterized
by A H NMR and high resolution mass spectroscopy.
79
2.6 Synthesis of BPD Derivatives with Hydroxyl Functionalities
2.6.1 Rationale
It has been concluded recently by Bonnett et. a l . 1 2 2 that photosensitizers with
amphiphilic character (having both hydrophilic and hydrophobic regions) are likely to
prove valuable as photosensitizing agents in the photodynamic therapy of cancer. This
conclusion is based upon some of the earliest attempts to design second-generation
photosensitizers with better tumour localizing properties that were begun by Bonnett and
Berenbaum in the mid-1980's. These pioneers tested over 100 modified porphyrins with
an in vivo model and determined, among other things, that compounds with amphiphilic
character were more effective sensitizers. Specifically, the para- and meta-hydroxy
substituted tetra(hydroxyphenyl)porphyrins (Figure 2.20 (a)) were found 1 2 3 to show good
selectivity for tumour versus muscle and skin tissue. Furthermore, on a mole to mole
basis, these two derivatives were found to be 25-30 times as potent as hematoporphyrin
derivative and Photofrin, as judged by the depth of tumour necrosis (death of tissue). The
resulting skin sensitivity after injection of these derivatives was quite low comparatively
which also points to some degree of tissue selectivity.
Further experimentation with the corresponding dihydroporphyrins (Figure
2.20(b)) showed that these derivatives were even more effective tumour photosensitizers.
In fact, these chlorins caused tumour necrosis at drug dose levels at which the porphyrin
derivatives were ineffective^. While this may be due to the larger extinction coefficient of
the irradiated absorption band of the chlorin derivatives, it appears that the hydroxy
functionalities are in part responsible for the improved results.
8 0
Figure 220 Porphyrins and Chlorins with Hydroxy Groups: a) Tetra(para-hydroxy-phenyl)porphyrin and Tetra(meta-hydroxyphenyl)porphyrin; b) Tetra(para-hydroxy-phenyl)chlorin and Tetra(meta-hydroxyphenyl)chlorin; c) Hydroxy Octaethylchlorin Derivatives
81
Bonnett and others have recently published the chemistry125 a n d biological
activity of several modified octaethylporphyrins with various hydroxy functionalities
(see Figure 2.20(c)). Al l of these compounds showed good anti-tumour activity as judged
by the depth of tumour necrosis in their in vivo model.
Boyle et. al.127 have synthesized zinc phthalocyanine derivatives with multiple
hydroxyl functionalities attached to the phenyl rings (Figure 2.21). The compounds with
the hydroxy groups directly attached to the phthalocyanine periphery showed poor tumour
response but the derivative with hydroxy groups bound via a three carbon chain showed a
low LD90 (the dose at which 90% of cells are destroyed) in vitro and good tumour cure
response in vivo.
where R= OH
(CH2)30H
(CH2)6OH
Figure 2.21 Zinc Phthalocyanine Derivatives with Hydroxy Groups
Several hydroxy functionalities were incorporated onto BPD in various fashions in
order to confer the above mentioned activity upon BPD.
82
2.6.2 Via Reduction of BPD Dimethyl Ester
It was felt that reduction of the various methyl ester groups on BPD dimethyl ester
would be a good way to incorporate alcohol groups. The synthetic sequence designed for
the dipiperidine and dimorpholine amines (Section 2.5.3) has already shown that the
selective reduction of the propionate ester chains to the corresponding BPD diol was
successful.
Further tic inspection of the side products generated in that reduction showed faster
and slower moving spots in relation to the Zn BPD diol. It was determined by mass
spectroscopy that the faster moving spots were the two possible mono alcohol/mono
methyl ester products. The slower moving spot appeared to have a different chromophore
than the starting compound or the various mono- and diols. The uv-vis spectrum of this
compound is overlaid with Zn BPD in Figure 2.22. The highest mass ion peak on the
mass spectrum was at 710 which is 28 mass units less than the Zn diol. The NMR
(Figure 2.22) showed the loss of the methyl ester resonance at 3.99 ppm (due to the methyl
ester at position 72)and a series of new peaks centered at 4.32 and 5.29 ppm. The first
resonance is a doublet of an AB quartet which integrated to two protons (the new CH?QH
moiety at C-72) and the second was a triplet (J=5.6Hz) integrating for 1 proton (the new
hydroxy proton). Using this data and high resolution mass spectroscopy (and later
microanalysis) this compound was characterized as the Zn BPD triol (28) depicted in
Figure 2.22 with the third reduction occurring at position-72.
The reduction had a large impact on the chemical shift of some of the proximate
resonances. The protons at 7 3 and 7 , normally overlapping or very close together in the
d6-DMSO l H NMR, were separated by almost 0.8ppm and the proton at position 7* was
shifted upfield 0.6ppm. The peak from the methyl ester at position 7* was not affected by
this reduction.
83
84
Attempts to synthesize the zinc triol selectively were much more successful than
with the diol. Utilizing the same reaction conditions but with slightly more than 6
equivalents of DIBAL-H, the Zn BPD triol (28) was synthesized in 81% yield from Zn
BPD (21) after quenching and chromatographic purification (Scheme 2.10).
HOCH 2 CH 2OH
Scheme 2.10 Synthesis of BPD Triol (29) from Zn BPD Dimethyl Ester (21)
85
The Zn BPD triol (28) was demetallated using trifluoroacetic acid followed by base
workup to produce the free-base BPD triol (29) in 92% yield. Because the methyl ester at
position 7^ was reduced, the electronic properties of the BPD triol are much different than
those of the other BPD derivatives. The uv-vis spectrum of BPD triol is shown in Figure
2.23. The absorption maximum is slightly blue-shifted to 686nm and the extinction
coefficient of this peak was determined to be only 2 1,600M*1 cm" 1 or roughly two-thirds
that of BPD dimethyl ester. It is clear that the conjugation of the methyl ester at position 72
is a crucial determinant of the electronic properties of the BPDs.
400 500 600 700 800 WAVELENGTH (nm)
Figure 2.23 UV-Vis Spectrum of BPD Triol (29) in CH2Cl2/MeOH
This compound is important from a synthetic standpoint for our lab. Until this
time, no compounds had been synthesized with modification of the substituents on the
'benzo' portion of BPD. This discovery will now allow the modification of BPD in a
completely new and selective way. Attempts are presently being made to synthesize the
BPD tetraol.
86
2.6.3 Via Transesterification
Given the success of the transesterification methodology detailed in section 2.3.3, it
was felt that it could be used to incorporate two equivalents of 1,2-ethanediol (ethylene
glycol) into BPD dimethyl ester. Treatment of BPD dimethyl ester (2) with a large excess
of this diol and concentrated sulfuric acid at room temperature for 2 days gave the BPD
di(ethylene glycol) ester (30) in 94% yield after workup and chromatography (Scheme
2.11).
Scheme 2.11 Synthesis of BPD Di(ethyleneglycol) Ester (30) from BPD Dimethyl Ester (2) Via Transesterification
2.6.4 Via Amidation
Referring to the discussion in section 2.3.4, there is some concern as to the stability
of ester linkages in biological systems. To provide a control for this, the ethanol
functionality was added to BPD via an amide linkage. Direct displacement was used to
provide the BPD di(ethanol)amide (32) in 75% yield (Scheme 2.12).
87
Scheme 2.12 Synthesis of BPD Di(ethanol)amide (32) By Direct Displacement
2.6.5 Via the BPD Vinyl Group
The hydrobromination-displacement methodology developed in section 2.3.2 was
used to add a hydroxy group at position 3 1 in a Markovnikov fashion. The BPD
hydrobromide was treated with water and heated overnight. The BPD vinyl hydrate (31)
was isolated as a mixture of diastereomers and characterized as such after chromatography
(Scheme 2.13).
(2) (31)
Scheme 2.13 Synthesis of BPD Vinyl Hydrate (31) Via Hydrobromination and Displacement
2.7 Synthesis of Phosphonate Ester Derivatives of BPD
2.7.1 Rationale
Boyle and others*28 have recently synthesized phosphonate ester substituted
phthalocyanines (Figure 2.24). The phosphonate moiety was chosen because of its
capacity to be hydrolyzed in base to the partially deprotected phosphonic acid derivative.
The researchers found that these anionic functionalities conferred water solubility on the
general hydrophobic phthalocyanines. These compounds were biologically tested and it
was found that the compound containing two of the phosphonic acid groups on the same
benzene moiety showed good in vitro and in vivo photodynamic activity while the tetra
substituted compound was completely inert in both systems. This result is further support
of the idea that amphiphilicity is an important feature of a photosensitizer.
Figure 2.24 Sodium Salts of Tetra- and Diphosphonic Acids of Zinc Phthalocyanine
In order to mimic the promising photodynamic behaviour exhibited by this compound,
BPD was substituted with these functionalities.
89
2.7.2 Via Iodide Displacement
The same synthetic sequence used to make the di(morpholine) and di(piperidine)
amines was employed here. The BPD diiodide was treated with a large excess of
triethylphosphite and refluxed in acetonitrile overnight. The solvent and excess
triethylphosphite were removed under reduced pressure and the crude residue was
chromatographed to provide the desired BPD diphosphonate ester (33) in 67% yield from
the BPD diol (Scheme 2.14). The product was characterized by A H and 3 1 P NMR (Figure
2.25) and high resolution mass spectroscopy. The assignments of the new ethoxy
resonances were confirmed by decoupling experiments.
Scheme 2.14 Synthesis of BPD Diphosphonate Ester (33) from BPD Diol (23)
Attempts to hydrolyze the phosphonate esters using aqueous sodium hydroxide
gave a mixture of compounds and the desired compound could not be isolated.
91
2.8 Summary
This chapter has dealt with the rationale behind and the synthesis of new
benzoporphyrin derivatives (BPD). Various functionalities (alkyl chains, hydroxy groups,
heterocyclic moieties, amines and phosphonate esters) bound through ester, amide, amine
and phosphorus-carbon bonds, have been incorporated onto the BPD structure. As
proposed, the bulk of these new compounds did not modify the BPD chromophore. In
addition, selected compounds all tested positive for singlet-oxygen production. All the
synthetic routes are general enough that any number of moieties can now be attached to the
BPD periphery.
In vitro and in vivo biological testing is currently being performed on compounds
(3)-(ll), (13)-(20), (23), (26), (27), (29)-(33).
Chapter 3 Fluorescence Quenching:
Results and Discussion
93
The theoretical basis for the fluorescence quenching experimentation was presented
in the introductory chapter. This chapter describes the selection of the fluorescent probe
and its incorporation into liposomes. The quenching experiments on five BPD derivatives
are detailed as are the practical considerations that had to be addressed. Stern-Volmer plots
are generated and the results are discussed.
3.1 Fluorescent Probe Selection
One of the most commonly used fluorescent probes is diphenylhexatriene (DPH).
Much of the reason for its use in studies of biological membranes is its high extinction
coefficient (ca. 80000cm" AM"1 at 355nm)l29 njgh quantum yield of fluorescence in
hydrophobic environments and its negligible fluorescence in water. Its excitation and
emission maxima are well separated from membrane absorbance.
However, DPH has been implicated in the three criticisms of fluorescence
quenching discussed in the previous section. In order to address these criticisms,
researchers have chemically joined the DPH moiety to various phospholipids and other
compounds. These compounds (Figure 3.1) are all designed so that they incorporate into a
liposome bilayer i.e. they have hydrophilic head groups and hydrophobic chain(s). The last
compound shown in Figure 3.1 is created by substituting one of the acyl chains of a
dihexadecanoyl phosphatidylcholine molecule with DPH to generate diphenylhexatriene
hexadecanoyl phosphatidyl choline (DPH-hPC).
D i p h e n y l h e x a t r i e n e ( D P H )
fH3
T r i m e t h y l a m m o n i u m D i p h e n y l h e x a t r i e n e
( T M A - D P H )
P r o p i o n i c A c i d D i p h e n y l h e x a t r i e n e
( P A - D P H )
2 - [ 3 - ( d i p h e n y l h e x a t r i e n y l ) p r o p a n o y l ] - l - h e x a d e c a n o y l -s n - g l y c e r o - 3 - p h c « p h c i e t J i a n o l a m i n e
( D P H - h P E )
2 - [ 3 - ( d i p h e n y I h e x a t r i e n y l ) p r o p a n o y l ] - l - h e x a d e c a n o y l -s n - g l y c e r o - 3 - p h o s p h o c h o l i n e
( D P H - h P C )
Figure 3.1 Various Diphenylhexatriene-Containing Fluorescent Probes
When this lipid-bound fluorescent probe is added to a solution of phospholipids dissolved
in organic solvent and the solvent is removed, the resulting film can be hydrated to provide
liposomes with a fluorescent probe imbedded in a defined way. Research has shown that
the DPH portion of DPH-hPC is oriented perpendicular to the bilayer plane (Figure 3.2)
and demonstrates restricted motion within the bilayer 130-131
Figure 3.2 Idealized Representation of the Orientation of Diphenylhexatriene
Hexanoyl Phosphatidylcholine (DPH-hPC) Fluorescent Probe in
a Dimyristoyl Phosphatidylcholine (DMPC) Bilayer
96
Furthermore, Morgan et.al. 1^ believe that the lipid-bound DPH is unlikely to exchange
rapidly between liposomes. This modification addresses two of the three aforementioned
problems in that the general location of the DPH moiety is well defined and the attachment
of the DPH to the phosphatidylcholine rmnimizes the exchange of the fluoroprobe between
liposomes. The third problem still remains but because of the intensity of DPH
fluorescence, very small amounts of it need be used. In fact, this experimentation
incorporates the lipid-bound DPH in a 14 to 10,000 ratio (mol/mol) to the lipids which is
below that used in similar experiments133. Because of the success of DPH-hPC in
addressing the above criticisms, it was used in this experimentation.
The following sections detail the incorporation of DPH-hPC into liposomes and
describes fluorescence quenching experiments performed on BPDMA and four newly
synthesized BPD derivatives: the methyl, propyl, pentyl and heptyl diesters (Figure 3.3).
Me02C P02Me
R= CH 3
C 3 H 7
R0 2C C0 2R
Figure 3.3 Structure of BPD Dimethyl, Dipropyl, Dipentyl, and Diheptyl Esters
97
3.2 Composition and Creation of Liposomes
The liposomes in which the DPH-hPC fluorescent probe was imbedded were
composed of the same lipids (and in the same proportions) as those used in the formulation
of BPDMA: Dimyristoyl phosphatidylcholine (DMPC: 61%) and egg phosphatidylcholine
(EPC: 39%) on a mole-to-mole basis. DMPC is a pure synthetic phospholipid composed
of a phosphatidylcholine head group with two fully saturated 14-membered (myristoyl)
acyl chains and EPG is a mixture of phospholipids with a phosphoglycerol head group and
various acyl chains ranging from 14-18 carbons (myristoyl-stearoyl) in length with up to 4
degrees of unsaturation per chain (Figure 3.4).
Dimyristoyl Phosphatidylcholine (DMPC)
Egg Phosphatidylglycerol (dimyristoyl component shown)
Figure 3.4 Dimyristoyl Phosphatidylcholine (DMPC) and Egg Phosphatidylglycerol (EPG)
The preparation of the DPH-bound liposomes is described in the experimental
section. Because of the concern that the introduction of a probe into a liposome may cause
98
changes in the local environment of the liposome, it was incorporated as only 14 parts in
10000 (0.14% on a mole DPH-hPC-to-mole total lipid basis).
The probe-embedded liposomes were reduced in size using an extrusion apparatus
and the populations of liposomes after each extrusion were sized using a Nicomp sub-
micron particle sizer. The extrusion apparatus and procedure is described in the
experimental section. The particle sizer utilizes the technique of dynamic light scattering
(also called quasi-elastic light scattering or photon correlation spectroscopy) to characterize
the size distribution of particles suspended in a solvent. Simply stated, a particle irradiated
by a laser gives rise to a scattered light wave, the intensity of which is dependent upon the
molecular weight or volume of the particle. What was interesting and deserves mention
here is the fact that a lOOnm diameter liposome solution that had remained in the
refrigerator for three weeks was sized and showed only a small amount of larger liposomal
material with the mean diameter of 136nm. The hardiness of this liposomal formulation is
remarkable because hydrated liposome suspensions are thermodynamically unstable and
tend to aggregate or fuse into larger species.
The final concentration of liposome-bound DPH-hPC after hydration was 68p.g per
mL aqueous buffer. This concentration was determined by measuring the fluorescence
intensity of hydrated liposome prior to and after extrusion. The intensities showed little or
no difference, supporting the assumption that no lipid was retained in the extruder. The
low concentration of liposomes was chosen to avoid unwanted light-scattering phenomena
which can affect the observed fluorescence emission.
3.3 Fluorescence Spectra of Liposome-Bound Diphenylhexatriene (DPH)
and BPD
The emission and excitation spectra of DPH-hPC/DMPC/EPG liposomes in
aqueous buffer and BPD dimethyl ester in N-methylpyrrolidinone are shown in Figure 3.5.
9 9
The excitation maximum of the lipid-bound DPH is at 362nm and the fluorescence
emission maximum is 433nm, which corresponds well with the absorbance maximum for
BPD at 430nm. BPD fluoresces at 694nm (with excitation at 430nm) in this solvent
Figure 3.5 Fluorescence Excitation and Emission Spectra of Liposome-Bound DPH-hPC in Aqueous Solution and BPD Dimethyl Ester in N-Methylpyrrolidinone
3.4 Photobleaching of Liposome-Bound DPH
Before the fluorescence quenching experiments could be performed, a number of
practical considerations had to be addressed. Initial experimentation on the liposome-
bound DPH with the continuous wave lamp showed that it was quite susceptible to
photobleaching, a light mediated process which changes or destroys the DPH
chromophore. Changing to the flash lamp significantly reduced the amount of
photobleaching but this was at the expense of the signal-to-noise ratio. Because we were
100
interested in determining the true quenching effects of various BPD derivatives on
liposome-bound probe, the flash lamp was chosen as the means of illumination because the
increased signal-to-noise ratio was not deemed necessary.
The amount of light hitting a sample is proportional to the monochromator bandpass
and several experiments were performed as oudined below to determine the parameters
which minimize the amount of photobleaching. The liposomes were irradiated at 362nm
and the fluorescence was detected at 433nm using a variation of the excitation and emission
bandpasses. The best combination was found to be an excitation bandpass of 2nm and an
emission bandpass of 16nm, where the amount of photobleaching was less than 1% over a
typical acquisition time and flash repetition rate (Table 3.1).
Excitation bandpass Emission bandpass % Fluorescence Photobieached
4nm 4nm 3.10±0.50
4nm 8nm 2.16±0.65
4nm 16nm 1.81±0.18
2nm 16nm 0.83±0.48
Table 3.1 %-Fluorescence Photobleaching of Liposome-Bound DPH-hPC at Various Excitation and Emission Bandpasses (EX 362nm, EM433,200 flashes)
3.5 Fluorescence Quenching and Stern-Volmer Plots
The four BPD diesters were dissolved in n-methylpyrrolidinone (NMP), a solvent
which is high boiling, water-miscible and an effective solubilizer of BPD's. The extinction
coefficients of each were determined at three different concentrations in the linear region of
the Beer-Lambert plot. The absorptivities agreed well for each compound but there were
some differences between compounds. In order to ensure the fluorescence quenching
101
results could be compared, the concentrations of the four solutions were standardized to a
common extinction coefficient (32,000 M'^cm" 1). This is reasonable because the
peripheral modifications to create the compounds did not involve modification of the BPD
chromophore. All four compounds have essentially the same optical spectra and should
have the same extinction coefficients. The differences between solutions is accounted for
by purity and/or weighing error.
To ensure that NMP was not responsible for the observed quenching of DPH
fluorescence, the fluorescence of the liposomes was measured before and after the addition
of 105fiL NMP (the largest amount of NMP used in the experiments; 9% by volume). Only
1.4% of the fluorescence was quenched after the addition.
A standard experiment proceeds as follows: A sample was placed in the
fluorometer and equipped with a teflon stir bar and a computer assisted acquisition was
established to measure the fluorescence immediately after irradiation by the xenon flash
lamp every 5 seconds. After pausing the acquisition, aliquots of each BPD were added to
the cuvette using a micropipet or Hamilton syringe. The experiment was resumed and the
next aliquot was added only after the fluorescence had leveled-off. The concentrations of
the BPD solutions was deliberately chosen so that they were large enough that a minimal
amount of NMP was added to the liposome solution, but small enough that several
graphical points could be determined.
The total concentrations were calculated and plotted against F Q / F (where F 0 is the
fluorescence of the probe prior to BPD addition and F is the fluorescence after each
addition of BPD) for each aliquot of BPD derivative added. Figure 3.6 shows the Stern-
Volmer plots for BPDMA and the four new BPD derivatives. Each of them is a
polynomial curve fit of two or three independent experiments for each BPD. All of the
Stern-Volmer plots are concave towards the y-axis, with this feature being more
pronounced for BPDMA amd the smaller alkyl chain length BPD derivatives. Note that as
the alkyl chain length decreases, the plots are increasingly more concave to the y-axis.
102
103
As was mentioned in the introduction, deviation from linearity for a Stern-Volmer
plot indicates that more than one type of quenching is occurring. At low concentrations of
quencher, the plots appear to be linear which may be reflective of diffusional (collisional)
quenching. As the concentration of quencher increases, the differences between the
various compounds are much more pronounced. The longer alkyl chain compounds are
much poorer quenchers than the shorter chain derivatives, for the same quencher
concentration.
These differences in quenching may be a result of the inability of the longer alkyl
chain length BPDs (the dipentyl and diheptyl esters) to approach sufficiently close to the
DPH moiety due to the steric bulk supplied by the alkyl chains. The shorter dimethyl and
dipropyl esters have much less steric bulk and would not be expected to cause this problem
and thus might interact with the probe more effectively.
Another plausible reason for the differences can be expressed in terms of where the
various derivatives are expected to be located within the liposome. For example, it is
presumed that the photosensitizer hematoporphyrin is located within liposomes in the way
shown in Figure 3.7. The carboxylic acid functionalities are expected to align with the
hydrophilic phospholipid headgroups and the predominantly hydrophobic porphyrin
backbone is expected to be imbedded within the hydrophobic side chains134
Figure 3.7 Probable Orientation of Hematoporphyrin DC Dipropionic Acid in Lipid Bilayers
104
The mono acid functionality of BPDMA is expected to confer this type of binding
within the liposome. The mono acid group interacts with the hydrophilic choline and
glycerol moieties of the phospholipids and thus forces the relatively hydrophobic remainder
of BPD to be aligned with the acyl chains (Figure 3.8 (a)). The Stern-Volmer plots show
the dimethyl ester quenches the DPH fluorescence in a way similar to BPDMA. This might
point to the fact that the BPD dimethyl ester derivative aligns itself predominandy so that
the relatively hydrophihc methyl esters interact with the hydrophilic portion of the liposome
bilayer (Figure 3.8 (b)). As the length of the ester increases, the most hydrophilic moieties
become the two methyl esters located at positions 7 2 and 7* on the benzene-like moiety of
BPD. This would then cause the longer alkyl chain BPD derivatives to orient themselves in
the bilayer so that the alkyl chains could be imbedded within the acyl chains (Figure 3.8
(c)). This would effectively invert the position of these derivatives in relation to the
dimethyl ester or BPDMA and lead to a different interaction and thus different quenching of
the liposome-bound probe.
As stated in the introduction, it is not known whether liposome-bound BPDMA is
free to diffuse into other liposomes. In order to determine this, the following experiment
was performed: A known amount of BPDMA was added to a known concentration of
DPH-hPC-bound liposomes and the fluorescence of the probe was measured prior to and
after the addition. To this solution was then added a small aliquot of liposomes (consisting
of the same lipids in the same ratio) in buffered aqueous solution which did not contain any
fluorescent probe. The amount of lipid added was identical to the initial amount of lipid
prior to addition of BPDMA. The fluorescence of the system was then measured.
If the BPDMA is free to diffuse out of the liposomes in which it is bound, the
fluorescence of the DPH probe should recover as the BPDMA diffuses into the newly
added liposomes. If the BPDMA is fixed within the initial liposome species, the
fluorescence would not be expected to change. The fluorescence measurement showed that
105
Figure 3.8(a) Proposed Orientation of
BPDMA Within a Lipid Bilayer
Figure 3.8(b) Proposed Orientation of
BPD Dimethyl Ester Within a Lipid Bilayer
Figure 3.8(c) Proposed Orientation of Longer
Alkyl Chain BPD Diesters (only BPD
Diheptyl Ester is Shown)
106
some of the fluorescence had recovered. The bulk of the fluorescence recovery occurred
almost immediately upon addition and attained a constant value after 400 seconds. Using
the Stern-Volmer plot of F Q / F versus concentration of BPDMA, the amount of BPDMA
remaining in the initial DPH-bound liposomes was interpolated. It was determined that
57% of the BPDMA had diffused out of the first species of liposomes into the newly added
species. Because the amounts of the two species of liposomes were the same (within
experimental error), if the BPDMA is not hindered in any way, it is expected statistically
that 50% of the BPDMA would diffuse out. Given that the error associated with this
procedure is, while not quantitatively assessed, probably quite high, it is reasonable to infer
that the BPDMA is free to diffuse and distribute amoung liposome populations.
Preliminary testing with liposome-bound BPD diheptyl ester has shown that it too
appears to be able to diffuse and distribute readily among liposome species. Although the
kinetics have not been investigated quantitatively, it appears that hydrophobic alkyl chains
incorporated into BPD do not have a major effect on the rate of intermembrane diffusion
but do affect the transverse position of BPD in the lipid bilayer.
3.6 Summary
Fluorescence quenching has been used to show that different benzoporphyrin
derivatives (BPD) interact differently with a liposome species. These differences are
manifest in Stem-Volmer plots which display the interaction of a quenching species with a
fluorescent probe-imbedded liposome. Some reasons for the generated results were
proposed. Furthermore, this methodology was used to determine that liposomal BPDMA
is able to freely diffuse into similar liposome species. This provides a clue in determining
the way in which BPDMA and other BPD's behave once they are injected into a human
body. Once the liposomal BPD contacts a lipoprotein, the BPD can immediately diffuse
107
across the liposome membrane to the lipoprotein and be carried through the blood stream to
various sites within the body.
3.7 Future Work
The above experimentation is a preliminary look at the interaction of BPD with a
liposome species. There is much more work that can be done in this area with regards to
varying the liposome compositions, solvents and aqueous buffers to determine their effect
on the incorporation and diffusion properties of BPD derivatives within liposomes.
Furthermore, others types of experimentation exist such as fluorescence lifetime
measurements, NMR studies, and kinetics determinations which might shed more light on
this interesting subject.
Chapter 4 Experimental
109
4.1 General Methods for BPD Derivatives
This general section covers the techniques and instruments used for the analysis and
purification of the products.
Elemental Analysis
Microanalyses were carried out in the microanalytical laboratory at the University of
British Columbia by Mr. Peter Borda using a Carlo Erba Elemental Analyzer 1106.
Analysis was attempted on all diester- and diamide-linked BPD derivatives as well as the
metallated BPD diol and BPD triol. In all cases, the BPD diesters and the metallated BPD
alcohols gave acceptable results. Unacceptable results were obtained for most of the BPD
diamides, even after further purification and recrystallization.
Nuclear Magnetic Resonance Spectroscopy
Proton nuclear magnetic resonance (*H NMR) spectra were obtained from samples
in deuteriochloroform ( C D C I 3 ) or hexadeuteriodimethylsulfoxide (d6-DMSO) on a Bruker
AC 200 (200MHz), a Varian XL-300 (300MHz) or a Bruker WH-400 (400MHz)
spectrometer. The chemical shifts are expressed in parts per million (ppm) on the 8 scale
with residual chloroform (8 = 7.24) or dimethylsulfoxide (8 = 2.49) as internal standards.
Signal multiplicities, coupling constants, integration ratios and assignment appear in
parentheses. Selective decouplings were performed on the same instruments.
The carbon-13 NMR ( l 3 c NMR) spectrum was obtained in C D C I 3 with a Varian
XL-300 (75MHz) instrument. The chemical shifts are also reported on the 8 scale using
C D C I 3 as internal standard.
110
Mass Spectroscopy
Low resolution mass spectra (LRMS) were recorded on a Kratos-AEI MS-50
spectrometer. High resolution mass spectra were recorded on a Kratos-AEI MS-50.
Electronic Spectroscopy
Electronic spectra were recorded on a Hewlett Packard Model 8452A diode array
spectrophotometer.
Chromatography
Column chromatography was performed on silica gel 60, 70-230 mesh, supplied
by E. Merck Co. Preparative thin layer chromatography was prepared on pre-coated
10x10cm 0.5mm or 1mm thick Whatman or Merck silica gel plates.
Reagents and Solvents
All chemicals and solvents were reagent or HLPC grade and purified using
literature methods when necessary.
Reaction Conditions
Due to the inherent light sensitivity of these compounds, in particular the vinyl
group, all reactions were performed in a blacked-out fume hood or surrounded by
aluminum foil.
Singlet Oxygen Determination
A lOmM stock solution of cholesterol in EtOAc/MeOH (1:1) and ethanolic solutions
of the appropriate new BPD derivative (20mM) were made. The two solutions were mixed
(1:1 v/v) and then saturated with air. The sample was exposed to a 500W tungsten/halogen
lamp fitted with a red glass filter (to provide light at >600nm) for 2 hours.
I l l
After irradiation, a small aliquot of the sample was transferred to a test tube and the
cholesterol hydroperoxide was reduced to the corresponding alcohol using NaBH4. The
solution was reduced under nitrogen and rediluted in C H 2 C I 2 . An aliquot was spotted on a
silica gel tic plate and developed twice in EtOAc/hexane (1:1). After drying, the plate was
immersed briefly in 5% sulfuric acid in ethanol and heated to 100°C. Production of a
bright blue spot running at Rfs=0.40 relative to the starting cholesterol was seen in every
compound tested. This compound cospotted with 5oc-hydroxycholest-6-en-3P-ol (the
singlet oxygen specific cholesterol by-product) obtained by literature methods.
4.2 General Methods for Fluorescence Quenching Experimentation
This general section covers the materials and instruments used in the fluorescence
quenching experiments
Reagents and Solvents
Dimyristoyl phosphatidylcholine (DMPC) and egg phosphatidylglycerol (EPG)
were purchased from Nippon Fine Chemicals (Osaka, Japan). 2-(3-
((diphenylhexatrienyl)propanoyl)-l-hexadecanoyl-sn-glycero-3-phospho-choline (DPH-
hPC) was purchased from Molecular Probes (Eugene, Oregon). [4-(2-hydroxyethyl)-l-
piperazineethanesulfonic acid] (HEPES) buffer, sodium chloride (NaCl) and hplc grade N-
methylpyrrolidinone (NMP) were purchased from Sigma Chemical. Hplc grade
chloroform ( C H C I 3 ) was purchased from Fischer Scientific and dried over activated 4A
mol sieves. The water used was triple distilled and free of fluorescent impurities.
Polycarbonate filters of defined pore size (100, 200, and 400nm) were purchased
from Nucleopore as were 0.22u.m filters.
112
Preparation of Liposomes
Stock solutions of the lipids and the fluorescent probe were made in HPLC grade
chloroform. Appropriate amounts of these solutions were pipetted into a 25 mL round
bottom flask and dissolved in 5mL chloroform. The flasks were applied to a rotary
evaporator at ca. 30mmHg and rotated at 130rpm. The flask was lowered into the water
bath which was kept at 39°C. The solvent was evaporated quickly with mild boiling of the
solvent. Once the solvent was removed, the flask was removed from the water bath and
kept under vacuum for 20 minutes. The flask was removed and placed within a lyophilizer
for 1 hour at 50^mHg to remove any traces of solvent The appearance of the lipid material
was a thin opaque white film lining the bottom of the flask
Hydration pf Liposomes
To the flask containing the deposited lipid film, lOmL aqueous 50mmolar
HEPES/150mmolar NaCl kept at 40°C were added. The flask was sealed with a ground
glass stopper and placed in a water bath that was maintained at 40°C. While keeping the
flask in the water bath, the flask was gently swirled by hand periodically to effectively
hydrate the lipid material. Each sample was left in the water bath for 1 hour. At this time
no material could be seen attached to the sides of the flask.
Extrusion of Liposomes
Size reduction of the liposomes is facilitated by the use of an extrusion apparatus
(Lipex Biomembrane Inc., Vancouver, B.C.). This apparatus is comprised of the
following components: 2 large 'O' rings, 1 small 'O' ring, filter support base,
113
thermobarrel, Extruder base with wingnuts and washers, inlet manifold, regulator
assembly, filter support disk, filter support mesh, Tygon tubing. Other necessary materials
are Tygon tubing, tube clamps, drain disk (25mm diameter PE, Nucleopore 230600),
400nm polycarbonate filter (25mm diameter PC, Nucleopore 110607), 200nm
polycarbonate filter (25mm diameter PC, Nucleopore 110606), lOOnm polycarbonate filter
(25mm diameter PC, Nucleopore 110605), nitrogen tank.
The extruder was assembled with a 400nm polycarbonate filter and the hydrated
solution was pipetted into the thermobarrel of the extruder which is kept at constant
temperature (40°C) using a circulating constant temperature water bath. The inlet manifold
was placed on the thermobarrel and fastened to the filter support base using wingnuts. The
regulator assembly was attached to the inlet manifold and nitrogen pressure was applied.
Extrusion at 40°C was facile and took place at lOOpsi. The liposomal solution was
extruded through the 400nm filter four more times for a total of 5 passes. This procedure
was repeated for the two remaining filter sizes with concomitant increase in the nitrogen
pressure needed for extrusion (200psi for 200nm filter and 350psi for lOOnm filter).
The extruded material was taken up in a lOcc syringe and filtered through a 220nm
filter into a centrifuge tube. The probe imbedded liposome material was used immediately
or stored in the dark at 4°C until needed. No samples were used that remained in the
refrigerator for more than 2 days. Samples that were stored in the refrigerator were filtered
through a 220nm filter immediately prior to use to ensure no liposome aggregates were
present. Measurement of the fluorescence of the refrigerated samples before and after
showed litde or no difference.
Size Determination of Liposomes
In order to ascertain whether the liposomes were indeed being sized-down by the
extrusion process, the solutions were sized after each filter size. This was performed using
114
the Nicomp Model 370 Submicron Particle Sizer (Particle Sizing Systems, Inc., Santa
Barbara, CA.). The particle sizing after the fifth pass through the 400nm filter the volume-
weighted analysis showed that 99% of the liposomes were less than 444nm. After the fifth
pass at 200nm, the volume-weighted analysis showed that 99% of the vesicles were below
306nm with 75% below 170nm. After the fifth pass at lOOnm, the volume-weighted
analysis showed that the mean diameter of the vesicles was lllnm with 99% of the
distribution below 184nm and 75% below 120nm. The same sample analyzed by intensity-
weighting showed the mean diameter to be 109nm with 99% of the distribution below
180nm. Number-weighting analysis for the post-lOOnm filter vesicles showed a mean
diameter of 90nm with 99% of the distribution below 145nm. These results show that the
extrusion process does size down the liposomes in the desired way.
Fluorescence Instrumentation
The fluorescence measurements were performed on a SLM-AMINCO AMINCO-
Bowman Series 2 Luminescence Spectrometer using a pulsed xenon lamp as the excitation
source. Excitation was at 362nm with 2nm bandwidth and emission was at 433nm with
16nm bandwidth. Fluorescence emissions were corrected for lamp fluctuation using the
reference signal from the excitation light source.
115
4.3 Synthesis of BPD Derivatives
BPD 1,4-diene dimethyl ester 1 3 5 (1). Ring B BPD 1,4-diene dimethyl ester
was acquired from Raylo Chemical and was found to have a large amount of the ring A
isomer as well as protoporphyrin IX dimethyl ester. Recrystallization from boiling ethyl
acetate (1 gram BPD/30mL solvent) gave the pure BPD 1,4-diene dimethyl ester as a
brown powder in 82% yield.: lU NMR (300 MHz, C D C I 3 ) 8 -2.47 (br. s,
2xpyrrolicNH), 2.09 (s, 3H, C H 3 - 7 ) , 3.16 and 3.19 (2t overlap, J=7.7, 7.7 Hz, 4H,
2 X R C H 2 C H 2 C O 2 R ) , 3.40, 3.47, 3.64, 3.65, 3.66 (5s, 15H, C H 3 - 2 , -12, -18 and methyl
ester -13 and -17), 3.60-3.72 (m, 1H, H-73), 3.92-4.03 (m, 1H, H-73), 3.88 and 3.98
(2s, 6H, methyl esters 71 and 72), 4.16 (t, J=7.8 Hz, 2H, R C H 2 C H 2 C O 2 R ) , 4.30 (t,
J=7.7 Hz, 2H, R C H 2 C H 2 C O 2 R ) , 6.15 (dd, J=1.4, 11.4 Hz, 1H, vinyl H-32), 6.35 (dd,
J=1.4, 17.9 Hz, 1H, vinyl H-32), 7.37 (dd, J=2.1, 6.7 Hz, 1H, H-7 4), 8.16 (dd,
J=11.6, 17.8 Hz, 1H, vinyl H-31), 9.20, 9.28, 9.64, 9.78 (4s, 4H, 4xmesoH); MS (EI)
mle calc'd for C 4 2 H 4 4 N 4 O 8 : 732.3159, found 732.3155; 732 (M+), 716 (M+-CH4),
673 (M+-C02CH3), 658 (M+-C02CH3-CH3).
116
BPD 1,3-diene dimethyl ester (2). In a 250mL round bottom flask were
placed a stirbar and BPD ring B 1,4-diene dimethyl ester (4.00g; 5.5xl0 - 3 mol).
Dichloromethane (180mL) was added and stirring was commenced for 5 minutes to
dissolve the solid. To the stirred solution was added 10 drops 1,5-
diazabicyclo[5.4.0]undec-7-ene (DBU) and the flask was sealed. The mixture was allowed
to stir for 26 hours in the dark after which time dc showed full conversion to the desired
compound. The solvent was evaporated in vacuo and the compound was chromatographed
on silica gel (silica gel 60, 70-230 mesh, l%MeOH/CH2Cl2 eluent). The pure 1,3-diene
fraction was evaporated in vacuo yielding 3.59g (90%) of a dark green crystalline solid.:
l H NMR (300 MHz, C D C I 3 ) 8 -2.31 (br. s, 2H, 2xpyrrolicNH), 1.78 (s, 3H, C H 3 - 7 ) ,
2.95 (s, 3H, methyl ester-71), 3.16 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 C H 3 ) , 3.20 (t, J=8
Hz, 2H, R C H 2 C H 2 C O 2 C H 3 ) , 3.42, 3.47, 3.63, 3.65, 3.67 (5s, 15H, C H 3 - 2 , -12, -18
and methyl esters 132 and 172), 4.17 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 C H 3 ) , 4.29 (t, J=8
Hz, 2H, R C H 2 C H 2 C O 2 C H 3 ) , 5.07 (s, 1H, H-7*), 6.17 (d, J=12 Hz, 1H, vinyl H-32),
6.37 (d, J=18 Hz, 1H, vinyl H-32), 7.46 (d, J=7 Hz, 1H, H-73), 7.84 (d, J=7 Hz, 1H,
H-7 4 ) , 8.12 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.15, 9.38, 9.68, 9.77 (4s, 4H,
4xmesoH); 13c NMR (75 MHz, C D C I 3 ) 8 11.16, 11.61, 12.40, 21.42, 21.79, 27.57,
29.68, 36.55, 36.96, 47.89, 51.54, 51.66, 51.74, 52.26, 52.56, 91.87, 93.38, 98.54,
117
99.76, 112.64, 1 2 1 . 1 1 , 121.88, 129.42, 131.13, 133.44, 134.00, 134.04, 136.21,
137.02, 137.15, 137.57, 138.39, 140.07, 151.28, 151.83, 152.39, 156.41, 165.46,
167.62, 170.66, 173.28, 173.70; UV-Vis (EtOH) XmSLX(e) 352 (48,000), 426 (78,200),
576 (17,200), 628 (9,400), 688 (31,400).; MS (electron impact(EI)) mle 732 (M+), 717
( M + - C H 3 ) , 701 ( M + - O C H 3 ) , 673 (M+-CC-2CH3), 658 ( M + - C O 2 C H 3 - C H 3 ) , 599 (M+-
2 X C O 2 C H 3 ) ; Analysis calc'd for C 4 2 H 4 4 N 4 O 8 : C, 68.84; H, 6.05; N, 7.65; found: C,
68.83; H, 6.21; N, 7.68. Positive test for singlet-oxygen production.
BPD dimethyl ester vinyl amine (3). BPD 1,3-diene dimethyl ester
(192.7mg; 2.63xl0"4 mol) was dissolved in dry dichloromethane (20mL) in a 50mL round
bottom flask containing a stirbar. The flask was sealed with a rubber septum and the
contents were left to stir in an ice bath for 10 minutes under an atmosphere of nitrogen. A
light stream of anhydrous hydrogen bromide (HBr) was then bubbled through the reaction
mixture for 10 minutes. After completion of the HBr addition, excess HBr was blown off
using a stream of nitrogen. The crude bromide was redissolved in dry dichloromethane
(20mL) and this was then evaporated under an atmosphere of nitrogen. Dry
dichloromethane (lOmL) was then added and the mixture was allowed to stir in an ice bath.
After several minutes, the contents of the flask were transferred by cannula into a flask
containing a stirbar and dry tetrahydrofuran (20mL) which had been previously saturated
118
with anhydrous ammonia. After stirring at 0°C for 10 minutes, the solution was brought
to room temperature and the solvent was evaporated in vacuo. Tic of the crude residue
showed some starting material and a dominant slower moving spot. The two compounds
were separated on silica gel (silica gel 60, 70-230 mesh) using a two step elution. Initially
the crude mixture was eluted using 20%EtOAc/CH2Cl2 to remove the starting material and
then the solvent polarity was raised to 5%MeOH/CH2Cl2- The appropriate fractions were
pooled and evaporated to yield 147.4mg (75% from the BPD dimethyl ester) BPD 1,3-
diene dimethyl ester vinyl amine as a mixture of two diastereomers.: A H NMR (200 MHz,
CDCI3) 5 -2.43 (br.s, 4H, 4xpyrrolicNH), 1.81 (s, 6H, 2xCH3-7), 2.09-2.21 (m, 6H,
2XCH3-31), 2.86 and 2.90 (2s, 6H, 2xmethyl ester-71), 3.15 and 3.20 (2t overlapping,
8H, 4XRCH2CH2CO2CH3), 3.41, 3.48, 3.61, 3.62, 3.63 (5s, 30H, 2x(CH3-2 -12 -18
and methyl esters-132 and -172)), 3.95 and 3.96 (2s, 6H, 2xmethyl ester-72), 4.18 and
4.33 (2t, 8H, 4XRCH2CH2CO2CH3), 5.09 and 5.11 (2s, 2H, 2XH-71), 5.76-5.92 (m,
2H, 2XH-31), 7.43 and 7.82 (2d, 4H, 2x(H-73 and H-7 4)), 9.15 (s, 1H, mesoH), 9.18
(s, 2H, 2xmesoH), 9.50 (s, 1H, mesoH), 9.69 (s, 2H, 2xmesoH), 9.73 (s, 2H,
2xmesoH); MS (EI) mle 749 (M+), 732 (M+-NH3); UV-Vis (CH2CI2) ?imax(peak ratio)
428 (2.49), 576 (0.54), 626 (0.29), 686 (1.00).
R0 2 C C0 2R
119
BPD dipropyl ester (4). A lOmL round bottom flask was fitted with a stirbar
and BPD 1,3-diene dimethyl ester (90.3mg; 1.23X10"4 mol) was added. Dry 1-propanol
(5mL; 6.69X10'2 mol) was added and the heterogeneous mixture was stirred at room
temperature for 2 hours, after which time 3 drops concentrated sulfuric acid were added.
The mixture turned bright green and became viscous so dichloromethane (lmL) was added.
The reaction mixture was stirred in the dark at room temperature for 44 hours after which
time a tic cospot showed full consumption of the starting material and a single, faster
moving spot. The entire reaction mixture was poured into 5% aqueous sodium acetate
(lOOmL) and extracted with equal volumes of dichloromethane. The organic layer was
washed several times with water and the solvent was evaporated to yield a residue which
was taken up in C H 3 C N and re-evaporated. The compound was redissolved in C H 2 C I 2
and an equal portion of hexanes were added to crystallize the product. Evaporation of
solvent gave 95.1mg (98%) of dark green crystalline product. : X H NMR (300 MHz,
C D C I 3 ) 6 -2.31 (br. s, 2H, 2xpyrrolicNH), 0.78 (t, J=8 Hz, 3H, RC02(CH2)2CH3),
0.81 (t, J=8 Hz, 3H, RC02(CH2)2CH3), 1.48-1.63 (m, 4H, 2 X R C H 2 C H 2 C H 3 ) , 1.76
(s, 3H, C H 3 - 7 ) , 2.94 (3, methyl ester-71), 3.15 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 R ' ) , 3.18
(t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 R ' ) , 3.41, 3.48, 3.62 (3s, 9H, C H 3 - 2 , -12 and -18), 3.97
(s, 3H, methyl ester-72), 4.02 (t, J=8 Hz, 2H, R C O 2 C H 2 C H 2 C H 3 ) , 4.07 (t, J=8 Hz,
2H, R C O 2 C H 2 C H 2 C H 3 ) , 4.17 (t, J=8 Hz, 2H, R C H ^ C ^ C C ^ R ' ) , 4.30 (t, J=8 Hz,
2H, R C H 2 C H 2 C O 2 R ' ) , 5.05 (s, 1H, H-71), 6.15 (d, J=12 Hz, 1H, vinyl H-32), 6.37
(d, J=18 Hz, 1H, vinyl H-32), 7.45 (d, J=8 Hz, 1H, H-73), 7.82 (d, J=8 Hz, 1H, H-74),
8.11 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.14, 9.35, 9.68, 9.75 (4s, 4H, 4xmesoH); MS
(EI) mle calc'd for C 4 6 H 5 2 N 4 O 8 : 788.3788, found 788.3781; 788 (M+), 773 (M+-
C H 3 ) , 729 ( M + - C O 2 C H 3 ) , 714 (M+-C02CH3-CH3); Analysis calc'd for
C 4 6 H 5 2 N 4 O 8 O . 5 H 2 O : C, 69.24; H, 6.70; N, 7.02; found: C, 69.07; H, 6.77; N,
6.75.
120
BPD dibutyl ester (5). To a lOmL round bottom flask containing BPD 1,3-
diene dimethyl ester (97.0 mg; 1.32X10-4mol) and a stirbar, 1-butanol (4mL; 4.37X10"2
mol) was added. Stirring was commenced and dichloromethane (lmL) was added to
solubilize the reagents. 6 drops concentrated sulfuric acid were added to the stirred
solution and the flask was left to stir in the dark for 5 days after which time tic showed full
conversion to the desired dibutyl ester. The reaction was quenched with 5% aqueous
potassium bicarbonate (lOOmL) and extracted with equal portions of dichloromethane.
The organic layer was washed several times with water and dried over sodium sulfate. The
yield after filtration and removal of solvent was 105.8mg (98%) BPD dibutyl ester as dark
green solid.: lU NMR (CDCI3) 5 -2.27 (br. s, 2H, 2xpyrrolicNH), 0.66-0.81 (m, 6H,
2xRC02(CH2)3CH3), 1.11-1.28 (m, 4H, 2 X R C H 2 C H 2 C H 3 ) , 1.42-1.55 (m, 4H,
2XRCH2CH2CH3), 1.81 (s, 3H, CH3-7), 2.93 (s, 3H, methyl ester-71), 3.14 (t, J=8 Hz,
2H, RCH2CH2CO2R') , 3.22 (t, J=8 Hz, 2H, RCH2CH2CO2R') , 3.45, 3.49, 3.62 (3s,
9H, CH3-2, -12 and -18), 3.98 (s, 3H, methyl ester-72), 4.01-4.09 (m, 4H,
2xRC02CH2(CH2)2CH3), 4.20 (t, J=8 Hz, 2H, RCH2CH2CO2R') , 4.32 (t, J=8 Hz,
2H, RCH2CH2CO2R') , 5.07 (s, 1H, H-71), 6.19 (d, J=12 Hz, 1H, vinyl H-32), 6.36
(d, J=18 Hz, 1H, vinyl H-32), 7.49 (d, J=8 Hz, 1H, H-73), 7.82 (d, J=8 Hz, 1H, H-74),
8.08 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.23, 9.46 (2s, 2H, 2xmesoH), 9.87 (br. s, 2H,
2xmesoH); MS (EI) mle calc'd for C48H56N4O8: 816.4101, found 816.4094; 816 (M+),
801 (M+-CH3), 774 (M+-CH3-C2H3), 757 (M+-CO2CH3), 742 (M+-CH3-CO2CH3);
Analysis calc'd for C48H56N4O8O.5H2O: C, 69.80; H, 6.95; N, 6.78; found: C,
69.98; H, 6.89; N, 6.61.
BPD dipentyl ester (6). BPD 1,3-diene dimethyl ester (107.4mg; 1.47X10"
4mol) was placed in a lOmL round bottom flask along with dry 1-pentanol (4mL;
3.68X10"2mol) and dichloromethane (lmL). The reagents were stirred for several minutes
at which time 4 drops concentrated sulfuric acid were added. The flask was stirred at room
temperature in the dark and the reaction progress was monitored by dc. After 5 days, a tic
121
cospot with the starting compound showed full conversion to the desired dipentyl ester.
The reaction was quenched using 5% aqueous potassium bicarbonate (lOOmL) and the free
base chlorin was extracted with dichloromethane (lOOmL). The organic layer was washed
twice more with bicarbonate (2xl00mL) and then with water (3xl00mL). After removal of
the dichloromethane in vacuo, the excess pentanol was removed by evaporation on the
vacuum line for two days and the residue was chromatographed on silica gel (silica gel 60,
70-230 mesh, 0.5%MeOH/CH2Cl2 eluent). The appropriate fractions were pooled and
evaporated to yield 121.2mg (98%) BPD 1,3-diene dipentyl ester.: lH NMR (400 MHz,
C D C I 3 ) 8 -2.29 (br. s, 2H, 2xpyrrolicNH), 0.61 (m, 6H, 2xRCC>2(CH2)4CH3), 0.97-
1.22 (m, 8H, 2 X R C H 2 C H 2 C H 3 ) , 1.38-1.46 (m, 4H, 2xRCH2(CH2)2CH3), 1.79 (s,
3H, C H 3 - 7 ) , 2.95 (s, 3H, methyl ester-71), 3.14 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 R ' ) ,
3.18 (t, J=8 Hz, 2H, RCH2CH2C02R'), 3.42, 3.47, 3.76 (3s, 9H, C H 3 - 2 , -12 and -18),
3.98 (s, 3H, methyl ester-72), 4.03 (t, J=7 Hz, 2H, R C O 2 C H 2 R ' ) , 4.04 (t, J=7 Hz, 2H,
R C O 2 C H 2 R ' ) , 4.18 (t, J=8 Hz, 2H, RCH2CH2e02R'), 4.30 (t, J=8 Hz, 2H,
R C H 2 C H 2 C O 2 R ' ) , 5.07 (s, 1H, H-71), 6.15 (d, J=12 Hz, 1H, vinyl H-32), 6.36 (d,
J=18 Hz, 1H, vinyl H-32), 7.45 (d, J=7 Hz, 1H, H-73), 7.82 (d, J=7 Hz, 1H, H-74),
8.11 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.14, 9.36, 9.68, 9.74 (4s, 4H, 4xmesoH); MS
(EI) mle calc'd for C 5 0 H 6 O N 4 O 8 : 844.4414, found 844.4419; 844 (M+), 829 (M+-
C H 3 ) , 785 (M+-C02CH3), 770 (M+-C02CH3-CH3), 757 (M+-C02CH3-2xCH3);
Analysis calc'd for C 5 0 H 6 O N 4 O 8 : C, 71.07; H, 7.16; N, 6.63; found: C, 70.97; H,
7.18; N, 6.66.
BPD dihexyl ester (7). BPD 1,3-diene dimethyl ester (81.7mg; 1.12X10"4
mol) and 1-hexanol (3mL; 2.39xl0"2 mol) were placed in a lOmL round bottom flask
which was equipped with a stirbar. After adding dichloromethane (lmL), stirring was
commenced and 4 drops concentrated sulfuric acid were added. The mixture was stirred at
room temperature in the dark for 5 days after which time tic showed full conversion to the
desired diester. The reaction was quenched using 5% aqueous potassium bicarbonate
122
(lOOmL) and extracted with dichloromethane (2xl00mL). After several water washings
(4xl00mL), the organic solvent was removed and the flask was placed on the vacuum line
to remove excess 1-hexanol. The final weight of dry B P D dihexyl ester was 94.2mg
(97%).: ! H NMR (300 MHz, C D C I 3 ) 8 -2.29 (s, 2H, 2xpyrrolicNH), 0.68-0.79 (m, 6H,
2xRC02(CH2)5CH3), 0.95-1.24 (m, 12H, 2xR(CH2)3CH3), 1.40-1.54 (m, 4H,
2 X O C H 2 C H 2 C 4 H 9 ) , 1.81 (s, 3H, C H 3 - 7 ) , 2.97 (s, 3H, methyl ester-71), 3.15 (t, J=8
Hz, 2H, R C H 2 C H 2 C O 2 R ' ) , 3.19 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 R ) , 3.42, 3.48, 3.63
(3s, 9H, C H 3 - 2 , -12 and -18), 3.99 (s, 3H, methyl ester-72), 4.01-4.11 (m, 4H,
2 X R C O 2 C H 2 C 5 H 1 1 ) , 4.18 (t, J=2 Hz, 8H, RCH2CH2C02R'), 4.31 (t, J=2 Hz, 8H,
R C H 2 C H 2 C O 2 R ' ) , 6.17 (d, J=12 Hz, 1H, vinyl H-32), 6.38 (d, J=18 Hz, 1H, vinyl H-
32), 7.46 (d, J=8 Hz, 1H, H-73), 7.83 (d, J=8 Hz, 1H, H-74), 8.12 (dd, J=12, 18 Hz,
1H, vinyl H-31), 9.15, 9.36, 9.69, 9.75 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C 5 2 H 6 4 N 4 O 8 : 872.4727, found 872.4724; 872 (M+), 857 (M+-CH3), 816 (M+-
C O 2 C H 3 ) , 798 (M+-CH3-C02CH3); Analysis calc'd for C 5 2 H 6 4 N 4 O 8 : C, 71.54; H,
7.39; N, 6.42; found: C, 71.23; H, 7.30; N, 6.30.
BPD diheptyl ester (8). B P D 1,3-diene dimethyl ester (91.1mg; 1.24X10"
4mol) and 1-heptanol (3mL; 2.12xl0"2 mol) were placed in a lOmL round bottom flask
containing a stirbar. Stirring was commenced upon addition of dichloromethane (lmL) and
after five minutes 4 drops concentrated sulfuric acid were added. The reaction was allowed
to proceed for 5 days after which time tic showed no starting material remained. The
reaction was quenched with 5% aqueous sodium acetate (lOOmL) and extracted with
dichloromethane (2xl00mL). The organic layer was washed several times with water
(3xl00mL) and after removal of the C H 2 C I 2 the flask was placed on the vacuum line for 3
days to remove the excess 1-heptanol. The weight of dry B P D diheptyl ester was 106.4mg
(92%).: i H NMR ( C D C I 3 ) 8 -2.92 (s, 2H, 2xpyrrolicNH), 0.54-0.84 (m, 6H,
2xRC02(CH2)6CH3), 0.93-1.31 (m, 16H, 2xRC02(CH2)2(CH2)4CH3), 1.38-1.56 (m,
4H, 2 X R C O 2 C H 2 C H 2 C 5 H 1 1 ) , 1.82 (s, 3H, C H 3 - 7 ) , 2.97 (s, 3H, methyl ester-71),
123
3.09-3.34 (m, 4H, 2 X R C H 2 C H 2 C O 2 R ' ) , 3.43, 3.49, 3.66 (3s, 9H, C H 3 - 2 -12 and -18),
3.99 (s, 3H, methyl ester-72), 4.00-4.10 (m, 4H, 2 X R C O 2 C H 2 C 6 H 1 3 ) , 4.19 (t, J=8 Hz,
2H, RCH.2CH2C02R'), 4.30 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 R ) , 5.08 (s, 1H, H-71),
6.18 (d, J=12 Hz, 1H, H-32), 6.39 (d, J=18 Hz, 1H, H-32), 7.47 (d, J=7 Hz, 1H, H-
73), 7.85 (d, J=7 Hz, 1H, H-74), 8.13 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.16, 9.36,
9.69, 9.75 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for C 5 4 H 6 8 N 4 O 8 : 900.5040, found
900.5030; 900 (M+), 885 ( M + - C H 3 ) , 841 (M+-C02CH3); Analysis calc'd for
C54H68N408-H20: C, 70.56; H, 7.68; N, 6.10; found: C, 70.60; H, 7.53; N, 5.94.
BPD dioctyl ester (9). BPD 1,3-diene dimethyl ester (59.7mg; 8.15xl0"5 mol)
was dissolved in dichloromethane (20rnL) in a 50 mL round bottom flask fitted with a
stirbar. To the stirring solution was added 1-octanol (1.7mL; 1.08xl0_2mol) and 8 drops
of concentrated sulfuric acid. The bright green solution was stirred at room temperature
and monitored by tic. After 48 hours the reaction was deemed complete by tic. The
contents of the flask were transferred to a separately funnel and neutralized with 0.1M
aqueous ammonium acetate (50mL). The organic layer was washed with water (3x50mL)
and organic solvent was removed and the residue evaporated under high vacuum for two
days to remove excess 1-octanol. The yield of dry BPD 1,3-diene dioctyl ester was
70.5mg (93%).: lH NMR (200 MHz, C D C I 3 ) 8 -2.30 (br. s, 2H, 2xpyrrolicNH), 0.70-
0.84 (m, 6H, 2xR(CH2)7CH3), 0.95-1.21 (m, 20H, 2xR(CH2)2(CH2)5CH3), 1.35-
1.52 (m, 4H, 2xRCH2CH2C6Hi3), 1.76 (s, 3H, CH3-7), 2.94 (s, 3H, methyl ester-71),
3.15 and 3.18 (2t overlapping, J=8, 8 Hz, 4H, 2 X R C H 2 C H 2 C O 2 R ' ) , 3.43, 3.49, 3.65
(3s, 9H, C H 3 - 2 , -12 and -18), 3.99 (s, 3H, methyl ester-72), 4.00-4.11 (m, 4H,
2 X R C O 2 C H 2 C 7 H 1 5 ) , 4.19 (t, J=8 Hz, 2H, R C H 2 C H 2 C O 2 R ' ) , 4.31 (t, J=8 Hz, 2H,
R C H 2 C H 2 C O 2 R ' ) , 5.07 (s, 1H, H-71), 6.17 (d, J=12 Hz, 1H, vinyl H-32), 6.37 (d,
J=18 Hz, 1H, vinyl H-32), 7.46 (d, J=8 Hz, 1H, H-73), 7.83 (d, J=8 Hz, 1H, H-74),
8.13 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.15, 9.36, 9.70, 9.75 (4s, 4H, 4xmeso H); MS
(EI) mle calc'd for C 5 6 H 7 2 N 4 O 8 : 928.5354, found 928.5343; 928 (M +), 869 (M+-
124
C O 2 C H 3 ) , 854 (M+-C02CH3-CH3), 830 ( M + - ( C H 2 ) 7 C H 3 ) , 815 ( M + - ( C H 2 ) 7 C H 3 -
C H 3 ) ; Analysis calc'd for C 5 6 H 7 2 N 4 O 8 : C, 72.39; H, 7.81; N, 6.03; found: C, 72.10;
H, 7 . 9 5 ; N, 5.82.
RNHCO CONHR
BPD dihexyl amide (10). BPD 1,3-diene dimethyl ester (220.9mg; 3.01x10-
4mol) and n-hexylamine (12mL; 9.08xl0_2mol) were placed in a 50mL round bottom
flask with a magnetic stirbar. The reagents were solubilized with freshly distilled
tetrahydrofuran (12mL) and the flask was fitted with a reflux condenser. The mixture was
then brought to reflux for 4 days after fitting the flask with a condenser. Tic of the mixture
showed conversion to a slower moving compound with substantial baseline material seen.
The reaction was stopped and the tetrahydrofuran was evaporated in vacuo. The oily
residue was dissolved in dichloromethane (lOOmL) and washed with dilute aqueous
hydrochloric acid (2xl00mL) and water (3xl00mL) and finally evaporated under high
vacuum. The crude compound was chromatographed on silica gel (silica gel 60, 70-230
mesh, 2-4%MeOH/CH2Cl2 as gradient eluent). The appropriate fractions were pooled and
dried to afford 128.5mg (49%) of BPD dihexyl amide with the faster moving fractions
starting material and the two monoamide/monoester compounds.: *H NMR (400 MHz,
C D C I 3 ) 5 -2.31 (s, 2H, 2xpyrrolicNH), 0.42 (t, J=7.0 Hz, 3H, R(CH2)5CH3), 0.52 (t,
J=6.7 Hz, 3H, R(CH2)5CH3), 0.57-0.76 (m, 4H, 2xR(CH2)4CH2CH3), 0.77-0.99 (m,
125
8H, 2xR(CH2)2(CH2)2C2H5), 1.06-1.29 (m, 4H, 2XRCH2CH2C4H9), 1.75 (s, 3H,
C H 3 - 7 ) , 2.89-3.10 (m, 11H, methyl ester-71 and 2xRCH2CH2CONHR' and
2 X R C O N H C H 2 C 5 H 1 1 ) , 3.40, 3.47, 3.62 (3s, 9H, CH3-2, -12 and -18), 3.98 (s, 3H,
methyl ester-72), 4.16 (t, J=7.3 Hz, 2H, RCH^C^CONHR') , 4.30 (t, J=7.6 Hz, 2H,
RCH2CH2CONHR') , 5.04 (s, 1H, H-71), 6.14 (d, J=11.5 Hz, 1H, vinyl H-32), 6.29 (t,
J=5.5 Hz, 1H, R(CH2)2CONHR'), 6.36 (d, J=17.8 Hz, vinyl H-32), 6.72 (t, J=5.4 Hz,
1H, R(CH2)2CONHR'), 7.43 (d, J=5.7 Hz, 1H, H-73), 7.81 (d, J=5.9 Hz, 1H, H-74),
8.10 (dd, J=11.6, J=17.8 Hz, 1H, vinyl H-31), 9.11, 9.32, 9.71, 9.81 (4s, 4H,
4xmesoH); MS (EI) mle calc'd for C52H66N6O6: 870.5044, found 870.5055; 871 (M+).
BPD dioctyl amide (11). BPD 1,3-diene dimethyl ester (220mg; 3.00X10"
4mol) and n-octylamine (15mL; 9.08X10"2mol) were placed in a 50mL round bottom flask
along with a stirbar. The reagents were solubilized with tetrahydrofuran (15mL) and the
flask was fitted with a condenser. The reaction mixture was brought to reflux and the
reaction progress was monitored by tic. After 5 days, much of the starting material had
been consumed and a predominant slower moving spot was noticed along with substantial
amounts of baseline material. The reaction was stopped and the tetrahydrofuran evaporated
in vacuo. The residue was then evaporated under high vacuum for 3 days to remove
excess n-octylamine. The crude product was chromatographed on silica gel (silica gel 60,
70-230 mesh, 2%MeOH/CH2Cl2 as eluent). The appropriate fractions were pooled,
evaporated and placed in a vacuum oven at 65°C for 2 days to remove traces of amine.
The final weight of the desired BPD dioctyl amide was 134.9mg (49%).: X H NMR (400
MHz, CDCI3) 8 -2.32 (s, 2H, 2xpyrrolicNH), 0.62-0.78 (m, 10H, 2xR(CH2)7CH3 and
2xR(CH2)6CH2CH3), 0.80-0.97 (m, 12H, 2xR(CH2)3(CH2)3C2H5), 0.98-1.16 (m,
4H, 2xR(CH2)2CH2C5Hn), 1.17-1.38 (m, 4H, 2XRCH2CH2C6H13), 1.76 (s, 3H,
CH3-7), 2.87-3.12 (m, 8H, 2xRCH2CH2CONHR' and 2XRCONHCH2C7H15), 2.95
(s, 3H, methyl ester-71), 3.37, 3.45, 3.59 (3s, 9H, CH3-2 -12 -18), 3.97 (s, 3H, methyl
ester-72), 4.14 (t, J=7.3 Hz, 2H, R C H 2 C H 2 C O N H R ' ) , 4.27 (t, J=7.6 Hz, 2H,
126
R C H 2 C H 2 C O N H R ' ) , 5.06 (s, 1H, H-71), 6.15 (d, J=11.5 Hz, 1H, vinyl H-32), 6.27-
6.40 (m, 2H, RCONHR' and vinyl H-32), 6.72 (t, J=5.4 Hz, 1H, RCONHR'), 7.42 (d,
J=5.7 Hz, 1H, H-7 3 ), 7.81 (d, J=5.7 Hz, 1H, H-7 4 ), 8.10 (dd, J=11.6, 17.8 Hz, 1H,
vinyl H-3 1), 9.12, 9.32, 9.69, 9.79 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C56H74N6O6: 926.5670, found 926.5689; 926 (M+).
BPD diacid (12). BPD 1,3-diene dimethyl ester (206.4mg; 2.82xl0"4mol) was
dissolved in 25% aqueous hydrochloric acid (15rnL) and stirring was commenced at room
temperature. After 45 minutes tic showed good conversion to the desired diacid with small
amounts of the 2 possible monoacid/monoester products. The mixture was transferred to a
separatory funnel and diluted to lOOmL with water. An equal portion of 10%
methanol/dichloromethane was added to extract the aqueous layer. After repeating 5 times,
the combined organic layers were pooled, evaporated to lOOmL total volume, washed with
water (3xl00mL), 5% aqueous potassium bicarbonate (3xl00mL), and again water
(lxlOOmL). The organic solvent was evaporated in vacuo and the residue was further
evaporated under high vacuum for 16 hours. Attempts to purify large quantities of this
compound were unsuccessful, but a small sample was isolated for characterization by
preparative chromatography on Whatman 0.5mm silica gel plates using
10%MeOH/CH2Cl2 with several drops acetic acid added to the developing chamber. The
127
crude BPD diacid was carried over to the various displacement reactions.: NMR (300
MHz, DMSO-d6) 8 -2.42 (br. s, 2H, 2xpyrrolicNH), 1.75 (s, 3H, C H 3 - 7 ) , 2.67-2.98
(m, 7H, 2xRCH2CH2C02H and methyl ester-71), 3.35, 3.46, 3.64 (3s, 9H, C H 3 - 2 -12
and -18), 3.91 (s, 3H, methyl ester-72), 3.99-4.12 (m, 2H, R C H 2 C H 2 C O 2 H ) , 4.18-4.32
(m, 2H, R C H 2 C H 2 C O 2 H ) , 5.21 (s, 1H, H-71), 6.17 (d, J=12 Hz, 1H, vinyl H-32),
6.43 (d, J=18 Hz, 1H, vinyl H-32), 7.79 (s, 2H, H-7 3 and -74), 8.37 (dd, J=12, 18 Hz,
1H, vinyl H-31), 9.35, 9.62, 9.84, 10.34 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C 4 0 H 4 0 N 4 O 8 : 704.2846, found 704.2842; 718 (M++14), 704 (M+).
RNHCO CONHR
BPD dimethyl amide (13). A 25mL 3 neck round bottom flask fitted with a
stirbar was flame-dried three times under N2- Upon cooling, crude BPD diacid (61.9mg;
8.8xl0"5mol) was added and dissolved in dichloromethane (12mL) which had been filtered
through neutral alumina prior to use. The middle septum was quickly replaced with a
condenser fitted with a drying tube and stirring was commenced . Under a nitrogen
atmosphere oxalyl chloride (0.50rnL; 5.73X10-3mol) was added to the reaction by syringe
and the mixture was brought to reflux on a water bath. After 30 minutes the water bath
was removed and the flask was cooled to room temperature. The condenser was replaced
with a rubber septum and the solution was evaporated under a strong nitrogen flow.
When dry, the crude diacid chloride was redissolved in dry dichloromethane (12mL) and
128
blown dry, again using a nitrogen flow. After repeating this procedure a third time, the
residue was taken up in dry dichloromethane (12mL) and with stirring and light nitrogen
flow maintained, 40% aqueous methyl amine (0.15mL; 1.74X10-3mol) was added by
syringe. The bright green protonated solution turned dull green immediately and the
mixture was allowed to stir at room temperature. After 40 minutes the septum was
removed and the solvent was evaporated in vacuo. The following day the crude mixture
was chromatographed on silica gel (silica gel 60, 70-230 mesh, 5%MeOH/CH2Cl2 eluent)
and the appropriate fractions were pooled and evaporated in vacuo to yield 43.0mg BPD
dimethyl amide (67% from the BPD 1,3-diene dimethyl ester).: *H NMR (300 MHz,
C D C I 3 ) 5 -2.41 (br. s, 2H, 2xpyrrolicNH), 1.78 (s, 3H, C H 3 - 7 ) , 2.49 (d, J=4.0 Hz, 3H,
R N H C H 3 ) , 2.55 (d, J=4.0 Hz, 3H, R N H C H 3 ) , 2.84-3.04 (m, 2xRCH2CH2COR' and
methyl ester-71), 3.39, 3.43, 3.60 (3s, 9H, C H 3 - 2 , -12 and -18), 3.97 (s, 3H, methyl
ester-7 2), 4.12 (t, J=8 Hz, 2H, RCH9CH?COR' ) . 4.25 (t, J=8 Hz, 2H,
RCH2CH2COR"), 5.06 (s, 1H, H-71), 6.17 (d, J=12 Hz, 1H, vinyl H-32), 6.35 (d,
J=18 Hz, 1H, vinyl H-32), 6.51 (m, 1H, R C O N H C H 3 ) , 6.86 (m, 1H, R C O N H C H 3 ) ,
7.45 (d, J=6.5 Hz, 1H, H-73), 7.82 (d, J=6.5 Hz, 1H, H-74), 8.09 (dd, J=12, 18 Hz,
1H, vinyl H-31), 9.15, 9.37, 9.74, 9.80 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C 4 2 H 4 6 N 6 O 6 : 730.3482, found 730.3470; 730 (M+), 715 (M+-CH3), 672 (M+-
C O N H C H 3 ) , 656 (M+-C02CH3-CH3).
BPD dibutyl amide (14). A three neck 50mL round bottom flask was fitted
with septa and a stirbar then flame-dried under nitrogen 3 times. Upon cooling, crude BPD
diacid (150mg; 2.13xl0-4mol) were quickly added and the septum was replaced. The
reagent was dissolved in dry dichloromethane (15mL) which was added by syringe via the
side septum. After a 5 minutes stirring, oxalyl chloride (0.50mL; 5.22xl0"3moi) were
added by syringe and the middle septum was replaced with a condenser fitted with a
septum. The mixture was brought to reflux for 20 minutes using a warm water bath.
Upon cooling the condenser was replaced with the septum and the nitrogen flow was
129
increased to blow off excess oxalyl chloride and the dichloromethane. The dry residue was
redissolved in dry dichloromethane (15mL) and again blown down to dryness. Another
portion of dichloromethane was added and stirring was commenced under a light flow of
nitrogen. After the bright green solution was solubilized, n-butyl amine (0.5mL; 5.06x10"
3mol) was added dropwise. The solution turned a dull green denoting the free-base
chlorin. The crude product was diluted with CH2CI2 to lOOmL and washed with water
(3xl00mL). The organic layer was then evaporated and the crude residue was
chromatographed on silica gel (silica gel 60, 70-230 mesh, 5%MeOH/CH2Cl2 as eluent).
The appropriate fractions were pooled and evaporated to provide 95.6mg of BPD dibutyl
amide (55% from the BPD 1,3-diene dimethyl ester).: lH NMR (400 MHz, CDCI3) 8
-2.30 (br. s, 2H, 2xpyrrolicNH), 0.32 (t, J=7.3 Hz, 3H, RCONH(CH2)3Qi3), 0.51 (t,
J=7.3 Hz, 3H, RCONH(CH2)3CH.3), 0.75-0.86 (m, 2H, RCONH(CH2)2CH2CH3),
0.87-1.04 (m, 4H, RCONH(CH2)2CH2CH3 and R C O N H C H 2 C H 2 C H 2 C H 3 ) , 1.10-
1.19 (m, 2H, R C O N H C H 2 C H 2 C H 2 C H 3 ) , 1.76 (s, 3H, CH3-7), 2.85-3.13 (m, 11H,
2xRCH2CH2CONHR and 2XRCONHCH2C3H7 and methyl ester-71), 3.41, 3.47, 3.61
(3s, 9H, CH3-2, -12 and -18), 4.17 (t, J=7.4 Hz, 2H, R C H 2 C H 2 C O N H R ) , 4.30 (t,
J=7.6 Hz, 2H, R C H 2 C H 2 C O N H R ) , 5.05 (s, 1H, H-71), 6.15 (dd, J=11.5, 1.0 Hz, 1H,
vinyl H-32), 6.27 (t, J=5.4 Hz, 1H, RCONHR), 6.34 (dd, J=17.8, 1.1 Hz, 1H, vinyl H-
32), 6.71 (t, J=5.4 Hz, 1H, RCONHR), 7.43 (d, J=5.7 Hz, 1H, H-73), 7.81 (d, J=5.7
Hz, 1H, H-74), 8.10 (dd, J=11.5, 17.8 Hz, 1H, vinyl H-31), 9.12, 9.33, 9.72, 9.83 (4s,
4H, 4xmesoH); MS (EI) mle calc'd for C48H58N6O6: 814.4418, found 814.4402; 814
(M+), 799 (M+-CH3); Analysis calc'd for C48H58N6O6O.5H2O: C, 69.96; H, 7.22; N,
10.20; found: C, 69.97; H, 7.33; N, 10.09. Positive test for singlet-oxygen production.
BPD di(4-(3-aminopropyl)morpholine) amide (15). A 3 neck round
bottom flask containing a stir bar was flame-dried three times under nitrogen. After cooling
to room temperature, crude BPD diacid (59.6mg; 8.45X10"5mol) was added to the flask.
The middle septum was at this time replaced with a condenser fitted with a septum. Dry
130
dichloromethane (20mL) was added by syringe through the side septum. Stirring was
commenced and the mixture was refluxed for 30 minutes to solubilize the reagent. Upon
cooling, oxalyl chloride (0.50mL; 5.73X10-3mol) was added dropwise and the mixture
was again brought to reflux for 40 minutes. Upon cooling, the nitrogen flow was increased
to blow off the solvent and the excess oxalyl chloride. When the residue was dry, another
portion of dry dichloromethane (15mL) was syringed in and subsequently blown off. The
mixture was stirred after adding dichloromethane (20mL) and 4-(3-aminopropyl)
morpholine (0.32mL; 2.19X10-3mol) was added and the solution was stirred at room
temperature for 40 minutes. The mixture was transferred to a separatory funnel and, after
adding methanol (5mL), washed with 5% aqueous potassium bicarbonate (3x50mL). The
organic layer was then washed with distilled water (3x50mL) and dried over sodium sulfate
overnight. This solution was filtered and evaporated in vacuo and chromatographed on
silica gel (silica gel 60, 70-230 mesh, 10%MeOH/CH2Cl2 eluent). The appropriate
fractions were collected to yield 56.8mg (70%) of the diamide.: lH NMR (300 MHz,
C D C I 3 ) 8 -2.33 (s, 2H, 2xpyrrolicNH), 1.02-1.13 (m, 2H, R C O N H C H 2 C H 2 C H 2 R ' ) ,
1.18-1.32 (m, 2H, R C O N H C H 2 C H_2C H 2 R ' ) , 1.35-1.48 (m, 4H,
2 X R C O N H C H 2 C H 2 C H _ 2 R * ) , 1.8.1 (s, 3H, C H 3 - 7 ) , 1.86-2.06 (m, 4H,
2XRCONHCH2CH2CH2R'), 2.93-3.28 (m, 23H, 2 X R C H 2 C H 2 C O R ' , methyl ester-71
and 2XRN(CH2CH2)20), 3.43, 3.50, 3.63 (3s, 9H, C H 3 - 2 -12 and -18), 4.00 (s, 3H,
methyl ester-72), 4.19 (t, J=8 Hz, 2H, R C H 2 C H 2 C O N H R ' ) , 4.33 (t, J=8 Hz, 2H,
R C H 2 C H 2 C O N H R ' ) , 5.08 (s, 1H, H-71), 6.18 (d, J=ll Hz, 1H, vinyl H-32), 6.34 (d,
J=18 Hz, 1H, vinyl H-32), 7.40 (m, 1H, RCONHR'), 7.45 (d, J=6 Hz, 1H, H-73), 7.60
(m, 1H, RCONHR"), 7.83 (d, J=6 Hz, 1H, H-74), 8.11 (dd, J=ll, 18 Hz, 1H, vinyl H-
31), 9.13, 9.35, 9.73, 9.80 (4s, 4H, 4XmesoH); MS (EI) mle calc'd for C 5 4 H 6 8 N 8 O 8 :
956.5164, found 956.5162; 956 (M+); Analysis calc'd for C54H68Ng08-MeOH: C,
66.78; H, 7.34; N, 11.33; found: C, 66.72; H, 7.33; N, 10.89.
131
BPD di(N,N-dimethylethylenediamine)amide (16). A three neck 50mL
round bottom flask was fitted with the appropriate septa and flame-dried under nitrogen
three times. The middle septum was briefly removed and crude BPD diacid (105.9mg;
1.50xl0-4mol) was added. The solid was dissolved in dichloromethane (15mL) and
briefly heated. After cooling, oxalyl chloride (0.40mL; 4.59xl0_3mol) were added
dropwise through a syringe. The mixture was brought to reflux for 20 minutes and which
time the nitrogen flow was increased to blow off the solvent and excess oxalyl chloride.
The residue was taken up in dry dichloromethane (15mL) and blown dry again. The
residue was dissolved in dichloromethane and N,N-dimethylethylene diamine (0.30mL;
2.73xl0"3mol) was added by syringe under nitrogen. The mixture was diluted to lOOmL
and washed with 0.1M aqueous hydrochloric acid (3xl00mL), 5% aqueous potassium
bicarbonate (lxlOOmL) followed by water (3xl00mL). The organic layer was dried over
sodium sulfate. The solvent was removed and the crude compound was chromatographed
on silica gel (silica gel 60, 70-230 mesh, 15%MeOH/CH2Cl2 eluent) to yield 65.7mg
(52%) of the desired diamide as a green powder.: ! H NMR (400 MHz, C D C I 3 ) 8 -2.40
(s, 2H, 2xpyrrolicNH), 1.75 (s, 3H, C H 3 - 7 ) , 1.85 (s, 6H, RNHCH2CH2N(CH_3)2),
1.97 (s, 6H, RNHCH2CH2N(CH3)2), 1.98 (t, J=7 Hz, 2H, R N H C H 2 C H 2 R ' ) , 2.12 (t,
J=7 Hz, 2H, R N H C H 2 C H 2 R ' ) , 2.89 (s, 3H, methyl ester-71), 2.92 (t, J=8 Hz, 2H,
R C H 2 C H 2 C O N H R 1 ) , 2.98 (t, J=8 Hz, 2H, R C H 2 C H 2 C O N H R ' ) , 3.02 (dt, J=5, 7 Hz,
2H, RNHCH2CH2NR'2), 3.10 (dt, J=5, 7 Hz, 2H, R N H C H ^ C ^ N R ^ ) , 3.38, 3.49,
3.65 (3s, 9H, C H 3 - 2 , -12 and -18), 3.91 (s, 3H, methyl ester-72), 4.06 (t, J=8 Hz, 2H,
R C H 2 C H 2 C O N H R ' ) , 4.25 (t, J=8 Hz, 2H, R C H 2 C H 2 C O N H R ' ) , 5.21 (s, 1H, H-71),
6.18 (d, J=12 Hz, 1H, vinyl H-32), 6.43 (d, J=18 Hz, 1H, vinyl H-32), 7.78 (br. s, 2H,
H-7 3 and H-74), 7.91 (t, J=5 Hz, 1H, RCONHR'), 7.98 (t, J=5 Hz, 1H, RCONHR'),
8.35 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.36, 9.59, 9.84, 9.86 (4s, 4H, 4xmesoH),
New peaks confirmed by decoupling; MS (EI) mle calc'd for C 4 8 H 6 O N 8 O 6 : 844.4636,
found 844.4638; 844 (M+), 799 (M+-C2H7N).
132
BPD di(N,N,N-trimethylethylenediamineammonium iodide) (17).
BPD di(N,N-dimethylethylenediarnine)arnide (16) (12mg; 1.42xl0"5mol) was dissolved
in dry acetone (5mL) and swirled to solubilize the reagent Methyl iodide (2mL; 3.21x10"
2mol) was added by pipet and the flask was swirled for 10 minutes. The solvent and
excess methyl iodide were evaporated in vacuo and the flask was placed under high
vacuum overnight. The weight of the desired diammonium iodide was 16mg (100%).: *H
NMR (400 MHz, DMSO-d6) 8 -2.3.9 (s, 2H, 2xpyrrolicNH), 1.77 (s, 3H, C H 3 - 7 ) , 2.16
and 2.66 (2, 18H, 2xRCONHCH2CH 2N(CH_3)3), 2.77-2.87 (m, 2H,
RCONHCH2CH_2N(CH3)3) , 2.89-3.07 (m, 9H, 2xRCH2CH_2CONHR\
R C O N H C H 2 C H _ 2 N ( C H 3 ) 3 and methyl ester-71), 3.21-3.38 (m, 4H,
2xRCONHCH2CH2N(CH3)3), 3.42, 3.53, 3.68 (3s, 9H, CH3-2-12 and-18), 3.92 (s,
3H, methyl ester-72), 4.10 and 4.29 (2t, J=8 Hz, 4H, 2xRCH2CH2CONHR), 5.25 (s,
1H,.H-71), 6.20 (d, J=12 Hz, 1H, vinyl H-32), 6.45 (d, J=18 Hz, 1H, vinyl H-32), 7.79
(br.s, 2H, H - 7 3 and -7 4 ), 8.08 and 8.16 (2t, J=7 Hz, 2H,
2xRCONHCH2CH2N(CH-3)3), 8.38 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.40, 9.62,
9.83, 9.90 (4s, 4H, 4xmesoH).
C H 3 0 2 C C 0 2 C H 3
133
BPD dimethyl ester (Eschenmoser) (18). BPD 1,3-diene dimethyl ester
(49.6mg; 6.77X10"5mol) and N^V-dimethylmethyleneammonium iodide (242.0mg;
1.31X10-3mol) were placed in a lOmL flask containing a stirbar. Dichloromethane (4mL)
was added and the mixture was allowed to stir at room temperature in the dark The
reaction progress was monitored by tic and after 68 hours a cospot of the reaction mixture
showed no starting material remained and a predominant slower moving compound had
appeared. The mixture was transferred to a separatory funnel, diluted with
dichloromethane (lOOmL) and washed three times with equal portions of water. The
organic layer was dried over sodium sulfate and, after filtration and evaporation, the
compound was chromatographed on silica gel (silica gel 60, 70-230 mesh,
6%MeOH/CH2Cl2 eluent). The appropriate fractions were pooled, evaporated,
redissolved in dichloromethane and crystallized by addition of an equal portion of hexanes.
The crystals were filtered to yield 47.2mg (88%) BPD 1,3-diene dimethyl ester
(Eschenmoser).: ! H NMR (400 MHz, DMSO-d6) 8 -2.40 (s, 2H, 2xpyrrolicNH), 1.77
(s, 3H, C H 3 - 7 ) , 2.70 (s, 6H, RCH2N(CH3)2), 2.88 (s, 3H, methyl ester-71), 3.12 (t,
J=7.3 Hz, 2H, R C H 2 C H 2 C O 2 C H 3 ) , 3.18 (t, J=7.2 Hz, 2H, R C H 2 C H 2 C O 2 C H 3 ) ,
3.33, 3.40, 3.52, 3.54, 3.62 (5s, 15H, CH3-2-12 and -18 and methyl ester-132 and
-172), 3.83-3.89 (m, 2H, RCH2N(CH3)2), 3.91 (s, 3H, methyl ester-72), 4.07 and 4.24
(2br.s, 4H, 2 X R C H 2 C H 2 C O 2 C H 3 ) , 5.16 (s, 1H, H-71), 6.88 (dt, J=16.1, 6.7 Hz, 1H,
vinyl H-32), 7.78 and 7.80 (ABq, 2H, H-73 and -74), 8.22 (d, J=16.1 Hz, 1H, vinyl H-
31), 9.31, 9.59, 9.74, 9.83 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for C 4 5 H 5 1 N 5 O 8 :
789.3740, found 789.3740; 789 (M+), 746 (M+-CH3N=CH2).
134
BPD dimorpholine amide (19). A 25mL round bottom flask was fitted with a
condenser, drying tube and stirbar. Under a nitrogen atmosphere, the flask was flame-
dried three times and after cooling crude BPD diacid (45.0mg; 6.4xl0'5mol) was added.
Dry dichloromethane (12mL) was added and the mixture was brought to reflux for 15
minutes. After cooling, oxalyl chloride (0.50mL; 5.73X10_3mol) were added by syringe
and the reaction mixture was brought to reflux for 30 minutes. After cooling to room
temperature, the condenser was replaced with a septum and the solvent was evaporated
under a strong nitrogen flow. When dry, the residue was taken up in dry dichloromethane
(15mL) and blown dry again. After drying, the crude diacid chloride was redissolved in
dichloromethane (lOmL). With a light nitrogen flow maintained, freshly distilled
morpholine (0.15mL; 1.72X10_3mol) was added dropwise via the septum. Hydrochloric
acid gas was evolved immediately upon addition of the amine. The mixture was stirred at
room temperature for 30 minutes after which time the solvent was evaporated in vacuo.
The crude compound was dissolved in 5% methanol/dichloromethane (50mL), washed
with 2% aqueous hydrochloric acid (2x50mL) followed by water (2x50mL) washes. The
organic phase was dried over sodium sulphate and the filtered solution was evaporated and
chromatographed on silica gel (silica gel 60, 70-230 mesh, 5%MeOH/CH2Cl2 eluent).
The desired fractions were evaporated and crystallized from dichoromethane/hexanes (1:1)
135
to provide 42.4mg (79% from the dimethyl ester) of the desired diamide as dark green
crystals.: *H NMR (400 MHz, CDCI3) 8 -2.28 (br. s, 2H, 2xpyrrolicNH), 1.79 (s, 3H,
CH3-7), 2.72-2.84 (m, 2H, 2H morpholine), 2.98-3.04 (m, 5H, methyl ester-71 and 2H
morpholine), 3.13-3.27 (m, 12H, 2XRCH2CH2COR' and 8H morpholine), 3.43 (s, 3H,
CH3-2), 3.45-3.54 (m, 5H, CH3-I2 and 2H morpholine), 3.59-3.70 (m, 5H, CH3-I8
and 2H morpholine), 4.00 (s, 3H, methyl ester-72), 4.22 (t, J=8 Hz, 2H,
RCH2CH2COR') , 4.34 (t, J=8 Hz, 2H, RCH2CH2COR'), 5.07 (s, 1H, H-71), 6.18 (d,
J=12 Hz, 1H, vinyl H-32), 6.36 (d, J=18 Hz, 1H, vinyl H-32), 7.46 (d, J=8 Hz, 1H, H-
l\ 7.83 (d, J=8 Hz, 1H, H-74), 8.11 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.15, 9.35,
9.68, 9.74 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for C48H54N4O8: 842.4006, found
842.3997, mle calc'd for C43H46N3O6: 728.3451, found 728.3441.842; 842 (M+), 728
(M+-CON(CH2CH2)20);
BPD di(piperidine) amide (20). A 50mL three neck round bottom flask was
fitted with the appropriate septa and flame-dried under nitrogen three times. Upon cooling,
crude BPD diacid (40.8mg; 5.79X10_5mol) was quickly added to the flask. The
compound was dissolved in dry dichloromethane (12mL) and the solution was stirred
under a stream of nitrogen. After a 5 minutes, 0.35mL oxalyl chloride (0.35mL; 4.01X10-
3mol) was added by syringe dropwise into the mixture. The middle septum was quickly
replaced with a condenser and the bright green mixture was brought to reflux for 30
minutes using a warm water bath. After cooling to room temperature, the condenser was
removed and replaced with a septum and the nitrogen flow was increased to blow off the
excess oxalyl chloride and the solvent. Dichloromethane (15mL) was added by syringe
and again the solvent was blown off. After this procedure was repeated a third time, the
acid chloride mixture was taken up in dry dichloromethane (15mL) and stirred at room
temperature. To the reaction vessel was added dry piperidine (0.20mL; 2.02xl0"3mol) and
the bright green solution turned a dull green denoting deprotonation of the pyrrolic
nitrogens. After stirring the flask for 30 minutes, the contents were diluted to 50mL with
136
dichloromethane in a separatory funnel and washed with dilute aqueous hydrochloric acid
(3x50mL) and by water (5x50mL). The organic layer was evaporated in vacuo and the
crude compound was chromatographed on silica gel (silica gel 60, 70-230 mesh, 2-
5%MeOH/CH2Cl2 gradient eluent). The appropriate fractions were pooled and evaporated
to afford 34mg (70% from BPD dimethyl ester) BPD di(piperidine) amide as a dark green
solid.: *H NMR (300 MHz, C D C I 3 ) 8 -2.30 (br. s, 2H, 2xpyrrolicNH), 1.09 (m, 2H,
RCON(CH2CH2)2CH2), 1.22 (m, 2H, RCON(CH2CH2)2CH_2), 1.28 (m, 8H,
2XRCON(CH2CH2)2CH2), 1.77 (s, 3H, CH>7), 2.97 (s, 3H, methyl ester-71), 3.04-
3.27 (m, 8H, 2 X R C H 2 C H 2 C O R ' and 4H piperidine), 3.40, 3.47 (2s, 6H, C H 3 - 2 and-
12), 3.49-3.61 (m, 4H, 4H piperidine), 3.63 (s, 3H, C H 3 - I 8 ) , 3.97 (s, 3H, methyl ester-
72), 4.17 (t, J=8.0 Hz, 2H, R C H 2 C H 2 C O R ' ) , 4.32 (t, J=8.0 Hz, 2H, R C H 2 C H 2 C O R ' ) ,
5.05 (s, 1H, H-71), 6.15 (d, J=12 Hz, 1H, vinyl H-32), 6.35 (d, J=18, 2 Hz, 1H, vinyl
H-32), 7.43 (d, J=7 Hz, 1H, H-73), 7.81 (d, J=7 Hz, 1H, H-74), 8.11 (dd, J=12, 18 Hz,
1H, vinyl H-31), 9.13, 9.35, 9.65, 9.74 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C 5 0 H 5 8 N 6 O 6 : 838.4421, found 838.4412, mle calc'd for C 5 1 H 6 2 N 6 O 7 : 870.4684,
found 870.4675; 870 (M+ + C H 3 O H ) , 838 (M+); Analysis calc'd for
C50H58N6O6-0.5MeOH: C, 70.94; H, 7.07; N, 9.82; found: C, 70.91; H, 7.00; N,
9.72;
137
Zn BPD dimethyl ester (21). A 500mL round bottom flask was fitted with a
stirbar, BPD 1,3-diene dimethyl ester (1.0152g; 1.4xl0-3mol) and dichloromethane
(80mL) and the mixture was stirred at room temperature. Anhydrous zinc(II) acetate
the stirring mixture. The flask was sealed and left for 16 hrs. at which time tic and uv-vis
spectroscopy showed full conversion to the desired compound. The solvent was removed
by evaporation in vacuo and the product was redissolved in dichloromethane, filtered
through a cotton plug, washed with H 2 O (3x) and evaporated to dryness. The compound
was dissolved in acetonitrile and evaporated to yield l.lOOOg (99%) Zn BPD 1,3-diene
dimethyl ester.: lH NMR (400 MHz, C D C I 3 ) 5 1.76 (s, 3H, C H 3 - 7 ) , 2.92-3.12 (m, 7H,
2 X R C H 2 C H 2 C O 2 C H 3 and methyl ester-71), 3.23, 3.25, 3.48, 3.55, 3.60 (5s, 15H,
C H 3 - 2 -12 -18 and methyl esters-13 and -17), 3.86-4.10 (m, 7H, 2 X R C H 2 C H 2 C O 2 C H 3
and methyl ester-72), 4.97 (s, 1H, H-71), 6.03 (d, J=12 Hz, 1H, vinyl H-3), 6.17 (d,
J=18 Hz, 1H, vinyl H-3), 7.37 (br. s, 1H, H-73), 7.76 (br. s, 1H, H-74), 8.03 (dd,
J=12, 18 Hz, 1H, vinyl H-3), 8.87, 9.10, 9.26, 9.46 (4s, 4H, 4xmeso H); MS (EI) mle
calc'd for C42H42N40gZn: 794.2296, found 794.2286; 794 (M+), 735 (M +-C02Me),
720 (M+-C02Me-CH3), 661 (M+-2xC02Me-CH3); Analysis calc'd for C42H42N40sZn:
C, 63.36; H, 5.32; N, 7.04; found: C, 63.71; H, 5.49; N, 6.92.; UV-Vis ( C H 2 C I 2 )
W (peak ratio) 358 (1.20), 442 (1.88), 620 (0.50), 672 (1.00).
(1.1363g; 6.2xl0"3mol) was dissolved in methanol (15mL) and this solution was added to
M e 0 2 C P 0 2 M e
(22)
H O C H 2 C H 2 O H
138
Zn BPD diol (22). A 50mL three neck round bottom flask was fitted with the
appropriate septa and a stirbar. The flask was flame-dried three times under nitrogen and
upon cooling, the top septum was removed and Zn BPD dimethyl ester (99.3mg; 1.3xl0-
4mol) was quickly added and the septum was replaced. Using a large syringe, 21mL
freshly distilled THF were added with a slow stream of nitrogen maintained. The reaction
vessel was immersed in an ice bath and stirring was continued for 15 minutes. An oven
dried lcc syringe was used to add 0.1M diisobutyl aluminum hydride (DIBAL-H) in
hexanes (0.50mL; 5.0xl0_4mol) dropwise to the cooled solution. The reaction progress
was monitored by tic and in 60 minutes the reaction was deemed complete. The top septum
was removed and lOmL saturated ammonium chloride was poured in to quench any
unreacted hydride. The vessel was removed from the ice bath and raised to room
temperature at which time the contents were transferred to a separatory funnel. 5%
methanol/dichloromethane (50mL) was added and the funnel was shaken to extract the
chlorin into the organic phase. The organic phase was washed with water (3xl00mL) and
dried over sodium sulfate. Upon filtration and evaporation in vacuo, the residue was
chromatographed on silica gel (silica gel 60, 70-230 mesh, 4%MeOH/CH2Cl2 eluent).
The appropriate fractions were pooled and evaporated to yield 47.4mg (52%) of the desired
Zn BPD diol.: 1H NMR (400 MHz, DMSO-d6) 6 1.71 (s, 3H, CH3-7), 2.20-2.31 (m,
4H, 2XRCH2CH2CH2OH), 2.92 (s, 3H, methyl ester-71), 3.32, 3.37, 3.52 (3s, 9H,
CH3-2 -12 and -18), 3.68-3.76 (m, 4H, 2XRCH2CH2CH2OH), 3.82-3.88 (m, 4H,
2XRCH2CH2CH2OH), 3.89 (s, 3H, methyl ester-72), 4.68 and 4.69 (overlapping t, 5.0
Hz, 2H, 2xR(CH2)30H), 5.06 (s, 1H, H-71), 5.99 (dd, J=1.4, 11.2 Hz, 1H, vinyl H-
32), 6.24 (dd, J=1.4, 18.0 Hz, 1H, vinyl H-32), 7.62 and 7.75 (2d, J=6.2 Hz, 2H, H-7 3
and -74), 8.17 (dd, J=11.7, 18.0 Hz, 1H, vinyl H-31), 8.96, 9.24, 9.58, 9.62 (4s, 4H,
4xmeso H); MS (EI) mle calc'd for C4QH42N406Zn: 738.2398, found 738.2402; 738
139
(M+), 679 (M+-C02CH3), 664 ( M + - C O 2 C H 3 - C H 3 ) ; Analysis calc'd for
C40H42N4O6Zn-1.5H"2O: C, 62.62; H, 5.91; N, 7.30; found: C, 62.60; H, 5.62; N,
7.14.
BPD diol (23). Zn BPD diol (57.5mg; 7.77xl0-5mol) was dissolved in 14%
methanol/dichloromethane (35mL), transferred to a separatory funnel and trifluoroacetic
acid (lmL) was added by pipet. After shaking, the organic layer was neutralized with 5%
aqueous potassium bicarbonate (50mL) followed by washing with distilled water
(3x50mL). The uv-vis spectrum showed only the free-base chlorin. The solvent was
evaporated in vacuo to yield 52.0mg (99%) of the non-metallated BPD diol.: NMR
(300 MHz, C D C I 3 ) 5 -2.37 (br. s, 2H, 2xpyrrolicNH), 1.80 (s, 3H, CH3-7), 2.33 (m,
4H, 2 X R C H 2 C H 2 C H 2 O H ) , 2.91 (s, 3H, methyl ester-71), 3.38, 3.42, 3.61 (3s, 9H,
C H 3 - 2 , -12 and -18), 3.78-4.15 (m, 11H, 2xRCH2CH2CH_2OH and
2xRCH2CH2CH20H and methyl ester-72), 5.06 (s, 1H, H-71), 6.16 (d, J=12 Hz, 1H,
vinyl H-32), 6.32 (d, J=18 Hz, 1H, vinyl H-32), 7.45 (d, J=7 Hz, 1H, H-73), 7.82 (d,
J=7 Hz, 1H, H-74), 8.11 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.15, 9.36, 9.76, 9.89 (4s,
4H, 4xmeso H), Hydroxy protons obscured; MS (EI) mle calc'd for C 4 0 H 4 4 N 4 O 8 :
140
676.3263, found 676.3253; 676 (M+), 658 (M+-H20), 602 (M+-C02CH3-CH3),
Positive test for singlet-oxygen production.
BPD ditosylate (24). BPD diol (5.6mg; 8.2xl0-°mol) was placed in a 5mL
round bottom flask containing a stir bar and the solid was dissolved in dry pyridine (lmL).
The mixture was stirred for a few minutes and p-toluene sulphonyl chloride (17mg;
8.92X10-5mol) was added. The mixture was immersed into a large insulated ice water
bath and left overnight. The next morning an aliquot was worked-up and tic showed
almost full conversion to the ditosylate with overlapping slower moving spots denoting the
two possible monotosylate/monoalcohol products. The mixture was transferred into a
separatory funnel containing dichloromethane (15mL) and washed with dilute aqueous
hydrochloric acid (3x25mL) followed by water (5x25mL). Column chromatography was
attempted but the residue was unstable and led to decomposition of the compound. The
crude residue was carried over to the iodination step.: MS (EI) mle 985 (M+), 849, 831.
BPD diiodide (25). Crude BPD ditosylate (6.5mg; 6.6xl0-°mol), sodium
iodide (13mg; 8.67xl0_5mol) and a stirbar were placed in a lOmL round bottom flask.
Acetonitrile (2mL) was added and the stirring mixture was slowly heated to a gende reflux.
After 10 minutes the oil bath was removed and a tic cospot (0.5%MeOH/CH2Cl2 eluent) of
the cooled mixture with the ditosylate showed a strong spot moving slightly faster than the
starting compound . This product, although slightly more stable than the ditosylate, was
difficult to purify but careful chromatography in the dark provided enough pure compound
for characterization.: lU NMR (300 MHz, C D C I 3 ) 8 -2.32 (br. s, 2H, 2xpyrrolicNH),
1.77 (s, 3H, CH3-7), 2.56-2.78 (m, 4H, 2 X R C H 2 C H 2 C H 2 I ) , 2.93 (s, 3H, methyl ester-
71), 3.34-3.81 (m, 13H, 2xRCH2CH2CH2l and C H 3 - 2 , -12 and -18), 3.86-4.24 (m,
7H, 2xRCH2CH2CH2l and methyl ester-72), 5.06 (s, 1H, H-71), 6.16 (d, J=12 Hz, 1H,
vinyl H-32), 6.37 (d, J=18 Hz, 1H, vinyl H-32), 7.45 (d, J=6 Hz, 1H, H-73), 7.82 (d,
J=6 Hz, 1H, H-74), 8.11 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.13, 9.36 (2s, 2H,
141
2xmesoH), 9.75 (s, 2H, 2xmesoH); MS (EI) mle 896 (M+), 837 (M+-CC-2CH.3), 822
(M+-C02CH3 - C H 3 ) , 768 (M+-HI), 640 (M+-2xHI).
BPD di(morpholine) amine (26). To a lOmL round bottom flask containing
BPD diiodide (29.8mg; 3.32xl0~5moi) dissolved in acetonitrile (3.5mL) was added dry
morpholine (3mL; 3.43xl0~2mol). The mixture was stirred for 24 hours after which time
tic showed full consumption of the diiodide. The mixture was diluted with
dichloromethane (50mL) and washed with 0.1N aqueous hydrochloric acid (3x50mL), 5%
aqueous potassium bicarbonate (lx50mL) and water (3x50mL). The organic solvent was
evaporated in vacuo and the crude compound was chromatographed on silica gel (silica gel
60, 70-230 mesh, 7%MeOH/CH2Cl2 eluent). The appropriate fractions were pooled and
evaporated in vacuo. The yield of BPD di(morpholine) amine was 17.4mg (64% from the
Zn BPD diol).: A H NMR (300 MHz, C D C I 3 ) 5 -2.32 (s, 2H, 2xpyrrolicNH), 1.77 (s,
3H, C H 3 - 7 ) , 2.26-2.71 (m, 16H, 2xRCH2CH2CH2NR'2 2xRCH2CH2CH2NR'2 and
2xRN(CH2CH2)20), 2.94 (s, 3H, methyl ester-71), 3.40, 3.47, 3.63 (3s, 9H, C H 3 - 2
-12 and -18), 3.69-3.80 (m, 8H, 2xRN(CH2CH2)20), 3.82-3.93 (m, 2H,
RCH2CH2CH2NR'2), 3.94-4.08 (m, 5H, RCH2CH2CH2NR'2 and methyl ester-72),
5.05 (s, 1H, H-71), 6.14 (d, J=12 Hz, 1H, vinyl H-32), 6.36 (d, J=18 Hz, 1H, vinyl H-
32), 7.44 (d, J=7 Hz, 1H, H-73), 7.82 (d, J=7 Hz, 1H, H-74), 8.12 (dd, J=12, 18 Hz,
1H, vinyl H-31), 9.13, 9.35, 9.63, 9.75 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C48H58N6O6: 814.4421, found 814.4412; 814 (M +), 714 (M+-C5HioNO).
BPD di(piperidine) amine (27). Into a 50mL round bottom flask containing
BPD diiodide (42.9mg; 4.79X10-^mol) dissolved in acetonitrile (25mL), dry piperidine
(5mL; 5.06X10-2mol) was added. The mixture was stirred at room temperature for 36
hours and monitored by tic. When the diiodide was fully consumed, the reaction mixture
was poured into dichloromethane (25mL) and washed with water (3x50mL), dilute
aqueous hydrochloric acid (3x50mL) and water (3x50mL). The organic layer was
evaporated and the crude residue was chromatographed on silica gel (silica gel 60, 70-230
142
mesh, 8%MeOH/CH2Cl2 eluent). The appropriate fractions were collected and evaporated
to yield 20.0mg (51% from the BPD diol) BPD di(piperidine)amine.: A H NMR (200
MHz, DMSO-d6) 5 -2.41 (s, 2H, 2xpyrrolicNH), 1.29-1.46 (m, 4H,
2xRN(CH2CH2)2CH2), 1.46-1.64 (m, 8H, 2xRN(CH2CH2)2CH2), 1.75 (s, 3H, C H 3 -
7), 2.18-2.71 (m, 19H, 2xRN(CH2CH2)2CH2, 2xRCH2CH2CH2NR'2 and methyl
ester-71), 3.38, 3.50, 3.65 (3s, 9H, C H 3 - 2 -12 and -18), 3.75-4.05 (m, 7H,
2xRCH2CH2CH2NR'2 and methyl ester-72), 5.25 (s, 1H, H-71), 6.19 (d, J=12 Hz, 1H,
vinyl H-32), 6.45 (d, J=18 Hz, 1H, vinyl H-32), 7.80 (s, 2H, H-7 3 and H-74), 8.40 (dd,
J=12, 18 Hz, 1H, vinyl H-31), 9.38, 9.60, 9.69, 9.87 (4s, 4H, 4xmesoH); MS (EI) mle
calc'd for C 5 0 H 6 2 N 6 O 4 : 810.4837, found 810.4823; 810 (M +), 712 (M+-C6H12N).
Zn BPD triol (28). A three neck round bottom flask was fitted with a stir bar
and the appropriate septa and flame-dried three times under nitrogen. The middle septum
was removed and Zn BPD dimethyl ester (180.0mg; 2.26X10~4mol) were quickly added
and the septum was replaced. The compound was dissolved in freshly distilled
tetrahydrofuran (25mL) added via the septum and the reaction flask was immersed in an ice
bath. After 15 minutes stirring under nitrogen, 1.0M DIBAL-H in hexanes (1.40mL;
1.40X10'3 mol) was added slowly by syringe. With nitrogen maintained, the reaction was
left to stir for 45 minutes. The reaction was quenched with saturated aqueous ammonium
CH2OH
H2OH
(28)
HOCH2
143
chloride (25mL) and extracted into 25% methanol/dichloromethane (50mL). After washing
with water (3x50mL), the organic layer was dried over sodium sulfate. Filtration, removal
of solvent and a final crystallization from dichloromethane/methanol/hexanes yielded
130.4mg of the desired Zn BPD triol (81%) as a blue-purple crystals.: A H NMR (400
MHz, DMSO-d6) 5 1.66 (s, 3H, CH3-7), 2.18-2.30 (m, 4H, 2 X R C H 2 C H 2 C H 2 O H ) ,
2.96 (s, 3H, methyl ester-71), 3.28, 3.30, 3.34, 3.47 (4s, 12H, CH3-2 -12 -18 and
methyl ester-72), 3.71 and 3.73 (2t overlap, J=6.0, 6.0 Hz, 4H, 2XRCH2CH2CH2OH),
3.78-3.86 (m, 4H, 2XRCH2CH2CH2OH), 4.32 (dAB quartet, 2H, RCH2OH), 4.50 (s,
1H, H-71), 4.68 and 4.69 (2t overlap, J=5.2, 5.3 Hz, 2H, 2xR(CH2)30H), 5.29 (t,
J=5.6 Hz, 1H, RCH2OH), 5.92 (d, J=11.5 Hz, 1H, H-32), 6.15 (d, J=17.9 Hz, 1H, H-
32), 6.58 and 7.36 (2d, J=5.1, 5.3 Hz, 2H, H-7 3 and -74), 8.06 (dd, J=17.8, 11.6 Hz,
1H, H-31), 8.64, 9.03, 9.47, 9.53 (4s, 4H, 4xmesoH); MS (EI) mle calc'd for
C39H42N405Zn: 710.2449, found 710.2437; 710 (M+); Analysis calc'd for
C39H42N405Zn-H20: C, 64.15; H, 6.07; N, 7.67; found: C, 64.53; H, 5.96; N,
7.76.; UV-Vis (MeOH) ^max (peak ratio) 436 (2.42), 538 (0.40), 582 (0.51), 604 (0.47),
658 (1.00).
H O C H 2 C H 2 O H
BPD triol (29). Zn BPD triol (17.0mg; 2.39xl0"5mol) was dissolved in 25%
methanol/dichloromethane (50mL) and transferred to a separatory funnel. Trifluoroacetic
acid (lmL) was then added dropwise and the solution was vigorously shaken. The organic
144
layer was neutralized with 5% aqueous potassium bicarbonate (3x50mL) and went from
bright green to brick red in colour. After water washings (3x50mL), the organic layer was
dried over sodium sulfate. Filtration and removal of solvent gave 14.3mg of the desired
BPD triol (92%) as a red-brown powder.: lH NMR (400 MHz, DMSO-d6) 5 -2.11 (br. s,
2H, 2xpyrrolicNH), 1.70 (s, 3H, CH3-7), 2.24-2.30 (m, 4H, 2 X R C H 2 C H 2 C H 2 O H ) ,
2.95 (s, 3H, methyl ester-71), 3.35, 3.45, 3.60 (3s, 9H, CH3-2 -12 and -18), 3.66-3.77
(m, 4H, 2xR(CH2)2CH20H), 3.82 (t, J=8 Hz, 2H, RCH2(CH2)20H), 4.02 (t, J=8 Hz,
2H, RCH2(CH2)20H), 4.30-4.46 (m, 1H, RCH2OH), 4.63 (s, 1H, H-71), 4.71 (t, J=6
Hz, 1H, R(CH2)30H), 4.76 (t, J=6 Hz, 1H, R(CH2)30H), 5.35 (t, J=6 Hz, 1H,
RCH2OH), 6.11 (d, J=12 Hz, 1H, vinyl H-32), 6.36 (d, J=18 Hz, 1H, vinyl H-32), 6.65
(d, J=7 Hz, 1H, H-73), 7.59 (d, J=7 Hz, 1H, H-74), 8.24 (dd, J=12, 18 Hz, 1H, vinyl
H - 3 1 ) , 9.04, 9.44, 9.72, 9.79 (4s, 4H, 4xmesoH); MS (LSIM) mle calc'd for
C39H44N4O5: 649.33925, found 649.33865; 649 (M++1); UV-Vis (MeOH) Xmax (e)
422 (71,200), 512 (8,300), 554 (12,700), 628 (6,400), 686 (21,600).
M e 0 2 C C 0 2 M e
C H 3 - \ \
(30)
CH3-<\ J
HO(CH2) 2 0 2 C C O ^ C H ^ O H
BPD di(ethyleneglycol) ester (30). BPD 1,3-diene dimethyl ester (125.0mg;
1.71X10-4mol) were placed in a lOmL round bottom flask equipped with a stir bar. Dry
145
ethylene glycol (6.9mL; 1.24X10-1mol) was then added and the heterogeneous mixture
was stirred at room temperature for 1.5hrs. The flask was cooled in an ice water bath and
4 drops concentrated sulfuric acid were added. The mixture was raised to room
temperature and stirred for 48hrs after which time a cospotted dc showed full consumption
of the starting dimethyl ester and a much slower moving product. The reaction solution
was dumped into 5% aqueous sodium acetate (30mL) and the bright green mixture turned a
dull green colour. The desired compound was extracted into chloroform (50mL) after
transferring to a separatory funnel. The organic layer was washed with water (3x50mL)
and evaporated in vacuo after addition of acetonitrile (30rnL) to remove any water. The
crude compound was chromatographed on silica gel (silica gel 60, 70-230 mesh, 1-
2%MeOH/CH2Cl2 gradient eluent). The appropriate fractions were pooled and dried to
afford 128.0mg (94%) BPD di(ethylene glycol) ester.: A H NMR (300 MHz, CDCI3) 5
-2.35 (br. s, 2H, 2xpyrrolicNH), 1.80 (s, 3H, CH3-7), 2.08 (br. s, 1H, RCH2OH), 2.38
(br. s, 1H, R C H 2 O H J , 2.94 (s, 3H, methyl ester-71), 3.22 (t, J=8 Hz, 2 H ,
R C H 2 C H 2 C O 2 R ) , 3.26 (t, J=8 Hz, 2H, RCH2CH2CO2R'), 3.42 (s, 3H, CH3-2), 3.45
(m, 5 H , CH3-12 and RCO2CH2CH_2OH) , 3.62 (m, 5 H , C H 3 - I 8 and
R C O 2 C H 2 C H 2 O H ) , 3.98 (s, 3H, methyl ester-72), 4.07-4.18 (m, 4 H ,
R C O 2 C H 2 C H 2 O H ) , 4.19 (t, J=8 Hz, 2H, R C H 2 C H 2 C 0 2 R ' ) , 4.32 (t, J=8 Hz, 2H,
R C H 2 C H 2 C O 2 R ' ) , 5.05 (s, 1H, H-71), 6.16 (dd, J=l, 12 Hz, 1H, vinyl H-32), 6.35
(dd, J=l, 18 Hz, 1H, vinyl H-32), 7.45 (d, J=6 Hz, 1H, H-73), 7.82 (d, J=6 Hz, 1H, H-
74), 8.08 (dd, J=12, 18 Hz, 1H, vinyl H-31), 9.15, 9.38, 9.75, 9.78 (4s, 4H, 4xmesoH);
MS (EI) mle calc'd for C44H48N4O10: 792.3372, found 792.3361; 792 (M+), 730 (M+-
2 x C H 2 0 H ) , 671 ( M + - 2 x C H 2 O H - C O 2 C H 3); Analysis calc'd for
C44H48N4O10O.5H2O: C, 65.91; H, 6.16; N, 6.99; found: C, 65.97; H, 6.19; N,
6.89.
146
BPD dimethyl ester vinyl hydrate (31). BPD dimethyl ester (175mg;
2.39xl0"4mol) was hydrobrominated in the same way described for BPD vinyl amine
(3). The crude hydrobromide was treated with water after being taken up in dry
tetrahydrofuran (20mL). The mixture was stirred at 50°C overnight. The crude product
was extracted into dichloromethane (50mL) and washed with water (3x50mL). After
removal of solvent, the solid was crystallized from dichloromethane/hexanes to afford the
BPD vinyl hydrate as blue-purple crystals (characterized as the diastereomeric mixture).:
lH NMR (400 MHz, cdci3) 5 -2.41 (br.s, 4H, 4xpyrrolicNH), 1.82 and 1.83 (2s, 6H,
ch3-7), 2.17 and 2.24 (2d, 5=6.1, 6.7 Hz, 6H, ch3-31), 2.89 and 2.90 (2s, 6H, methyl
ester-71), 3.09-3.21 (m, 8H, 4xrch2ch2co2ch3), 3.35, 3.36, 3.42, 3.43, 3.56, 3.60
(6s, 18H, 2xCH3-2 -12 and -18), 3.63 and 3.65 (2s, 6H, 2xmethyl ester-132 and 172),
3.98 (s, 6H, 2xmethyl ester-72), 4.06-4.14 (m, 4H, 2xrch2ch2co2ch3), 4.20-4.28
(m, 4H, 2xrch2ch2co2ch3) , 5.08 and 5.09 (2s, 2H, H-7*), 6.46 and 6.55 (2q,
J=6.7, 6.7 Hz, 2H, H-31), 7.45 and 7.83 (2d, J=5.5, 5.5 Hz, 2H, H-7 3 and -74), 9.27,
9.33, 9.34, 9.45, 9.60, 9.62, 9.66, 9.68 (8s, 8H, 8xmesoH); MS (EI) mle calc'd for
c42h46n4o9: 750.3265, found 750.3258; 750 (M+), 732 (M+-H20).
HO(CH2) 2NHCO CONH(CH2) 2OH
BPD diethanol amide (32).BPD 1,3-diene dimethyl ester (46.8mg; 6.39x10"5
mol) was added to a solution of freshly distilled tetrahydrofuran (25mL) and dry
ethanolamine (lmL; 1.66X10-2mol) in a 50mL round bottom flask. A stirbar was added to
the flask which was fitted with a condenser and a drying tube. The mixture was stirred at
56°C for two days at which time tic showed no remaining starting material, a slower
moving spot and baseline material. The reaction was stopped and the solvent was removed
under vacuum. The residue was then taken up in dichloromethane (50mL), transferred to a
separatory funnel, washed with distilled water (3x50mL) and the organic layer was dried
over sodium sulfate. The solution was filtered and tic of the compound showed the slower
moving compound with no baseline material. The product was indeed the desired diamide
and weighed 37.9mg (75%).: lH NMR (400 MHz, DMSO-d6) 5 -2.38 (s, 2H,
2xpyrrolicNH), 1.74 (s, 3H, CH3-7), 2.90 (s, 3H, methyl ester-71), 2.93 (t, J=7 Hz, 2H,
RCH2CH2CONHR'), 2.99 (t, J=7 Hz, 2H, RCH2CH2CONHR'), 3.06 (dt, J=6, 5 Hz,
2H, RNHCH2CH2OH), 3.11 (dt, J=6, 5 Hz, 2H, RNHCH2CH2OH), 3.20-3.35 (m,
4H, 2XRCH2CH2OH), 3.36, 3.49, 3.64 (3s, 9H, CH3-2 -12 and -18), 4.05 (t, J=7 Hz,
2H, RCH2CH2COR'), 4.23 (t, J=7 Hz, 2H, RCH2CH2COR'), 4.48-4.57 (m, 2H,
2xR(CH2)20H), 5.22 (s, 1H, H-71), 6.18 (d, J=12 Hz, 1H, vinyl H-3), 6.43 (d, J=18
Hz, 1H, vinyl H-3), 7.78 (br. s, 2H, H-73 and H-74), 7.97 (t, J=5 Hz, 1H, RCONHR'),
7.99 (t, J=5 Hz, 1H, RCONHR'), 9.37, 9.59 (2s, 2H, 2xmesoH), 9.85 (br. s, 2H,
148
2xmesoH); MS (El)m/e calc'd for C 4 4 H 5 0 N 6 O 8 : 790.3693, found 790.3694; 790 (M+),
772 (M+-H20), 7 5 4 (M+-2H20).
BPD diphosphonate ester (33). Crude BPD diiodide (59.6mg; 6.65x10'5
mol) dissolved in acetonitrile (7mL) was placed in a 25mL round bottom flask containing a
stirbar. Stirring was commenced in the dark at room temperature and diethyl phosphite
(4mL; 2.33xl0-2mol) was added. A condenser was fitted and the flask was brought to
reflux. Reflux was maintained for 24 hours at which time tic showed conversion to a much
slower moving compound than the diiodide. The acetonitrile was removed and the flask
was placed on high vacuum to remove the excess triethyl phosphite. After 2 days, on the
vacuum line, no trace of triethyl phosphite was noticed. The crude compound was
chromatographed on silica gel (silica gel 60,70-230 mesh, 4%-5%MeOH/CFf2Cl2 gradient
eluent). The appropriate fractions were pooled and evaporated to yield 41.1mg (67% from
the starting BPD diol) of the BPD diphosphonate ester.: lH NMR (400 MHz, DMSO-d6)
5 -2.39 (br. s, 2H, 2xpyrrolicNH), 1.16 and 1.18 (2t overlapping, J=6.7 Hz, 12H,
4 x R P O ( C H 2 C H _ 3 ) 2 ) , 1.76 (s, 3H, C H 3 - 7 ) , 2.04-2.17 (m, 4H,
2xRCH2CH2CH2PO(OEt)2), 2.22-2.40 (m, 4H, 2xRCH2CH2CH2PO(OEt)2), 2.89 (s,
3H, methyl ester-71), 3.39, 3.50, 3.66 (3s, 9H, C H 3 - 2 -12 and -18), 3.91 (s, 3H, methyl
ester-72), 3.93-4.06 (m, 10H, RCH2CH2CH2PO(OEt)2, 2xRPO(OCH2CH3)2), 4.14 (t,
149
J=7 Hz, 2H, RCH2CH2CH2PO(OEt)2), 5.22 (s, 1H, H-7 1), 6.18 (d, J=11.7 Hz, 1H,
vinyl H-32), 6.44 (d, J=18.2 Hz, 1H, vinyl H-32), 7.78 (s, 2H, H-7 3 and -74), 8.36 (dd,
J=11.7, 18.2 Hz, 1H, vinyl H-31), 9.38, 9.61, 9.84, 9.88 (4s, 4H, 4xmesoH); 3 1 P
NMR (CDCI3) 5 32.0, 32.5; MS (EI) mle calc'd for C48H62N4O10P2: 916.3945, found
916.3923; 916 (M+).
150
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