Gold(III) macrocycles: Nucleotide-specific unconventional catalytic inhibitors of human topoisomerase i

Article abstract of DOI:10.1021/ja412350f

Topoisomerase IB (Top1) is a key eukaryotic nuclear enzyme that regulates the topology of DNA during replication and gene transcription. Anticancer drugs that block Top1 are either well-characterized interfacial poisons or lesser-known catalytic inhibitor compounds. Here we describe a new class of cytotoxic redox-stable cationic Au³⁺ macrocycles which, through hierarchical cluster analysis of cytotoxicity data for the lead compound, 3, were identified as either poisons or inhibitors of Top1. Two pivotal enzyme inhibition assays prove that the compounds are true catalytic inhibitors of Top1. Inhibition of human topoisomerase IIα (Top2α) by 3 was 2 orders of magnitude weaker than its inhibition of Top1, confirming that 3 is a type I-specific catalytic inhibitor. Importantly, Au³⁺ is essential for both DNA intercalation and enzyme inhibition. Macromolecular simulations show that 3 intercalates directly at the 5′-TA-3′ dinucleotide sequence targeted by Top1 via crucial electrostatic interactions, which include π-π stacking and an Au?O contact involving a thymine carbonyl group, resolving the ambiguity of conventional (drug binds protein) vs unconventional (drug binds substrate) catalytic inhibition of the enzyme. Surface plasmon resonance studies confirm the molecular mechanism of action elucidated by the simulations.

Full text of DOI:10.1021/ja412350f

Article
pubs.acs.org/JACS
Open Access on 04/02/2015
Gold(III) Macrocycles: Nucleotide-Specific Unconventional Catalytic
Inhibitors of Human Topoisomerase I
Kate J. Akerman,† Alexander M. Fagenson,‡ Vidusha Cyril,‡ Michael Taylor,‡ Mark T. Muller,‡
Matthew P. Akerman,† and Orde Q. Munro*,†,§
†School of Chemistry and Physics, University of KwaZulu-Natal, Private Bag X01, Scottsville, Pietermaritzburg, 3209, South Africa
‡College of Medicine, Biomedical Research, University of Central Florida, 6900 Lake Nona Boulevard, Orlando, Florida 32827,
United States
S
* Supporting Information
ABSTRACT: Topoisomerase IB (Top1) is a key eukaryotic
nuclear enzyme that regulates the topology of DNA during
replication and gene transcription. Anticancer drugs that block
Top1 are either well-characterized interfacial poisons or lesser-
known catalytic inhibitor compounds. Here we describe a new
class of cytotoxic redox-stable cationic Au3+ macrocycles
which, through hierarchical cluster analysis of cytotoxicity
data for the lead compound, 3, were identified as either
poisons or inhibitors of Top1. Two pivotal enzyme inhibition
assays prove that the compounds are true catalytic inhibitors of
Top1. Inhibition of human topoisomerase IIα (Top2α) by 3 was 2 orders of magnitude weaker than its inhibition of Top1,
confirming that 3 is a type I-specific catalytic inhibitor. Importantly, Au3+ is essential for both DNA intercalation and enzyme
inhibition. Macromolecular simulations show that 3 intercalates directly at the 5′-TA-3′ dinucleotide sequence targeted by Top1
via crucial electrostatic interactions, which include π−π stacking and an Au···O contact involving a thymine carbonyl group,
resolving the ambiguity of conventional (drug binds protein) vs unconventional (drug binds substrate) catalytic inhibition of the
enzyme. Surface plasmon resonance studies confirm the molecular mechanism of action elucidated by the simulations.
the DNA−enzyme complex both in its unpoisoned23 and
INTRODUCTION
■
poisoned23−26 states.
Monomeric human topoisomerase 1B (Top1) regulates DNA
CICs may operate by blocking two key steps in the enzyme’s
topology throughout key cellular events such as DNA
catalytic cycle: substrate binding or covalent cleavage complex
formation. Compounds inhibiting the first step (CIC1) are
either conventional competitive inhibitors (binding to Top1)
replication and gene transcription.1−3 Eukaryotic Top1 relaxes
both positively and negatively supercoiled DNA5 and is an
established anticancer drug target6 (its inhibition initiates
or unconventional competitive inhibitors (binding to DNA).
apoptosis7,8 and hence tumor regression). Recent single-
Examples of the former are unknown, but unconventional
molecule nanomanipulation3,9,10 and molecular dynamics11,12
competitive inhibitors exist and are either DNA intercala-
studies of Top1 in the presence and absence of inhibitors and
tors27,28 or minor groove binders29−31 or both.32 Step 2
the originally proposed13 catalytic cycle of Top1 may be used to
catalytic inhibitors (CIC2) are currently rather obscure; one
construct a current view of the enzyme’s four-step cycle (Figure
recently described indolizinoquinoline-5,12-dione derivative,
1).
CY13I, possibly fits this descriptor.33
Drugs that block Top1 fall into two distinct classes: (1) well-
Of relevance to this study, DNA-binding Au3+ porphyrins34
characterized interfacial poisons (IFPs) and (2) less common
were classified as Top1 catalytic inhibitors,35 while other Au3+
catalytic inhibitor compounds (CICs).6 Currently, all DNA-
complexes were initially misassigned as Top1 IFPs35 and hence
intercalating IFPs arrest DNA strand religation by non-covalent
reclassified as catalytic inhibitors.36 The correct assignment of a
binding at the nick site of the Top1−DNA cleavage complex,4
poisoning the enzyme midcycle. Known IFPs include
compound’s mechanism of action (MOA) with Top1 is not
camptothecin (CPT) and its analogues and synthetic14
straightforward. The inherent problem is that inhibition of
compounds, e.g., indolocarbazoles,14,15 indenoisoquino-
supercoiled DNA relaxation by Top1 alone does not distinguish
lines,15−17 dibenzonaphthyridones,18,19 and aromathecins.2,20
between the actions of CICs and IFPs nor does it distinguish
between conventional and unconventional competitive inhib-
Some minor-groove binders that engage Top1’s DNA substrate
below the nick site, e.g., Hoechst 33258 and 33342,21 prevent
ition.
strand religation and are non-interfacial Top1 poisons. The
rational design22 of new IFPs and conceptual understanding of
how established IFPs work4 is underpinned by X-ray data for
Received: December 4, 2013
Published: April 2, 2014
© 2014 American Chemical Society
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Figure 1. (a) Illustration of key events in the catalytic cycle of human Top1. Step 1: Top1 binds supercoiled DNA (scDNA; 5′-TA-3′ dinucleotide
pair as target). Step 2: nucleophilic attack of the 3′-phosphate linking the TA pair (scissile strand) by Y723 (catalytic tyrosine residue) affords a
covalent DNA−Top1 cleavage complex and nicked strand. Step 3: the intrinsic torque stored in scDNA drives ratchet-like rotation about the non-
scissile strand until strand religation occurs with concomitant release of Y723. The turnover frequency1 is up to 6000 min−1. The relaxed DNA
(rDNA) is then released4 by the enzyme in step 4. Interfacial poisons (IFPs) such as camptothecin (CPT) bind the nick site via 5′-TA-3′
intercalation and H-bonding to Top1 to form a ternary drug·scDNA−Top1 adduct, poisoning the enzyme. CICs operate differently by either
blocking substrate recognition by Top1 (type 1 competitive inhibitor, CIC1) or, in principle, preventing the formation of the covalent cleavage
complex by blocking the nucleophilic attack of the scissile strand by Y723 (type 2 competitive inhibitor, CIC2). (b) Structures of new cytotoxic
pyrrole-based Au3+ macrocycles 1−5, free base macrocycle 6, and the Ni2+ analogue of 1, compound 7.
Since CIC’s do not trap Top1-nicked DNA, DNA damage by
this class of compounds is likely to be lower than that caused by
IFPs.37 The paucity of Top1 CICs coupled with their
anticipated reduced genotoxicity38,39 relative to IFPs creates
significant opportunities for drug discovery. Here we report on
a new class of cytotoxic macrocyclic Au3+ Top1 CICs (Figure
1) and precise delineation of the MOA of the lead compound,
3.
discussion is illustrated with selected, typical ion pairs from
the asymmetric unit of 3 (Figure 2). The Au3+ ions in salts 1,
RESULTS AND DISCUSSION
■
Compounds 1−5 reflect a design evolution over our recently
patented class of cytotoxic bis(pyrrolide-imine) Au3+ chelates.40
Specifically, we have employed macrocycles to enhance the
redox and chemical stability of the metal ion in conjunction
with a quinoxaline ring to augment DNA intercalation.
Macrocycles for 1−3 and 5 were synthesized by a literature
method.41 Direct metalation of the macrocycle (Route A,
Figure S2) was only successful for 1 and 5 (i.e., those
macrocycles bridged by a propyl chain). For compounds 2−4
with slightly more elaborate alkyl groups linking the imine
nitrogen atoms, a metal-templated cyclization had to be
employed (Route B).40 More specifically, the diamine bridge
required to effect cyclization of the macrocycle was added after
Au3+-binding by the bis(pyrrole-aldehyde) precursor.
(Although uncommon, Au3+ ions reportedly template aldehyde
and amine condensations.42)
Figure 2. View of the X-ray structure (100 K) of two of six
independent ion pairs from salt 3 (50% thermal ellipsoids for non-H
atoms; H atoms are shown as capped cylinders). Solvent molecules
have been omitted for clarity; selected atom labels are shown. Atom
color code: gray, C; lilac, N; orange, P; green, F; gold, Au; pale blue,
H. Crystallographic details for 1, 2b, and 3 are given in the Supporting
Information. Selected mean bond distances (Å) and angles (°) for all
six independent cations of 3: Au−Npyrrole, 1.98(1); Au−Nimine, 2.02(2);
CNimine, 1.30(3); Npyrrole−Au−Npyrrole, 97.1(8); cis-Npyrrole−Au−
Nimine, 81.9(8); trans-Npyrrole−Au−Nimine, 177(1); Nimine−Au−Nimine
,
99.2(4). The mean plane separation (3.4 Å) reflects notable π-stacking
for both the head-to-tail and oblique dimers of 3.
X-ray Structures. We were able to elucidate the structures
of compounds 1, 2b, and 3 by single crystal X-ray diffraction
(Table S1) despite the ordinarily challenging morphology of
the crystals (fine, brick-red needles). Because of the high degree
of similitude in the structures of 1−3, particularly the Au3+ ion
coordination geometry and macrocycle conformation and the
appreciable cytotoxicity of 3 (vide infra), the following
2b, and 3 are nominally square planar with Au−N bond
distances spanning the range 1.97−2.06 Å. The Au−Npyrrole
distances average 1.98(3) Å for all nine cations and are 1.5%
shorter than the Au−Nimine bonds, which average 2.01(6) Å
(Table S2). This reflects the fact that the pyrrole groups are
anionic σ-donors with more acute C−N−C angles than the
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corresponding CN−C angles of the imine donors. The
coordination group distances are comparable to those of other
Au3+ complexes (Au−N: 1.928−2.216 Å).43 The average
Npyrrole−Au−Npyrrole and Nimine−Au−Nimine bond angles for
the propyl-bridged systems 1 and 2b are 99.6(8)° and 96.5(5)°,
respectively. For the six independent cations of 3, these angles
average 97.1(9)° and 99.2(6)°. The larger seven-membered
chelate ring of 3 accounts for the somewhat more obtuse mean
Nimine−Au−Nimine bond angle. The average Npyrrole−Au−Nimine
bond angle of 82(1)° is effectively invariant for 1−3.
The cation conformations of 1−3 deviate mildly from
planarity mainly in response to crystal packing constraints. In
each structure, the cations form π-stacked dimers with head-to-
tail (Figure 2) or oblique geometries (Figures S8−S14)
characterized by interplanar spacings of ∼3.4 Å, as found in
other π-stacked polyaromatic compounds.44,45 There are no
aurophilic Au···Au contacts between cations (Au···Au distances
>5 Å). The Au3+ macrocycle pairs within dimers of 1−3 exhibit
overlaps ranging from 74% in 3 to 89% in 1. Importantly, the π-
stacking proclivity of the cations highlights their potential as
DNA intercalators.
DNA Binding and Intercalation. The affinity constants,
KA, for non-covalent binding of 1−5 to calf-thymus DNA
(ctDNA) were determined by fluorometric titrations involving
the displacement of intercalated ethidium bromide (EB, Figure
3a). From the loss of EB fluorescence (614 nm) with increasing
concentration of 1−5, KA values ranging from 2 × 106 to 4 ×
106 M−1 bp were obtained and are similar in magnitude to that
reported for a gold(III) porphyrin (3 × 106 M−1 bp).34 The KA
values for 1−5 showed no independent linear correlation with
either the steric bulk of the macrocycle’s alkyl bridge or the
lipophilicity of the cation. However, analysis of KA as a function
of both variables concurrently (Figure 3b) gives a significant
three-dimensional bivariate linear correlation and confirms that
cations with a small alkyl bridge and high lipophilicity, e.g. 5,
exhibit the highest KA values. The correlation itself suggests that
1−5 are DNA intercalators since not only are the KA values
determined by competitive displacement of a known DNA
intercalator (EB)46 but also the monotonic increase in KA with
increasing lipophilicity of the compounds clearly reflects
binding principally within the relatively non-polar intrahelical
space of the duplex DNA target (i.e., between π-stacked bases).
To confirm the intercalation data and examine possible roles
for the metal ion, electrophoretic mobility shift assays
(EMSAs), thermal denaturation, and DNA unwinding experi-
ments were carried out. Regarding the first of these tests, Figure
3c shows that 1 induces a greater mobility shift for a plasmid
substrate (pHOT1, lanes 5−9) than the archetypal reference
compound for DNA intercalation (EB, lanes 2−4), despite the
comparable ctDNA affinity constants for the two compounds
(EB, KA = 5(1) × 106 M−1 bp; compound 1, KA = 4.0(4) × 106
M−1 bp). The large mobility shift induced by 1 reflects marked
unwinding of the DNA duplex, consistent with intercalation.
Note that supercoiled plasmid DNA is affected most upon
binding of 1 (Figure 3d), as evidenced by the 11% mobility
shift relative to nicked-open circular DNA (NOC DNA, 5%
shift) and very marked band broadening apparent at a
concentration of only 5 μM (lane 7). In the case of a
negatively supercoiled plasmid with intact double strands, the
linking number describing the DNA topology, Lk, must remain
constant irrespective of the extent of local unwinding (i.e.,
change in twist, Tw) induced by the intercalator. To keep Lk
Figure 3. (a) Displacement of intercalated EB from ctDNA by 3
studied by emission spectroscopy (298 K, pH 7.0, 15% DMSO-TRIS/
HCl buffer, 15 μM ctDNA, 15 μM EB). Inset: determination of [3] at
50% loss of EB fluorescence (C50); C50 is used to determine the
ctDNA affinity constant, KA, of 3. (b) Graph of KA for 1, 2a, and 3−5
as a function of the steric bulk of the macrocycle’s alkyl bridge and the
hydrophobicity of the Au3+ complex, log(Po/w) (Table S3). The
surface is the best-fit bivariate linear regression function to the data.
(c) EMSA of selected compounds with supercoiled (SC, form I)
pHOT1 plasmid DNA (DNA, 31.3 ng/well; TBE buffer, pH 7.8);
some nicked-open circular (NOC, form II) DNA is present. The lanes
contain pHOT1 plasmid DNA (lane 1), increasing concentrations of
EB (lanes 2−4), 1 (lanes 5−9), 6 (lanes 10−12), and 7 (lanes 13−15).
The data prove that Au3+ is essential for DNA binding. (d) Two- and
three-dimensional deconvolution of the DNA bands in lanes 5−8 of
the EMSA gel shown in Part (c).
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Figure 4. (a) HRM curves for a linear 291-bp dsDNA fragment (pH 8.4) derived from the human ACTN3 gene as a function of the concentration of
1. The plot reflects the negative first derivative of the fluorescence intensity (−dF/dT, 510 nm) from the DNA-intercalating reporter dye (CYBR
Green) used to monitor strand separation. (NDC; no drug control, DMSO.) (b) Plot of the melting temperatures, Tm, against the concentrations of
three Au3+ macrocycles and the metal-free macrocycle 6. The zero-slope fit for 6 has an intercept, Tm, of 83.0(1) °C and reflects the mean value of
Tm for the amplicon (since 6 does not interact with the DNA). The dose−response function for 1 is: Tm = Tm0 + (Tmmax − Tm0)/(1 + (C/EC50)p),
where C is the molar concentration of 1. The fit parameters were: Tm0 = 82.95(9) °C; Tmmax = 88.1(4) °C; EC50 = 9(2) × 10−6 M; p = 0.9(1); χ2 =
0.013; R2 = 0.996. (c) DNA-unwinding assay to prove intercalative binding by 1. Lane 1, supercoiled pHOT1 plasmid DNA; lane 2, relaxed pHOT1
plasmid (effected by incubation with 10 units of Top1 for 30 min at 37 °C); lanes 3−5, increasing concentrations of the intercalator control m-
AMSA added to relaxed pHOT1; lanes 6−14, increasing concentrations of compounds 1, 7, and 6 added to relaxed pHOT1 as indicated.
Abbreviations: NOC, nicked-open circular DNA; SC, supercoiled DNA; RX, relaxed; TI, DNA topoisomer bands; m-AMSA, 4′-(9-
acridinylamino)methanesulfon-m-anisidide.
constant, Wr, which reflects supercoiling of the plasmid, must
cationic DNA·1 adduct (as would be required for reverse
migration of the DNA) is physically highly improbable.
Collectively, the EMSA data for 1 reflect intercalative binding
of dsDNA by the Au3+ macrocycle. This conclusion was further
tested by concurrent analysis of the metal-free macrocycle
(compound 6) and the Ni2+ analogue of 1 (compound 7);
neither induces a DNA mobility shift nor in fact displaces EB
from DNA in solution. The Au3+ ion is thus obligatory for
DNA intercalation by 1−5. That association depends on the
presence of Au3+ reflects electrostatic binding of the intercalator
to the DNA (as neither 6 nor 7 are charged) and a pivotal ion···
dipole interaction formed between the Au3+ ion and a carbonyl
oxygen atom of thymine (vide infra).
adjust according to eq 1:47,48
(1)
Lk = Tw + Wr
For Au3+ macrocycle 1, intercalation of negatively supercoiled
pHOT1 is expected to reduce the number of supercoils and
thus the macromolecule’s supercoil density, thereby signifi-
cantly impeding its mobility in the gel matrix (as observed).
The three-dimensional plot displayed in Figure 3d further
highlights the DNA distribution with increasing [1] in lanes 5−
8 and specifically emphasizes the appearance of a third, non-
migratory form of DNA (form III, lanes 7 and 8). Two possible
explanations for the appearance of this immobile DNA species
at higher concentrations of 1 are: (i) aggregation of the DNA·1
adduct(s) occurs, trapping the DNA in the well, or (ii) full
charge-neutralization of the DNA takes place. However, the
latter explanation seems unlikely since each adjacent dinucleo-
tide pair in dsDNA is bridged by two phosphate groups, each
with a formal charge of −1. Intercalation of a monocationic
intercalator can thus only neutralize 50% of the total charge on
the DNA duplex even if every adjacent dinucleotide pair non-
covalently binds to one Au3+ macrocycle, which is improbable.
Finally, the apparent “disappearance” of the DNA in lane 9 ([1]
= 50 μM) signals either reverse migration of all DNA·1 adducts
or complete saturation of the EB binding sites by 1 so that
staining of the gel for DNA visualization is ineffective. The
latter explanation is the more likely given that formation of a
Because the mode of interaction of the Au3+ macrocycles
with DNA has to be established unambiguously to determine
the mechanism of action underpinning the cytotoxicity of 2a
and 3 (vide infra), we performed two additional tests that
unequivocally support an intercalative DNA binding mode for
the compounds (Figure 4). In the first experiment, thermal
denaturation (melting) of a linear 291-bp DNA duplex (an
amplicon from exon 15 of the human ACTN3 gene)49−51 was
studied as a function of the identity and concentration of the
added compound (Table S12). From Figure 4a,b, the melting
point, Tm, increased in sigmoidal fashion with increasing
concentration of 1−3, consistent with the enhanced thermal
stability that is expected to accompany uptake of an
intercalator.52,53 Significantly, no change in Tm was observed
for 6 over the full concentration range. The values of ΔTm (the
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change in melting point relative to the control) were +4.65,
+3.90, and +4.40 °C for compounds 1−3, respectively, when
measured at their highest concentrations (∼50 μM). From
in lane 8 merit further reflection. As discussed in the
literature48,62 and Supporting Information, the method used
is incapable of determining whether the scDNA product
obtained at high concentrations of the intercalator is: (1)
positively or negatively supercoiled or (2) the result solely of
intercalator-induced torque or, alternatively, inhibition of Top1.
From lane 8 in Figure 4c, it is therefore possible that 1 may be
an inhibitor of Top1, but this cannot be concluded with
certainty in the absence of definitive mechanistic tests (e.g.,
tests that can discriminate between an IFP and a catalytic
inhibitor of the enzyme).
Cytotoxicity. Compounds 1−5 were prescreened by the
National Cancer Institute (NCI, Bethesda, MD) against their
panel of 60 human cancer cell lines; 2a and 3 were sufficiently
cytotoxic to warrant full five-dose screens and 3 proceeded to in
vivo hollow-fiber studies (Figures S35−S43, Table S6). Table 1
Figure 4b, the dose−response function for 1 gives the
max
saturating value of the melting point increase, ΔTm
=
+5.1(4) °C. The ΔTm values determined here are in good
agreement with those reported for the metallointercalators
[Co(pic)2(dppz)] (+4.4 °C), where pic = picolinate and dppz =
dipyrido[3,2-a:2′,3′-c]phenazine,54 and [Ru(NH3)4(dppz)]2+
(+5.2 °C)55 as well as organic protoberberine derivatives
(+2.9 to +6.4 °C).56 Other dppz-based metallointercalators,
however, exhibit ΔTm values as large as +14 °C.52
Several factors will determine the measured value of ΔTm
(beyond the identity of the primary intercalating group). These
include the nature of the ancillary ligands, the base sequence of
the dsDNA substrate (since this governs the density of binding
sites for base pair-specific intercalators), the solution conditions
(pH, buffers, salts57), and the experimental method. The
Table 1. Summary of Cytotoxicity Parameters for
method used here evidently affords reliable data over a wide
Compound 3 from a Five-Dose Screen against the NCI’s
Panel of 60 Human Cancer Cell Lines
a
max
concentration range, permitting measurement of ΔTm
.
However, as shown in Figure 4a, some caution in evaluating
the data is required at higher compound concentrations when
the signal is weak (−dF/dT < 0.1). For 1−3, the signal vanishes
above doses of ∼50 μM. The loss of signal intensity in the
experiment reflects two main processes: (1) The release of
CYBR Green (an intercalating dye that binds AT base pairs in
dsDNA via minor groove entry58) upon strand separation and,
consequently, quenching of its emission by torsional motion in
its non-intercalated state.57 This is the physical basis under-
pinning melt analysis with the high-resolution melt (HRM)
method available in commercial real-time PCR machines. (2)
Competition between the dye and intercalator for binding sites
along the length of the dsDNA substrate. This is evident from
the dose-dependent decrease in the emission signal from the
dye at a fixed temperature in Figure 4a and accounts for the
vanishing signal at high concentrations of 1−3. These points
noted, our conclusion that 1−3 are DNA intercalators remains
unambiguous. Further, from the data for 6, the irremissibility of
the Au3+ ion for DNA intercalation is again evident.
cancer
leukemia
N
GI50, μM
IC50, μM
LC50, μM
3
9
7
6
9
7
8
2
6
1(1)
6(5)
5(1)
>100
97(6)
non-small cell lung
colon
15(13)
10(6)
7(5)
89(16)
50(32)
>100
CNS
3(2)
10(8)
melanoma
ovarian
8(5)
27(20)
19(14)
25(21)
16(11)
17(14)
16(7)
11(8)
14(12)
7(4)
94(11)
89(24)
>100
renal
prostate
breast
5(4)
57(38)
86(19)
b
average
57
7(4)
a
Abbreviations: N, number of cell lines within each cancer category;
GI50, compound concentration effecting 50% growth inhibition; IC50,
compound concentration effecting 100% growth inhibition; LC50,
compound concentration that induces 50% cell death. Total number
of cell lines used. Estimated standard deviations are given in
parentheses; large values indicate variable susceptibility of a specific
group of cell lines to the test compound (see Figure S41).
b
In the second experiment, we used a standard DNA
unwinding assay59 designed to identify intercalators from
their interaction with DNA in the presence of Top1 (Figure
4c). The method was first described by Hsiang et al. in 198560
and elaborated on by Pommier et al. in 1987.48 In our
experiment, pHOT1 plasmid DNA was first relaxed by
incubation with 10 units of Top1 for 30 min at 37 °C (lane
2). Increasing concentrations of the test compounds were then
added, and the solutions incubated for a further 30 min interval
at 37 °C prior to workup and analysis. The DNA intercalator
control m-AMSA gave the expected Gaussian distribution of
DNA topoisomers (characterized by a linking number change,
ΔLk, of 1)48,61 at a concentration of 50 μM after workup. The
same topoisomer band distribution was observed at a
concentration of only 5 μM in the reaction with 1 (lane 7),
consistent with 1 being a more potent DNA intercalator than
m-AMSA. At a higher concentration (50 μM), 1 returns the
plasmid to its fully supercoiled state (lane 8). The reactions
with the Ni2+ analogue of 1 (compound 7) and the metal-free
macrocycle 6 confirm that neither intercalate DNA since the
signature dose-dependent Gaussian distribution of topoisomer
bands is noticeably absent in lanes 9−14. Similar results were
obtained running the reaction the other way48 (i.e., starting
with supercoiled pHOT1; Figure S54). The observations for 1
lists average cytotoxicity parameters for 3 grouped by cancer
type. The mean IC50 value across all cell lines for the Au3+
macrocycle is 16(7) μM; the large standard deviation reflects
considerable variation in the cytotoxicity of the compound
across each group of cell lines (as highlighted by the radar plot
of Figure 5). Leukemia, central nervous system (CNS), and
colon cancer were the most sensitive cancer types to 3.
Importantly, a total of 19 (out of 57) cancer cell lines had IC50
values for compound 3 that were <5 μM, accounting for
selection of 3 by the NCI for in vivo hollow-fiber assays (Table
S7, Figures S42 and S43). The five most sensitive cell lines were
NCI-H522 (non-small lung cancer, IC50 = 280 nM), RXF-393
(renal cancer, IC50 = 1.3 μM), SF-268 (CNS cancer, IC50 = 1.4
μM), SW-620 (colon cancer, IC50 = 1.5 μM), and LOX-IMVI
(melanoma, IC50 = 1.7 μM). In the NCI’s non-tumored animal
toxicity assay with female athymic nude mice (Figure S42),
compound 3 was remarkably well tolerated as reflected by a
100% survival rate after 17 days for doses ranging from 100 to
400 mg kg−1 dose−1 (intraperetoneal injection). Despite these
encouraging data, 3 was insufficiently cytotoxic in the majority
of the cell lines tested in the NCI’s hollow fiber cytotoxicity
assay to warrant progression to the next investigational phase
(tumor xenograft studies in live mice). Notwithstanding the
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of this study were both redox stable and essentially non-
aggregating under biologically pertinent conditions (Figures
S15 and S16), which might at least partly explain the fact that 3
was particularly well-tolerated in mice and essentially non-toxic
even at the highest test concentration of 400 mg kg−1.
From the graph of log(GI50) values for 3 against all 60 tumor
cell lines (Figure 5a), compound 3 is comparable to cisplatin
and midway between the limits of highly and mildly cytotoxic
compounds such as topotecan (a Top1 IFP) and 5-fluorouricil
(a clinically deployed antimetabolite), respectively. In order to
identify the MOA and hence the cellular target(s) of 3, the GI50
data for 3 were compared with equivalent data for 26
compounds of known MOA from the NCI’s database using
hierarchical cluster analysis (Figure 5b). The cytotoxicity profile
of 3 is clearly akin to that of CPT, the archetypal Top1 IFP.
Although the statistical data strongly suggest that 3 targets
Top1 in tumor cell cultures, they do not prove that 3 is an IFP
of Top1. Unambiguous determination of the MOA is essential
to definitively assign 3 as a Top1 IFP or Top1 CIC.
Enzyme Targeting. Cyril and Muller’s recent solid-phase
enzyme inhibition assay64 capable of distinguishing between
Top1 IFPs and CICs was used to elucidate the MOA of 3
(Figure 6a). The assay employs Top1 immobilized in Ni2+-
coated wells. Added IFPs trap DNA covalently bound to Top1
during enzyme turnover, which is subsequently detected by an
enhanced fluorescence signal from the DNA-sensing dye
PicoGreen. CICs of Top1 do not result in covalently trapped
DNA, favoring diminished emission from the reporter dye with
increasing compound concentration. At concentrations of 100
μM, 1−4 all give a negative readout (PicoGreen relative
fluorescence) akin to the signal obtained from the CIC control
(mitoxantrone, MTX). From this assay, 1−4 are clearly
catalytic inhibitors of Top1 at higher concentrations.
This result was confirmed for 3 using a novel assay designed
to trap Top1−DNA complexes using ultra-high enzyme:DNA
ratios65 (Figure 6b). In lanes 1−5, increasing [Top1] at a fixed
[DNA] in the absence of NaCl (ensuring a high DNA affinity
for Top1) increases the yield of NOC DNA in the reaction.
This is the expected result as we previously reported66 that
endogenous Top1 may “cluster” at catalytic sites on genomic
DNA in situ. Importantly, at high [Top1] and in the presence of
an IFP like CPT, multiple single-strand cleavages afford double-
strand breaks and thus linear DNA after workup. An IFP of
Top1 is therefore identified by its ability to generate linear
DNA in the assay. Lanes 11−16 confirm that 3 is not an IFP of
Top1 (no linear DNA); rather, the dose-dependent drop in
[NOC DNA] reflects catalytic inhibition of Top1 by 3.
Figure 5. (a) Radar plot of −log(GI50) values vs cancer cell line from
the NCI-60 cytotoxicity screen for 3 against their panel of 60 human
cancer cell lines. Comparative data for cisplatin, camptothecin (CPT),
and 5-fluorouricil (5FU) are shown to illustrate the cell-line dependent
response to the drugs as well as their effective cytotoxic range [mean
−log(GI50) values are given in the legend]. The graphical inset
illustrates the dose−response function for 3 with the CNS cancer cell
line U251 (the relevant GI50 value is indicated by the orange arrow on
the radar plot). (b) Hierarchical cluster analysis (group average
method, Minkowski distances) of the GI50 data for 3 with analogous
data for 26 anticancer drugs with known mechanisms of action (data
taken from the NCI database). Compound 3 clusters with
camptothecin (a Top1 IFP).
From Figure 6a, both 1 and MTX give a similar (positive)
readout to CPT at doses of 0.1 and 1 μM, suggesting that they
are Top1 IFPs at low concentrations. The switch to catalytic
inhibition of Top1 occurs at higher doses (∼1−10 μM). This
was confirmed for 1 by integrating the dose-dependent linear
DNA band intensities in a gel equivalent to that shown in
Figure 6b. From Figure S56, the concentration of linear DNA
increases with increasing [1] up to 500 nM, thereafter
exhibiting a sigmoidal decrease consistent with catalytic
inhibition of Top1 at higher doses. Dual-mode inhibition of
topoisomerases is well-known for Top2α65,67,68 and is equally
feasible, though rarely observed, for Top1. Compounds such as
doxorubicin target the DNA nick-sites in the enzyme−DNA
covalent cleavage complex at low doses, poisoning the enzyme,
but intercalate the DNA substrate at higher concentrations,
engendering catalytic inhibition.67 Mechanistically, a large
lower than desired activity of 3 in the hollow fiber assays, we
believe that 3 and its structural congeners represent an
important group of compounds that have the potential to be
developed into novel metallodrug lead candidates with
relatively low side-effects because of the in vivo stability of
the macrocyclic Au3+ complex. Going forward, absorption,
distribution, metabolism, and excretion studies are clearly
warranted in the case of 3 and may well delineate the factors
that reduce the efficacy of the compound in live animals.
Since release of an exogenous metal ion by a ligand system in
a live animal is often the cause of acute toxicity,63 an important
question to answer prior to determining the MOA of a
metallodrug candidate is how stable the compound is in
aqueous solution, especially in the presence of cellular reducing
agents such as glutathione. We found that the Au3+ macrocycles
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activity of the DNA intercalator control m-AMSA (lanes 13−
15), compound 3 engenders equivalent inhibition of the
enzyme at one-tenth of the concentration. Collectively, the data
in Figure 6 demonstrate that the Au3+ macrocycles are Top1
CICs. Furthermore, we estimate that the method-dependent
IC50 values for 3 are 49(2) nM and 9.2(5) μM from the assays
in Figure 6c,b, respectively (Figures S44 and S45). Notably,
these assays neither distinguish between conventional (drug
binds enzyme) and unconventional (drug binds substrate)
catalytic inhibition of Top1 nor between type 1 and 2 CICs of
the enzyme. However, since the DNA affinity of 1−5 has been
established with certainty, the compounds may be tentatively
assigned as unconventional CICs of Top1. (It is important to
note that concrete assignment of the above mechanism requires
an independent experiment to prove that the compounds do
not bind to the enzyme itself; vide infra.)
Finally, since hierarchical cluster analysis of the NCI-60 data
for 3 indicated a moderately close link between the in vitro
cellular target(s) of the Au3+ macrocycle and zorubicin (a
Top2α poison), we evaluated the compound’s ability to inhibit
Top2α. The results (Figure S55) indicate that 3 is a weak
catalytic inhibitor of the enzyme, as evidenced by an inhibitory
effect commencing only at a relatively high compound dose of
50 μM. Since no linear DNA was detected in the assay,
compound 3 is not an IFP of the enzyme. Evidently, 3 does not
target type II topoisomerases to the same extent as Top1. From
this, we conclude that 3 is a type I-specific CIC.
Binding Site Determination. If Top1’s DNA substrate is
targeted by 3, then where does intercalative binding of 3 occur?
One cannot naively assume that because 3 is a type 1 CIC of
Top1 (Figure 1), it binds to precisely the same nucleotide
sequence as the enzyme. We have used a conformational search
strategy with an empirical force field parametrized for
macromolecular simulations to accurately locate the binding
site of 3.
Because most molecular mechanics (MM) simulation
programs lack parameters for metal complexes, the X-ray data
for 1−3 (nine independent experimental structures) were first
used to develop a force field for the Au3+ macrocycles. We
augmented the SP4 force field (within AMMP)69 with
parameters to model 1−5 and related Au3+ chelates. The new
force field rivals density functional theory (DFT) in structural
accuracy but at a fraction of the computational cost (Table S2;
Figures S5 and S6). We then devised a conformational search
strategy (Figures 7a and S7) to determine the energetically
preferred intercalation point for 3 along the length of the 22-bp
DNA duplex typically employed for X-ray structure determi-
nations of Top1 with its DNA substrate (Figure 7d).23 (In this
respect it is noteworthy that classical force fields perform
surprisingly well when compared with computationally
expensive high-level quantum chemical simulations for
calculations of both the structure of DNA and, importantly,
base-pair stacking energies.70)
Figure 6. Enzyme target and mechanism assignment. (a) Relative
fluorescence intensity from DNA-bound PicoGreen (525 nm) vs the
concentration of two drug standards (CPT and mitoxantrone, MTX)
and 1−4. The assay measures covalent Top1−DNA trapped by the
test compound, distinguishing between IFPs and CICs. The data are
corrected for background emission (range indicated by the dashed
lines). (b) Proof of catalytic inhibition of Top1 by compound 3 using a
gel-based product-trapping assay (high enzyme:DNA ratio, 0 mM
NaCl, [pHOT1] = 31.3 ng/well in all reactions). Lanes 1−5: enhanced
yield of nicked-open circular DNA (NOC DNA) with increasing
[Top1]. Lanes 6−10: effect of increasing [CPT] from 0.5 to 50 μM
(100 units of Top1). Lanes 11−16: catalytic inhibition of Top1 (100
units) with increasing [3] attenuates the yield of NOC DNA (linear
DNA, L, is essentially absent above a dose of 1 μM). (c) Standard
scDNA relaxation assay for 3 and the DNA-intercalator m-AMSA.
Lanes 1−3: pHOT1 DNA alone, DMSO control, and Top1-mediated
relaxation of the substrate (no inhibitor present), respectively. Lanes
4−12: inhibition of Top1 with increasing [3] (complete inhibition
occurs at 5 μM). m-AMSA only completely inhibits Top1 at 50 μM
(lanes 13−15).
association constant, KA1, exists for formation of the ternary
drug·DNA−enzyme covalent cleavage complex, while a smaller
association constant, KA2, exists for intercalation of the
enzyme’s DNA substrate.65 Because the two assays used here
each permit a distinction to be made between Top1 IFPs and
catalytic inhibitors in a single experiment, they are ideal for
detecting dual-mode inhibitors. Consequently, 1 can be firmly
assigned as a dual-mode (IFP-CIC) Top1 inhibitor. Com-
pounds 2a−4, in contrast, are simple CICs.
In the conventional Top1 DNA relaxation assays (Topo-
GEN, Inc.; Figure 6c), 3 showed marked inhibition of the
enzyme at 500 nM and complete inhibition of the enzyme at a
dose of 5 μM (lanes 9 and 10). When compared with the
The lowest-energy conformation (Figure 7b, Table S5) has 3
intercalated at a 5′-TA-3′ site (T10-A11) via the major groove.
The butyl chain of the macrocycle juts out into the major
groove, oriented downstream (5′ to 3′). The next lowest-
energy conformation (Erel = 45.6 kJ mol−1, Figure 7c) has 3
intercalated via the minor groove (butyl chain oriented
downstream) at the same dinucleotide pair (T10-A11).
Minor groove intercalation yields a less stable non-covalent
adduct than major groove intercalation at a TA step for several
reasons, as depicted in Figure 8 (and Figures S46 and S47) and
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Figure 7. Binding site determination for 3. (a) Conformational search
strategy with a 22-bp DNA duplex sequence [Part (d)] favored by
Top1 and a modified SP4 force field for macromolecular simulations.
A macrocycle such as 3 may intercalate between an adjacent base pair
in one of four energetically distinct ways. The search proceeds in a
manner akin to stepping down the rungs of a ladder. (b) Structure of
the lowest-energy conformation of DNA·3. The compound
intercalates at the 5′-TA-3′ site (T10-A11) via major groove entry.
(c) Structure of the next-lowest energy conformation (minor groove
T10-A11 intercalation adduct) of DNA·3. The higher energy of this
adduct reflects, in part, the non-planar conformation for the DNA-
bound Au3+ macrocycle relative to the global minimum. (d) Graph of
thermodynamic stability vs base pair (left) for the 22-bp DNA duplex.
ΔGhelix is the empirical free energy penalty for strand separation at the
specified base pair. Right: graph of the relative energies of all simulated
DNA·3 intercalation adducts vs ΔGhelix. The best-fit straight line for
the major groove adducts is given by: Erel = 154(6) (ΔGhelix) − 83(20)
kJ mol−1; R2 = 0.985.
Figure 8. (a) View (roughly perpendicular to the AuN4 plane) of the
AMMP/SP4-simulated structure of the binding pocket (T10-A11) of
the lowest-energy intercalation adduct of 3 with the 22-bp DNA
duplex given in Figure 7d. The π-stacking between the bases and the
pyrrole and quinoxaline rings of 3 is highlighted along with the
juxtaposition of a T10 carbonyl oxygen atom (red sphere) and the
Au3+ ion (yellow sphere) of the intercalator. The Au···O distance is
3.13 Å. (b) Top view of the minor groove intercalation adduct
highlighting partial intercalation of the Au3+ macrocycle and the
practically negligible π−π overlap of the pyrrole rings with the bases of
the binding pocket. This view explains why the minor groove T10-A11
intercalation adduct is 45.6 kJ mol−1 higher in energy than the lowest-
energy conformation. In both structures, the DNA is rendered as a
stick model in shades of gray and lilac, H atoms are omitted for clarity,
and 3 is shown as a ball and stick model (C, green; N, blue; Au, gold).
The two inset illustrations show edge-on projections viewed from the
major to minor groove of the upper structure in both cases (left image,
van der Waals radii including H atoms; right image, ball and cylinder
model with H atoms omitted for clarity).
enumerated at this juncture. (1) In contrast to the major
groove intercalation adduct where both pyrrole rings and parts
of the quinoxaline ring of 3 are involved in π−π stacking with A
and T, only the edges of the quinoxaline ring of 3 interact with
these bases in the minor groove intercalation adduct. (2) The
extent of insertion of the macrocycle into the intrahelical space
is markedly less in the minor groove intercalation adduct. (3)
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The Au3+ macrocycle is forced to adopt a somewhat more
distorted (non-planar) conformation in the minor groove
intercalation adduct, which leads to a 5.58 kJ mol−1 increase in
the torsional strain energy relative to the conformer that best-
fits the binding pocket by major groove entry. (The total
energy difference between the two Au3+ macrocycle con-
formations is 4.94 kJ mol−1, with the conformer intercalated via
major groove entry lowest in energy.) (4) Electrostatic
interactions between the Au3+ ion and heteroatoms (O, N)
of either A or T in the minor groove adduct are absent (Figure
8b). Significantly, when embedded within the T10-A11 binding
pocket the lowest-energy major groove intercalation adduct of
3 exhibits a short Au···O contact (3.13 Å) involving the nearest
carbonyl oxygen of T10 and the Au3+ ion. This electrostatic
interaction (primarily an ion−dipole attraction) evidently
makes a vital contribution to the overall stability of the DNA·
3 non-covalent complex. Perhaps most significantly, the
conformational search gives the base pair specificity of 3
when the unique intercalation adducts are ordered from lowest
energy (highest affinity) to highest energy (lowest affinity):
(shown in Figure 7a) as a conformational reference. The data
are available in Tables S9−S11. Figure 9 graphically depicts the
TA(j) > TA(i) > AT(i) > AT(j) ≫ GA(i) > TT(i) > AC(j) > CT(j)
> AG(j) > AG(i)
where superscripts (j) and (i) are major and minor groove
insertions, respectively.
Independent verification of the above prediction that the
intercalation site for 3 (T10-A11) precisely matches Top1’s
target site for strand scission is mandatory. The experimental
thermodynamic stabilities of the 10 possible nearest-neighbor
interactions (e.g., AA/TT, AT/TA, etc.) present in a DNA
duplex are known71 and underpin the empirical algorithm used
to calculate the free energy penalty for helix unwinding
(ΔGhelix) by the program WEB-THERMODYN.72 We used this
method with a sliding window of two adjacent nucleotides to
analyze the 22-bp DNA duplex of interest (Figure 7d). As
shown by the bar graph, the stability of the adjacent nucleotide
pairs follows the order: TA < AT < AC < AG ≅ CT < AA = TT
< GA. The free energy penalty for separating the nucleotide
pair 5′-TA-3′ is thus the smallest (<1 kJ mol−1). The fact that
Top1 has evolved to effect single-strand scission of duplex
DNA at a 5′-TA-3′ site, a thermodynamic weak point in the
double helix, is noteworthy. It is also clear that a metal-
lointercalator such as 3 similarly targets a 5′-TA-3′ site in
duplex DNA and that because 3 and Top1 share the same DNA
target (substrate), compound 3 will necessarily function as a
type 1 unconventional catalytic inhibitor of the enzyme.
The scatter graph (Figure 7d) compares the relative energies
of the DNA·3 adducts with ΔGhelix for dinucleotide pair
separation. A good linear relationship exists for adducts formed
by binding/entry of the intercalator via the major groove (but
not the minor groove). Clearly, the stability order (base pair
specificity) for the major groove DNA intercalation adducts of
3 parallels the thermodynamic stability of the dinucleotide pairs
along the 22-bp sequence because the intercalator binds via the
major groove and has to part the bases, thereby locally
unwinding24 the helix at the intercalation point.
Figure 9. Analysis of key conformational parameters for the two
lowest-energy DNA·3 adducts formed by major and minor groove
intercalation of the Au3+ macrocycle at the T10-A11 step of the 22-bp
duplex depicted in Figure 7. (a) Graph of propeller twist (π) as a
function of base pair index for bases 4−19. (b) Graph of buckling
angle (κ) as a function of base pair index for bases 4−19. (c) Bar
graphs of base-pair opening (σ, left) and rise (Δz, right) for selected
base pairs close to the intercalation point.
most significant of the metallointercalator-induced conforma-
tional perturbations. Relative to the reference B-form 22-bp
duplex, which has a mean propeller twist, π, of −11.9(2)°, the
binding of 3 by either major or minor groove intercalation at
the T10-A11 step shifts the propeller twist from the range −30°
to −15° to between −5 and 0° immediately after the
intercalation point, after which π returns to ca. −25° (Figure
9a). The reduction in propeller twist reflects the more regular,
coplanar arrangement of the DNA bases which come into direct
DNA Conformation. Both major and minor groove
intercalation of 3 at the energetically favored T10-A11 step in
the 22-bp duplex lead to a number of characteristic conforma-
tional perturbations of the DNA. We used the program
W3DNA73,74 to quantify these effects at the level of individual
steps (i.e., complementary base pairs) using the AMMP-
calculated structure of the intercalator-free 22-bp DNA duplex
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contact with the aromatic intercalator through π-stacking; the
local ordering of the bases evidently extends from A11 to G12,
but no further. Figure 9b highlights the marked buckling of the
base pairs induced by intercalation of 3 at the T10-A11 step
through either the major or the minor groove. The mean
buckling parameter, κ, for the reference B-form 22-bp duplex
measures 0(3)°, as expected for an unperturbed B-form DNA
conformation.75 The intercalation of 3, in contrast, leads to
substantial buckling of the base pairs up- and downstream of
the intercalation point with κ averaging −3(8)° over the full 22-
bp duplex for both intercalation adducts. The swing or reversal
in κ from ca. −12° immediately upstream of the intercalation
point to ca. +12° immediately downstream of the T10-A11 step
is pronounced; such a conformational perturbation has been
previously noted for the structure of adriamycin-intercalated
DNA.76 The intercalation of 3 at the T10-A11 step is further
characterized by reverse opening (σ) of the 10-TA and 11-AT
base pairs (σmax = +10.1° for the 10-TA pair of the minor
groove adduct) relative to the reference B-form 22-bp duplex
for which σ = −10(1)° (Figure 9c). Finally, as expected the rise
between adjacent bases in the stack, Δz, increases at the
intercalation point because the intercalator assumes the
position of a hydrogen-bonded base pair in the stack.75,77
This is depicted graphically in Figure 9c where the 11-AT pair
clearly exhibits Δz values of 7.2 and 7.0 Å for the major and
minor groove intercalation adducts, respectively. The calculated
Δz values are consistent with the experimental values for base
rise that accompany the presence of a DNA-bound intercalator
(ca. 6.1−7.3 Å).46,76−78
Metallointercalation at TA Sites. Do other metal-
lointercalators bind at 5′-TA-3′ steps in duplex DNA? Although
a range of base pair specificities seem to exist for metal-
lointercalators and are probably ligand and metal ion
dependent, the recently determined X-ray structure of DNA-
bound Λ-[Ru(phen)2(dppz)]2+,79 where phen is 1,10-phanan-
throline, showed that the most symmetric (of many)
intercalative binding modes for this cationic minor groove
metallointercalator involved a 5′-TA-3′ site in a palindromic
duplex DNA sequence that was characterized by deep
intromission of the dppz ligand into the DNA intrahelical
space in a manner akin to the major groove intercalation adduct
calculated for 3 (Figure 8a). (Note that the quinoxaline ring of
3 is essentially two-thirds of the phenazine ring system in dppz,
so the intercalative components of the two ligand systems are
to a substantial extent comparable.) Interestingly, Λ-[Ru-
(phen)2(dppz)]2+ did not intercalate at a 5′-AT-3′ site in an
analogous palindromic duplex DNA sequence.79
ancillary ligand pairs (phen vs bpy), the non-innocence of the
co-ligands in directing (at least partly) the site of dppz
intercalation is highlighted. The intercalation mode calculated
here for 3 is therefore distinct, more closely parallels that
observed for major groove TA intercalation by Δ-[Rh-
(bpy)2(chrysi)]+ (where chrysi = chrysene-5,6-quinone dii-
mine),81 and evidently reflects the fact that the Au3+ ion is
square planar and housed within a macrocycle with practically
no steric bulk orthogonal to the mean plane of the quinoxaline
and pyrrole ring systems.
Molecular Basis of Enzyme Inhibition. Figure 8a gives a
detailed view of the dinucleotide binding site of the lowest-
energy structure of DNA·3. As noted above, in addition to π−π
stacking interactions of adenine and thymine with the pyrrole
rings of 3, a significant 3.13-Å O···Au contact involving a
carbonyl oxygen atom of T10 and the Au3+ ion of 3 indicates
that the binding pocket interacts electrostatically with the metal
ion. The simulations therefore explain why Au3+ is required for
DNA intercalation by the compounds. That 3 explicitly targets
a TA rather than an AT site is also apparent. Specifically, if the
order of the bases in the duplex is mutated to A10-T11 in the
simulation, formation of the key thymine O···Au3+ contact is
obviated (Figure S48). This A10-T11 intercalation adduct is,
furthermore, 82.0 kJ mol−1 higher in energy than the global
minimum energy structure (T10-A11 intercalation via the
major groove).
If 3 binds at Top1’s T10-A11 target site, can the enzyme still
recognize its substrate and form a covalent cleavage complex?
We answered these questions by simulating the structure of
wtTop1 non-covalently bound to DNA·3 (T10-A11 inter-
calation). From Figure 10a,b, the quinoxaline ring of 3
protrudes into the minor groove and sterically blocks the
enzyme’s probe residue R364 from hydrogen bonding to N3 of
G12. Steric repulsion between the quinoxaline ring and D533 is
also evident. Furthermore, as shown by the overlay of the
simulated structure of Top1·DNA·3 with the X-ray structure23
of the Top1 Y723F mutant (mTop1) bound to the 22-bp DNA
duplex (Figure 10b),23 intercalation of 3 engenders partial
unwinding of the DNA helix downstream of the intercalation
point. The strand shift measured by the displacement of G12 is
3.53 Å, consistent with π-stacked 3 assuming the position
normally occupied by a nucleobase (A11) and the magnitude of
the base rise (Δz = 7.2 Å; Figure 9c). Importantly, the scissile
strand’s T10-A11 phosphodiester link shifts 1.52 Å down-
stream, potentially thwarting attainment of the transition state
for covalent Tyr−O−P bond formation. Our macromolecular
simulations clearly predict that Top1 will neither bind its DNA
substrate nor form a cleavage complex in the presence of 3.
Mechanism Substantiation: SPR Studies. The foregoing
predictions were experimentally verified by determining
whether or not mTop1, a catalytically inactive mutant analogue
of Top1, binds to an oligonucleotide target fixed to a surface
plasmon resonance (SPR) chip in the presence and absence of
3 (Figure 10c). The mutant enzyme (mTop1) contains a point
mutation at the active site tyrosine (Y723F) which does not
appreciably alter overall protein structure or DNA binding
affinity but destroys the ability of the enzyme to engage DNA
in a cycle of cleavage and religation (which would complicate
our analysis of DNA binding). In this experiment, the surface of
the SPR chip was derivatized with a biotinylated 20-bp DNA
duplex before passing a solution of mTop1 over the chip. An
SPR response function commensurate with protein uptake
(40−100 s) followed by saturation (100−160 s) and desorption
Despite the different trajectories preferred for DNA
intercalation by 3 and Λ-[Ru(phen)2(dppz)]2+, the foregoing
experimental observation of TA over AT specificity parallels the
thermodynamic DNA base pair specificity determined here for
3 using macromolecular simulations. Note that the coordina-
tively saturated metal ion in Λ-[Ru(phen)2(dppz)]2+ does not
interact with any DNA bases and that the phen ligands
seemingly direct the trajectory of intercalation (minor groove
to major groove) through non-covalent interactions with the
minor groove nucleotides at the binding site. Interestingly, the
X-ray structure of DNA-bound Δ-[Ru(bpy)2(dppz)]2+, where
bpy is 2,2′-bipyridine, exhibits similar minor groove inter-
calation, but with a rather different dinucleotide pair specificity
to the former Ru2+ complex (binding at central and terminal 5′-
AT-3′ and 5′-CG-3′ steps, respectively, being favored).80 Since
the two Ru2+ complexes under discussion differ only in their
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desorption of 3 (Figure S50). The data reflect the presence
of the three 5′-TA-3′ sites along the synthetic 20-bp DNA
duplex and the fact that each has a unique microscopic affinity
constant for 3. The macroscopic KD values for 1 and 3 (Table
S4 and Figure S53) were 2.8 and 3.4 μM, respectively, broadly
in accord with their ctDNA affinity constants (Figure 3a).
Importantly, 3 did not bind to mTop1 (Figure S52), indicating
that the enzyme is neither a primary nor a secondary target for
the compound.
Based on the above SPR data, DNA binding experiments,
and enzyme targeting assays, the lead compound of this study
may be confidently assigned as an unconventional type 1
catalytic inhibitor of Top1. Going forward, it will be interesting
to ascertain whether the in vitro mechanism of action for 3
applies in vivo (i.e., in a chromatin setting). Although such
assays exist for IFPs of Top1,82 non-emissive Top1 CICs such
as 1−5, which are unsuitable for confocal microscopy, currently
present several unmet challenges regarding detection of
endogenous Top1 inhibition within cell nuclei.
CONCLUSIONS
■
In summary, we have synthesized and characterized a new class
of nominally planar cationic Au3+ macrocycles that incorporate
two pyrrole-imine units linked to a quinoxaline moiety on one
side and an alkyl chain bridge on the opposite side. From
inception, the compounds were designed to be cytotoxic DNA
intercalators. Physical measurements of DNA binding by the
compounds indicate that they are intercalators with high affinity
constants (KA > 106 M−1 bp) for ctDNA that correlate
interdependently with the lipophilicity of the salt and the steric
bulk of the alkyl chain bridge within the macrocycle.
Hierarchical cluster analysis of NCI-60 cytotoxicity data for
the most active compound (salt 3) indicated that 3 correlates
most closely with the topoisomerase IB (Top1) poison
camptothecin. Several topical enzyme inhibition assays were
used to prove that 3 is a catalytic inhibitor (and not a poison)
of Top1. Since catalytic inhibition of human topoisomerase IIα
(Top2α) by 3 was 2 orders of magnitude weaker than its
inhibition of Top1, compound 3 is a type I-specific agent.
New MM parameters were developed for the SP4 force field
for macromolecular simulations from the nine independent X-
ray structures of the Au3+ macrocycles determined herein for
parametrization. A conformational search strategy was devised
to locate the lowest energy intercalation site (adjacent
nucleobase pair) within a 22-bp DNA duplex commonly used
as a Top1 substrate. The simulations showed that 3 intercalates
DNA at the enzyme’s 5′-TA-3′ dinucleotide target sequence via
major groove entry (the minor groove adduct being >45 kJ
mol−1 higher in energy) and that a crucial Au···O electrostatic
interaction accounts for the observed base pair specificity.
Macromolecular simulations of a ternary non-covalent 3·DNA·
Top1 complex suggested that the molecular mechanism of
action of DNA-bound 3 is to block substrate recognition by the
enzyme through steric repulsion. Surface plasmon resonance
studies confirmed (1) that Top1 fails to bind its DNA substrate
in the presence of 3, (2) that 3 does not bind to Top1 itself,
and (3) that the base specificity of 3 deduced by the
macromolecular simulations (TA) is correct.
Figure 10. DNA intercalation by 3, the ensuing structural
perturbations, and their impact on DNA binding by Top1. (a)
Lowest-energy simulated structure of the wild type Top1·DNA·3
ternary complex illustrating steric displacement of Arg-364 and Asp-
533 by the quinoxaline ring of 3 protruding out the minor groove.
These “probe residues” of the enzyme are key to DNA substrate
recognition. (b) Overlay of the simulated structure displayed in Part
(a) with the X-ray structure of the Y723F Top1 mutant (mTop1)
bound to DNA (pdb code: pdb1a36). Key structural perturbations of
the enzyme’s substrate are highlighted. (c) DNA binding by mTop1 in
the presence and absence of 3 determined by SPR (surface plasmon
resonance; 37 °C, 41 μL min−1 flow rate, pH 7.4, 10% DMSO). The
on-chip duplex DNA sequence is illustrated beneath the graph; the
enzyme’s TA dinucleotide target is highlighted. The biotinylated 20-bp
duplex was anchored to avidin bound to the sensor chip surface. A
conceptual illustration of the relevant equilibrium is given above the
graph of SPR response functions.
(>160 s) was obtained. After flushing the chip with buffer, a
solution of mTop1 and 3 (50 nM) was passed over the chip as
before. No SPR response was detected (baseline signal in
Figure 10c), consistent with fast uptake of 3 by the DNA and
inhibition of enzyme binding by the immobilized DNA·3
intercalation adduct. The experiment therefore confirms the
MOA elucidated by the macromolecular simulations.
Finally, we garnered experimental proof that 3 binds to the
TA sites of the 20-bp oligonucleotide. Specifically, a solution of
3 passed over the DNA-embellished SPR chip afforded three
stepwise association response functions prior to discrete
The overarching conclusion of this multifaceted study is that
the most cytotoxic Au3+ macrocycle, lead compound 3, is an
unconventional type 1 catalytic inhibitor of human Top1.
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Journal of the American Chemical Society
Article
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ASSOCIATED CONTENT
* Supporting Information
Compound synthesis, spectroscopic characterization data,
molecular biology and methods, NCI cytotoxicity screens,
molecular simulation methods, tables of data, additional figures,
crystallographic data (CCDC: 928389−928391, www.ccdc.cam.
ac.uk/data_request/cif). This material is available free of charge
via the Internet at http://pubs.acs.org.
■
S
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AUTHOR INFORMATION
Corresponding Author
munroo@ukzn.ac.za
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Notes
The authors declare the following competing financial
interest(s): The authors wish to acknowledge that M.T.M. is
a member of the Scientific Advisory Board for TopoGEN, Inc.
§Visiting Fulbright Scholar to the University of Central Florida
ACKNOWLEDGMENTS
■
This work was supported by AuTEK Biomed (Advanced
Materials Division of MINTEK, RSA), the National Research
Foundation (RSA), the University of KwaZulu-Natal, Topo-
GEN Inc., Port Orange FL-32128, the NIH (indirectly through
the DTP), and the University of Central Florida. The authors
thank Dr. Bongyong Lee (postdoc to M.T.M., UCF) for
synthesis of the 20-bp biotinylated DNA duplex used for the
SPR studies. The authors are indebted to Dr. Holger Ott
(Application Scientist, Chemical Crystallography, Bruker AXS
GmbH, Karlsruhe, Germany) for collecting the X-ray data for 3
and for generating the initial structure model as part of a Bruker
X8 Prospector X-ray diffractometer test demonstration. We
thank Craig Grimmer for his help with recording NMR spectra,
Caryl Janse Van Rensburg for recording mass spectra, Sandipa
Bhikraj and Teshica Chatturgoon for checking the synthesis
and characterization of 2b, and Dr. Greg Watson for collecting
the HRM data. O.Q.M. thanks the Fulbright Foundation for a
2011/2 Visiting Professor’s Research Scholarship to the USA
and UCF.
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