Development and application of a novel acridinium ester for use as a chemiluminescent emitter in nucleic acid hybridisation assays using chemiluminescence quenching

Article abstract of DOI:10.1039/b811947c

Chemiluminescent acridinium esters (AEs) permit the development of high sensitivity ligand binding assays due to a combination of high intensity light emission and very low backgrounds. Here these advantages are exploited for use in homogeneous nucleic acid hybridisation assays using quenched chemiluminescence. AE chemiluminescence is conventionally initiated at highly alkaline pH. Novel "active" AEs were designed that permit initiation under conditions compatible with maintenance of nucleic acid hybrids (i.e. pH less than 9). Methyl red was found to be a dark quencher species capable of functioning at this pH. Practical application of the chemiluminescence quenching assay system has been demonstrated using two model nucleic acid hybridisation assays based on intra- and intermolecular emitter/quencher pairs.

Full text of DOI:10.1039/b811947c

View Article Online / Journal Homepage / Table of Contents for this issue
PAPER
www.rsc.org/obc | Organic & Biomolecular Chemistry
Development and application of a novel acridinium ester for use as a
chemiluminescent emitter in nucleic acid hybridisation assays using
chemiluminescence quenching
Richard C. Brown,a Zhaoqiang Li,a,b Andrew J. Rutter,a Xiaojing Mu,b Owen H. Weeks,a Keith Smith†b and
Ian Weeks*a
Received 14th July 2008, Accepted 21st October 2008
First published as an Advance Article on the web 24th November 2008
DOI: 10.1039/b811947c
Chemiluminescent acridinium esters (AEs) permit the development of high sensitivity ligand binding
assays due to a combination of high intensity light emission and very low backgrounds. Here these
advantages are exploited for use in homogeneous nucleic acid hybridisation assays using quenched
chemiluminescence. AE chemiluminescence is conventionally initiated at highly alkaline pH. Novel
“active” AEs were designed that permit initiation under conditions compatible with maintenance of
nucleic acid hybrids (i.e. pH less than 9). Methyl red was found to be a dark quencher species capable of
functioning at this pH. Practical application of the chemiluminescence quenching assay system has
been demonstrated using two model nucleic acid hybridisation assays based on intra- and
intermolecular emitter/quencher pairs.
the AE/quencher labelled probe induced by hybridisation with the
Introduction
analytical target nucleic acid will have the effect of either separating
Acridinium esters (AEs) have been used extensively as end-point
or bringing closer the emitter/quencher pair such that the emitted
labels in ligand binding assays such as immunoassays and nucleic
signal intensity proportionately reflects the degree of hybridisation
acid hybridisation assays.1 Conventionally, AE chemiluminescence
and thus the quantity of analyte nucleic acid present. However,
is measured as a rapid flash type emission (typically one second
simple substitution of a chemiluminescent AE label for a fluores-
duration) of visible light (wavelength approximately 430 nm).
cent label as presently used in analogous fluorescence quenching
Ligand binding assays can be “heterogeneous” or “homogeneous”
assays will not result in a workable system since, in contrast to
depending respectively on whether or not the binding complexes
formed between the target molecule and binding reagent need
fluorescence, AE chemiluminescence requires an altered chemical
environment (e.g. elevated pH) to effect signal emission. This will
to be physically separated from non-bound reagent, for example
impact on the components of the assay system for several reasons.
by solid-phase adsorption, prior to end-point measurement. AEs
Firstly the hybridised nucleic acid duplex cannot survive exposure
to elevated pH (>9)3 because of dissociation. Secondly the desired
optical absorption characteristics of the chosen matched quencher
molecule must be retained under the conditions used to initiate the
chemiluminescent reaction.
have been used successfully in both types of assay but have
not thus far been widely applied to homogeneous assay systems
based on energy transfer quenched chemiluminescence. The more
commonly encountered quenched fluorescence methods use a
matched fluorescent emitter/quencher pair to monitor ligand-
binding reactions. However, the performance of simple, solution-
phase fluorescence-based assays is limited by the low sensitivity
of detection of the emitted signal as a consequence of the
background due to scattered incident light and endogenous sample
fluorescence.2 In contrast, quenched AE chemiluminescence offers
greatly increased sensitivity as a result of its inherently lower
background.
The general rules for the use of AE as a biomolecular labelling
species are that the chemiluminescent signal is easily triggered,
that the AE can be readily coupled to ligand and that the
chemiluminescent and biochemical properties of such AE–ligand
conjugates are essentially the same as the individual components.
Theuseof N-hydroxysuccinimideester derivatives ofAE havebeen
shown to permit efficient coupling with selected amine groups in
proteins or modified oligonucleotides using established labelling
methods and post-labelling purification techniques.1 This allows
production of high specific activity labelled species that can be used
at low concentrations (1–10 femtomoles per reaction) in sensitive
ligand binding assays.
We were particularly interested in exploiting the potentially
high sensitivity of detection of AE chemiluminescence quenching
systems in nucleic acid probe hybridisation assays. Depending on
the nature of the probe/target design, conformational change of
For application to nucleic acid chemiluminescence quenching
assays several other, more specific, criteria also need to be achieved.
Firstly the AE chemiluminescence must be sufficiently intense to
permit high sensitivity measurements over a conveniently short
time period when initiated under conditions compatible with
the maintenance of the labelled probe-nucleic acid target duplex
which, evidence suggests, is less than or equal to a pH of 9.3
aMolecular Light Technology Research Limited (A subsidiary of Gen-Probe
Incorporated, 10210 Genetic Center Drive, San Diego, CA 92121, USA), 5
Chiltern Close, Cardiff Industrial Park, Cardiff, Wales, UK CF14 5DL
bDepartment of Chemistry, University of Wales, Swansea, Wales, UK SA2
8PP
† Present address: School of Chemistry, Cardiff University, Cardiff, Wales,
UK, CF10 3AT.
386 | Org. Biomol. Chem., 2009, 7, 386–394
This journal is
The Royal Society of Chemistry 2009
©

View Article Online
Secondly, the AE labelled probe must also display sufficient
stability under conditions of storage and use such that there is
no appreciable loss of activity. This is of particular relevance in
the context of assay incubation conditions where the ambient pH
may itself be alkaline. Finally the chemiluminescent reaction must
allow the emitting species to remain in sufficiently close proximity
to the quencher for energy transfer to occur. This combination
of features has not previously been demonstrated nor applied to
AE quenched chemiluminescence assays by commonly used AEs.
Thus, achievement of a functional chemiluminescence quenching-
based system has necessitated the design and synthesis of a novel
AE compound that has this desired combination of properties.
Additionally, a further requirement was to ensure that the
proposed quencher was capable of quenching the emission under
the chemical conditions used to bring about the chemiluminescent
reaction.
and thus renders the molecule more “active”. Although presently
established AEs do not yield chemiluminescence at pH ≤9 (which
is necessary to preserve nucleic acid duplexes), we reasoned
that “activated” AEs would be chemiluminescent under such
conditions. However, such substitutions are also likely to render
the phenyl ester bond more prone to hydrolysis and thus unstable
in aqueous solution and so a compromise would be needed
in this regard. It has also been shown that some stabilisation
towards hydrolysis can be achieved by incorporating bulky groups
at the 2 and 6 positions of the phenol.5d Our strategy was to
examine structures where electron withdrawing groups were also
used to create some element of steric inhibition of hydrolysis by
incorporating them at these positions. Several structures were
investigated and three candidate structures (compounds 1 to 3)
were selected for further study.
Here we describe the design and synthesis of a novel
AE/quencher pair and demonstrate the successful application of
the chemiluminescence quenching principle to model assay sys-
tems. Molecular “hairpin probes”4 were used to demonstrate the
application of this principle to intramolecular emitter/quencher
pairs and complementary oligonucleotide pairs were used to
demonstrate the principle as applied to intermolecular hybridi-
sation, in this example as a model helicase substrate. These
systems demonstrate that AE chemiluminescence quenching can
be successfully used as the basis of highly sensitive nucleic acid
hybridisation assays and their associated applications.
Selection of optimum AE
Fig. 1 shows typical chemiluminescent emission kinetics of
compounds 1–3 when the chemiluminescent reaction is initiated
(here at pH10).
Results and discussion
Design and synthesis of the chemiluminescent emitter and quencher
Fig. 1 Kinetics of chemiluminescence of compounds 1, 2 and 3 initiated
at pH 10 using a 1 second measurement time. Chemiluminescence signal
is expressed as arbitrary relative light units (RLU).
It can be seen that compound 2 exhibits the fastest kinetics
and so appears to be a good candidate to exhibit lower pH
chemiluminescence emission. However, studies of the stability
of buffered aqueous acetonitrile solutions of the compounds
(Table 1) showed that compounds 2 and 3 are inherently less stable
than compound 1 and therefore less likely to be suitable for routine
use.
A good compromise between the reactivity and stability of the
substituted phenyl ester was achieved with the 2, 6-dibromophenyl
AE labels used in existing chemiluminescent ligand binding assays
are exposed to highly alkaline hydrogen peroxide to produce
chemiluminescence. Most such labels produce little or no emission
at pH ≤95a and thus cannot satisfy the criteria required for
the present application. It has previously been established that
the chemiluminescent reaction rate of the AE can be varied by
changing the substituents of the phenyl ring.
Stabilisation of the phenolate anion leaving group, which is
produced during the chemiluminescent reaction, with electron-
withdrawing substituents (accompanied by the reduction in pKa
of the corresponding phenol5c), increases the rate of the reaction
Table 1 Stability of substituted phenyl AE compounds 1–3a
Compound
0 daysb
4 daysb
6 daysb
1
2
3
100
100
100
98
95
74
100
71
57
a Hydrolysis of the phenyl acridinium ester results in the formation of a
non-chemiluminescent product which is reflected by loss of chemilumines-
cent activity. b Data are RLU expressed as a percentage of the emission
intensity of the freshly prepared solution.
This journal is
The Royal Society of Chemistry 2009
Org. Biomol. Chem., 2009, 7, 386–394 | 387
©

View Article Online
ester (compound 1). Compound 4 is an analogue of compound
1 in which the chemiluminescent reaction results in the acridone
emitter no longer being covalently attached to the ligand. This
property has been utilised in the past with established AEs such
as compound 5 in heterogeneous assays to minimise unwanted
loss of quantum yield of the primary emitter due to possible non-
specific quenching effects exerted by the attached ligand1 and to
allow maximum sensitivity of detection to be achieved.
both selected for further investigation as oligonucleotide labels for
a chemiluminescence quenching assay, which permitted side-by-
side comparison of the effect of the geometry of attachment to
oligonucleotides on the quenching efficiency. However, it was first
necessary to identify a suitable quencher of the chemiluminescent
emission.
Selection of optimum quencher
However, for the present purpose, where specific quenching is
the basis of the assay, such a property represents a disadvantage
since maximal chemiluminescence quenching is required. Fig. 2
and 3 show typical chemiluminescent kinetic profiles, produced at
pH 14 and 9, respectively, of generic oligonucleotides labelled with
compounds 1, 4 and 5.
The key requirement for a suitable quenching molecule is that it
must act as a dark quencher, which causes the electronically excited
state of the primary emitter of the chemiluminescent reaction
to reach its ground state by radiationless decay rather than by
photonic emission. In the present situation, this property must be
a feature of the quencher molecule under the chemical conditions
that exist at the point of excited state product formation during the
chemiluminescent reaction. By contrast, this is not a consideration
for the design of fluorescence quenching systems where there is no
net chemical change during the photophysical process. Accord-
ingly, molecules known to quench fluorescence may be ineffective
as chemiluminescence quenchers. It is known from fluorescence
studies6 that quenching can occur by either resonance energy
transfer or contact-mediated quenching depending on the system
concerned. The latter has less constraint in terms of the overlap of
the emission spectrum of the donor and the absorption spectrum
of the quencher compared to the former. However, in general
terms, it is desirable to maximize spectral overlap with a quencher
having a high extinction coefficient in the event that resonance
energy-transfer is an important contributor to quenching in a
given system. Fig. 4 shows an overlay of the emission spectrum
of N-methylacridone and the absorption spectra of dabcyl and
methyl red (as terminal 3¢ polyA(10) oligonucleotide conjugates)
obtained under conditions corresponding to those required for
the chemiluminescent reaction (pH 9). The emission spectrum is
representative of the acridone emitters of the chemiluminescent
reactions studied here and displays a relatively narrow emission
bandwidth of 415 nm to 470 nm with a maximum at approximately
430 nm. It can be seen that, under the chemical conditions
used here for the chemiluminescent reaction, the wavelength
overlap of the acridone emission with the absorption of a methyl
red oligonucleotide conjugate (lmax = 460 nm) was marginally
Fig. 2 Kinetics of chemiluminescence of compounds 1, 4 and 5 initiated
at pH 14.
Fig. 3 Kinetics of chemiluminescence of compounds 1, 4 and 5 initiated
at pH 9.
As can be seen from Fig. 2, all three compounds readily
undergo a chemiluminescent reaction at pH 14. However, as
shown in Fig. 3, although compounds 1 and 4 yielded greater
than 95% of the total chemiluminescent signal within 5 seconds
at pH 9, compound 5 was inactive at this pH and is thus
unsuitable for use in situations where the integrity of ligand
binding complexes must be preserved. Compounds 1 and 4
displayed similar chemiluminescent kinetics, though compound
1 exhibited slightly lower light emission intensity when coupled to
the oligonucleotide ligands used here. This was of little practical
significance in this study and so compounds 1 and 4 were therefore
Fig. 4 Absorption spectra of polyA(10 )dabcyl (B) and polyA(10)methyl red
(C) both at approximately 20 mM (normalised by OD 260 nm) in Tris buffer
(0.2 M, pH 9.0). (A) corresponds to the fluorescence emission spectrum of
N-methylacridone in arbitrary fluorescence units.
388 | Org. Biomol. Chem., 2009, 7, 386–394
This journal is
The Royal Society of Chemistry 2009
©

View Article Online
greater than that with a dabcyl oligonucleotide conjugate (lmax
=
primary emitter from the rest of the molecular complex during the
chemiluminescent reaction, which reduces the efficiency of specific
quenching. By contrast, this does not occur with compound 1 since
the primary emitter remains attached to the oligonucleotide. Thus
the background signal for compound 1 labelled probe was less than
1% of the signal observed when a 10-fold molar excess of target
was present. Accordingly, an approximate sensitivity of detection
of less than 2 fmol was achieved.
The dose-response curve of the assay of the target using the
compound 1 labelled probe was also compared using pH 14 and
pH 9 chemiluminescence conditions. At pH 14 the zero target
signal was much higher than that obtained at pH 9 (Fig. 6) and
the assay exhibited a narrower dynamic range.
480 nm). In practice, both compounds appeared suitable though
here methyl red was selected as the candidate quencher for further
investigation.
Use of AE compound 1 and methyl red quencher derivative 6 as an
intramolecular emitter/quencher pair demonstrated by a model
chemiluminescence quenching hairpin probe
In order to demonstrate the use of the selected emitters and
quencher a labelled “hairpin” oligonucleotide probe sequence
5¢CCGGTCCAGGTGGAGCAATGATCTTGATCTTCAT-
¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯
GACCGG 3¢ (sequence 1) was used. The model hairpin
probe was a conventional stem-loop structure4 consisting of a
28-nucleotide target binding sequence (underlined) flanked by
6-nucleotide complementary stem-binding sequences (italics).
The oligonucleotide was initially created with an incorporated
3¢ tritylthio group and a free 5¢ amino group. The probe was
labelled with either compound 1 or compound 4 at the 5¢ amino
group and both probes were then de-protected to reveal the free
thiol group and labelled with quencher 6 at the 3¢ position. In its
normal state the labelled probe (sequence 1) adopts a “hairpin”
configuration as a result of intramolecular duplex formation of the
stem-binding regions, resulting in juxtaposition of the emitter and
quencher. Consequently, when the chemiluminescent reaction was
initiated, quenching occurred as demonstrated by the absence of
emitted light. However, following incubation with its complemen-
tary target, 5¢ATGAAGATCAAGATCATTGCTCCACCTG 3¢
(sequence 2), emission of light occurred when the chemilu-
minescent reaction was initiated. This is consistent with the
assumption that the target-binding region of sequence 1 forms an
intermolecular duplex with sequence 2 which disrupts the existing
stem-binding (intramolecular) duplex of sequence 1. This, in turn,
results in separation of the emitter and quencher. Fig. 5 shows
dose-response curves of the binding of the target (sequence 2)
with sequence 1 labelled with quencher 6 and either compound 1
or compound 4. Although both displayed a target dose-related
increase in chemiluminescence intensity, the zero target signal
of the compound 1 probe was much lower than that of the
compound 4 probe. The high signal observed with the compound
4 labelled probe in the absence of target (i.e. the background
signal) is probably the result of the covalent dissociation of the
Fig. 6 Dose-response of the binding of target (sequence 2) to the
model hairpin probe (sequence 1) labelled with compound 1 using
chemiluminescence conditions at pH 9 and pH 14.
This observation is consistent with the assertion that with
conventional AE chemiluminescence at pH 14, the stem-loop
structure is “de-hybridised” non-specifically with the resultant loss
of emitter/quencher proximity.
In order to demonstrate the use of the selected chemilumine-
scent emitter and quencher in an intermolecular emitter/
quencher pair we first obtained a 44mer oligonucleotide,
5¢CGCGAAGCGTTCGCACTTCGTCCCGCCTTCCTGCG-
¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯
CCTTCCTGT 3¢ (sequence 3), and a 36mer oligonucleotide,
¯
5¢CATGCTCGCAGCGGGACGAAGTGCGAACGCTTCGC-
¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯ ¯
G 3¢, (sequence 4), part of which is complementary to part of
¯
sequence 3 (underlined). The first 26 nucleotides of the 5¢ end of
the 44mer are complementary to the first 26 nucleotides of the 3¢
end of the 36mer and the remainder of the nucleotides comprises
a non-complementary ragged end with a 3¢overhang of the longer
strand. The 44mer oligonucleotide was designed with a 5¢ NH2
moiety for subsequent AE conjugation whereas the 3¢ end of the
36mer oligonucleotide was provided with the quencher already
attached. The 5¢ end of the 44mer (sequence 3) was labelled with
either compound 1 or 4. When hybridised to each other, these
oligonucleotides form a duplex with both blunt and ragged ends,
consistent with the secondary structure that enables the duplex
to act as a substrate for the 5¢ DNA helicase enzyme from B.
stearothermophilus7 (Cambio Ltd., UK). The emitter/quencher
moieties are thus juxtaposed at the blunt end of the duplex.
In order to study the quenching of the emission of compound
1 labelled sequence 3 by quencher labelled sequence 4, we firstly
Fig. 5 Dose-response of the binding of target (sequence 2) to the model
hairpin probe (sequence 1) labelled with either compound 1 or compound
4. Data points are mean SD (n = 3).
This journal is
The Royal Society of Chemistry 2009
Org. Biomol. Chem., 2009, 7, 386–394 | 389
©

View Article Online
incubated the former with a 1000-fold molar excess of the latter
at 43 C for 10 minutes following which chemiluminescence was
measured at pH 9 and pH 14.
sponding compound 4 labelled oligonucleotide. Accordingly, the
duplex formed by the association of compound 1 labelled sequence
3 and quencher labelled sequence 4 was subsequently used as a
substrate for helicase enzyme activity studies in which the action
of the enzyme is to unwind nucleic acid duplexes.
◦
Fig. 7 shows that co-incubation of the two complementary
oligonucleotides resulted in loss of chemiluminescent signal when
measured at pH 9. This quenching was not observed using
chemiluminescence measurement at pH 14 nor with replacement
of sequence 4 with a non-complementary (irrelevant) quencher
labelled oligonucleotide with chemiluminescence measured at
pH 9, indicating that the loss of signal was due to specific,
hybridisation-mediated chemiluminescence quenching.
Initial studies of AE-labelled sequence 3 (performed in the
absence of quencher labelled sequence 4) indicated that the
recovery of signal following the prescribed incubation was only
43% and 47% (relative to the RLU obtained at 0 minutes) for
compound 4 and compound 1, respectively. This suggested that
the combination of even slightly alkaline pH (here pH 7.5) and
elevated temperature (50 ◦C) resulted in a loss of signal due to
partial hydrolysis of the AE. In order to overcome the potential
loss of specific chemiluminescence activity due to AE hydrolysis
under the above conditions, the production of substrate was
undertaken in mildly acidic buffer (annealing reagent), which was
used with extended incubation times at lower temperature. This
yielded a substrate comprising a duplex with a quenching efficiency
of greater than 99% as measured by comparison of the chemilu-
minescence yield obtained using pH 9 (maximally hybridised) and
pH 14 (maximally single stranded) chemiluminescence conditions.
This substrate was thus expected to demonstrate low background
and high yield when used in helicase enzyme activity assays.
The suitability of the above duplex as a substrate for helicase
enzyme was demonstrated by incubating mixtures of various
concentrations of the enzyme with the substrate thence measuring
chemiluminescence at pH 9 of 10 mL samples taken from the
reaction mixture at defined times. The enzyme incubation was
performed in the presence of the required co-factors at pH 6.8
which is optimal for this particular enzyme. Fig. 9 shows the
reaction time course for helicase activity, which demonstrates a
progressive increase in chemiluminescent signal consistent with
the predicted duplex unwinding and consequent separation of
the respective emitter/quencher labelled oligonucleotide strands.
At maximal unwinding there was an approximately 40-fold in-
crease in chemiluminescent emission relative to the corresponding
background signal in the absence of enzyme. Omission of either
magnesium or ATP co-factors from the incubation buffer resulted
in loss of enzyme activity (data not shown) confirming that the
change in chemiluminescence was mediated specifically by enzyme
activity.
Fig. 7 Intermolecular hybridisation induced chemiluminescence quench-
ing. Data are normalised to allow for increased efficiency of the chemilu-
minescent reaction initiated at pH 14 (shaded bars) relative to pH 9 (black
bars). The response axis is expressed as the RLU as a percentage of the
signal obtained with no quencher oligonucleotide present (“no oligo”).
In order to permit comparison of the relative performance of
compound 1 and compound 4 labelled sequence 3, a 12.5 fmol
aliquot of each labelled probe respectively was incubated with
various amounts of the quencher labelled sequence 4 for 60 min-
◦
utes at 50 C. Fig. 8 shows that increased duplex formation was
associated with a quenched chemiluminescent signal at pH 9 in
both cases.
Fig. 8 Comparison of the chemiluminescence emission intensity of
compound 1 and 4 labelled sequence 3 following hybridisation with various
doses of quencher labelled sequence 4.
However, in common with the hairpin probe described above,
the maximum degree of quenching of the compound 1 labelled
oligonucleotide was greater than that observed with the corre-
Fig. 9 Time course of helicase-dependent formation of single stranded
compound 1 labelled 44mer (sequence 3).
390 | Org. Biomol. Chem., 2009, 7, 386–394
This journal is
The Royal Society of Chemistry 2009
©

View Article Online
Synthesis of AEs 1–5 and quencher 6
Conclusion
The various substituted phenyl acridine ester synthetic intermedi-
ates of compounds 1–5 were synthesised as described previously.8
Briefly, the acridine-9-carboxylic acid was converted to the acid
chloride by the action of thionyl chloride. A suspension of the acid
chloride in anhydrous pyridine was then stirred under anhydrous
conditions with the required substituted phenol to yield the ester.
Compounds 4 and 5 were prepared as described previously.8
Fluorescence quenching has been widely used as a means of
detecting and quantifying nucleic acids in solution-based ligand-
binding assays, but suffers from relatively poor sensitivity of
detection in this context. By contrast, chemiluminescent molecules
can form the basis of simple, highly sensitive detection sys-
tems. However, we have demonstrated that the simple view of
taking a known fluorescence quenching system and replacing
the fluorescent molecule with a chemiluminescent molecule is
an untenable one since other factors need to be considered.
Acridinium esters (AE) are widely recognised as highly successful
chemiluminescent labels and offer many advantages over other
labels. Despite this fact, no workable consideration has been
given previously as to how AE might be used in a quenched
chemiluminescence, ligand-binding assay since the highly alkaline
conditions used to initiate AE reactions are not compatible with
maintaining the integrity of the binding complex, which is crucial
for modulating the quenching process. We have demonstrated this
by showing non-specific loss of quenching when chemilumines-
cence quenched binding complexes are exposed to the elevated
pH normally associated with conventional AE chemiluminescent
systems. Moreover, many fluorescence quenchers are potentially
incompatible with the change in chemical conditions required to
initiate AE chemiluminescence such that performance predicted
from fluorescent systems may be reduced under the conditions
required for certain chemiluminescence reactions. In fact, we have
found it necessary to design a novel AE/quencher pair capable
of functioning within the restrictive chemical boundaries required
for maintenance of the integrity of biological binding complexes
which is crucial for modulation of specific quenching and which is
sufficiently stable for routine use.
Chemiluminescence of oligonucleotide probes labelled with
compound 1 was empirically optimised to allow an acceptably
rapid flash (5 seconds duration) at a pH that allowed maximal
signal generation but with minimal disruption of the secondary
structure of the hybridised probe. Under the conditions employed
here, 0.2 M Tris pH 9.0 was found to give a rapid signal with
minimal apparent disruption of the duplex. We also found that
derivatives of compound 6 were highly effective quenchers of the
chemiluminescence emission under these conditions. Increasing
the chemiluminescent reaction pH above 9 showed a progressive
increase in background signal consistent with loss of secondary
structure and concomitant non-specific loss of quenching.
We have demonstrated the ability of the optimised quenched
chemiluminescence system to form the basis of highly sensitive,
homogeneous detection systems for oligonucleotide binding pro-
cesses and related applications in which the intensity of chemi-
luminescence emission is modulated by either intramolecular or
intermolecular duplex formation.
Synthesis of 9-(2,6-dibromophenoxycarbonyl)-10-(3-succinimi-
dyloxycarbonylpropyl)acridinium iodide (Compound 1). A solu-
tion of 4-iodobutanoic acid (480 mg, 2.24 mmol), N-hydroxy-
succinimide (247 mg, 2.14 mmol) and dicyclohexylcarbodiimide
(523 mg, 2.53 mmol) was stirred in tetrahydrofuran (12 ml) under
◦
anhydrous conditions at 0 C for 3 h then at room temperature
overnight. Following filtration of the mixture, the filtrate was
concentrated on a rotary evaporator and chromatographed on
a silica column eluted with toluene–ethyl acetate (4 : 1 v/v).
Fractions at Rf = 0.48 were combined and evaporated to yield
succinimidyl 4-iodobutanoate as a yellow solid (567 mg, 84% yield,
mp 86–87 ◦C; nmax/cm-1 1739 and 1657 (CO); dH (400 MHz,
CDCl3, Me4Si) 3.22 (2 H, t, J 7, ICH2CH2), 2.78 (4 H, s,
COCH2CH2CO), 2.71 (2 H, t, J 7, CH2CH2COO), 2.18 (2 H,
quintet, J 7, CH2CH2CH2); dC (400 MHz, CDCl3, Me4Si) 4.2,
26.0, 28.5, 32.2, 168.0, 169.4; m/z (EI) 311 (M+, 1%), 197 (100), 184
(29), 169 (52)). A portion of this material (51.6 mg, 0.17 mmol) was
mixed with 2,6-dibromophenyl acridine-9-carboxylate (57.2 mg,
0.13 mmol) in nitrobenzene (2 mL) and heated in a glass tube
on an oil bath at 150 ◦C for 0.5 h. Acetonitrile (0.5 mL) was
added and the crude product precipitated by addition of diethyl
ether (2 mL) followed by repeated washing with ethyl acetate (10 ¥
2 mL). Compound 1 was obtained in low yield (approximately 3%)
as a red powder (nmax/cm-1 1816, 1746, 1721 (CO); dH (400 MHz,
acetonitrile-d3, Me4Si) 8.93 (2 H, d, J 9, 2 ¥ NCCHCH), 8.62
(2 H, d, J 8, 2 ¥ CCCHCH) 8.40 (2 H,m, 2 ¥ NCCHCH), 8.01
(2 H, m, CCCHCH), 7.73 (2 H, d, J 8, C(Br)CHCHCHC(Br)),
7.20 (1 H, t, J 8, C(Br)CHCHCHC(Br)), 5.33 (2 H, t, J 8,
NCH2CH2CH2CO), 3.02 (2 H, t, J 7, NCH2CH2CH2CO), 2.69
(4 H, s, COCH2CH2CO), 2.45 (2 H, m, NCH2CH2CH2CO); m/z
(ES+) 638.9769 (M+ C28H2179Br2N2O6) requires 638.9761.
Synthesis of 9-(2,6-dinitrophenoxycarbonyl)-10-(10-succinimi-
dyloxycarbonyldecyl)acridinium trifluoromethanesulfonate (Com-
pound 2). An anhydrous mixture of 11-bromoundecanoic acid
(2.192 g, 8.27 mmol) and sodium iodide (3.04 g, 20.3 mmol) was
stirred in acetone (20 mL) for 20 h. The mixture was then poured
into water (20 mL) and extracted with diethyl ether (4 ¥ 25 mL).
The combined ether extracts were washed with saturated sodium
thiosulfate solution (20 mL), dried over anhydrous magnesium
sulfate, filtered and evaporated to yield a pale pink solid which
was purified on a silica column eluted with hexane–ether–formic
acid (100 : 100 : 0.1 v/v). Fractions at Rf = 0.49 were combined
and evaporated to yield 11-iodoundecanoic acid as a white solid
Experimental
◦
◦
(2.23 g, 83% yield, mp 63–64 C (lit. 64–65 C); nmax/cm-1 1693
(CO); dH (400 MHz, CDCl3, Me4Si) 3.12 (2 H, t, J 7, ICH2CH2),
2.23 (2 H, t, J 7, CH2CH2CO), 1.75 (2 H, quintet, J 7, ICH2CH2),
1.56 (2 H, quintet, J 7, CH2CH2CH2CO), 1.28–1.18 (12 H, m,
ICH2CH2 (CH2)6CH2CH2CO); dC (400 MHz, CDCl3, Me4Si)
7.5, 25.0, 28.9, 29.4, 29.7 (3 lines), 30.8, 33.9, 34.3, 179.9; m/z
All reagents were of the highest grade, purchased from estab-
lished commercial sources, and were dried where required using
recommended methods. Oligonucleotides designed to our own
specifications were purchased from Eurogentec (Southampton,
UK) and were certified 95% pure.
This journal is
The Royal Society of Chemistry 2009
Org. Biomol. Chem., 2009, 7, 386–394 | 391
©

View Article Online
(EI) 313 ([M + H]+, 3%). A portion of this material (556 mg,
1.78 mmol) was mixed with N-hydroxysuccinimide (440 mg,
3.83 mmol) and dicyclohexylcarbodiimide (770 mg, 3.76 mmo◦ l)
in tetrahydrofuran (20 mL) under anhydrous conditions at 0 C
for 3 h then at room temperature overnight. Following filtra-
tion of the mixture, the filtrate was concentrated on a rotary
evaporator and chromatographed on a silica column eluted with
toluene–ethyl acetate (4 : 1 v/v). Fractions at Rf = 0.59 were
combined and evaporated to yield a white solid which was
recrystallized from hexane + ethyl acetate to yield succinimidyl
11-iodoundecanoate (387 mg, 53% yield, mp 84 ◦C; nmax/cm-1
1725 (CO); dH (400 MHz, CDCl3, Me4Si) 3.16 (2 H, t, J 7,
ICH2CH2), 2.81 (4 H, s, COCH2CH2CO), 2.57 (2 H, t, J 7,
CH2CH2COO), 1.79 (2 H, quintet, J 7, ICH2CH2), 1.71 (2 H,
quintet, J 7, CH2CH2CH2COO), 1.41–1.22 (12 H, m, ICH2CH2
(CH2)6CH2CH2COO); dC (400 MHz, CDCl3, Me4Si) 7.7, 25.0,
26.0, 28.9, 29.1, 29.4, 29.6, 29.7, 30.9, 31.3, 34.0, 169.1, 169.6; m/z
0.17 mmol) and 1,2-dichloroethane (1.6 mL) was refluxed under
nitrogen for 21 h. The yellow residue formed upon evaporation of
the solvent was extracted with dichloromethane (1 mL) and the ex-
tract chromatographed on silica gel eluted with dichloromethane–
acetonitrile (3 : 1 v/v). The fractions having Rf = 0.19 were pooled
and evaporated to give compound 3 as a yellow solid in low yield
(15.9 mg, 10.7%, mp 164–165 ◦C; nmax/cm-1 1815, 1779, 1733 (CO);
dH (400 MHz, CDCl3, Me4Si) 8.78 (2 H, d, J 9, 2 ¥ NCCHCH),
8.71 (2 H, d, J 9, 2 ¥ CCCHCH), 8.49 (2 H, dd, J 9, 7, 2 ¥
NCCHCHCH), 8.06 (2 H, d, J 8, 2 ¥ CF3CCHCH), 7.98 (2 H,
dd, J 9, 7, 2 ¥ CCCHCHCH), 7.12 (1 H, t, J 8, CF3CCHCHCH),
5.60 (2 H, t, J 8, NCH2CH2), 2.76 (4 H, s, COCH2CH2CO), 2.51
(2 H, t, J 7, CH2CH2COO), 2.29–2.18 (2 H, m), 1.84–1.69 (4 H, m)
and 1.51–1.26 (10 H, m) (NCH2(CH2)8CH2COO). dC (400 MHz,
CDCl3, Me4Si) 24.9, 26.0, 26.9, 29.0, 29.3, 29.4, 29.5, 29.6, 30.1,
31.3, 53.3, 119.8, 124.3, 125.7, 126.0, 128.5, 129.0, 130.0, 132.4,
140.6, 141.7, 144.9, 161.8, 169.1, 169.7; dF (400 MHz, CDCl3)
-78.4 (CF3SO3 ), -59.8 (CF3); m/z (ES+) 717.2390 (M+ – CF3SO3 )
+
-
-
(CI) 427 (M + NH4 , 15%), 301 (15), 225 (16), 149 (30) 132 (100).
This material (188 mg, 0.460 mmol) was mixed with silver triflu-
oromethanesulfonate (214 mg, 0.836 mmol) in benzene (1.5 mL)
under anhydrous conditions for 18 h at room temperature and the
mixture chromatographed on a short (2 g) silica column eluted
with dichloromethane. Fractions at Rf = 0.48 were combined
and evaporated to yield a white gum consisting of succinimidyl
11-(trifluoromethanesulfonyloxy)undecanoate (133 mg, 67%);
(100%) C37H35F6N2O6 requires 717.2394), 519 (17), 505 (8), 491 (9).
Synthesis of 2-(4-dimethylaminophenylazo)-N-[2-(2-iodoacetyl-
amino)ethyl]benzamide (Compound 6). 2-[4-(Dimethylamino)-
phenylazo]benzoic acid (1.251 g, 4.65 mmol) in THF (30 mL)
was cooled to 0 ◦C and dicyclohexylcarbodiimide (1.103 g,
5.3 mmol) was added, followed by N-hydroxysuccinimide (0.568 g,
4.94 mmol). The mixture was stirred at 0 ◦C for 2 h, thence
overnight at room temperature. The mixture was filtered and the
filtrate evaporated to dryness to give a red solid which was recrys-
tallised from acetone–diethyl ether and dried under vacuum to give
succinimidyl 2-[4-(dimethylamino)phenylazo]benzoate (0.849 g,
50%, mp 124–126 ◦C; FAB-MS m/z 361 [(M + H)+, 23%], 252 [17],
133 [100]). A portion of this (74 mg, 0.20 mmol) in dioxane (2 mL)
was mixed with t-butyl N-(2-aminoethyl)carbamate (99 mg,
0.63 mmol) in methanol (1 mL). The mixture was shaken at room
temperature in the dark for 4 h. Tlc (toluene–EtOAc 1 : 1) showed
that all the starting material (Rf = 0.85, violet) was converted
into a new component (Rf = 0.60, orange). Combined fractions
from chromatography on a silica gel column eluted with toluene–
EtOAc 1 : 1 yielded N-[2-(t-butoxycarbonylamino)ethyl]-2-[4-
(dimethylamino)phenylazo]benzamide as an orange solid (75 mg,
n
max/cm-1 1819, 1787, 1725 (CO); dH (400 MHz, CDCl3, Me4Si)
4.44 (2 H, t, J 7, SOCH2CH2), 2.74 (4 H, s, COCH2CH2CO),
2.51 (2 H, t, J 7, CH2CH2COO), 1.73 (2 H, quintet, J 7,
SOCH2CH2), 1.65 (2 H, quintet, J 7, CH2CH2CH2COO), 1.37–
1.17 (12 H, m, SOCH2CH2(CH2)6CH2CH2COO); dC (400 MHz,
CDCl3, Me4Si) 24.9, 25.4, 26.0, 29.1, 29.2, 29.3, 29.4, 29.5, 29.6,
31.3, 78.1, 119.0, 169.0, 169.6; dF (400 MHz, CDCl3) -74.8. A
portion of the above product (57 mg, 0.13 mmol) was mixed
with 2,6-dinitrophenyl acridine-9-carboxylate (53 mg, 0.13 mmol)
in 1,2-dichloroethane (2 mL) and refluxed under nitrogen for
20 h. The solvent was evaporated and the residue dissolved in
dichloromethane and chromatographed on a small silica column
(0.5 g) eluted with dichloromethane followed by dichloromethane–
acetonitrile (3 : 1 v/v). Fractions having Rf = 0.17 were pooled
and evaporated to yield compound 2 in low yield (15.8 mg, 14%)
as a yellow oil. nmax/cm-1 1812, 1778, 1735 (CO); dH (400 MHz,
CDCl3, Me4Si) 8.71–8.60 (4 H, m, 2 ¥ NCCHCH, 2 ¥ CCCHCH),
8.41 (2 H, dd, J 8, 7, 2 ¥ NCCHCHCH), 8.34 (2 H, d, J 8, 2 ¥
NO2CCHCH), 7.92 (2 H, dd, J 9, 7, 2 ¥ CCCHCHCH), 7.78
(1 H, t, J 8, NO2CCHCHCH), 5.48 (2 H, t, J 8, NCH2CH2),
2.69 (4 H, s, COCH2CH2CO), 2.45 (2 H, t, J 7, CH2CH2COO),
2.08 (2 H, m), 1.68–1.53 (4 H, m) and 1.35–1.10 (10 H, m)
(NCH2(CH2)8CH2COO). dC (400 MHz, CDCl3, Me4Si) 24.9,
26.0, 26.9, 29.0, 29.2, 29.4, 29.5, 29.6, 30.1, 31.3, 53.1, 119.6,
124.1, 128.9, 129.7, 130.2, 131.1, 136.6, 140.7, 141.6, 143.9, 144.8,
◦
91%, mp 131–134 C; FAB-MS m/z 434.2 [(M + Na)+, (6%),
412 [(M + H)+, 50%], 252 (82), 133 (100). A portion of this
material (70 mg, 0.17 mmol) in dioxane (2 mL) was treated
with concentrated hydrochloric acid (0.6 ml). The orange solution
turned to violet. The mixture was stirred at room temperature for
15 min and then evaporated to dryness. The residue was washed
with dioxane (2 ¥ 3 mL) and diethyl ether (3 mL) and was then
dissolved in MeOH (4 mL). Aqueous saturated sodium bicar-
bonate solution (2 mL) was added followed by dichloromethane
(20 mL) and then water (20 mL). The organic layer was isolated,
dried over anhydrous magnesium sulfate and evaporated to give
N-(2-aminoethyl)-2-[4-(dimethylamino)phenylazo]benzamide as
a red solid (42 mg, 79%), mp 119–124 ◦C; FAB-MS m/z
312 [(M + H)+, 13%), 252 (31), 133 (100). To this material
(60 mg, 0.19 mmol) in dichloromethane (5 mL) was added
triethylamine (20 mg, 0.20 mmol). The mixture was cooled to
-
161.1, 169.1, 169.7; m/z (ES+) 671.2352 (M+ – CF3SO3 ) (100%)
C35H35N4O10 requires 671.2348), 519 (15), 505 (23), 491 (42), 361
(77), 246 (44).
Synthesis of 9-(2,6-bis(trifluoromethyl)phenoxycarbonyl)-10-
(10-succinimidyloxycarbonyldecyl)acridinium trifluoromethanesul-
fonate (Compound 3).
A
mixture of succinimidyl 11-
0
◦C and iodoacetyl chloride (40 mg, 0.20 mmol) was added.
(trifluoromethanesulfonyloxy)undecanoate (75 mg, 0.17 mmol),
2,6-bis(trifluoromethyl)phenylacridinium-9-carboxylate (75 mg,
After 5 min at 0 ◦C, tlc (CH3Cl–MeOH, 4 : 1) showed that most
of the starting material (Rf = 0.4) was converted into a new
392 | Org. Biomol. Chem., 2009, 7, 386–394
This journal is
The Royal Society of Chemistry 2009
©

View Article Online
component (Rf = 0.6). Stirring at 0 ◦C was maintained for
another 5 min and the resulting mixture was then evaporated to
dryness. The residue was redissolved in dichloromethane (3 mL)
and loaded onto a silica gel column, which was then eluted
with CH3Cl–MeOH, 7 : 1. The desired fractions were combined
and evaporated to yield 2-(4-dimethylaminophenylazo) N-[2-(2-
acetate buffer (0.1 M, pH 7.0)/acetonitrile. Absorbance was mon-
itored at 260 nm and 0.5 mL fractions were collected into aqueous
lithium lauryl sulfate (10 mL, 10%, w/v). Chemiluminescence
(1 mL/fraction) was measured at pH 14 and the peak fractions
were pooled. The labelled oligonucleotide was re-precipitated
by sodium acetate/chilled ethanol as above, then taken up in
probe storage reagent and stored at -70 ◦C. AE conjugated
oligonucleotide yields and concentrations were determined by con-
ventional absorption spectroscopy using a GenQuant spectropho-
tometer (Pharmacia) as recommended by the manufacturer.
◦
iodoacetylamino)ethyl]benzamide (25 mg, 27%, mp 140–142 C;
dH (400 MHz, CDCl3, Me4Si) 9.52 (exch.), 8.35 (1 H, dd, J 8, 2),
7.80–7.73 (3 H, m), 7.54 (1 H, td, J 8, 2), 7.48 (1 H, td, J 8, 2),
6.79 (2.h, d, J 9), 3.70 (2 H, m), 3.61 (2 H, s), 3.53 (2 H, m), 1.70
(exch.); FAB-MS m/z 480 [(M + H)+, 75%)], 252 (52), 107 (100).
Conjugation of quencher 6 to AE-labelled oligonucleotides
Chemiluminescence kinetics and stability measurements
Oligonucleotides for dual-labelling were synthesised by established
methods and obtained commercially with an amine linker to
facilitate AE labelling and a trityl-protected thiol to facilitate
coupling of the quencher 6. The desired quantity of AE-labelled
oligonucleotide prepared as described above (typically 10 nmol
in 20 ul probe storage reagent) was diluted with an aqueous
solution of triethylammonium acetate (200 mL, 0.1 M, pH 7.0).
The protective trityl group was removed by the addition of
aqueous silver nitrate solution (6.7 mL, 0.01 M) and the mixture
incubated for 30 minutes at room temperature. An aqueous
solution of dithiothreitol (10 mL, 0.01 M) was added and the
mixture incubated for 5 minutes at room temperature with shaking
Compounds 1, 2 and 3 were dissolved in acetonitrile (50 mL)
and diluted in 1 mM hydrochloric acid to yield a concentration
between 1 and 10 ng/mL. The chemiluminescence of 10 mL
samples of these solutions was measured in a luminometer
(“Lumino”, Stratec, Pforzheim, Germany) equipped with in situ
addition of initiating reagents. Chemiluminescence was activated
by sequential dispensing of 80 mM H2O2 (200 mL) and 0.2 M
glycine pH10 (200 mL) with signal measurement for 1 s (following
the second reagent addition) at a 50 ms sample rate. In order
to obtain an approximate, relative measure of stability of the
compounds, the above acidic aqueous solutions of compounds 1,
2 and 3 were held at 4 ◦C and further 10 uL samples removed and
measured at intervals of 4 and 6 days. AEs 1, 4 and 5 (1–10 ng) were
compared at pH 9 and pH 14. Here the second reagent addition
in the luminometer was either respectively 0.2 M Tris pH 9.0 or
0.5 M NaOH.
¯
following which it was centrifuged at 10000 rpm for 2 minutes.
The supernatant was carefully removed and transferred to a fresh
test tube to which an aqueous solution of sodium bicarbonate
(195 mL, 0.1 M, pH 9.0) was then added. Quencher 6 (1 mg) was
dissolved in DMSO (30 mL) and 6.6 mL of this was added to
the AE-labelled oligonucletide solution with mixing. Following
incubation for 90 minutes at room temperature, the mixture
Comparison of the absorption spectra of quenchers
¯
was clarified by centrifugation at 10000 rpm for 2 minutes. The
Initial experiments investigated the overlap between the known
fluorescence emission spectrum of N-methylacridone (which ap-
proximates to the chemiluminescence emission spectra of the AEs
being studied here) and the absorption spectra of polyA10 dabcyl
(17 mM) and polyA10 methyl red (21 mM). Absorption spectra were
recorded in Tris buffer (0.2 M, pH 9) over the range from 250 nm
to 600 nm using a Cecil 3000 spectrophotometer.
supernatant was removed and the oligonucleotide recovered by
sodium acetate/ethanol precipitation as described above. The
pellet was dissolved in probe storage reagent and stored at -70 ◦C.
Intramolecular dual-labelled probe experiments
For hybridisation experiments using the hairpin probe, AE
quencher labelled sequence 1 (10 fmol) in hybridisation buffer
[50 mL, aqueous lithium succinate (0.1 M, pH 5.1) containing
lithium lauryl sulfate (8.5% w/v), LiOH (125 mM), EDTA
(1.5 mM) and EGTA (1.5 mM)] was mixed with various known
quantities of oligonucleotide target (sequence 2) in water (50 mL)
and incubated at 60 ◦C for 15 min followed by room temperature
incubation for 15 min. Chemiluminescence of an aliquot of the
hybridisation assay solution (10 mL) was then measured in the
luminometer. For pH 14 measurement the luminometer reagents
were (1) aqueous H2O2 (0.2 mL, 80 mM) followed immediately by
(2) aqueous NaOH (0.2 mL, 0.5 M). Measurement duration was 1
to 2 seconds as appropriate. For pH 9, a single addition of aqueous
Tris/HCl (0.2 mL, 0.2 M) containing H2O2 (80 mM) was used,
with a measurement duration of 1 to 5 seconds as appropriate.
Conjugation of AE to oligonucleotides
Oligonucleotides for labelling with AEs were synthesised with a
terminal 5¢ or 3¢ NH2. The oligonucleotide (10 nmol) in labelling
buffer (10 mL, HEPES 0.125 M, pH 8.0) was added to a solution
of the AE in DMSO (10 mL). The contents were gently mixed
and incubated for 20 min at 37 ◦C, following which excess N-
hydroxysuccinimide ester on the AE was deactivated by addition
of lysine (5 mL, 0.125 M in labelling buffer). The oligonucleotide
was precipitated by adding sequentially aqueous sodium acetate
(30 mL, 3 M), water (245 mL) and aqueous glycogen (5 mL,
40 mg/mL) and chilled (-20 ◦C) ethanol (640 mL), then was
incubated on dry ice for 10 min prior to centrifugation at
15000 rpm in a 0 ◦C centrifuge. The supernatant was carefully
¯
aspirated and the pellet taken up in probe storage reagent
(20 mL, sodium acetate, 0.1 M, containing lithium lauryl sulfate,
0.1%, w/v).
Intermolecular dual-labelled probe experiments
The labelled oligonucleotide was further purified by conven-
tional gradient RPLC (C18), using aqueous triethylammonium
Initial experiments were designed to investigate the optimal
conditions for maximal annealing and to demonstrate that the
This journal is
The Royal Society of Chemistry 2009
Org. Biomol. Chem., 2009, 7, 386–394 | 393
©

View Article Online
intermolecular quenching worked in practice. These experiments
used dilutions of the two labelled single stranded oligonucleotides
(AE labelled sequence 3 and quencher labelled sequence 4) in
aqueous HEPES buffer (0.1 M, pH 7.4) containing sodium
chloride (10 mM) and glycerol (10% v/v). The AE labelled
sequence 3 was incubated with excess quencher labelled sequence
4. It was determined that the optimal annealing protocol for
helicase substrate generation consisted of incubation, in annealing
reagent [aqueous sodium succinate (20 mM), sodium chloride
(50 mM), magnesium chloride (1 mM), Triton-X100 (0.1% w/v),
pH 5.1)], of quencher labelled sequence 4 (3 mM) with the AE
labelled sequence 3 (2 mM) for 18 hours at room temperature,
which yielded 200 pmol of substrate. Aliquots of the substrate
solution were stored at -80 ◦C. The enzyme assay buffer was
aqueous HEPES (0.1 M, pH 6.8), containing sodium chloride
(10 mM), magnesium chloride (5 mM), bovine serum albumin
(500 mg/mL), adenosine triphosphate (ATP, 1 mM). A mixture
of various concentrations of enzyme (0, 40 and 200 nM) and
substrate (2.5 nM) diluted in enzyme buffer was incubated at
room temperature and the reaction stopped by the addition
of one volume of stop reagent (enzyme buffer plus Triton
X-100, 1% w/v). Chemiluminescence was measured at pH 9 in
a 10 mL sample aliquot of the incubation solutions using the
method described above for the intramolecular labelled probe
experiments.
References
1 L. J. Arnold, P. W. Hammond, W. Wiese and N. C. Nelson, Clin.
Chem., 1989, 35, 1588–1594; N. C. Nelson, M. A. Reynolds and
L. J. Arnold, in Non-Isotopic Probing, Blotting, and Sequencing, ed.
L. J. Kricka, Academic Press, San Diego, 1995, pp. 391–427; I. Weeks,
M. Sturgess, R. C. Brown and J. S. Woodhead, Methods Enzymol., 1986,
133, 366–387; N. C. Nelson, A. B. Cheikh, E. Matsuda and M. M.
Becker, Biochemistry, 1996, 35, 8429–8438; I. Weeks, Chemiluminescence
Immunoassay, Elsevier, Amsterdam, 1992.
2 J. Comley, Drug Discovery World, 2003, 4, 91–97.
3 M. C. Williams, J. R. Wenner, I. Rouzina and V. Bloomfield, Biophys. J.,
2001, 80, 874–881; D. Y. Lando, S. G. Haroutiumian, S. M. Ku’lba, E. B.
Dalian, P. Orioli, S. Mangani and A. A. Akhrem, J. Biomol. Struct. Dyn.,
1994, 12, 355–366.
4 S. Tyagi and F. R. Kramer, Nat. Biotechnol., 1996, 14, 303–308.
5 (a) R. C. Brown, I. Weeks, M. Fisher, S. Harbron, C. J. Taylorson and J. S.
Woodhead, Anal. Biochem., 1998, 259, 142–151; (b) J. S. Woodhead, I.
Weeks and S. Batmanghelich, US Patent 5,656,207, 1997; N. C. Nelson,
J. S. Woodhead, I. Weeks and A. B. Cheikh, US Patent 5,827,656, 1998;
(c) D. D. Perrin, B. Dempsey and E. P. Sergeant, pKa prediction for
organic acids and bases, Chapman and Hall, London, 1981; (d) S. J.
Law, T. Miller, U. Piran, C. Klukas, S. Chang and J. Unger, J. Biolum.
Chemilum., 1989, 4, 88–98.
6 S. E. Marras, F. R. Kramer and S. Tyagi, Nucleic Acids Res., 2002, 30,
e122.
7 L. E. Bird, J. A. Brannigan and D. B. Wigley, Nucleic Acids Res., 1998,
26, 2686–2693.
8 K. Smith, Z. Li, J. J. Yang, I. Weeks and J. S. Woodhead, J. Photochem.
Photobiol. A: Chem., 2000, 132, 181–191; K. Smith, J. J. Yang, Z.
Li, I. Weeks and J. S. Woodhead, J. Photochem. Photobiol. A: Chem.,
submitted.
394 | Org. Biomol. Chem., 2009, 7, 386–394
This journal is
The Royal Society of Chemistry 2009
©