U.S. patent application number 09/946829 was filed with the patent office on 2002-05-23 for detection of binding factors with fluorescence polarization.
Invention is credited to Le, Xiao-Chun (Chris).
Application Number | 20020061531 09/946829 |
Document ID | / |
Family ID | 26923875 |
Filed Date | 2002-05-23 |
United States Patent
Application |
20020061531 |
Kind Code |
A1 |
Le, Xiao-Chun (Chris) |
May 23, 2002 |
Detection of binding factors with fluorescence polarization
Abstract
This invention relates to a simple and quick method for the
detection, identification and/or quantitation of binding factors
using fluorescence techniques. A fluorescent probe is incubated
with a factor or group of factors, and the presence of a factor
capable of binding the probe can be detected by fluorescence
polarization. When coupled with a separation step, this invention
allows on-line monitoring of binding complex formation.
Inventors: |
Le, Xiao-Chun (Chris);
(Edmonton, CA) |
Correspondence
Address: |
Gerald F. Swiss
BURNS, DOANE, SWECKER & MATHIS, L.L.P.
P.O. Box 1404
Alexandria
VA
22313-1404
US
|
Family ID: |
26923875 |
Appl. No.: |
09/946829 |
Filed: |
September 4, 2001 |
Related U.S. Patent Documents
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Application
Number |
Filing Date |
Patent Number |
|
|
60230060 |
Sep 1, 2000 |
|
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Current U.S.
Class: |
435/6.14 ;
435/7.1; 436/516 |
Current CPC
Class: |
G01N 33/536
20130101 |
Class at
Publication: |
435/6 ; 435/7.1;
436/516 |
International
Class: |
C12Q 001/68; G01N
033/53; G01N 033/561 |
Claims
We claim:
1. A method for detecting a binding factor for a probe, comprising:
(a) labeling the probe with a fluorophore; (b) incubating the
labeled probe with a factor or a group of factors which may bind
the labeled probe to form a binding complex; (c) separating the
binding complex and the free probe into different fractions; and
(d) subjecting each fraction from step (c) to fluorescence
polarization measurement under conditions wherein the binding
complex produces a fluorescence pattern different from that of the
free probe, thereby allowing detection of the binding complex.
2. The method of claim 1 wherein the free probe and the complex are
separated by using capillary electrophoresis.
3. The method of claim 1 wherein the group of factors comprises a
chemical compound library.
4. The method of claim 4 wherein the chemical compound library is a
combinatorial library.
5. The method of claim 1 wherein the group of factors comprises a
mixture of natural products.
6. The method of claim 6 wherein the mixture of natural products
comprises a cell lysate.
7. The method of claim 1 wherein the group of factors comprises
nucleic acid.
8. The method of claim 7 wherein the nucleic acid is genomic
DNA.
9. The method of claim 8 wherein the probe is capable of binding to
modified DNA.
10. The method of claim 10 wherein the modified DNA is a DNA
adduct.
11. The method of claim 1 wherein the probe is selected from the
group consisting of protein and nucleic acid.
12. The method of claim 1 wherein the probe has a molecular weight
of less than about 10,000 daltons.
13. The method of claim 1 wherein the probe has a molecular weight
of less than about 5,000 daltons.
14. The method of claim 1 wherein the probe has a molecular weight
of less than about 3,000 daltons.
15. The method of claim 1 further comprising the step of
determining binding affinity and/or stoichiometry between the probe
and the binding factor.
16. The method of claim 1 wherein the fluorophore is
fluorescein.
17. A method for detecting a nucleic acid damage in a nucleic acid
sample, comprising: (a) incubating the sample with (i) a
polypeptide which is capable of binding the damaged nucleic acid;
and (ii) a fluorophore-labeled probe which is capable of forming a
complex with the polypeptide to compete with the damaged nucleic
acid for the polypeptide; and (b) analyzing the incubation mixture
under conditions wherein the complex formed between the probe and
the polypeptide produces a fluorescence pattern different from that
of a free probe.
18. The method of claim 17 wherein the polypeptide is an antibody
which is capable of binding damaged DNA.
19. The method of claim 17 wherein the DNA damage is a covalent
modification.
20. The method of claim 17 wherein the DNA damage is a benzopyrene
addition.
21. The method of claim 17 wherein the DNA sample is genomic
DNA.
22. A method for detecting a fluorophore labeled probe, comprising:
(a) incubating a probe with a fluorophore under conditions which
allow labeling of the probe by the fluorophore; and (b) subjecting
the incubation mixture to fluorescence polarization under
conditions wherein the fluorophore labeled probe produces a
fluorescence pattern which is different from that of a free probe
which is not labeled by the fluorophore.
23. The method of claim 22 further comprising the step of
fractionating the incubation mixture.
Description
RELATED APPLICATIONS
[0001] This application claims the benefit of U.S. application Ser.
No. 60/230,060, filed Sep. 1, 2000, which is hereby incorporated by
reference in its entirety.
FIELD OF THE INVENTION
[0002] This invention relates to methods of detection,
identification and quantitation of binding factors with
fluorescence techniques.
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[0067] All of the above publications, patents and patent
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to the same extent as if the disclosure of each individual
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individually indicated to be incorporated by reference in its
entirety.
BACKGROUND OF THE INVENTION
[0068] Affinity binding complex formation is an essential step in
biological or pharmaceutical phenomena. For example, the binding of
proteins to DNA underlines many cellular activities including the
control of gene expression, site-specific recombination,
replication and repair of DNA damage. Enzyme-substrate interactions
involve the recognition and binding of substrate by the enzyme as
the first step. Hormones, neurotransmitters, lymphokines and other
effector molecules bind to their receptors to initiate the cellular
process which ultimately lead to achievement of their
functions.
[0069] Consequently, affinity binding complexes are also important
tools in biological or pharmaceutical research. For example, drug
discovery often involves identification of binding factors of a
particular target which mediates a disease. A variety of methods
have been employed to detect affinity binding complex formation in
order to identify the binding factors. For example, the gel
electrophoresis mobility shift assay (EMSA) is the most commonly
used method in the study of protein-DNA interactions. This method
is based on the observation that binding of a protein to DNA
fragments leads to a reduction in the electrophoretic mobility of
the DNA fragment in non-denaturing polyacrylamide or agarose gels.
While used extensively, EMSA requires relatively large amounts of
sample and lengthy analysis time. Moreover, the assay is not
suitable when dissociation of protein-DNA complex occurs during gel
electrophoresis.
[0070] As another example, capillary electrophoresis (CE) combined
with affinity recognition has gained a tremendous growth in recent
years, with increasing biochemical, clinical, and pharmaceutical
applications. A key element of the technique is the use of a
molecular recognition agent, typically a protein that binds to a
target molecule with high specificity and affinity. The complex
formation can occur either before or during the electrophoretic
separation, depending on the stability of the resultant complex. In
applications such as CE-based immunoassays, however, tight binding
of the analyte to the protein is essential to achieve a high degree
of sensitivity and reproducibility. Ideally, the affinity complex
thus formed should remain intact throughout the electrophoretic
separation.
[0071] In current practice of affinity CE, the formation and
stability of the complex are usually established by titration
experiments, in which a series of solutions containing the
substrate and its binding protein in various ratios are analyzed.
The emergence of a new peak upon addition of the binding protein to
the substrate is taken as the evidence for complex formation and
the relative intensities corresponding to the complex and the free
substrate are used for quantitation. The titration experiments have
proved to be very useful in studies of binding interactions;
however, they are time-consuming and unable to provide unequivocal
identification of the complex when the complex is not well
separated from the unbound molecules. Therefore, there remains a
need for a simple and sensitive method to detect affinity complex
formation.
SUMMARY OF THE INVENTION
[0072] This invention is directed to a simple method based on
laser-induced fluorescence polarization (LIFP) detection of an
affinity complex of a fluorescent probe and its binding factor. The
affinity complexes are readily distinguished from the unbound
molecules on the basis of their fluorescence polarization, which is
sensitive to changes in the rotational diffusion characteristics
arising from molecular association or dissociation. The relative
increase in fluorescence polarization upon complex formation varied
with the molecular size of the binding pairs. A small molecule
rotates fast in solution and exhibits a low value of polarization
whereas a large molecule exhibits a higher polarization because of
its slower motion under the same conditions. Thus, changes in
fluorescence polarization can reflect the association or
dissociation status between molecules of interest. When combined
with capillary electrophoresis or other suitable separation
procedures, this method allows for on-line monitoring of affinity
complex formation.
[0073] Accordingly, an aspect of this invention is directed to a
method for detecting a binding factor for a probe, comprising:
[0074] (a) labeling the probe with a fluorophore;
[0075] (b) incubating the labeled probe with a factor or a group of
factors which may bind the labeled probe to form a binding
complex;
[0076] (c) separating the binding complex and the free probe into
different fractions; and
[0077] (d) subjecting each fraction from step (c) to fluorescence
polarization measurement under conditions wherein the binding
complex produces a fluorescence pattern different from that of the
free probe, thereby allowing detection of the binding complex.
[0078] The separation step may be performed simultaneously with, or
prior to, the fluorescence polarization detection step. The free
probe and the bound probe may be separated by any method which is
compatible with fluorescence polarization. Preferably, the
separation method is capable of being performed in liquid phase.
The separation method is more preferably liquid chromatography or
electrokinetic chromatography, and most preferable capillary
electrophoresis or capillary gel electrophoresis.
[0079] In another aspect of the present invention, this method can
be applied to screen a chemical compound library, such as
combinatorial library. Thus, in order to identify a compound which
is capable of binding to a molecule of interest, the molecule of
interest is used as a probe and labeled with a fluorophore. The
labeled probe is then incubated with a compound library and the
whole mixture can be analyzed by fluorescence polarization.
Alternatively, the mixture is separated with a suitable method. The
fractions are monitored on-line with fluorescence polarization
which can distinguish the free probe from the bound probe, thereby
identify whether there is a binding compound in the particular
fraction.
[0080] Similarly, this method can also be applied to screen a
mixture of natural products, such as a cell lysate or a homogenate
of tissue. The natural products may come from any source, including
animals, plants and microorganisms.
[0081] In another aspect of this invention, the method can be used
to determine if a particular sequence or modification exists in a
DNA. For example, exposure to certain carcinogens induce alkylation
or other kinds of modification of DNA, which may result in
mutations and abnormal gene expression to cause cancer formation.
By using a specific probe which binds to a certain DNA sequence or
modification, one can detect if this sequence or modification
exists in the genomic DNA for diagnosis purposes.
[0082] The probe can be a protein (including peptides),
particularly antibodies, enzymes and cell surface receptors. The
probe can also be a nucleic acid, carbohydrate or carbohydrate
derivatives, small organic or inorganic compounds, or any molecule
which can be labeled with a fluorophore or is naturally
fluorescent. The probe is preferably less than about 15,000 daltons
in molecular weight, more preferably less than about 10,000
daltons, yet more preferably less than about 5,000 daltons, still
more preferably less than about 3,000 daltons, and most preferably
less than about 1,500 daltons.
[0083] In yet another aspect, this invention provides a method to
determine the binding affinity or stoichiometry of an affinity
complex.
[0084] In addition, this invention can also be used to monitor the
formation of fluorescently labeled molecules, which may be used as
probes in the present method but are not limited to such use.
Labeling of molecules with a fluorophore is often required for high
sensitivity quantitation and detection of the molecules. However,
it is difficult to identify whether the desired molecule is labeled
or what the labeling efficiency is. This invention provides a fast
method to monitor the labeling process since the labeled molecule
can be readily differentiated from the free fluorephore because of
their difference in fluorescence polarization.
BRIEF DESCRIPTION OF THE DRAWING
[0085] FIG. 1
[0086] Electropherograms showing vertically and horizontally
polarized fluorescence of the complex formed between
fluorescein-labeled vancomycin and its antibody. A 35 cm long, 20
mm i.d., fused silica capillary was used for separation with a 25
mM disodium tetraborate (pH 9.1) as the running buffer. The
separation voltage was 25 kV. Fluorescence detection was
post-capillary with vertically polarized excitation at 488 nm. Iv
and Ih correspond to vertically and horizontally polarized
fluorescence intensity, respectively, measured at 515 nm.
DETAILED DESCRIPTION OF THE INVENTION
[0087] This invention relates to a simple and quick method for the
detection, identification and/or quantitation of binding factors
using fluorescence techniques. A fluorescent probe is incubated
with a factor or group of factors, and the presence of a factor
capable of binding the probe can be detected by fluorescence
polarization. When coupled with a separation step, this invention
allows on-line monitoring of binding complex formation.
[0088] Prior to describing the invention in further detail, the
terms used in this application are defined as follows unless
otherwise indicated.
[0089] Definitions
[0090] As used herein, a "probe" can be any molecule or substance
for which it is desired to find a binding factor. Examples of a
probe include proteins, peptides, nucleic acids, carbohydrates or
carbohydrates derivative, and small organic or inorganic
compounds.
[0091] As used herein, a "fluorophore" is any fluorescent
substance, for example, fluorescein.
[0092] As used herein, "a factor or a group of factors" means a
substance or a mixture of substance of any nature. A factor may be
a protein, a peptide, a nucleic acid, a carbohydrate or
carbohydrate derivative or any other organic or inorganic compound.
A group of factors may be a mixture of factors of the same nature,
or a mixture of factors of different natures. For example, a cell
lysate or homogenate is a group of factors which contains proteins,
carbohydrates, lipids, nucleic acids, and any other substance
contained in a cell or cells.
[0093] As used herein, a nucleic acid "modification" refers to any
change in the structure of the nucleic acid sequence. Changes in
the structure of a nucleic acid sequence include changes in the
covalent and non-covalent bonds in the nucleic acid sequence.
Illustrative of these changes are mutations, mismatches, strand
breaks, as well as covalent and non-covalent interactions between a
nucleic acid sequence, which contains unmodified and/or modified
nucleic acids, and other molecules. Illustrative of a covalent
interaction between a nucleic acid sequence and another molecule
are changes to a nucleotide base (e.g., formation of thymine
glycol) and covalent cross-links between double-stranded DNA
sequences which are introduced by ultraviolet radiation or by
cis-platinum. Yet another example of a covalent interaction between
a nucleic acid sequence and another molecule includes covalent
binding of two nucleic acid sequences to psoralen following
ultraviolet irradiation. Non-covalent interactions between a
nucleic acid sequence and another molecule include non-covalent
interactions of a nucleic acid sequence with a molecule other than
a nucleic acid sequence and other than a polypeptide sequence.
Non-covalent interactions between a nucleic acid sequence with a
molecule other than a nucleic acid sequence and other than a
polypeptide sequence are illustrated by non-covalent intercalation
of ethidium bromide or of psoralen between the two strands of a
double-stranded deoxyribnucleic acid sequence.
[0094] As used herein, the term "mutation" refers to a deletion,
insertion, or substitution. A "deletion" is defined as a change in
a nucleic acid sequence in which one or more nucleotides is absent.
An "insertion" or "addition" is that change in a nucleic acid
sequence which has resulted in the addition of one or more
nucleotides. A "substitution" results from the replacement of one
or more nucleotides by a molecule which is different molecule from
the replaced one or more nucleotides. For example, a nucleic acid
may be replaced by a different nucleic acid as exemplified by
replacement of a thymine by a cytosine, adenine, guanine, or
uridine. Alternatively, a nucleic acid may be replaced by a
modified nucleic acid as exemplified by replacement of a thymine by
thymine glycol.
[0095] The term "mismatch" refers to a non-covalent interaction
between two nucleic acids, each nucleic acid residing on a
different nucleic acid sequence, which does not follow the
base-pairing rules. For example, for the partially complementary
sequences 5'-AGT-3' and 5'-AAT-3', a G-A mismatch is present.
[0096] The term "strand break" when made in reference to a double
stranded nucleic acid sequence includes a single-strand break
and/or a double-strand break. A single-strand break refers to an
interruption in one of the two strands of the double stranded
nucleic acid sequence. This is in contrast to a double-strand break
which refers to an interruption in both strands of the double
stranded nucleic acid sequence. Strand breaks may be introduced
into a double stranded nucleic acid sequence either directly (e.g.,
by ionizing radiation) or indirectly (e.g., by enzymatic incision
at a nucleic acid base).
[0097] As used herein, a "sample" may be a biological sample or an
environmental sample. Environmental samples include material from
the environment such as soil and water. Biological samples may be
animal (e.g., human), fluid (e.g., blood, plasma and serum), solid
(e.g., stool), tissue, liquid foods (e.g., milk), and solid foods
(e.g., vegetables). A biological sample may comprise a cell, tissue
extract, body fluid, chromosomes or extrachromosomal elements
isolated from a cell, genomic DNA, RNA, cDNA and the like.
[0098] Methods
[0099] In the present invention, fluorescence polarization is used
to distinguish between a fluorescently-labeled probe and a complex
containing both the probe and a factor which binds the probe. The
complex exhibits higher polarization than the probe because a small
molecule, such as the probe, rotates freely in solution and tends
to yield no polarization. However, when the probe is bound by
another factor and the size of the complex is significantly larger
than the free probe, the complex rotates much less freely,
resulting in significantly higher polarization.
[0100] Accordingly, the molecular weight of the probe useful in the
present invention should be less than about 20,000 daltons. A
larger molecule will generate sizable polarization, making any
increase in polarization more difficult to detect. Furthermore,
since the size increase upon forming a complex to such a large
probe is relatively less significant, it is also harder to further
increase the polarization. Preferably, the probe has a molecular
weight of less than 15,000 daltons. The molecular weight is more
preferably less than about 10,000 daltons, yet more preferably less
than about 5,000 daltons, still more preferably less than about
3,000 daltons, and most preferably less than about 1,500
daltons.
[0101] The probe can be a protein, nucleic acid, carbohydrate,
lipid, or any molecule which can be labeled with a fluorophore. In
particular, peptides, oligonucleotides and oligosaccharides are
good probes due to their small sizes and easy synthesis. If an
antibody is a candidate for a probe, it is preferable to use a
fragment of the antibody, such as an Fab fragment, rather than the
entire antibody.
[0102] The present method can be practiced with or without
separation of the binding complex and the probe. Thus, the entire
incubation mixture can be subjected to fluorescence polarization
without separation, and the polarization pattern is compared to
that of the probe alone. An increase in polarization would indicate
the presence of at least one binding factor in the sample used to
bind the probe.
[0103] Alternatively, the incubation mixture can be separated
according to any method known in the art, and then subjected to
polarization. In this case, the free probe and the complex will
appear in two different fractions, and the identification of each
fraction can be determined according to the polarization
measurement (i.e, the complex has significantly higher polarization
than the free probe). It is also possible to run the free probe
alone by the same separation method as an indicator to determine
which fraction contains the free probe. Once the position of the
free probe is known, the complex can simply be detected and
quantitated by fluorescence measurement, such as laser-induced
fluorescence (LIF), without polarization.
[0104] It is preferable to separate the free probe from the complex
in the present invention. One reason is that the sensitivity with
which to detect the complex is much higher if the method contains a
separation step. When the entire incubation mixture is subjected to
polarization, the signal from the free probe is mixed with that
from the complex, and the net increase in polarization may not be
very conspicuous. This is particularly a problem if the probe is
relatively large, resulting in a substantial background
polarization. If the free probe is separated from the complex, the
background noise arising from the free probe is eliminated, and
polarization due to complex formation can be detected with higher
sensitivity.
[0105] A common problem with separating binding complexes is that
weak binding complexes tend to dissociate during separation. For
example, in the mobility shift method of detecting binding
complexes, an incubation mixture is separated by gel
electrophoresis, and the position of the complex is then located by
detecting the label contained in the probe. However, due to the pH,
temperature or electronic field to which a complex is exposed
during electrophoresis, a weak complex may dissociate during
electrophoresis. If so, the location of the probe would be
mistakenly interpreted as the location of the complex.
[0106] This problem is not a concern in the present invention.
Since the fluorescence polarization measurement is very different
between a complex and a free probe, it can be determined with
significant certainty whether a fraction contains a complex or a
free probe. Furthermore, the present invention can be used to
monitor complex formation on-line during the course of the
separation. Therefore, the presence of a complex can be detected
before it dissociates.
[0107] A particular interesting application of the present
invention is library screening. It is common to screen a large
number of chemical compound libraries for a binding factor. Since
the present invention enables one to quickly determine if a binding
factor exists in a library or not, without having to separate the
free probe and the complex, it is very useful in the initial
screening. Once it has been determined which libraries contain
binding factors, the incubation mixtures containing the libraries
of interest can be separated using an appropriate method, and the
present invention can again be used to locate the fractions having
the complex, thereby allowing identification of the binding
factors.
[0108] Another particular application of the present invention is
to detect nucleic acid damages. Nucleic acid damages, such as
mismatches, breaks or the formation of DNA adducts, often occur
after the nucleic acid is exposed to carcinogens. The present
invention can be used to detect nucleic acid damages using a factor
which is capable of binding damaged nucleic acids. Thus, a sample
suspected of having damaged nucleic acids can be incubated with a
binding factor as well as a fluorescently labeled oligonucleotide
probe harboring the specific nucleic acid damage. The binding
complex between the probe and the binding factor can be detected
using the present invention, with or without separation. If the
sample contains the damage of interest, it would compete with the
probe for the binding factor to result in a decrease in binding
complex formation between the probe and the factor.
[0109] Any factor capable of binding specifically to a damaged
nucleic acid can be used in the present invention, such as any
antibody which recognizes a specific DNA adduct; the UvrA and UvrB
proteins which bind UV dimers, polycyclic aromatic hydrocarbon
adducts, cis-platinum adducts, aflatoxin adducts, psoralen adducts,
anthramycin adducts, mitomycin C adducts, N-acetoxy-2-aminofluorene
adducts, and N-hydroxy-2-aminofluorene adducts; the DNA-dependent
protein kinase which specifically bind to DNA double strand ends;
poly(ADP-ribose) polymerase which binds to single-strand breaks and
double-strand breaks; and the MutS protein which binds to several
different mismatches (see, e.g. U.S. Pat. No. 6,132,968).
[0110] The following examples are offered to illustrate this
invention and are not to be construed in any way as limiting the
scope of the present invention.
EXAMPLES
[0111] In the examples below, the following abbreviations have the
following meanings. Abbreviations not defined have their generally
accepted meanings.
[0112] .degree.C.=degree Celsius
[0113] hr=hour
[0114] min=minute
[0115] .mu.M=micromolar
[0116] mm=millimolar
[0117] M=molar
[0118] ml=milliliter
[0119] .mu.l=microliter
[0120] mg=milligram
[0121] .mu.g=microgram
[0122] PAGE=polyacrylamide gel electrophoresis
[0123] rpm=revolutions per minute
[0124] FBS=fetal bovine serum
[0125] DTT=dithiothrietol
[0126] PBS=phosphate buffered saline
[0127] CE=capillary electrophoresis
[0128] LIFP=laser induced fluorescence polarization
[0129] LIF=laser induced fluorescence
[0130] PMT=photomultiplier tube
[0131] SSB=single-stranded DNA binding protein
[0132] FITC=Fluorescein isothiocyanate
[0133] SEA=staphylococcal enterotoxin A
[0134] FPIA=fluorescence polarization immunoassay
[0135] IAF=5-iodoacetamidofluorescein
[0136] RT=reverse transcriptase
[0137] TBE=Tris-borate-EDTA
[0138] FAM=carboxyfluorescein
[0139] AMV=avian myeloblastosis virus
[0140] MMLV=Moloney murine leukemia virus
[0141] BPDE=benzo[a]pyrene diol epoxide
[0142] TMR=tetramethylrhodamine
[0143] EOF=electroosmatic flow
[0144] LED=light-emitting diode
[0145] DMEM=Dulbecco's modified Eagle's medium
[0146] DMSO=dimethylsulfoxide
Section A: Examples 1-6
Application of the Present Invention in Complex Formation Between
Various Biological Molecules
[0147] Instrumentation. A laboratory-built capillary
electrophoresis with laser induced fluorescence polarization
(CE/LIFP) detection system was used (Ye, et al., 1998).
Electrophoresis was performed using a high voltage power supply
(Model CZE 1000R, Spellman, Plainview, N.Y.) and a fused silica
capillary (Polymicro Technologies, Phoenix, Ariz.). The detection
end of the capillary was inserted in a sheath flow cuvette (NSG
Precision Cells, Farmingdale, N.Y.). A plane polarized laser beam
from an argon ion laser (Model 2014-65ML, Uniphase, San Jose,
Calif.) was filtered through a laser line filter (488 nm, 10-nm
band width, Newport, Fountain Valley, Calif.) and was used for
excitation. The light emitted during fluorescence was collected
with a 60.times. microscope objective lens (0.7 NA, Universe
Kogaku, Oyster Bay, N.Y.), filtered with a narrow bandpass filter
(515 nm, 10-nm band width, Newport), and passed through a pinhole.
The emitted light was subsequently split with a broadband
polarizing beamsplitter cube (Melles Griot, Irvine, Calif.) into
vertically and horizontally polarized components, which were
detected with two photomultiplier tubes (PMT1 and PMT2, R1477,
Hamamatsu, Japan). The operation of the power supply and the
acquisition of data were controlled by a Power Macintosh computer
with an application software written in LabView (National
Instruments, Austin, Tex.).
[0148] There is no fundamental difference between this apparatus
and the conventional fluorescence detectors that are also capable
of anisotropy measurements. The only difference is that our cuvette
(flow cell) is much smaller (0.2.times.0.2 square) than
commercially available cells and that our detector is capable of
handling the small volumes suitable for CE separation.
[0149] CE/LIFP Analysis. Unless otherwise stated the CE/LIFP were
run as follows. A capillary of 20 mm i.d, 148 mm o.d. and 35 cm in
length was used in conjunction with 25 mM Na.sub.2B.sub.4O.sub.7
(pH 9.1) as a typical running buffer. Prior to sample analysis, the
capillary was preconditioned periodically by successive rinsing
with 0.1 M NaOH, deionized water and the running buffer to ensure
reproducibility of the separation. Samples were electrokinetically
injected into the capillary by applying an electric field of 143
V/cm for 5 s. Separation was carried out under an electric field of
714 V/cm. The electrophoretic mobility (m) of a solute was
calculated using the following equation (Haugland, 1996; Schantz,
et al., 1972).
m=L(1/t.sub.eo-1/t)/E (1)
[0150] where L and E are the capillary length and applied electric
field strength; t.sub.eo and t are the migration times of the
solvent and solute, respectively.
[0151] Horizontally and vertically polarized fluorescence
intensities measured by the LIFP detector were optimized by
aligning a tightly focused laser beam with a small-diameter sample
stream and by balancing signals from the two PMTs. An aqueous
solution of disodium fluorescein (10.sup.-9 M) was passed through a
capillary inserted in a sheath flow cuvette. The sheath fluid,
identical to the CE run buffer, was introduced into the cuvette
hydrodynamically by keeping the inlet reservoir of the sheath
buffer 1 cm higher than the outlet reservoir. The vertically
polarized laser beam was focused onto a spot about 20 mm below the
tip of the capillary. The angle and position of the cuvette
relative to the detection optical path were adjusted so that
roughly equal signals with maximum outputs from both PMTs were
achieved. The values of fluorescence anisotropy (A) were calculated
according to (Shimura, et al., 1994; Schultz, et al., 1993;
Lakowicz 1983).
A=(I.sub.v-I.sub.h/(I.sub.v+2I.sub.h) (2)
[0152] where I.sub.v and I.sub.h are the fluorescence intensities
of vertically and horizontally polarized components,
respectively.
[0153] The values of fluorescence polarization, P, were calculated
according to (Lakowicz 1983; Perrin 1926; Weber, 1953; Dandliker,
et al. 1970)
P=(I.sub.v-GI.sub.h)/(I.sub.v+GI.sub.h) (3)
[0154] where I.sub.v and I.sub.h are the fluorescence intensities
of the vertically and horizontally polarized components,
respectively; G is an empirical constant that corrects for the
polarization bias introduced by the optics and the detection
system. The G value was determined as the intensity ratio of
vertical to horizontal polarization components of fluorescein. For
a well-balanced system, the polarization bias is relatively small
(G=0.98-1.0) and therefore, is negligible.
[0155] It is possible to have unequal transmission of the two
orthogonal polarizations through the emission optical trains and,
therefore, unequal sensitivity of the PMT detectors for vertical
and horizontal polarized emission. To correct for this potential
bias, the PMT voltage was adjusted until the fluorescence
intensities from the two PMT's (the vertically and horizontally
polarized fluorescence) were identical for dilute fluorescein
(10.sup.-9 M), which is assumed to have negligible anisotropy.
[0156] In the first set of examples, laser induced fluorescence
polarization (LIFP) detector was used in conjunction with capillary
electrophoresis to demonstrate the utility of CE/LIFP. Changes in
the electrophoretic mobility and fluorescence polarization of the
fluorescent probe upon complex formation with the binding partner
were measured simultaneously, thereby providing complementary
information on the binding interaction. This information could not
be obtained with either CE or LIFP used alone. Unless otherwise
noted, the smaller molecule of a binding pair was labeled with a
fluorophore such as fluorescein. The complex was formed by mixing
the fluorescent substrate with the corresponding binding partner
and was electrophoretically separated from the unbound substrate
followed by on-line detection with LIFP. Our results showed
expected increases in fluorescence polarization upon complex
formation, demonstrating the usefulness of the technique in binding
studies involving a wide variety of biomolecules.
[0157] For binding systems that have low affinity, dissociation may
take place during separation. We overcame this problem by including
the binding reagents in the CE separation buffer to stabilize the
complex as demonstrated in system in DNA-protein binding studies. A
fluorescently labeled oligonucleotide or SSB protein was used as a
probe and the binding interactions with its partner were studied in
either of two formats, depending on the stability of the complexes
formed. For weak binding interactions, the binding partner was
included in running buffer to stabilize the complexes during CE
separation. The electrophoretic mobility and fluorescence
anisotropy of the fluorescent probe were measured as a function of
the concentration of its binding partner in the running buffer.
Both the electrophoretic mobility and fluorescence anisotropy were
used to determine the binding constants and cooperativity. For high
affinity interactions, mixtures containing the fluorescent probe
and its binding partner at varying ratios were incubated prior to
separation. The complexes formed off-column were then separated by
CE with a running buffer free of the binding components. The
electrophoretic mobility and fluorescence anisotropy measurements
were used for the identification of the complexes and for the study
of binding stoichiometry.
[0158] Materials and Reagents. Disodium fluorescein of purified
grade was obtained from Fisher Scientific (Fair Lawn, N.J.) and was
used for instrument alignment. Fluorescein isothiocyanate (FITC)
isomer I, L-tryptophan, staphylococcal enterotoxin A (SEA),
polyclonal (rabbit) antibody to SEA, SSB protein, d(pT).sub.18,
single-stranded M13mp8 phage DNA and FPIA (fluorescence
polarization immunoassay) dilution buffer (pH 7.4) containing
phosphate, bovine protein and sodium azide, were obtained from
Sigma (St. Louis, Mo.).
[0159] Fluorescein-dUTP was obtained from Molecular Probes (Eugene,
Ore.). A 5'-oligolabeling kit containing T4 polynucleotide kinase,
ATPS and 5-iodoacetamidofluorescein (IAF) was obtained from
Amersham Pharmacia Biotech (Buckinghamshire, England).
[0160] Fluorescein labeled oligonucleotides 11-mer
(5'-CGCGATACGCC-3'; SEQ ID NO:1) and 37-mer
(5'-CCTTAAGCTTCCTCAACCACTTACCATACTCGAGATT-3'; SEQ ID NO:2) were
provided by J. Lee of Cross Cancer Institute and T. Carnelley of
Department of Public Health Sciences, University of Alberta.
[0161] The fluorescein labeled vancomycin and polyclonal (sheep)
antibody to vancomycin were from a Sigma diagnostics reagent set.
The actual compositions and concentrations of these solutions were
not available. The trp repressor protein, trp operator DNA and trp
binding buffer (pH 7.6) were obtained from PanVera (Madison, Wis.)
as a tip repressor-DNA binding kit. The trp operator was a
5'-fluorescein labeled, 25 base pair, oligonucleotide with
sequence:
[0162] (5'-ATCGAACTAGTTAACTAGTACGCAA-3')
[0163] (3'-TAGCTTGATCAATTGATCATGCGTT-5')
[0164] Fluorescent Labeling of SEA. SEA was labeled with FITC and
the extent of modification was estimated according to the methods
described by Brinkley, et al., 1992. A 10-fold molar excess of FITC
was added to a solution of SEA (0.1 mg/mL) in 25 mM
Na.sub.2B.sub.4O.sub.7 (pH 9.1). The reaction was allowed to
proceed for 1 h at room temperature and then terminated by adding
excess of hydroxylamine. The fluorescently labeled SEA was
transferred into a disposable dialyzer tube (Spectra/Por CE Sterile
DispoDialyzer, molecular weight cut-off 10,000 Da) and purified by
dialysis against 10 mM sodium phosphate buffer (pH 7.4) at
4.degree. C. for 2 days. The degree of fluorescent labeling was
determined by analyzing the fluorescently labeled SEA and the free
dye in 25 mM Na.sub.2B.sub.4O.sub.7 (pH 9.1) using a
Hewlett-Packard (Palo Alto, Calif.) Model 1040A diode array
detector. A Gilson (Villies le Bel, France) Model 307 HPLC pump was
used to introduce the sample solution. The molar ratio (R) of the
fluorophore to SEA protein was calculated according to the
following equation:
R=A.sub.490, pe.sub.p/[A.sub.277, p-A.sub.490, p(A.sub.277,
d/A.sub.490, d)]e.sub.d (4)
[0165] where A.sub.277, p and A.sub.490, p are the absorbance of
fluorescently labeled SEA protein at 277 and 490 nm; A.sub.277, d
and A.sub.490, d are the absorbance of the free dye at 270 and 490
nm; e.sub.p and e.sub.d are the extinction coefficients of SEA at
277 nm and the free dye at 490 nm, respectively.
[0166] Fluorescent Labeling of SSB protein and d(pT).sub.18. The
SSB protein was labeled with FITC and the extent of modification
was estimated according to the methods described by Brinkley, et
al., 1992. A 10-fold molar excess of the dye was added to a
solution of the SSB protein (1.4 .mu.g/mL) in 25 mM
Na.sub.2B.sub.4O.sub.7 (pH 9.1). The reaction was allowed to
proceed for 1 h at room temperature and then stopped by adding
excess of hydroxylamine. The FITC-labeled protein was purified
using a prepacked bio-spin column (Bio-Gel P-6, Bio-Rad, Hercules,
Calif.). The absorbance of FITC (280 nm) and FITC-SSB (490 nm) in
25 mM Na.sub.2B.sub.4O.sub.7 (pH 9.1) was measured using a
Hewlett-Packard (Palo Alto, Calif.) Model 1040A diode array
detector equipped with a Gilson (Villies le Bel, France) Model 307
HPLC pump. The molar ratio of the fluorophore to protein was
calculated from the absorbance measurements using the following
extinction coefficients: FITC (Fey, et al., 1984), e.sub.490=73000
cm.sup.-1 M.sup.-1; and SSB (Thompson, et al., 1986),
e.sub.280=120000 cm .sup.-1 M.sup.-1. Absorbance of FITC at 280 nm
was corrected for in the calculation of the molar ratio of FITC to
protein. The labeling of d(pT).sub.18 at the 5'-end with
5-iodoacetamidofluorescein was accomplished by following a protocol
provided by Amersham.
[0167] Formation of the Complexes. Various volumes (0, 2, 4, 6, 8
mL) of antibody solution from a test kit for vancomycin were mixed
with 10 mL aliquots of fluorescein-labeled vancomycin solution in
0.5 mL microcentrifuge tubes. FPIA dilution buffer was added to
each tube to a final volume to 200 mL. The tubes were vortexed for
30 s and the mixture was allowed to incubate at room temperature
for 15 min. Binding studies for SEA-antibody and trp
repressor-operator systems were conducted similarly, with
appropriate amounts of the binding partners. The samples were
analyzed by CE/LIFP.
[0168] For weak interactions, protein-DNA complexes were formed
on-column with excess amounts of the protein. Buffer solutions
containing various concentrations of the SSB protein or
oligonucleotide were used as CE running buffers. The fluorescently
labeled DNA was injected into the capillary for CE/LIFP analysis.
For strong interactions, protein-DNA complexes were formed
off-column. Various volumes (0, 2.5, 5.0, 10, 15 mL) of the M13
phage DNA solution ( 43 nM) were mixed with 1.0-mL aliquots of
FITC-SSB protein solution (12.5 mM) in 0.5-mL microcentrifuge
tubes. FPIA dilution buffer was added to each tube to a final
volume of 50 mL. The tubes were vortexed for 30 s and the mixtures
incubated at room temperature for at least 15 min prior to CE/LIFP
analysis.
Example 1
[0169] Peptide-Protein Interaction. Binding of vancomycin to its
antibody was chosen as an example because of the therapeutic
importance of vancomycin and because of the availability of its
antibody as an affinity agent. Vancomycin is a water soluble,
tricyclic glycopeptide and is strongly bound to its antibody in
solution as has been demonstrated in a homogenous immunoassay
(Schenzer, et al., 1983).
[0170] The complex, formed as described above, and the unbound
vancomycin can be resolved by CE and can be readily identified
based on their differential fluorescence polarization values. Two
electropherograms were obtained from a single CE separation of a
sample containing fluorescein-labeled vancomycin and
anti-vancomycin antibodyand are shown in FIG. 1. Both vertically
(I.sub.v) and horizontally (I.sub.h) polarized fluorescence
components were measured simultaneously. Fluorescein was added as a
reference compound to correct for any possible polarization bias of
the instrument. The fluorescein-labeled vancomycin and the
fluorescein dye rotate rapidly in solution and exhibit little
fluorescence polarization. Thus, the fluorescence intensities
corresponding to the two polarized components are nearly equal. The
binding of vancomycin to its antibody results in a substantial
increase in the molecular size and a slower rotation of the
molecule. The complex exhibits significant fluorescence
polarization. The intensity of the vertically polarized
fluorescence (I.sub.v) was significantly higher than that of the
horizontal component (I.sub.h) for the complex. The same trend was
observed with the complex formed at various vancomycin to antibody
ratios. A mean fluorescence polarization was found to be
0.28.+-.0.02. This value represents the intrinsic polarization of
the complex, which depends on rotational diffusion of the molecule
but is independent of the amounts of the drug and antibody
added.
[0171] The increase of fluorescence polarization upon complex
formation can be expected from the fluorescence polarization
principle (Lakowicz 1983; Perrin 1926; Weber, 1953; Dandliker, et
al. 1970). A fluorescent molecule, when excited by a polarized
light, emits fluorescence with its polarization (P) controlled by
rotational correlation time (f) and fluorescence lifetime (t) as
shown by the Perrin equation (Perrin, 1926)
(1/P-1/3)=(1/P.sub.0-1/3)(1+t/f) (5)
[0172] where P.sub.0 is the intrinsic polarization in the absence
of rotational diffusion. When the rotational correlation time is
small relative to the fluorescence lifetime, the fluorescence is
depolarized. When the fluorescence lifetime is constant for a given
fluorophore (e.g., 4 ns for fluorescein), an increase in
polarization may be observed with increasing rotational correlation
time. The rotational correlation time can be estimated according to
the Debye-Stokes-Einstein equation (Gottfried, et al., 1999)
f=Mh(v+h)/RT (6)
[0173] where M is the mass of the molecule, h is the viscosity of
the solution, v is the specific volume of the molecule, h is the
degree of hydration, T is the absolute temperature, and R is the
ideal gas constant. Using a typical specific volume (v) of 0.735
cm.sup.3/g and a typical value of hydration (h=0.2 cm.sup.3/g)
(Gottfried, et al., 1999), we estimated the rotational correlation
time of vancomycin (0.7 ns) and its complex with the antibody (58
ns). The reduced rotational diffusion of vancomycin when bound to
the antibody resulted in an increase in fluorescence polarization
from 0.08 for the free vancomycin (1838 Da) to 0.28 for the
antibody-bound vancomycin (.about.152,000 Da). The molecular
weight, the estimated rotational correlation time and the observed
fluorescence polarization are summarized in Table 1.
1TABLE 1 Molecular weight, estimated rotational correlation time
and observed fluorescence polarization of FITC labeled substrates
and their protein complexes Estimated Molecular Rotational Observed
Weight Correlation Time Fluorescence Substrate and Complex (Da)
(ns).sup.a Polarization FITC labeled substrates Vancomycin 1,838
0.7 0.08 .+-. 0.02 Trp Operator 15,000 5.8 0.11 .+-. 0.02 SEA
28,000 11 0.14 .+-. 0.02 Complexes Vancomycin-Antibody 152,000 58
0.28 .+-. 0.02 Trp Operator-Repressor 30,000 12 0.25 .+-. 0.02
SEA-Antibody 180,000 69 0.14 .+-. 0.02 .sup.aCalculated from eq
(4), with T = 293K, .nu. = 0.735 cm.sup.3/g and h = 0.2 cm.sup.3/g,
(see ref 15).
[0174] As intrinsic property of a molecule, the characteristic
fluorescence polarization provides evidence for the binding of the
substrate to its antibody without the need of tedious titration
procedures, thereby promising considerable savings on time and
reagents. This feature makes the CE/LIFP approach particularly
suitable for applications such as screening specific monoclonal
antibodies where the speed and convenience are the major concerns
when choosing a screening method (Barret, 1994).
Example 2
[0175] Protein-Protein Interaction. Binding of SEA to its antibody
was studied with an intention of developing CE based immunoassays
for this natural toxin. SEA has been known for many years to cause
food poisoning (Marrack, et al., 1990) and, therefore, it is of
considerable public health interest to develop rapid and sensitive
analytical methods. Radioimmunoassays and enzyme-linked
immunosorbent assays for this toxin have been described in the
literature (Johnson, et al., 1973; Fey, et al., 1984; Thompson, et
al., 1986).
[0176] Experiments were run to characterize the FITC labeled SEA
and to examine the formation and stability of its complex with the
corresponding antibody. Using equation (1) and literature values of
extinction coefficients for the dye (e.sub.d=73,000 cm.sup.-1
M.sup.-1) and the protein (e.sub.p=40,900 cm.sup.-1
M.sup.-1),(Haugland, 1996; Schantz, et al., 1972) we estimated the
molar ratio of the dye to the protein to be 5.9.+-.0.3 for the
labeled SEA.
[0177] Fluorescently labeled SEA exhibits an appreciable
polarization (0.14), making it readily identifiable even in the
presence of the residual dyes. This finding suggests that the
CE/LIFP may be used to monitor the progress of labeling reactions
and to verify the purity of the reaction products. The broadness of
the peak is attributed to the heterogeneity of the protein
(Schantz, et al., 1972) as well as to the presence of multiply
labeled products (Craig, et al., 1998).
[0178] The electropherograms from a CE/LIFP analysis of a mixture
of FITC-SEA and its antibody show a new peak attributable to the
complex between fluorescently labeled SEA and its antibody. The
complex exhibited measurable fluorescence polarization (0.14) as
seen with the free SEA. However, there was no net increase of
polarization upon complex formation. This is not surprising given
the relatively high molecular weight of SEA itself (28,000 Da). In
an aqueous solution at room temperature, the rotational correlation
time (f) of SEA is estimated using equation (6) to be approximately
11 ns, which is about 2.5 lifetimes of fluorescein (t, less than 4
ns). With such a long rotational correlation time, the t/f term
(<0.4) in equation (5) contributes little to the observed
fluorescence polarization (P). The polarization may approach to
P.sub.0, the intrinsic value for the molecule because it is known
that fluorescence polarization generally approaches saturation with
molecular weight beyond 20,000 Da (Guo, et al., 1998). As a result,
further increase in correlation time due to the binding of antibody
did not lead to any significant increase in polarization. The
polarization of 0.14 for the complex is well below the theoretical
limit of 0.5, indicating that local rotation of the fluorophore may
occur within the SEA molecule.
[0179] Because of no increase in polarization noted in this case,
additional evidence is needed to confirm the complex formation,
which can be obtained by titrating a fixed amount of FITC-SEA with
varying amounts of the antibody. As expected, the complex peak
increases at the expense of the unbound SEA peak (Wan, et al.,
1999)
Example 3
[0180] Protein-DNA interaction. The interaction between trp
operator (DNA) and trp repressor protein of Escherichia coli serves
as a well-characterized system for gene expression and regulation
(Crawford, et al., 1980). In the presence of tryptophan, the
repressor protein binds with high affinity to the operator sequence
found within the promoter region of the trpEDCBA operon and
represses transcription of those genes whose protein products are
responsible for the synthesis of tryptophan. In the absence of
tryptophan, trp repressor is inactive and the trp operon is
expressed, resulting in the biosynthesis of tryptophan. The binding
of the trp operator to trp repressor has been studied extensively
(Carey, 1988; Otwinowski, et al., 1988;Lawson, et al., 1988;
LeTilly, et al., 1993; Zhang, et al., 1994; Stebbins, et al.,
1996).
[0181] Two formats were used to explore the potential of CE-LIFP in
the study of DNA-protein interactions in the trp operator (DNA) and
trp repressor protein system. In the first format, the affinity
complex was formed dynamically in the separation capillary and
maintained at equilibrium with the free protein during the
electrophoresis. In the second format, the complex was preformed by
incubation and then separated from the unbound protein
molecule.
[0182] The electropherograms of fluorescently labeled trp operator
oligonucleotide obtained with and without the trp repressor protein
in the running buffer were compared. The I.sub.v, and I.sub.h,
representing vertically and horizontally polarized fluorescence,
respectively were measured. In the presence of the trp repressor, a
tailed peak with migration time of 4.6 min was observed. This peak
corresponds to the complex formed between the trp operator and trp
repressor judging from the fluorescence polarization that increased
from 0.11 (without the trp repressor in the running buffer) to 0.25
(with the inclusion of the trp repressor in the running
buffer).
[0183] It is noted that fluorescently labeled DNA displays multiple
peaks in both cases and that these peaks do not disappear even in
the presence of excess trp repressor. The persistence of the
multiple peaks suggests the presence of multiple DNA structures
(Stebbins, M. A , et al., 1996; Hamdan, et al., 1998), some of
which are not recognized by the repressor protein. Experiments have
established the dynamic formation of the DNA-protein complex (Wan,
et al., 1999).
Example 4
[0184] SSB protein/ssDNA. The SSB protein plays an important role
in the DNA replication, recombination and repair proces
(Bandyopadhyay, et al., 1978; Krauss, G., et al., 1981; Chase, et
al., 1986; Lohman, et al., 1994) although mechanisms for its
functions in these processes have not yet been elucidated. The SSB
protein exists as a tetramer in solution with a subunit weight of
about 20000. It binds cooperatively to single-stranded DNA
(approximately 32-60 nucleotides per protein), keeping the DNA in
an extended configuration and protecting it from nuclease
digestion. The results shown in the SSB/DNA binding experiments
demonstrate the resolving power and identification capabilities of
CE/LIFP in these systems.
[0185] CE/LIFP may be used in binding studies through the
measurement of changes of either electrophoretic mobility or
fluorescence anisotropy of a binding component upon affinity
interactions. Usually, the binding component of lower molecular
weight is used as a probe so as to induce greater mobility or
anisotropy changes upon binding. In this example, both
fluorescently labeled DNA fragments and binding protein were used
as probes for the study of protein-DNA interactions.
[0186] In this example, a fluorescein labeled 11-mer (F-11-mer) was
evaluated as a probe for examining its binding with the SSB
protein. Electropherograms of F-11-mer in the absence and presence
of the SSB protein in the running buffer were run and the vertical
(I.sub.v) and horizontal (I.sub.h) components of fluorescence were
measured simultaneously for each. Fluorescein labeled dUTP (F-dUTP)
was used as a reference compound to correct for possible
fluctuations in electroosmotic flow and unequal detection
sensitivity between the two detection channels of the instrument.
In the absence of the binding protein, the fluorescent probe has an
electrophoretic mobility of m=3.99.times.10.sup.-4 cm.sup.2
V.sup.-1 s.sup.-1 and a fluorescence anisotropy of A=0.05. In the
presence of 0.7 mM of the SSB protein in the running buffer, the
electrophoretic mobility of the oligonucleotide probe is reduced to
1.93.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1 whereas the
anisotropy is increased to 0.25. The decrease in electrophoretic
mobility and increase in anisotropy are due to the binding of the
F-11-mer with the SSB protein. In contrast, the mobility and
anisotropy of F-dUTP are essentially unchanged, consistent with the
fact that the SSB protein has very low binding affinity for the
mononucleotide.
[0187] The electrophoretic mobility and anisotropy changes observed
in the experiments arise from the effects of the binding protein on
the molecular motion of the probe. In the absence of the binding
protein, the fluorescently labeled oligonucleotide probe migrates
with a mobility similar to that of F-dUTP in the free zone
electrophoresis mode. It has a low fluorescence anisotropy because
of its small molecular size (MW 4000) and random motion in
solution. When bound to the SSB protein (MW 80000), the size of the
fluorescent molecule is markedly increased, resulting in a slower
molecular motion in the solution. Therefore, it is not surprising
that binding of the SSB protein to the oligonucleotide probe gives
rise to a marked increase in anisotropy. The mobility and
anisotropy of the complex approach to those of the binding protein
(m=1.07.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1 and A=0.23 for
FITC-SSB. While the electrophoretic mobility of a compound is
proportional to its charge to mass ratio, (Chu, et al., 1995;
Baker, 1995) the fluorescence anisotropy is mainly determined by
its molecular size, shape, and fluorescence lifetime (Lakowicz,
1999).
[0188] The mobility and anisotropy of the F-11-mer were measured
with the running buffer containing various concentrations of the
SSB protein. Nanoliter amounts of the F-11-mer (10.sup.-9 M) were
injected into the capillary that was filled with the running buffer
containing 10.sup.-7-10.sup.6 M SSB protein. Thus, the
concentration of the binding protein in the running buffer was in
large excess and was not affected significantly by its binding with
the F-11-mer. Under this condition, the quantitative interpretation
of the binding profiles can be carried out using a standard
four-parameter logistic equation (Motulsky, 1999):
y=(a+bK.sup.nx.sup.n)/(1+K.sup.nx.sup.n) (7)
[0189] where y is the observed response such as electrophoretic
mobility or fluorescence anisotropy of the oligonucleotide probe at
a given concentration of the binding protein, x; K is the apparent
binding constant; a and b are the responses of the free and bound
probe, respectively; and superscript n is the Hill coefficient
describing the steepness of the curve. The experimental data were
fitted to the above equation using nonlinear regression analysis
(SigmaPlot, version 4, SPSS Inc.). Binding constant (K)
measurements based on mobility and anisotropy of the F-11-mer were
similar, which were approximately 4.4.times.10.sup.6 M.sup.-1 and
5.times.10.sup.6 M.sup.-1, respectively. These values are
comparable to previous measurements by other methods. For example,
Molineux (Molineux, et al, 1975) reported that SSB has an affinity
of about 2.times.10.sup.6 M.sup.-1 for d(pT).sub.8 and Krauss
(Krauss, et al, 1981) reported an affinity of 1.4.times.10.sup.6
M.sup.-1 for d(pT).sub.16 using fluorescence quenching methods.
Binding constants and fitting parameters from nonlinear regression
analysis are summarized in Table 1.
[0190] Furthermore, binding interactions involving protein-DNA
complexes that differ in stoichiometry can be accomplished with
this instrumentation. Fluorescein labeled 37-mer (F-37-mer) was
chosen as a DNA probe since its complexes with the SSB protein are
of higher stability, allowing examination of a distribution of
different species in the binding interactions. Electropherograms of
F-37-mer with the absence and presence of the SSB protein in the CE
running buffer were collected. Changes in electrophoretic mobility
and fluorescence anisotropy upon formation of complexes between
F-37-mer and SSB protein are observed as expected. It is noted that
the initial single peak of F-37-mer was split into two when bound
to the SSB protein. Both complex peaks display strong fluorescence
isotropy (A=0.23). These are likely 2:1 (peak 2) and 1:1 (peak 1)
protein-DNA complexes. For the 1:1 binding interaction, variations
of the m and A values for the F-37-mer with the binding protein
concentration were compared, from which the apparent binding
constants were obtained. Again, both mobility and anisotropy
measurements gave very similar results, with binding constants of
approximately 2.times.10.sup.7 M.sup.-1 (see Table 1). This is
approximately 5-fold increase in binding affinity of SSB for the
37-mer compared to its binding with the 11-mer. This is consistent
with the contribution of cooperativity to the binding strength. The
SSB protein is a tetramer and the number of the binding sites of
the SSB protein varies with the length of the oligonucleotides
because each of the four subunits of the protein covers 6-8
nucleotides (Krauss, et al, 1981; Chase, et al., 1986). While one
subunit may bind to the 11-mer, all the four subunits of the SSB
tetramer could bind to the 37-mer, resulting in the corresponding
increase in binding constant. Because the two complex species were
not well resolved particularly at low protein concentrations, there
was a relatively large uncertainty associated with mobility and
anisotropy measurements for the 2:1 complex. Consequently, we were
unable to precisely determine the corresponding binding constant
for the 2:1 complex.
Example 5
[0191] Fluorescein Labeled DNA Binding Protein as a Probe.
Fluorescently labeled DNA oligonucleotides have exclusively been
used as probes in the analysis of protein-DNA interactions by gel
retardation, CE or FP techniques. However, many bioanalytical
applications require the use of a fluorescently labeled binding
protein for its ability to recognize and bind to specific
structures of DNA. In this example, FITC-labeled SSB protein
interacts with a synthetic oligonucleotide and a single-strand DNA.
The FITC-labeled SSB protein was prepared as described in the
experimental section and the dye to protein molar ratio was 5.3. In
a solution of 25 mM disodium tetraborate with pH 9.1, the labeled
SSB protein displayed an electrophoretic mobility of
1.07.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1 and a fluorescence
anisotropy of 0.23.
[0192] Electropherograms of FITC-SSB protein were obtained with
running buffers containing varying amounts of oligonucleotide,
d(pT).sub.18. Both vertically and horizontally polarized
fluorescence emissions were acquired simultaneously. There was only
a slight increase in fluorescence anisotropy (DA>>0.02) of
the protein probe upon binding to the oligonucleotide. This is in
accordance with the fact that fluorescence anisotropy of a probe
generally approaches saturation when the probes molecular weight
exceeds 20000 (Lakowicz, 1999; Wan, et al., 1999; Guo, et al.,
1998).
[0193] It is noted that formation of the complex gives rise to some
significant changes in the mobility and peak shape of the
FITC-labeled SSB protein. As the concentration of d(pT).sub.18 in
the running buffer increases, the electrophoretic mobility of the
SSB protein increases with the peak becoming increasingly
dispersed. The mobility increase of the low mobility species (the
FITC-SSB protein, m=1.07.times.10.sup.-4 cm.sup.2 V.sup.-1
s.sup.-1) is due to its binding to a high mobility DNA
(m=2.19.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1 for fluorescein
labeled d(pT).sub.18). The peak broadening suggests that the
formation of multiple protein-DNA complexes is possible in the
presence of increasing amount of DNA. A single protein molecule may
bind several DNA molecules in the presence of excess DNA. The
complexes of varying protein to DNA ratios co-migrate in the
separation capillary as a broad band. To clarify this point, we
chose a DNA fragment much longer than d(pT).sub.18 to form stable
complexes with the SSB protein. Because of increased stability, the
multiple complexes formed off-column can be separated without the
need for adding the binding partner to the running buffer.
Example 6
[0194] Detection of labeled protein binding to ssDNA. In the
following embodiment, single-stranded M13mp8 phage DNA (7229 bases)
was selected as a binding partner for the labeled SSB protein.
Varying amounts of the phage DNA were incubated with a series of
binding solutions containing a fixed amount of the FITC-labeled SSB
protein. The mixtures were then analyzed by CE/LIFP with a running
buffer free of the binding components. Separations of FITC-SSB
protein and its complexes with the DNA at various molar ratios of
DNA to protein show that as the amount of DNA in the reaction
mixture increases, new peaks emerge and become increasingly
retarded and broadened. The broad and multiple peaks with
increasing migration times showed strong fluorescence anisotropy
(A=0.25), indicating the presence of multiple protein-DNA
complexes. Three major peaks with increasing mobilities (peak 2,
2.16.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1; peak 3,
2.66.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1; and peak 4,
3.16.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1) corresponded to
the complexes with increasing DNA to protein ratios.
[0195] In the absence of the DNA, peak 1 corresponds to the SSB
protein probe which has a m=1.07.times.10.sup.-4 cm.sup.2 V.sup.-1
s.sup.-1. Addition of the DNA to the protein (with DNA-to-protein
ratio of 0.008 and 0.016) causes a decrease in peak 1 and the
appearance of peak 2 (m=2.16.times.10.sup.-4 cm.sup.2 V.sup.-1
s.sup.-1 ), indicating the formation of DNA-protein complex. With
further increase of DNA-to-protein ratio, the mobilities of the
complexes (2.66.times.10.sup.-4 cm.sup.2 V.sup.-1 s.sup.-1 for peak
3; and 3.16.times.10.sup.-1 cm.sup.2 V.sup.-1 s.sup.-1 for peak 4)
shift towards that of the DNA (m=3.50.times.10.sup.-4 cm.sup.2
V.sup.-1 s.sup.-1)..sup.27 Because the amount of the protein was
fixed in this series of experiments, increasing amounts of DNA in
the reaction mixture favor the formation of complexes of increasing
DNA:protein ratio. This example demonstrates an application of
CE/LIFP to study multiple-complexes.
Section B: Examples 7-10
Complex Formation Between HIV-RT and Aptamers
[0196] In the next few examples the detection of human
immunodeficiency virus type 1 reverse transcriptase was
accomplished using aptamers as probes in affinity capillary
electrophoresis and laser induced fluorescence polarization. CE
determination of HIV-1 RT using a noncompetitive affinity assay has
several advantages in terms of vastly decreasing analysis time and
involves much simpler chemical procedures. A fluorescently-labeled
aptamer such as RT 12 or RT 26 eliminates the need for the use of
radio-labeled materials, and provides the first direct assay for
HIV-1 RT. Since the aptamers were evolved to bind selectively to
HIV-1 RT, interferences from RTs of other species was eliminated or
greatly attenuated. Used in conjunction with other laboratory
procedures correlating HIV-1 RT activity to viral loads, the assay
could prove useful in the determination of HIV-1 viral load.
[0197] Apparatus: The CE/LIF instrument used in this work is the
same as described above except that a 543.5 nm green He--Ne laser
(Melles Griot, Irvine, Calif., USA) with a 5 mW maximum out put was
used as the excitation source.
[0198] Reagents: All solutions were prepared using 18.2 MW
distilled, deionized water (DDW) from a Milli-Q Gradient 10 Water
System (Millipore, Nepean, Canada). Tris-borate-EDTA (TBE) (0.089 M
tris, 0.089 M boric acid, 0.0025M EDTA, pH 8.3), tris-glycine
(0.025 M tris, 0.192 M glycine, pH 8.3) and disodium tetraborate
buffers (0.1 M, pH 9.1) were prepared using reagent-grade materials
and diluted to desired concentrations with DDW prior to being
filtered through a 0.22 .mu.m filter to remove particulate matter.
The RT 12 aptamer (5'-ATCTACTGGATTAGCGATACTCGATTAGGTC-
CCCTGCCGCTAAACCATACCGCGGTAACTTGAGCAAAATCACCACTGCAGGGG-3'; SEQ ID
NO:3) and the RT 26 aptamer
(5'-ATCCGCCTGATTAGCGATACTTACGTGAGCGTGCTGTCCCCTAAAGGTGAT-
ACGTCACTTGAGCAAAATCACCTGCAGGGG-3'; SEQ ID NO:4) were labeled with
5'-FAM (5'-carboxyfluorescein) at the University Core DNA Services,
University of Calgary, Canada. HIV-1 RT was obtained from
Worthington Biochemicals (Lakewood, N.J.). RTs from the enhanced
avian myeloblastosis virus (AMV) and the Moloney murine leukemia
virus (MMLV) were obtained from Sigma (Mississauga, Canada). Cell
culture media (RPMI with 10% fetal bovine serum (FBS)) was obtained
from the Cross Cancer Institute at the University of Alberta.
[0199] Capillary Electrophoresis/Laser-Induced Fluorescence:
Uncoated fused silica capillaries (20 .mu.m I.D., 150 .mu.m O.D.)
were cut to a length of 40 cm and inserted into the sheath flow
cuvette where the laser beam was focused. Samples were injected for
5 s at a voltage of 15 kV (375 V/cm), and electrophoresis was
carried out at a running voltage of 20 kV (500 V/cm). The running
buffer utilized for all experiments was 1.times.tris glycine. The
laser power was set at 4 mW throughout. Periodically, the
capillaries were treated by running 0.1 M NaOH through the system
at an running voltage of about 100 V/cm for 30 mintues, followed by
the running buffer (1.times.tris glycine) at 500 V/cm, to remove
protein material adsorbed on the capillary wall.
[0200] Affinity Complex Formation: The RT 12 aptamer and RT 26
aptamer were received in 0.020 .mu.g and 0.040 .mu.g quantities,
respectively. These were diluted to 60 .mu.L in 1.times.TBE in 600
.mu.L microcentrifuge tubes and stored in a freezer at 20.degree.
C. when not in use, as were the RTs of HIV-1, AMV and MMLV. Stock
solutions of 80 nM for the RT 12 aptamer and 170 nM for the RT 26
aptamer were prepared in 1.times.TBE in 600 .mu.L microcentrifuge
tubes. A 1000 nM stock solution of HIV-1 RT was similarly prepared
in DDW. Complex formation was carried out in an incubation buffer
of 1.times.TBE. The desired concentration of aptamer and protein
was obtained by pipeting the appropriate volumes of aptamer and
protein stock solutions into a 60 .mu.L volume in 600 .mu.L
microcentrifuge tubes. The tubes were then vortexed for 30 s and
put on ice for about 5 minutes prior to sample injection into the
capillary. All stock solutions were stored at 20.degree. C. when
not in use and all samples were kept on ice during the course of
experimentation.
[0201] Interference Studies: To determine the degree to which the
aptamers would bind with RTs from AMV and MMLV, experiments were
conducted in which complex formation experiments, as described
above, were undertaken with AMV and MMLV RTs substituted for HIV-1
RT. Furthermore, complex formation experiments were conducted with
RTs of HIV-1, AMV and MMLV mixed together, with the AMV and MMLV
RTs at the same or higher concentration than HIV-1 RT. To determine
the degree to which matrix effects from cell culture media would
interfere with complex formation, aliquots of RPMI containing 10%
FBS were added to samples containing both HIV-1 RT and aptamer, as
well as aptamer alone.
Example 7
[0202] Detection of Affinity Complex Formation after CE Separation.
The affinity complex was formed by adding increasing concentrations
of aptamer to a fixed concentration of HIV-1 RT (50 nM). In the
absence of HIV-1 RT, the aptamer peak is sharp with a migration
time of around 4.2 minutes. Tailing, a characteristic of the
aptamer peak, was observed even at the lowest aptamer
concentrations used (1.7 nM). This most likely results from
impurities in the DNA, as has been previously observed (German, et
al., 1998). These DNA impurities likely contain multiple DNA
structures, which cannot be recognized by the HIV-1 RT protein
(Wan, et al., 1999; Stebbins, et al., 1996; Hamden, et a.,
1998).
[0203] The RT-26-HIV-1 affinity complex was formed. The peak
corresponding to the complex had a migration time of about 3.3
minutes and was well resolved and Gaussian in appearance. The lack
of features such as bumps or shoulders suggested that the affinity
complex was primarily of single stoichiometry. At concentrations of
50 nM HIV-1 RT and 17 nM RT 26, the complex peak migrated as a
doublet or a shoulder appeared, indicating that RT 26 and HIV-1 RT
were forming a complex of two stoichiometries. The peak area of the
HIV-1 RT-aptamer affinity complex increases with increasing aptamer
concentration.
[0204] Experiments were also undertaken using RT 12 at 8, 15, and
20 nM concentrations added to a solution containing a fixed
concentration of 50 nM of HIV-1 RT. The results parallel those
described above, in that the CE peak area of the affinity complex
increased with increasing aptamer concentration. However, the RT
12-HIV-1 RT affinity complex exhibited two distinct peaks, one
forming at the same migration time as the RT 26-HIV-1 RT complex,
while an additional early peak formed at about 2.9 minutes. HIV-1
RT is a heterodimer of total molecular weight (120 kDa) with two
sub-units of molecular weight 51 kDa and 66 kDa. Although it is
possible that the dimeric forms of HIV-1 RT may be binding to
different sites on the differently-structured RT 12 aptamer, a more
likely explanation is that the RT 12 is being incorporated into the
affinity complex in such a way as to produce a complex of two
different stoichiometries (Wan, et al., 1999). Later experiments,
in which the RT 12-HIV-1 RT affinity complex was observed to
migrate as a single peak, confirm the belief that an affinity
complex of different stoichiometries was formed in these
experiments. Because of the higher binding constant of the RT 26
aptamer (81-mer), it was chosen for all further work.
Example 8
[0205] Use of CE/LIFP to create Calibration Curves: Calibration
curves for HIV-1 RT were constructed using aptamer concentrations
of 17 nM and 60 nM, preferebly, 17 nM of the RT 26 aptamer. The RT
26 probe peak area decreases with increasing HIV-1 RT
concentration, reaching a limiting value at 100 nM and disappearing
completely at 800 nM of HIV-1 RT. That the aptamer was completely
incorporated into the affinity complex indicates the aptamer was at
its preferred orientation in the TBE incubation buffer, without the
need for heat denaturing and the presence of Mg.sup.2+ salts, as
was found in another assay in which DNA aptamers were used to bind
IgE and thrombin (German, et al., 1998). Most sample solutions were
stable for about two weeks if immediately frozen after use, after
which both the probe and affinity complex peak areas were
significantly diminished. At HIV-1 RT concentrations of 7 nM or
lower, it was necessary to perform experiments within 30-40 minutes
because aptamer and complex peak area began to deteriorate. Because
of practical considerations such as these, calibration curve
samples were prepared sequentially, and samples were prepared fresh
daily.
[0206] The calibration curve for the bound complex showed an
initial steep increase in fluorescence intensity with HIV-1 RT
concentration, followed by leveling off, indicative of binding
saturation, beginning at about 100 nM of HIV-1 RT. For the aptamer
probe peak, this was mirrored by a similar steep loss in
fluorescence intensity, followed by a leveling off at about 100 nM.
It is incorporated in the affinity complex, ultimately completely,
at 800 nM of HIV-1 RT. The steeply rising sections of the curves
were then investigated to determine analytical utility. The linear
dynamic range for both probe and complex peaks extends to 50 nM of
HIV-1 RT. In the case of the aptamer probe, least-squares linear
regression provides a best-fit line having a correlation
coefficient (r.sup.2) of 0.985 and a slope of 0.612, whereas the
same fit to the bound complex provided an r.sup.2 value of 0.986
and a slope of -0.938. Relative standard deviations range from a
high of 7.1% to majority of 1.9%-2.5% for both probe and complex
peaks.
Example 9
[0207] Use of CE/LIFP in Specificity determination: Experiments
were performed in which all or some of the RTs were added together
with HIV-1 RT and 17 nM of RT 26 aptamer. AMV RT is present at a
concentration comparable to that of HIV-1 RT, whereas, MMLV RT was
an order of magnitude more concentrated and was increased to over
two orders of magnitude above that of HIV-1 RT. The peak areas as
measured by CE of the unbound aptamer did not decrease, remaining
essentially identical to that in the presence of HIV-1 RT alone.
These results indicate that the presence of other RT proteins, such
as AMV-RT and MMLV-RT, does not affect the determination of HIV-1
RT.
[0208] The RTs of AMV and MMLV can be shown not to cross-react with
the aptamer and to be specific for HIV-1 RT. Using high
concentrations of RT (4-4000 units/.mu.L) and aptamer probe (140
nM), AMV-RT concentrations of 4 units/.mu.L, and MMLV-RT
concentrations of 4000 units/.mu.L, The peak area of the unbound
aptamer as measured by CE is essentially the same in the absence
and presence of AMV-RT and MMLV-RT.
Example 10
[0209] Use of CE/LIFP to show Effects of Sample Matrix:
Electropherograms from the analysis of mixtures containing 20 nM
HIV-1 RT and 17 nM RT 26 aptamer in TBE buffer, in RPMI cell
culture medium and in 100-fold dilute culture medium were measured.
Undiluted culture medium clearly affected the formation and CE/LIF
analysis of the complex. This is not surprising because the RPMI
culture medium was supplemented with 10% FBS. This protein is known
to affect HIV-1 RT (Lee, et al., 1987). When the cell culture
medium was diluted 100-fold, the matrix interference on the complex
formation and CE/LIF analysis was minimal. The analysis of mixtures
of the RT 26 aptamer and HIV-1 RT in TBE buffer and in 100-fold
dilute culture media show similar electropherograms.
Section C: Examples 11-21
Detection of Damaged DNA
[0210] In the next set of examples, the detection of DNA adducts of
benzo[a]pyrene using immuno-electrophoresis with laser-induced
fluorescence is demonstrated on the analysis of A549 cells.
Benzo[a]pyrene belongs to a class of compounds called Polycyclic
aromatic hydrocarbons (PAHs), which are known exhibit strong
carcinogenic properties, presumably as a result of the damage that
they or their metobolites cause cause to DNA. In vivo, B[a]P is
converted to benzo[a]pyrene diol epoxide (BPDE). Because of the
biological significance of DNA damage and repair, many techniques
have been developed for the determination of DNA damage (Pfeifer,
1996).
[0211] Synthetic BPDE-DNA adduct was used as a standard probe in a
competitive assay to determine the levels of BPDE-DNA adducts in a
human lung carcinoma cell line exposed to BPDE. A fluorescently
labeled BPDE-DNA adduct standard and a BPDE-specific antibody were
added to a sample containing unknown amount of unlabeled BPDE-DNA
adduct. The unlabeled BPDE-DNA adduct and the labeled BPDE-DNA
adduct compete to form complexes with the antibody. CE separation
of the bound and unbound adducts allows determination of the bound
concentration, which in turn is related to the amount of BPDE-DNA
adduct in the sample. In contrast to other methods of performing
immunoassays, CE-LIF allows rapid analysis, excellent mass
sensitivity and potential for automation. The popularity of this
technique in immunoassays is well reflected in numerous reports,
primarily for the determination of therapeutic drugs (Schulz, et
al., 1995; Schmalzing, et al., 1995; Chen, et al., 1994;
Evangelista, et al., 1994; Chiem, et al., 1998).
[0212] Preparation of BPDE-DNA adduct standard. BPDE powder
(benzo[a]pyrene-r-7, t-8-dihydrodiol-t-9, 10-epoxide (+/-) (anti)
was obtained from Midwest Research Institute (Kansas City, Mo.,
USA. MRI 0477; Lot CSL-98-775-17-16). The BPDE powder was dissolved
in dimethyl sulfoxide (DMSO) to a stock solution of 3 mM. A 16-mer
oligonucleotide, 5'-CCCATTATGCATAACC-3' (SEQ ID NO:5), was treated
with BPDE at a molar ratio of 1:5 (oligonucleotide: BPDE), using a
protocol similar to that described by Cosman (Cosman, et al.,
1990). The oligonucleotide was reconstituted in a buffer containing
20 mM phosphate/1.5% triethylamine at pH 11. To the
oligonucleotide, a BPDE solution was added to a final concentration
of 270 mM. The final mixture was incubated in the dark at ambient
temperature overnight with gentle shaking. Purification of the
BPDE-modified oligonucleotide was carried out in two separate
rounds of HPLC elution using a preparative column (Phenomenex,
Torrance, Calif., USA. LUNA Su C18(2); 250.times.10 mm 5 mm
particle size). In the first round, an isocratic elution using 70%
methanol and 30% of 20 mM phosphate, pH 7 was used to purify the
BPDE-oligonucleotide by separating the oligonucleotides from the
unreacted BPDE. The eluent containing the BPDE-oligonucleotide and
the unmodified oligonucleotide was freeze dried and subjected to a
second round of HPLC purification using a gradient elution of
methanol/20 mM phosphate at pH 7. This purification step separates
the BPDE-oligonucleotide from the unmodified oligonucleotide. The
freshly purified BPDE-oligonucleotide was subjected to a standard
kinase reaction to facilitate subsequent ligation to 5 other
oligonucleotides to form a BPDE-DNA duplex of 90 base pairs. The
BPDE-DNA duplex was gel purified using a 7.5% native polyacrylamide
gel, and subsequently subjected to UV and fluorescence scanning to
measure DNA concentration as well as to confirm the presence of
BPDE moiety on the 90-mer.
[0213] Specific monoclonal antibodies. Monoclonal antibodies 8E11
and 5D11 were obtained from BD PharMingen (San Diego, Calif., USA).
Both antibodies were derived from BALB/c mice immunized with
racemic anti-BPDE modified guanosine conjugated with bovine serum
albumin (Santella, et al., 1984).
[0214] Preparation of BPDE-DNA adducts from A549 cells. A human
lung carcinoma cell line (A549) was incubated with BPDE to produce
DNA adducts in genomic DNA. Briefly, the cell line was maintained
in DMEM/F12 medium (Gibco BRL, Gaithersburg, Md., USA) supplemented
with 10% fetal bovine serum. The cells were seeded at
1.times.10.sup.5 cells per plate and maintained at 95% humidity and
5% CO.sub.2 for 20 hours prior to the addition of BPDE. Treatment
of BPDE was carried out in duplicate sets of A549 cells. Old
culture media were removed from each culture plate and the cells
were washed twice with phosphate buffered saline (PBS). Media
containing BPDE at various concentrations (9.4, 18.8, 37.5, 75,
150, and 300 mM final concentration) were added accordingly to the
designated plates. The cells were further incubated in the media
containing BPDE for 2 hours. The cells were then washed with PBS
prior to the addition of DNAzol lysis reagent (Gibco BRL) to
facilitate cell lysis. Subsequent steps involved a standard 99.9%
ice cold ethanol precipitation and a 70% cold ethanol wash to
purify the genomic DNA. The final DNA pellet was dissolved in
distilled deionized water (ddH.sub.2O) and DNA concentration was
measured at OD.sub.260 using ddH.sub.2O as a blank.
[0215] CE-LIF Instrumentation. The instrument for capillary
electrophoresis with laser induced fluorescence detection is
described above, with one modification. A 543.5 nm green He--Ne
laser (Melles Griot, Irvine, Calif., USA) with a 5 mW maximum
output was used as the excitation source. In addition, in place of
the polarizing beam splitter, a 580DF40 band-pass filter was used
before the transmitted light is collected by the PMT.
[0216] CE separation. Capillary electrophoresis of the sample was
performed using a 29-cm long, 20 .mu.m i.d., 150 .mu.m o.d.
fused-silica capillary (Polymicro Technologies, Phoenix, Ariz.,
USA). Electrophoresis buffer was a Tris-glycine mixture containing
25 mM Tris and 192 mM glycine at pH 8.3. The injection end of the
capillary was set at a positive polarity and the other end
installed inside the sheath-flow cuvette was grounded. Sample
introduction was performed by electrokinetic injection at 10 kV for
5 to 10 s unless otherwise indicated. Separation was performed with
an electric field of 330 to 830 V/cm.
[0217] Immuno-complex of BPDE-DNA adducts. The incubation
conditions were optimized for short reaction time and stable
complex between the BPDE-DNA adduct and its antibody. The
incubation was carried out at room temperature for 10 min in the
dark. The incubation buffer was identical to the separation buffer
except at half the ionic strength. The effect of buffer ionic
strength on complex stability was studied by using the Tris-glycine
buffer at various concentrations.
[0218] Competitive binding of BPDE-DNA adducts. Two
oligonucleotides, a 16-mer and a 90-mer, were used as probes for
competitive immunoassay. They each contained a single BPDE adduct
in the middle and both were fluorescently labeled at a 5' end with
a tetramethylrhodamine (TMR). Another adduct standard carrying an
identical BPDE-DNA adduct was prepared. This adduct standard is 16
bases in length and not fluorescently labeled. This BPDE-16 mer
competes with the TMR-labeled BPDE-90 mer or TMR-labeled BPDE-16
mer to form complexes with the BPDE antibody. To determine the
levels of BPDE-DNA adduct in the A549 cells exposed to BPDE, the
purified genomic DNA from these cells was analyzed and the adducts
in the DNA competed with the TMR-labeled BPDE-90 mer standard for
binding with the antibody 8E11.
Example 11
[0219] Using CE/LIFP to determine Free solution mobility of DNA
adducts. Under free zone electrophoretic conditions, DNA fragments
are not separable when driven by electro-osmotic flow alone because
of the similar mass-to-charge ratio between DNA fragments. The
immunoassay presented here makes use of an antibody to specifically
form a complex with DNA adducts so that the complex can be
separated from the free DNA. The antibody-bound DNA adduct migrates
out first and then the unbound DNA. This migration behavior can be
expected from equation (8) (Karger, et al., 1989). Eq. (8) predicts
the influence of the effective charge (Q), solution viscosity (h)
and the radius of an analyte (r) on electrophoretic mobility
(m.sub.ep). Both the antibody-bound and unbound DNA adducts are
negatively charged. The direction of their electrophoretic mobility
is opposite to that of the electroosmatic flow (EOF). The decrease
in total charge-to-mass ratio after antibody binding decreases the
mobility of the bound DNA adduct moving back to the injection end
(positive polarity). The net result is a faster migration directed
towards the detector end (direction of EOF).
m.sub.ep=Q/6phr (8)
[0220] Comparing the two antibodies for their affinity to the BPDE
-90 mer, antibody 8E11 formed more complex than the antibody 5D11
formed. This may reflect the fact that 8E11 was raised against BPDE
mononucleotides and 5D11 was raised against BPDE modified calf
thymus DNA. Thus, the 5D11 might be expected to have a higher
affinity for long stretches of DNA.
[0221] The longer the capillary column, the more likely that the
complexes may dissociate during electrophoresis. At a capillary
length of 60 cm, the amount of detectable antibody-bound DNA
adducts was reduced by approximately 5-fold relative to an
identical mixture separated on a 30-cm capillary. This reduction
may be due to the instability of the complexes during
electrophoretic separation, or may be caused by adsorption of the
complexes on the capillary wall. These problems could be avoided by
using a shorter column to carry out the separation without losing
resolution.
[0222] Separation at high field strength also helps to improve
resolution. In this Example, at a field strength of approximately
830 V/cm (25 kV for 30-cm capillary), the antibody-bound and
unbound DNA adducts were baseline resolved in less than 2 minutes,
with a significant improvement in separation efficiency for the
antibody-bound DNA adduct. Using Tris-glycine as the separation
buffer, Joule heating was not excessive at this high field strength
as the current generated was very low (.about.2.2 mA).
[0223] Buffer strength may be varied, with the optimum separation
at 0.5.times.Tris-glycine (12.5 mM Tris/96 mM glycine) in this
Example. The plate count for the bound and unbound adducts using
this buffer condition was calculated to be 6.times.10.sup.5 and
1.times.10.sup.6 plates per meter respectively. Incubation time and
temperature were also investigated. We observed antibody binding to
the DNA adduct at incubation time as short as 1 min and temperature
of incubation as low as 0.degree. C. We found that an incubation
between 5 and 10 min at ambient temperature was suitable for the
formation of complex and for rapid sample analysis.
[0224] To ensure that the DNA remains in its denatured form,
formamide was added to the incubation buffer to prevent the
complementary DNA strands from being renatured during
electrophoresis. Between 2.5 and 12.5% (v/v) formamide, the ratio
of the bound to unbound adducts was relatively constant. Th
concentration of formamide should be kept below 82.5% (v/v).
Example 12
[0225] Using CE/LIFP in a Competitive assay. Because antibodies are
bidentate, each antibody molecule is able to bind with up to two
antigen molecules. One peak in the electropherogram corresponds to
the complex between one antibody and one DNA adduct. A second peak
corresponds to the complex of one antibody with two DNA adduct
molecules. The 1:1 and 1:2 complexes between the antibody and DNA
adducts are well separated, demonstrating high resolution of the CE
system. A competitive assay was performed using the TMR-labeled
BPDE-16 mer as a probe and the unlabeled BPDE-16 mer as a
competitor. As is characteristic of competitive assays, an increase
of BPDE-16 mer (unlabeled competitor) corresponds to the decrease
of the complexes between the fluorescent BPDE-16 mer and the
antibody.
[0226] The BPDE-90 mer that was fluorescently labeled with TMR was
also used as a probe to demonstrate competitive immunoassay
response with the unlabeled BPDE-16 mer. A similar competitive
response was obtained, suggesting that the antibody binds to the
BPDE whether it is present in the 16-mer or the 90-mer
oligonucleotides.
Example 13
[0227] Using CE/LIFP to Determination of BPDE-DNA adducts in A549
cells.
[0228] The competitive immunoassay was applied to the determination
of BPDE adducts in A549 cells that were treated with various doses
of BPDE. The TMR-labeled BPDE-90 mer was used as the probe and the
DNA from A549 cells was heat denatured. Increasing amounts of
BPDE-DNA adducts were formed as the cells were incubated with
increasing concentrations of BPDE for 2 hrs. The BPDE-DNA adducts
compete with the TMR labeled BPDE-90 mer probe for the antibody
binding, resulting in the corresponding decrease of antibody
complexes (peaks 1 and 2) of the fluorescent BPDE-90 mer. As
expected one peak corresponding to the 1:1 complex between the
antibody and the TMR-labeled BPDE-90 mer was observed. A second
peak attributed to the 1:2 complex of antibody with the TMR-labeled
BPDE-90 mer and the DNA adducts from A549 cells was also
observed.
[0229] Using the synthetic BPDE adduct 90-mer as a fluorescent
probe and specific monoclonal antibodies to BPDE-DNA adducts, we
demonstrated a rapid assay for BPDE-modified DNA in a human lung
carcinoma cell line. This approach requires less than 4 min per
separation and has excellent resolving power to separate the bound
and unbound DNA adducts. The same approach may be extended to
assays for other types of DNA damage.
[0230] Described in the next series of examples is an assay that
combines immunological recognition of damaged DNA, capillary
electrophoresis separation, and laser-induced fluorescence
detection (Le, et al., 1998; Xing, et al., 2001). A primary
(1.degree.) mouse monoclonal antibody specific for the DNA lesion
was used to bind to the DNA lesion. A secondary (2.degree.)
anti-mouse IgG antibody that was labeled with a fluorescent dye,
tetramethylrhodamine (TMR), was used to bind with the primary
antibody. The resulting complex of 2.degree.
antibody+1.degree.antibody+damaged DNA was separated using
free-zone capillary electrophoresis and detected with laser-induced
fluorescence. The assay was used to measure thymine glycol, a
typical DNA damage induced by ionizing radiation, and to study DNA
repair (Le, et al., 1998). Subsequently, the assay was extended to
a study of BPDE adducts in DNA from human lung carcinoma cells
(A549) that were incubated with nanomolar concentrations of BPDE
(Xing, et al., 2001).
[0231] Reagents. Unmodified oligonucleotides were synthesized by
the Department of Biochemistry DNA synthesis laboratory, University
of Alberta, or by Integrated DNA Technologies (Coralville, Iowa).
All oligonucleotides were purified by sequencing polyacrylamide gel
electrophoresis prior to use. Purity of the oligonucleotides was
confirmed by .sup.32P-radiolabeling and gel electrophoresis.
Tetramethylrhodamine (TMR)-labeled oligonucleotide was synthesized
by University Core DNA Services, (University of Calgary, AB).
(.+-.)-r-7,t-8-dihydroxy-t-9,10-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene
(anti) [(.+-.)-anti-BPDE] was supplied by the National Cancer
Institute Chemical Carcinogen Reference Standard Repository
(Midwest Research Institute, Kansas City, Mo.). Premixed
polyacrylamide/bisacrylamide (19:1) solution was purchased from
BioRad Laboratories (Cambridge, Mass.). Enzymes were supplied by
Amersham Pharmacia Biotech (Piscataway, N.J.). Monoclonal
antibodies 5D11 and 8E11 were purchased from BD PharMingen (San
Diego, Calif.). Cell supernatant containing monoclonal antibody E5
(Baan, et al., 1988) was kindly provided by Dr. William Watson,
Shell International Chemicals BV, Shell Research and Technology
Center, Amsterdam, Netherlands, and was prepared as described by
Booth (Booth, et al., 1994). Polyclonal mouse IgG antibody was
purchased from Calbiochem (La Jolla, Calif.). Solvents and other
biochemicals were supplied by Sigma Chemical (St. Louis, Mo.),
Fisher Scientific (Pittsburgh, Pa.), or VWR Canlab (Mississauga,
ON, Canada).
[0232] Design of probe. In order to imitate DNA damage as it occurs
naturally in cellular DNA, we designed a 90-base pair
double-stranded oligonucleotide. The desired characteristics of
this oligonucleotide were that it be fluorescently-labeled, contain
a known amount of damage, and be long enough to be recognized by a
variety of antibodies and other DNA-binding proteins. The
oligonucleotide consists of six overlapping, complementary
oligonucleotides of varying lengths that were annealed and ligated
to form a complete double-stranded 90-mer. The oligonucleotide
sequences used in the current study were: oligonucleotide 1:
5'-TMR-labeled-CCTTAAGCTTCCTCAACCACTTACCATACTCGAGATT-3' (SEQ ID
NO:2); oligonucleotide 2: 5'-GAGTAT-GGTAAGTGGTTGAGGAAGCTTAAGG-3'
(SEQ ID NO:6); oligonucleotide 5:
5'-GTCATATGCCGCCTCTGA-CCTTCCTAGAATTCCATCC-3' (SEQ ID NO:8);
oligonucleotide 6: 5'-GGATGGAATTCTAGGAAGGTCAG-AGGCGG-3' (SEQ ID
NO:9). The sequence of oligonucleotide 3 and its complementary
strand (oligonucleotide 4) may be changed to create a variety of
desired damage types, typically with a single damaged nucleotide in
the middle of oligonucleotide 3. In the current study the sequences
used were: oligonucleotide 3: 5'-CCCATTATGCATAACC-3' (SEQ ID NO:5);
oligonucleotide 4: 5'-CATATGACGGTTATGCATAATGGG-AATCTC-3' (SEQ ID
NO:7). The fluorescent Label (oligonucleotide 1) and damaged
nucleotide (oligonucleotide 3) are on the same strand to allow both
double- and single-stranded DNA studies.
[0233] Synthesis of damaged oligonucleotide. (.+-.)-anti-BPDE was
used as the model carcinogen for synthesis of the damaged
oligonucleotide. A 16-mer with the sequence 5'-CCCATTATGCATAACC-3'
(SEQ ID NO:5) was synthesized to encourage maximum yield of the
BPDE-N.sup.2 deoxyguanosine (dG) adduct (Margulis, et al., 1993;
Funk, et al., 1997). The BPDE-oligonucleotide reaction was based on
the procedure described by Margulis (Margulis, et al., 1993), with
slight modifications. The 16-mer was diluted in 20 mM phosphate
buffer, (pH 11), containing 1.5% triethylamine, to a concentration
of 60 mM in a volume of 400 mL. A fresh 3 mM solution of
(.+-.)-anti-BPDE in DMSO was prepared, and 40 mL was added to the
oligonucleotide solution. This corresponded to a
BPDE:oligonucleotide ratio of 5:1. The reaction was carried out at
room temperature for 20 hours in the dark with gentle shaking.
[0234] Purification of BPDE-oligonucleotide. The components in the
BPDE-oligonucleotide reaction mixture were separated using
reversed-phase HPLC. The HPLC system consisted of a Dionex
(Sunnyvale, Calif.) AGP1 advanced gradient pump with
online-degassing module, either an analytical or preparative C18
column, and a Waters (Milford, Mass.) 484 tunable absorbance
detector in series with a Shimadzu RF-551 fluorescence HPLC monitor
(Columbia, Md.). The detectors were connected to a Hewlett Packard
Model 35900 multichannel interface (Palo Alto, Calif.), which
converted the signals for use by a computer running ChemStation
software (Hewlett Packard, Palo Alto, Calif.). Preparative
separation was carried out on a 10.0.times.250 mm, 5 mm Luna C18(2)
preparative column (Phenomenex, Torrance, Calif.). The reaction
products were initially assessed on the analytical column using a
protocol described previously (Margulis, et al., 1993; Cosman, et
al., 1990). This procedure employed a linear 0-90% methanol
gradient in 20 mM sodium phosphate buffer (pH 7.0) in 60 min, with
a flow rate of 0.75 mL/min. To reduce separation times for large
volumes of the reaction mixture, HPLC purification of the
BPDE-16-mer was carried out in two steps. The first separation was
under isocratic conditions, using a mobile phase of 70%
methanol/30% 20 mM sodium phosphate, pH 7.0 buffer and a flow rate
of 0.75 mL/min and 3.5 mL/min for the analytical and preparative
columns, respectively. Elution of products were monitored in series
by the absorbance detector (wavelength=260 nm for DNA) and the
fluorescence detector (excitation wavelength=343 nm, emission
wavelength=400 nm for BPDE). This first separation removed
unreacted BPDE as well as the tetrol hydrolysis products. DNA
fractions were collected, dried using a centrifugal evaporator, and
redissolved in distilled deionized water (ddH.sub.2O). The second
separation consisted of a linear 10-40% methanol/20 mM sodium
phosphate, pH 7.0 buffer gradient in 7.5 min (4%/min) followed by
an additional 5 minutes at 40% methanol. This separated the
BPDE-oligonucleotide from unreacted oligonucleotide.
BPDE-oligonucleotide fractions were collected, dried to remove
methanol and redissolved in ddH.sub.2O. The samples were desalted
using Sep-Pak C18 reversed-phase columns (Waters). The sample was
applied to a prepared Sep-Pak cartridge, then washed with 10 mL of
the following solutions: 25 mM ammonium bicarbonate (pH 8.0); 25 mM
ammonium bicarbonate/5% acetonitrile; H.sub.2O/5% acetonitrile;
H.sub.2O/5% acetonitrile. The BPDE-oligonucleotide was then eluted
with 4.times.1 mL of H.sub.2O/30% acetonitrile, dried and
redissolved in ddH.sub.2O.
[0235] Synthesis and purification of 90-mer oligonucleotides. Prior
to ligation with the other 5 oligonucleotides, it was necessary to
phosphorylate the freshly purified BPDE-16-mer at the 5'-end.
Reaction mixtures included: .about.200 pmol of BPDE-16-mer or
control 16-mer, 4 mL of 100 mM ATP (400 pmol), 1.2 mL of 10.times.
polynucleotide kinase reaction buffer, and ddH.sub.2O to a total
volume of 12 mL. T4 polynucleotide kinase (PNK) was added (1 mL,
6.1 units/mL), then samples were mixed and incubated at 37.degree.
C. for 1 hour. After complete reaction, the excess PNK was heat
denatured at 70.degree. C. for 10 minutes. The 16-mers were then
mixed with the TMR-labeled 37-mer and the other 4 oligonucleotides
so that all would be in 2:1 excess over the 16-mers. 5.times.DNA
ligase buffer was added to a final concentration of 1.times. and
the mixture was heated in a water bath to 70.degree. C. for 10
minutes, then allowed to cool over several hours to room
temperature. DNA ligase was added (2 mL, 8.5 Weiss units/mL) and
the sample incubated overnight at 16.degree. C.
[0236] Purification of the BPDE and control ligation products was
achieved using preparative, 7.5% native polyacrylamide gel
electrophoresis (PAGE). Blectrophoresis was carried out at 600 V
for 6 hours with a water cooling core to prevent denaturation of
the ligation products. The bands were visualized by brief exposure
to ultraviolet light, causing the TMR label to fluoresce, and cut
from the gel. The gel slices were crushed and soaked to elute the
products overnight in 0.3 M sodium acetate, pH 5.2 on a rotary
shaker protected from light. After elution, polyacrylamide
fragments were removed from solution using filter units prepared in
the lab. The solution was passed through silanized glass wool
followed by GF/C glass microfibre filter paper (Whatman). The
samples were then extracted and back-extracted with equal volumes
of phenol/chloroform/isoamyl alcohol (25:24:1) followed by
chloroform/isoamyl alcohol (24:1). Oligonucleotides were
precipitated by adding MgCl.sub.2 to 10 mM and 3 volumes of
ice-cold 95% ethanol and, then placed at -20.degree. C. overnight.
The following day samples were centrifuged for 45 minutes at 14000
rpm and 4.degree. C., supernatant was removed, and the pellets were
washed once with 95% ethanol. Samples were again centrifuged for 10
minutes, dried and redissolved in ddH.sub.2O. UV-Vis absorbance
scans were performed on the resulting oligonucleotide solutions to
determine concentration as well as to confirm the presence of the
TMR dye and BPDE moiety.
[0237] Instrumentation for analysis of ligation products, was the
same as described in examples 11-13. with the following
modification. The system was equipped with an auxiliary microscope
to assist in the alignment of the optics. The microscope was used
to visualize the position of the laser beam with respect to both
the sample flow through the capillary and the collection optics,
represented by a light-emitting diode (LED) positioned behind the
pinhole in the collection assembly. Alignment was achieved by
initially fixing the position of the collection assembly, then
adjusting the capillary and laser-focusing objective using X-Y-Z
translation stages. The angle of the fluorescence-collecting
objective and the position of the collection assembly were also
adjustable for optimization of alignment.
[0238] Samples were electrokinetically injected into the capillary
by applying an injection voltage of 10000 V for 5 seconds. The
separation was carried out at room temperature with a separation
voltage of 20000 V. The running buffer used was
1.times.Tris-glycine (25 mM Tris, 250 mM glycine), pH 8.3. The
capillary was washed approximately every 5-10 injections with 0.1 M
NaOH (applied by syringe for 1 min) followed by electrophoresis
using 1.times. Tris-glycine, pH 8.3 for 7 minutes. The initial
voltage was kept low to prevent excessive joule heating in the
capillary. As the running buffer replaced the NaOH in the
capillary, current decreased allowing the running voltage to be
gradually increased to 20000 V for the final 5 minutes of the
reconditioning period. All capillary electrophoresis data were
analyzed using Igor Pro software (version 3.1, WaveMetrics Inc.,
Lake Oswego, Oreg.).
[0239] Characterization of BPDE and control 90-mers. Prior to
analysis, 90-mer samples were diluted to appropriate concentrations
in running buffer (1.times. Tris-glycine, pH 8.3). The 90-mer
products were analyzed either in their native form or their
denatured, single-stranded form. Denaturation of the 90-mers was
achieved by heating the samples at 100.degree. C. for 10 minutes in
a heating block, then transferring directly to ice to prevent
reannealing. After cooling, the samples were briefly centrifuged in
a microcentrifuge to collect condensation from the side of the
tube, then gently mixed to ensure a homogenous solution. Total
sample volume was typically 20 mL, which allowed for convenient
injection into the capillary. For experiments involving antibodies,
fresh dilutions of antibody stock solutions were prepared
immediately before analysis and kept on ice. After addition of
antibody to the 90-mer solution, the sample was gently vortexed to
ensure complete mixing.
[0240] Treatment of A549 cells with BPDE. A human lung carcinoma
cell line (A549) was incubated with BPDE to produce DNA adducts in
genomic DNA. The cell line was maintained in DMEM/F12 medium (Gibco
BRL, Gaithersburg, Md.) supplemented with 10% fetal bovine serum.
The cells were seeded at 1.times.10.sup.5 cells per plate and
maintained at 95% humidity and 5% CO.sub.2 for 20 hours prior to
the addition of BPDE. Old culture media were removed from each
culture plate and the cells were washed twice with phosphate
buffered saline (PBS). Media containing BPDE at various
concentrations (0, 2.5, 5, and 10 mM final concentration) were
added to the designated plates. The cells were further incubated in
the media containing BPDE for 2 hours. The cells were then washed
with PBS prior to the addition of DNAzol lysis reagent (Gibco BRL)
to facilitate cell lysis and DNA extraction. Subsequent steps
involved a 99.9% ice cold ethanol precipitation and a 70% cold
ethanol wash to purify the genomic DNA. The final DNA pellet was
dissolved in distilled deionized water (ddH.sub.2O) and DNA
concentration was measured at OD.sub.260 using ddH.sub.2O as a
blank.
[0241] Competitive assay for BPDE-DNA adducts. The DNA samples from
the A549 cells were analyzed for BPDE-DNA adducts by competitive
assay using the TMR-labeled 16-mer or 90-mer oligonucleotides as
probes. Mixtures containing 60 nM of the oligonucleotide probe, 0.4
mg/mL of mouse monoclonal antibody 8E11, and 80 mg/mL of the DNA
from A549 cells were incubated in 20 mL of tris-glycine buffer (25
mM tris and 200 mM glycine, pH 8.3) at room temperature for 30 min.
These were subjected to CE/LIF analysis to detect both
antibody-bound and unbound fluorescent probes.
Example 14
[0242] Using CE/LIF to determine Affinity interactions of
BPDE-90-mers with a monoclonal antibody. The purification of
BPDE-16-mer oligonucleotide, the synthesis and purification of BPDE
90-mer ligation products, and the characterization of the BPDE
90-mer ligation products were carried out as known to those skilled
in the art.
[0243] Preliminary experiments using monoclonal antibody 8E11
demonstrated that the specific antibody bound to the BPDE-90 mer,
not the control 90 mer. These results were obtained by first
denaturing the 90-mers (5.times.10.sup.-9 M), then adding 8E11
antibody to a final concentration of 20 mg/mL and incubating for 10
min at room temperature (21.degree. C.). The same fluorescence
intensity scale was used for both 90-mers for ease of comparison.
For the mixture of the BPDE 90-mer and 8E11, an additional peak was
present in the electropherogram with a migration time of
approximately 3.0 min. This peak represented the complex between
the antibody and single-stranded BPDE-DNA, and was well-resolved
from the denatured 90-mer peak at 4.1 min. When comparing the
fluorescent signals between runs, the total area of the two peaks
for the mixture of BPDE 90-mer and 8E11 was very similar to the
area of the peak for the BPDE 90-mer alone. The formation of an
antibody-DNA complex was not observed with the control 90-mer,
indicating a specific interaction of the antibody with the BPDE
90-mer.
[0244] The effect of incubation time and temperature on complex
formation were investigated using the same concentrations of BPDE
90-mer (5.times.10.sup.-9 M) and 8E11(20 mg/mL). For incubations
carried out at both room temperature and on ice, the interaction
did not change significantly between 1 min and 20 min. At room
temperature, the complex was stable after 45 min. For incubation on
ice, the complex decreased slightly after 45 min when compared to
the 20 min incubation. In general, room temperature incubations
with 8E11 resulted in more stable and reproducible complex
formation than incubations on ice. This result is expected since
the recommended temperature for conventional immunoassays using
8E11 is 37.degree. C. (Santella, et al., 1984; Hsu, et al., 1995),
and most inumunochemical procedures require incubation temperatures
of either 37.degree. C. or room temperature. Based on these
results, further experiments with 8E11 antibody were carried out at
room temperature. An incubation time of 5 min was chosen for ease
of sample preparation and analysis.
[0245] Overnight incubations resulted in a decrease of the
DNA-antibody complex, as well as a reversion to the doublet shape
for the free 90-mer peak. This result suggests that 90-mer samples
left overnight tended to re-anneal to the double-stranded form,
causing dissociation of the DNA-antibody complex. This also implies
that the affinity of 8E11 for double-stranded BPDE-DNA is less than
for single-stranded BPDE-DNA.
[0246] The difference in affinity of 8E11 antibody between single-
and double-stranded BPDE-modified DNA was further confirmed by
comparing its binding with heat-denatured BPDE 90-mer and native
BPDE-90 mer (20 mg/mL 8E11). Antibody-oligonucleotide complex
formation was approximately 6 fold higher for the denatured
single-stranded 90-mer than the native form. Thus, denaturation of
samples by heat before incubation with antibodies was retained for
further experiments.
Example 15
[0247] Using CE/LIF to Determination of specific antibody using
BPDE-90 mer as a probe. An application of the fluorescent BPDE-90
mer probe was demonstrated for the determination of anti-BPDE
antibody. Calibration from the analyses of mixtures containing
different amounts of 8E11 and a constant concentration of the
detatured BPDE-90 mer probe (5.times.10.sup.-9 M) were made. A
DNA-antibody complex peak was observed with 8E11 concentrations as
low as 0.1 mg/mL. This concentration corresponds to
0.7.times.10.sup.-9 M (or 0.7 nM) assuming a molecular weight of
approximately 150,000 for the antibody 8E11. The concentration of
BPDE-90 mer (5 nM) was in excess and the formation of its complex
with the antibody was not complete. The amount of the complex
increased at higher concentrations of 8E11, up to 10 mg/ml (7 nM).
At this concentration complex formation appeared to reach
saturation, since further increase of antibody concentrations did
not increase the proportion of 90-mer bound to 8E11.
Example 16
[0248] Using CE/LIF for Screening for anti-BPDE antibodies using
the fluorescent BPDE-90mer probe. The fluorescent BPDE-90 mer probe
was further used to screen for specific binding proteins, with 3
antibodies as model protein analytes. Monoclonal antibodies 8E11,
5D11 and E5 are all specific for BPDE-modified DNA. A comparison
between these antibodies was conducted to determine differences in
their reactivity to the BPDE 90-mer standard as well as their
behavior in the capillary electrophoresis system. Conditions used
for sample preparation were identical to earlier experiments: heat
denaturation of the 90-mer at 100.degree. C. for 10 min, cooling on
ice, then incubation with antibody at room temperature for 5 min
before injection. Polyclonal mouse IgG was used as a negative
control since it is essentially the same molecular structure
(isotype) as the monoclonal antibodies but is not expected to react
with the BPDE 90-mer. The BPDE 90-mer probe concentration was fixed
at 5.times.10.sup.-9 M and the antibodies were added in varying
amounts. All three monoclonal antibodies reacted with the 90-mer
probe, with 8E11 giving the highest formation of complex. The
negative control showed a very slight reactivity but was
insignificant compared to the other antibodies, even at
concentrations up to 40 mg/mL.
[0249] Antibodies 8E11 and E5 were found to bind specifically to
the BPDE adduct. No cross-reactivity with the unmodified control
90-mer was observed for either 8E11 or E5. The antibody 5D11 showed
slight cross-reaction with undamaged DNA. When incubated with 20
.mu.g/mL 5D11, the control 90-mer formed a peak corresponding to
antibody complex, with about 2.1% of total peak areas as compared
with the BPDE 90-mer. This non-specific interaction between 5D11
and undamaged DNA is in agreement with previous studies (Santella,
et al., 1984) that have demonstrated cross-reactivity, and is a
result of its being raised against a full-length BPDE-DNA antigen.
Both 8E11 and E5 were raised against BPDE-guanosine monomers
conjugated to carrier proteins (Baan, et al., 1988; Santella, et
al., 1984) and therefore do not recognize undamaged DNA.
[0250] The incomplete binding of the DNA damage probe with the
antibodies (up to 50% of binding) is probably because the probe is
a mixture of several BPDE-90 mer isomers. The stereochemistry of
the BPDE-N.sup.2-dG adduct could be important to its binding with
specific antibodies. In the preparation of the BPDE-modified
16-mer, (.+-.)-anti-BPDE was reacted with the oligonucleotide. The
covalent bond that forms between BPDE and guanosine may be either
cis- or trans-relative to the hydroxyl group on the adjacent carbon
atom. Therefore, there may be as many as four different
configurations of the BPDE 16-mer: (+)-trans, (+)-cis, (-)-trans,
and (-)-cis (2). The reaction protocol was designed to minimize the
formation of cis-adducts (Funk, et al., 1997), but a mixture of
(+)-trans and (-)-trans adducts with a small amount of cis adducts
would be expected in the BPDE 16-mer reaction products (Cosman, et
al., 1990). Because these stereoisomers were pooled together after
purification by HPLC and before the ligation reaction, the 90-mer
product would also contain these configurations. The advantage of
this mixture is that it more accurately represents the spectrum of
damage that would occur in human DNA samples. The disadvantage is
that BPDE-DNA antibodies exhibit different affinities for these
stereoisomers (Hsu, et al., 1995). In competitive inhibition
studies using BPDE-modified 11-mers, Hsu (Hsu, et al., 1995)
demonstrated a lower affinity for the
(-)-trans-anti-BPDE-N.sup.2-dG adduct than for the
(+)-trans-anti-BPDE-N.sup.2-dG adduct. For antibodies 8E11 and 5D11
this lower affinity was 66% and 20% of the (+)-trans adduct,
respectively. Both antibodies exhibited much lower affinities for
the cis adducts compared to the (-)-trans adduct. Since the 90-mer
contained a combination of both trans adducts, the stereospecific
difference in affinity may in part be responsible for the
differences in complex formation observed for these antibodies. The
presence of different BPDE-90mer isomers may also contribute to the
observed incomplete binding. The other possible reason for the
incomplete binding is the presence of residual oligonucleotides
that do not contain BPDE and therefore, do not bind to the
antibodies.
[0251] In addition to the isomer-specific reactivities, Hsu (Hsu,
et al., 1995) showed a difference in affinity between 8E11 and 5D11
when considering only the (+)-trans adduct. 8E11 was approximately
7 times more sensitive than 5D11 for the very short 11-mer
oligonucleotide. For full-length heat-denatured BPDE-DNA, the two
antibodies were almost identical. This difference is likely due to
the antigens against which these antibodies were raised:
BPDE-N.sup.2-dG mononucleotide for 8E11, full-length BPDE-DNA for
5D 11. 5D 11 may require a longer sequence of DNA surrounding the
damaged site for binding which would not be present in the 11-mer.
Given these results one might predict that for DNA of intermediate
length (90 bases), 8E11 would still have a higher affinity than 5D
11, but to a lesser extent. These results are consistent with
previous findings, which indicates that monoclonal antibody 8E11 is
likely the best choice for detecting BPDE-damaged DNA using the
capillary electrophoresis/laser-induced fluorescence assay.
Example 17
[0252] Using CE/LIF and the BPDE-DNA probe in a competitive assay
for BPDE-DNA adducts in cells. The 90-mer probe described herein
has many potential uses in DNA damage research. It enables the
investigation of alternative assay methods, including CE-based
competitive immunoassays (Tao, et al., 1996; Ye, et al., 1998; Lam,
et al., 1999; Wan, et al., 1999) using the probe as a fluorescent
probe (competitor). This approach is based on competition between
damaged DNA and the fluorescent probe for the binding sites of a
limited amount of antibody. With little or no damaged DNA in a
sample, the probe achieves maximum complex formation with the
antibody. As the amount of damaged DNA in the sample mixture
increases, the probe is displaced from the antibody. This would
result in an increase in the free probe peak and a decrease in the
probe-antibody complex peak. This method has been demonstrated by
using oligonucleotide and genomic DNA containing BPDE-damaged sites
(Tan, et al., 2001). Electropherograms from the analysis of
BPDE-DNA adducts in A549 cells that were incubated with 2.5, 5, and
10 mM BPDE for 2 hr were collected. Again, increasing amounts of
BPDE-DNA adducts were formed as the cells were incubated with
increasing concentrations of BPDE. The BPDE-DNA adducts compete
with the TMR labeled BPDE-DNA adduct probe for the antibody
binding, resulting in the corresponding increase of the unbound
probe (peak 3) and decrease of antibody complexes (peaks 1 and 2)
of the fluorescent probe. This analysis requires less than 4 min
per separation and has excellent resolving power to separate the
bound and unbound DNA adducts. The same approach may be extended to
assays for other types of DNA damage.
[0253] Another important aspect of the probe's design is the
flexibility to substitute different damage types in the molecule
with relative ease. The sequences of the two center
oligonucleotides may be changed depending on the desired
modification. By inserting these different damaged oligos, a
variety of DNA damage detection systems can be investigated using
the corresponding damage probe and CE/LIF. The technique itself
combines specific recognition with high sensitivity detection,
minimal sample preparation, and fast analysis times (5 minutes per
run).
Example 18
[0254] Using CE/LIF to Determine Stoichiometry of antibody binding
with TMR-BPDE-16-mer (16mer*) oligonucleotide. In this example
CE/LIF is used to determine the binding stoichiometry of DNA
adducts with antibodies. Advantage is taken of the fact that both
size and charge of the molecules contribute to CE separation. If
additional charges can be introduced to the complex due to binding,
then the separation of the multiple complexes becomes possible.
Fluorescent oligonucleotide probes that contain a single adduct
which can be recognized by an antibody were designed. These probes
introduce large mobility changes to the antibody when bound to the
probe because of the highly negative charge of the probe. With
these probes, we are able to study the binding stoichiometry
between oligonucleotides and the antibody. DNA adducts of
benzo[a]pyrene diol epoxide (BPDE) were looked at in this example.
Available monoclonal IgG antibody has a high affinity for BPDE-DNA
adducts, allowing detailed information on binding stoichiometry
between the antibody and the DNA adducts to be obtained. This
example provides direct information on antibody binding
stoichiometry.
[0255] Reagents: Oligonuleotides were synthesized by the Department
of Biochemistry DNA synthesis laboratory, University of Alberta, or
by Integrated DNA Technologies (Coralville, Iowa). All
oligonucleotides were purified by sequencing polyacrylamide gel
electrophoresis prior to use. Purity of the modified
oligonucleotieds was confirmed by gel electrophoresis and
.sup.32P-postlabeling. Tetramethylrhodamine (TMR)-labeled
oligonucleotide was synthesized by University Core DNA Services,
(University of Calgary, AB). (.+-.)-r-7,t-8-dihydroxy-t-9,10-ep-
oxy-7,8,9,10-tetrahydrobenzo[a]pyrene [(.+-.)-anti-BPDE] was
supplied by the National Cancer Institute Chemical Carcinogen
Reference Standard Repository (Midwest Research Institute, Kansas
City, Mo.). Mouse monoclonal antibody 8E11 was purchased from BD
PharMingen (San Diego, Calif.). Polyclonal rabbit IgG antibody was
purchased from Calbiochem (La Jolla, Calif.). Solvents and other
biochemicals were supplied by Sigma (St. Louis, Mo.), Fisher
Scientific (Pittsburgh, Pa.), or VWR Canlab (Mississauga,
Ontario).
[0256] Synthesis of BPDE-DNA adducts: Two 16-mers with the sequence
5'-CCCATTATGCATAACC-3' (SEQ ID NO:5) were synthesized and reacted
with BPDE to yield the BPDE-N.sup.2 deoxyguanosine (dG) adduct
(Margulis, et al., 1993; Funk, et al., 1997). One of the 16-mer
oligonucleotides was labeled with TMR at the 5' end, and the other
was not labeled. The formation of BPDE-oligonucleotide was based on
the procedure described by Margulis (Margulis, et al., 1993) with
slight modifications. The 16-mer was diluted in 20 mM phosphate
buffer (pH 11) containing 1.5% triethylamine, to a concentration of
60 mM in a volume of 400 mL. To the oligonucleotide solution was
added 40 mL 3 mM BPDE in DMSO. This corresponded to a
BPDE:oligonucleotide ratio of 5:1. The reaction was carried out at
room temperature for 20 hours, in the dark with gentle shaking. The
double stranded TMR-BPDE-90-mer and BPDE-90-mer were constructed
through ligation of BPDE-16-mer with five other
oligonucleotides.
[0257] Purification of TMR-BPDE-oligonucleotide: The components in
the TMR-BPDE-oligonucleotide reaction mixture were separated using
reversed-phase HPLC. The HPLC system consisted of a Dionex
(Sunnyvale, Calif.) AGP1 advanced gradient pump with online
degassing module, an analytical C18 column, a Waters (Milford,
Mass.) 484 tunable absorbance detector in series with a Shimadzu
(Tokyo, Japan) RF-551 fluorescence detector. The detectors were
connected to a Hewlett Packard (Palo Alto, Calif.) Model 35900
multichannel interface, which converted the signals for use by a
computer running ChemStation software (Hewlett Packard). The
analyses were performed using a Luna C18(2) analytical column
(4.6.times.250 mm, 5 mm, Phenomenex, Torrance, Calif.). The
reaction products were purified initially using a gradient elution.
This procedure employed 10 mM sodium phosphate buffer (pH 7.0) and
acetonitrile. The acetonitrile content was initially 10%, linearly
ramped to 15% during the first 20 min, and then kept at 15% for 5
min. After the BIPDE-modified oligonucleotides were eluted and
collected, the column was washed using 50% acetonitrile for 20 min
to remove unreacted BPDE and its metabolites. The flow rate was 1.0
mL/min. Elution of products were monitored in series by an
absorbance detector (wavelength=260 nm for DNA) and a fluorescence
detector (excitation wavelength=535 nm, emission wavelength=580 nm
for TMR). The fractions containing the BPDE-oligonucleotide were
pooled and further purified to remove any unmodified
oligonucleotides. This second HPLC purification step was carried
out using an isocratic elution with 10 mM phosphate buffer (pH
7.0), containing 12.5% acetonitrile as mobile phase. Other HPLC
conditions were the same as described above. The collected
fractions were concentrated by pressurized air under room
temperature and dissolved in distilled deionized water
(ddH.sub.2O). The purified BPDE-oligonucleotides were evaluated
using HPLC under isocratic elution conditions, and showed good
purity of above 95%. The concentration of the final TMR-BPDE-16-mer
was estimated using absorbance at 260 nm.
[0258] Instrumentation for analysis of the DNA-BPDE adducts:
Analysis and characterization of the DNA-BPDE adducts was carried
out using a laboratory-built capillary electrophoresis laser
induced fluorescence (CE/LIF) system as described in examples 13
on.
[0259] Samples were electrokinetically injected into the capillary
by applying an injection voltage of 15 kV for 5-10 seconds. The
separation was carried out at room temperature with a separation
voltage of 15 kV. The running buffer were either 1.times.
tris-glycine (25 mM Tris, 192 mM glycine, pH 8.3) or
0.5.times.tris-glycine (12.5 mM tris and 96 mM glycine, pH 8.3).
The capillary was washed approximately every 3 injections with 0.02
M NaOH electrophoretically at 15 kV for 7 min followed by
electrophoresis using water and the running buffer for 7 min each.
All capillary electrophoresis data were analyzed using Igor Pro
software (version 3.1, WaveMetrics Inc., Lake Oswego, Oreg.).
[0260] Complex formation of TMR-labeled BPDE-DNA adducts and
antibody: TMR-BPDE-16-mer and TMR-BPDE-90-mer samples were diluted
to appropriate concentrations in running buffer (tris-glycine, pH
8.3). The double stranded BPDE-90-mer was denatured by heating at
95.degree. C. for 5 min in a heating block. It was then placed on
ice to prevent reannealing. After cooling, the samples were briefly
centrifuged in a microcentrifuge (Z233M, Hermle) to collect
condensation from the side of the tube, then gently mixed to ensure
a homogenous solution. TMR-BPDE-16-mer was synthesized as a
single-stranded oligonucleotide. Appropriate dilutions of antibody
stock solutions were prepared immediately before use and kept on
ice. After addition of antibody to the BPDE-oligonucleotide
solutions, the samples were gently vortexed to ensure complete
mixing and incubated at the room temperature for 5-10 min, then
analyzed by CE/LIF. The total sample volume was typically 20
mL.
[0261] Simultaneous binding of two BPDE-DNA adducts to the
antibody: Both TMR-labeled and unlabeled BPDE-adducts were allowed
to bind with the antibody. The freshly diluted BPDE-16-mer (16mer)
and TMR-BPDE-16-mer (16mer*) solutions were mixed together before
addition of the antibody. Concentrations of the TMR-BPDE-16-mer and
the antibody were kept constant at 9.6 nM and 33.3 nM,
respectively, and the unlabeled BPDE-16-mer was varied from 2.5 nM
to 5.0 mM. The sample mixtures were incubated for 5 min at room
temperature, then analyzed by CE/LIF. Similarly, the antibody was
added to mixtures of the TMR-BPDE-16mer and BPDE-90mer to study the
competitive binding and ligand exchange.
[0262] A series of electropherograms from CE/LIF analyses of
mixtures containing 24 nM TMR-BPDE-16-mer (16mer*) and varying
concentrations of mouse monoclonal antibody (Ab) to BPDE, from 0.5
mg/mL to 16.0 mg/mL were collected. In these mixtures, 2 complexes
between the Ab and the TMR-labeled BPDE-16-mer (16mer*) can be
expected as follows:
Ab+16mer*=Ab(16mer*)+Ab(16mer*).sub.2 (9)
[0263] The free (unbound) 16mer* gives a peak that was well
resolved from the two complexes. The unbound 16mer* oligonucleotide
and the antibody-bound 16mer* complexes are negatively charged.
Under the free zone CE separation conditions, the direction of
their electrophoretic mobility (.mu..sub.ep) is opposite to that of
the electroosmotic flow (EOF). Among the three fluorescent species
the unbound 16mer* has the highest negative effective charge. It
has the highest electrophoretic mobility towards the positive
(injection) end, and thus the longest migration time (3.4 min).
When the 16mer* binds to an antibody molecule, the reduction of the
effective charge results in a smaller electrophoretic mobility and
thus a shorter migration time (2.1 min). When the Ab binds to two
16mer* molecules, the charge density of the complex is between
those of the Ab(16mer*) complex and the unbound 16mer*. Therefore,
a smaller mobility shift is expected. Indeed, we observed the
Ab(16mer*).sub.2 complex at 2.4 min. The two complexes are well
resolved from each other, (resolution of 1.55).
[0264] It is evident that formation of the two complexes depends on
the relative concentrations of the TMR-BPDE-16-mer oligonucleotide
(16mer*) and the antibody. The intensity of Ab(16mer*) complex (1:1
stoichiometry, peak 1) increases with increasing concentration of
the antibody, whereas the Ab(16mer*).sub.2 complex (1:2
stoichiometry, peak 2) reaches a maximum at an antibody
concentration of 1.0 .mu.g/mL and then decreases gradually with
increasing concentrations of the antibody.
[0265] The primary complex of the 1:1 stoichiometry increases with
increasing concentration of antibody. When the concentration of the
antibody is 8-16 ,.mu.g/mL, approximately 80-85% of the total
16mer* oligonucleotide is present as the primary complex with the
antibody (1:1 stoichiometry). Formation of the secondary complex,
Ab(16mer*).sub.2 (1:2 stoichiometry) is favored at lower
concentrations of the antibody, when the 16mer* oligonucleotide is
in excess. The amount of the secondary complex reaches a maximum at
the antibody concentration of 1 .mu.g/mL, when 50% of the total
16mer* is present as the secondary complex. As the molecular weight
of the mouse monoclonal antibody IgG is approximately 150,000 Da,
an antibody concentration of 1 .mu.g/mL is approximately 7 nM.
Although the Ab(16mer*).sub.2 complex is observed throughout the
entire antibody concentration range studied, from 0.02 .mu.g/mL
(.about.0.14 nM) to 16 .mu.g/mL (.about.100 nM), it is predominant
only when the antibody concentration is below 2 .mu.g/mL (.about.14
nM) and is lower than the concentration of the 16mer* (24 nM). At
similar concentrations of antibody (4 .mu.g/mL or 27 nM) and the
16mer* (24 nM), the primary complex dominants, suggesting that the
complex with one binding site is preferred.
[0266] To further study the secondary complex, varying
concentrations of the 16mer* (2.4-19.2 nM) were mixed with a fixed
concentration of the antibody (0.4 .mu.g/mL or .about.2.7 nM), and
the mixtures were analyzed using CE/LIF. Fluorescence intensity of
the secondary and primary complexes as a function of the
concentration of the fluorescently labeled BPDE-16-mer (16mer*)
were measured. At a lower concentration of the 16mer* (2.4 nM) than
the antibody concentration (2.7 nM), the peak intensity of the
primary complex is comparable to that of the secondary complex.
When the 16mer* concentration is higher (4.8-19.2 nM) than the
antibody concentration (.about.2.7 nM), the secondary complex
dominates. The fluorescence intensity of the secondary complex
increases with increasing amounts of the 16mer* until it reaches a
plateau at approximately 17 nM when the amount of the antibody
presumably becomes the limiting factor.
[0267] Most previous studies on immunoassays were not able to
address the binding stoichiometry although multiple complexes might
have formed. The present study clearly shows the formation of
primary and secondary complexes. The behavior may be qualitatively
explained by a two binding site model. The mouse monoclonal
antibody is an IgG, which has two specific binding sites for
antigen (hapten), in this case, BPDE-16-mer oligonucleotide. When
an antibody is in excess of antigen, there are more available
binding sites for antigen, thus most complex exists as the primary
complex (1:1 stoichiometry). Only when the binding sites are
limited as in the case of lower antibody concentration, the
secondary complex (1:2 stoichiometry) dominates. These results
suggest that binding to the second sites of antibody is less
favored probably because of possible steric hindrance by the
binding on the first site.
Example 19
[0268] Using CE/LIF to Determine the Binding of the antibody with
TMR-BPDE-90-mer (90mer*): To confirm the preferential binding to
the first site and the possible hindrance to the secondary binding,
we further compared binding of the antibody with a TMR-BPDE-90-mer
(90mer*) oligonucleotide. The 90mer* is approximately 28,000 Da,
about five times higher than the 16mer* (.about.5,000 Da).
Eelectropherograms from CE/LIF analyses of samples containing 50 nM
90mer* incubated with varying concentrations of mouse monoclonal
antibody, 0-20 .mu.g/mL (0-140 nM) were collected. In the absence
of antibody, the 90mer* gave a single peak at migration time of 3.2
min. After incubation with the antibody, two additional peaks are
present in the electropherogram with migration times of 2.41 and
2.60 min. These peaks represent the complexes between the antibody
and single stranded 90mer* oligonucleotide. Because of the
electrophoretic mobility shift, they are well resolved from the
unbound 90mer*. Despite the small difference in migration time
between the two complexes (0.19 min), these two complexes are
baseline resolved. The first peak is due to the primary complex of
the antibody and DNA adduct with 1:1 stoichiometry, and peak 2 is
due to the secondary complex of 1:2 stoichiometry.
[0269] The binding of the anti-BPDE antibody with TMR-BPDE-90-mer
appears to be weaker than the binding with TMR-BPDE-16-mer.
Approximately 60% of the total 90mer* (50 nM) formed complex with
the antibody when the concentration of antibody was .about.140 nM.
This percentage is lower than that for the 16mer* where 80-85% of
the total 16mer* (24 nM) complexed with the antibody (50-100 nM).
This is not surprising considering that the antibody (mouse
monoclonal, 8E11) was raised against BPDE-adduct of mononucleotide
and perhaps has a higher affinity to shorter stretches of DNA.
[0270] The ratio of the primary and secondary complexes was also
found to depend on the relative concentrations of the antibody and
the 90mer*, similar to that shown in the 16mer* experiments
described above. However, the secondary complex of the antibody
with the 90mer* is less stable than the secondary complex of the
same antibody with the 16mer*. The primary complexes of the
antibody with both the 90mer* and the 16mer* are much more stable.
The results support our suggestion that the primary binding is
stronger than the secondary binding. Despite the identical sequence
and structure of the two binding sites of the monoclonal IgG
antibody, it is likely that the secondary binding is affected by
the primary binding on the first site. This may be caused by a
conformational change of the antibody structure after binding to
one site. Alternatively, steric hindrance of the molecule bound to
one site of the antibody may affect the binding of another molecule
on the second site of the same antibody molecule. A comparison of
our results on the 16mer* and 90mer* indicates that the secondary
binding with the larger 90mer* is less favored. These results
suggest that steric hindrance plays an important role in the
formation of the secondary complex between an antibody and an
antigen.
Example 20
[0271] Using CE/LIF to measure competitive binding of antibody with
TMR-BPDE-16-mer and unlabeled BPDE-16-mer: The commonly used
competitive immunoassays are based on competitive binding of two
ligands to a limiting amount of antibody. Typically, one ligand is
labeled and is used as a detection probe. The target analyte is
usually the unlabeled ligand. In the traditional competitive assay
format, the analyte (unlabeled ligand) can only be indirectly
determined through monitoring the relative intensity of signals
produced from the labeled ligand. In this study, It was found that
the primary and secondary complexes of the antibody with the
ligands can be separated. Thus, we decided to further study the
binding of multiple ligands to the antibody, with a possibility of
developing new approaches to binding assays.
[0272] Electropherograms from CE/LIF analyses of mixtures
containing 33 nM antibody, 35 nM TMR-BPDE-16-mer (16mer*), and
varying concentrations (0, 0.05, 0.5 and 5.0 mM) of unlabeled
BPDE-16-mer (16mer) were collected. In the absence of the unlabeled
BPDE-16-mer, the primary complex [Ab(16mer*)] dominates and its
fluorescence intensity is much higher than that of the secondary
complex [Ab(16mer*).sub.2].
[0273] While the primary complex decreases with increasing
concentrations of the competing unlabeled 16mer, a typical
competitive immunoassay behavior, the secondary complex increases
initially with increasing concentrations of the 16mer. The latter
behavior is unique and has not been reported previously with
competitive immunoassays. This observation can be explained as
following.
[0274] The present CE/LIF technique allows the separation of two
antibody complexes when a labeled ligand (L*) (e.g., 16mer*) is
mixed with the antibody.
Ab+L*=AbL*+AbL*.sub.2 (10)
[0275] To allow for detection of an unlabeled ligand (L),
competitive immunoassay approaches rely on the competition of
unlabeled ligand (L) with the labeled ligand (L*) for the limiting
amount of antibody (Ab). Following species may be formed:
AbL*+L=AbL+L* (11)
AbL*+L=AbL*L (12)
AbL*.sub.2+L=AbL*L+L* (13)
AbL*.sub.2+2L=AbL.sub.2+2L* (14)
[0276] The species that are fluorescent and can be detected include
the free ligand L*, the primary complex AbL*, and the secondary
complexes AbL*L and AbL.sub.2*. It is clear that the end products
are a redistribution of L* and L in the complexes although L* and L
compete for the same antibody.
[0277] The relative changes of the primary and secondary complexes
with varying concentrations of unlabeled 16mer when it is incubated
with 9.6 nM TMR-labeled 16mer* and 33 nM (5 .mu.g/mL) antibody were
measured. When the concentration of the unlabeled 16mer is below 10
nM, the primary complex [Ab(16mer*)] dominates and its amount
remains almost constant with increasing concentration of the
unlabeled 16mer up to 10 nM. The total ligand concentration (16mer*
and 16mer) is lower compared with that of the antibody, and there
are excess antibody binding sites available. Thus, no significant
competition between the two ligands takes place. Increasing the
competing 16mer concentration from 10 nM to 200 nM result in the
reduction of the primary complex in a manner similar to that
commonly observed for conventional competitive assay. Further
increase of the unlabeled 16mer concentration to above 250 nM
results in disappearance of the primary Ab(16mer*) complex,
probably due to the displacement of the 16mer* by the excess
16mer:
Ab(16mer*)+16mer=Ab(16mer)+16mer* (15)
[0278] The secondary complexes initially increases with increasing
concentration of the unlabeled 16mer, and reaches a maximum when
the concentration of the 16mer is 100 nM. This behavior is
different from that of traditional competitive immunoassays where
only the mixture of antibody complexes are commonly detected and
the separation of the 1:1 and 1:2 complexes is not available for
examination. Further increase of the 16mer concentration (100-1000
nM) results in a gradual decrease of the secondary complex. Only at
this high concentration range (100-10000 nM) of the competing 16mer
was the competitive immunoassay behavior observed.
[0279] When the secondary complex and the unlabeled BPDE-16-mer are
plotted using logarithm scales, two linear lines are observed. A
linear correlation coefficient of 0.97 and a positive slope are
observed between the secondary complex and the concentration of the
16mer below 100 nM. This region probably corresponds to
redistribution of the ligands. Above 100 nM, a linear correlation
of 0.997 and a negative slope are observed. This may indicate
competition with limited amount of antibody. These two linear lines
clearly distinguish the competition and redistribution, and can be
used as quantitative curves for different concentration zones.
[0280] It is expected that two types of the secondary complexes
could be formed: one antibody bound to two TMR-BPDE-16-mer
[Ab(16mer*).sub.2] and one antibody bound to one TMR-BPDE-16-mer
and one BPDE-16-mer [Ab(16mer*)(16mer)]. These two secondary
complexes of 16-mer cannot be separated since there is only slight
difference between the TMR-labeled and unlabeled 16-mer
oligonucleotides. However, the two secondary complexes can be
observed using a different competing oligonucleotide as described
below.
Example 21
[0281] Using CE/LIF in competitive binding studies between
TMR-BPDE-16-mer and unlabeled BPDE-90-mer: To observe two types of
secondary complexes, we further devised a binding system using
unlabeled BPDE-90-mer to compete with the labeled TMR-BPDE-16-mer
for a limiting amount of the antibody. In the absence of the
90-mer, the antibody forms two complexes with the 16mer*;
Ab(16mer*) and Ab(16mer*).sub.2. In the presence of BPDE-90-mer,
the BPDE-90-mer competes with the 16mer* for the antibody binding
sites. Several complexes containing the 90-mer may be formed,
including Ab(90mer), Ab(90mer).sub.2, and Ab(16mer*)(90mer). Among
these complexes, only Ab(16mer*)(90mer) is fluorescent and can be
detected. The observation of Ab(16mer*)(90mer) and the accompanying
decrease of the Ab(16mer*).sub.2 in the presence of the 90-mer
clearly demonstrate the redistribution of the two ligands and the
binding stoichiometry. The redistribution of ligands between two
binding sites of the antibody cannot be observed in traditional
binding assays that are based on measurement of total bound
ligands.
[0282] In this study, we demonstrated that the DNA adduct and the
antibody formed two complexes with the 1:1 and 2:1 stoichiometry.
Binding of the antibody with a mixture of the TMR-labeled and
unlabeled BPDE-16-mer showed a typical competitive binding behavior
when a high concentration of the unlabeled BPDE-16-mer was used.
With a lower concentration of the unlabeled BPDE-16-mer, the TMR
labeled BPDE-16-mer was partially distributed into the secondary
complex. The results suggest that the two binding sites of the
mouse monoclonal antibody are dependent upon each other, and that
the primary binding has a higher affinity than the secondary
binding.
[0283] Scatchard plot is commonly used to characterize molecular
binding events (Scatchard, 1949). For polyclonal antibody,
Scatchard plot is curved because of the heterogeneous population of
the antibody molecules. For monoclonal antibody, Scatchard plot is
considered linear because monoclonal antibody is homogeneous with
the same affinity and has two identical antigen-binding sites.
However, this does not take into account the differences between
the secondary and the primary binding. When the secondary binding
is affected by primary binding, the Scatchard plot could deviate
from linearity. In fact several authors have observed non-linear
Scatchard plots for monoclonal antibody although reasons for the
non-linearity was not discussed. It is possible that secondary
binding and the redistribution may contribute to the non-linear
nature of calibration curves for traditional competitive assays
where the primary and secondary complexes are not separated. In
traditional competitive inmmunoassay, the dose-response curve
usually is sigmoid in shape with two asymptotes and one point of
inflection (Nix, et al., 2000).
* * * * *