This application is a continuation-in-part of U.S. application Ser. No. 08/805,411, filed Feb. 24, 1997.
The invention of this application was conceived with support from the United States Government, and specifically the National Institutes of Health.
The present invention is directed to methods for determining a nucleic acid in a sample with a probe molecule using electron transfer from an electron donor to an electron acceptor. The invention is further directed to compounds useful in such methods and to compounds useful for the preparation of the probe molecules.
Determinations of nucleic acids is becoming increasingly important as a tool for diagnosis in the health fields. For example, the presence of nucleic acids from organisms, like viruses, usually not present in the human body, can be determined using probes for the infecting nucleic acids. Further, any changes in the genome which may have a potential influence on the metabolism and the state of health of the individual can be determined. Such changes may have occurred by mutation or other means. Nucleic acid determination has made further progress with the introduction of nucleic acid amplification procedures, like the polymerase chain reaction (PCR).
The presently known nucleic acid assays can be divided into two types, the heterogeneous and the homogeneous assays. In heterogeneous assays, the nucleic acid is determined by binding to a nucleic acid probe which is labeled for detection or by incorporation of labeled mononucleoside triphosphates and subsequent immobilization of the so-labeled nucleic acid to a solid phase. This is preferably done by using a solid phase bound capture probe, a format which provides the advantage that any excess amount of labeled probes or mononucleotides can easily be separated from the solid phase bound labeled nucleic acid. The homogeneous type of nucleic acid assay uses the interference between two labels. In a first method, the two labels are linked together and the event of hybridization initiates cleavage of the linkage between the two labels. (The labels are chosen such that they elicit a signal as soon as they are separated.) In a second method, the distance between the labels is changed by hybridization events. In this case, the labels may be located on one probe or on two separate probes having the capability of hybridizing to the analyte nucleic acid such that the labels can interact with each other.
Electron transfer between donors and acceptors is to be subdivided into two categories. In the first category, the donors (Do) and acceptors (Ac) are bound to the DNA duplex by non-covalent forces, such as van der Waals', electrostatic and hydrogen bonding. In the second class are systems where Do and Ac are covalently linked to the DNA. The earliest demonstration of the first approach was reported by Fromherz and Rieger in 1986, who studied photoinitated electron transfer (PET) from intercalated ethidium to surface-associated methyl viologen (Fromherz, P.; Rieger, B. J. Am. Chem. Soc. 1986, 108, 5361). Electron transfer products were demonstrated by direct observation of the reduced viologen acceptor. However, no special effect of the DNA, other than to provide a high effective concentration of the donor and acceptor, was observed. In 1992, Harriman and Brun reported PET from ethidium and acridine donors to diazapyrenium acceptors under conditions where the redox components were intercalated (Brun, A. M.; Harriman, A. J. Am. Chem. Soc. 1992, 114, 3656). Multiexponential electron transfer kinetics were attributed to Do-Ac separations of 3, 4, and 5 base pairs. The B value derived in that study (0.88 Å−1) is comparable to that determined for Do-Ac systems in proteins, where stacked π-electron systems are not available for mediating electron transfer. Barton, Barbara and co-workers studied transition metal complex donors and acceptors which are intercalated into DNA and found that quenching of the donor fluorescence as well as recovery of the ground state absorption proceeds at rates which are independent of the number of bound acceptors, suggesting a very shallow distance dependence for electron transfer through the DNA duplex (β<0.2 Å−1) (Arkin, M. R.; Stemp, E. D. A.; Holmlin, R. E.; Barton, J. K.; Hörmann, A.; Olson, E. J. C.; Barbara, P. F. Science 1996, 273, 475). However, cooperative binding of the donor and acceptor molecules, which would account for the loading-independent kinetics, could not be completely ruled out in that system.
One of the problems associated with the use of non-covalently bound donor and acceptor molecules in these studies is the inability to control precisely the location of the redox components relative to one another when they are bound to the DNA. In one extreme, the intercalation locations will be controlled statistically, leading to a distribution of Do-Ac separation distances. At another extreme, binding will be cooperative, leading to short distances between Do and Ac over a wide range of concentrations.
Covalent linkage of Do and Ac to the 5′-ends of complementary oligonucleotides has led to systems with better defined Do-Ac separation distances. Barton, Turro and co-workers reported fluorescence quenching that occurs in less than one nanosecond for a system containing linked Do and Ac metal complexes intercalated near the ends of a 15 base pair duplex (Murphy, C. J.; Arkin, M. R.; Jenkins, Y.; Ghatlia, N. D.; Bossmann, S.; Turro, N. J.; Barton, J. K. Science 1993, 262, 1025). A rate this fast indicates that the distance dependence of electron transfer through DNA is extremely shallow, but this interpretation must be regarded with caution pending a clear demonstration of redox products. In contrast, there is a report of a covalently linked system having Do and Ac metal complexes at the opposite ends of an 8 base pair duplex which shows electron transfer on a microsecond time scale (Meade, T. J.; Kayyem, J. F. Angew. Chem. Int. Ed. Engl. 1995, 34, 352). In this case the redox components were not intercalated within the helix so the rate of electron transfer may simply reflect the time required to orient the donor and acceptor in order to obtain sufficient electronic coupling through the π-electron stack before long distance electron transfer can occur.
At this time, there are many unresolved questions regarding the ability of duplex DNA to mediate electron transfer. None of the systems cited above has unambiguously demonstrated the rate or efficiency of electron transfer between donor and acceptor moieties held at a fixed distance of separation in a DNA/DNA duplex.
In a modification designed to detect hybridization of nucleic acids in homogeneous solution, Tyagi and Kramer (Tyagi, S.; Kramer, F R. Nature Biotechnology 1996, 14, 303) describe a doubly substituted single-stranded DNA construct that possesses a stem-loop (i.e. hairpin) structure. The construct contains a fluorescer covalently linked to one terminus of the strand and an energy transfer quencher of the fluorescer at the opposite terminus. When unconjugated, this single stranded chain exists predominantly in hairpin conformation that constrains the fluorescer and quencher to be relatively close in space. When in this structural form, excitation of the fluorescer with actinic light leads to reduced emission because the fluorescing excited state transfers its energy to the nearby quencher. However, when this single-stranded structure hybridizes with a second strand complementary to its loop region, the distance between the fluorescer and quencher is increased and, consequently, the efficiency of fluorescence increases. The change in fluorescence intensity is an indicator that hybridization has occurred.
The modification described by Tyagi and Kramer offers several advantages for homogeneous real-time assays for hybridization. However there are certain disadvantages to the system they report. First, the indication of hybridization relies on energy transfer quenching of the fluorescer. This requires that the quencher have a lower excited singlet energy than the fluorescer, and this can cause difficulties in selecting a quencher whose absorption spectrum does not overlap with that of the fluorescer. Second, the nature of the hairpin structure requires that a portion of the single-stranded probe molecule be self-complementary. In general, this self-complementary portion will not hybridize with the target strand of the nucleic acid to be determined. This requirement will reduce the association constant of the hybrid duplex DNA. A further disadvantage of the modification described by Tyagi and Kramer is that covalent linkage of the fluorescer and quencher at the terminal positions of the single-stranded DNA probe is cumbersome synthetically and far from ideal for an assay. The long chain of atoms used to bind the fluorescer and donor to the single-stranded DNA is flexible and, consequently, the fluorescer and quencher will exist in many confirmations, even in the stem-loop structure, some of which may be ineffective at quenching the emission of the fluorescer. This will contribute to a high background fluorescence in assays for hybridization. Finally, a further disadvantage of covalent linkage of the fluorescer and quencher at the terminal positions is that unraveling of the stem structure at these positions, which is commonly to be expected, will increase the distance between the fluorescer and quencher, and increase the number of available conformations. Both of these effects will lead to an increase in background emission.
In a further attempt to modify DNA Shimidzu and co-workers reported the synthesis and characterization of a modified DNA oligomer containing an acridine moiety covalently linked at an internal position (Fukui, K.; Morimoto, M.; Segawa, H.; Tanaka, K.; Shimidzu, T. Bioconjugate Chem. 1996, 7, 349). Hybridization with a complementary oligomer containing either a thymine or an abasic site at the appropriate position opposite the acridine yields a 1:1 duplex with the acridine, apparently, intercalated within the helix. Electron transfer to the acridine moiety was not reported.
Described in WO 95/15971 is the conjugation of oligonucleotides with intercalators that can act as electron donors or electron acceptors. The resulting complexes represent a series of derivatives that are bimolecular templates whose use as probe molecules relies on duplex DNA to provide a path for the transfer of electrons over very large distances at extremely fast rates. In this role the DNA duplex is described as and must function as a “bioconductor”. In WO 95/15971 there is disclosure of a method wherein oligonucleotides are labeled at each end with different electron transfer moieties and it is demonstrated that these moieties are capable of electron transfer through the duplex under certain conditions. These electron transfer moieties are complexes of ruthenium and other heavy metal ions with organic ligands which can change electronic state during electron transfer. Further in WO 95/15971, there is a suggestion that the phosphodiester bonds in an oligonucleotide can be replaced by peptide bonds thus using peptide nucleic acids (PNA) as bioconductors.
The subject of the present invention is a method for determining a nucleic acid in a sample comprising binding a probe having a polymeric backbone different from the natural sugar phosphate backbone of DNA or RNA to the nucleic acid by means of base-mediated hydrogen bonding, wherein this probe has an electron acceptor or an electron donor, or both, bound covalently either at terminal positions or at internal positions of the probe molecule. Further, stimulation of the electron donor or electron acceptor by any one of several means elicits a different outcome depending upon whether the probe molecule is bound to the nucleic acid. In particular, the present invention is a method for inducing an electron or hole transfer from an electron donor to that electron acceptor or a hole transfer from said electron acceptor to an electron donor and determining the occurrence of the electronic transfer as a measure of said nucleic acid. It is an object of the present invention to provide a method for reliable detection of electron or hole transfer.
A further object of the invention is to modify the efficiency or rate of electron (hole) transfer in bioinsulators and bioconductors.
In another aspect, the invention is directed to a method of controlling the rate and efficiency of electron (hole) transfer with high specificity for the indication of duplex formation.
From this prior art it is not apparent to one skilled in the art that DNA/DNA, DNA/PNA and PNA/PNA hybrids will be bioconductors under all conditions of electron transfer driving force and time scale. In particular, electron conduction (electron transfer or hole transfer) from an electronically excited state, or other electron donor or electron acceptor species not having an essentially infinite lifetime, must compete with return of the electronically excited state to its ground state, or consumption by some other means of the electron donor or acceptor with a limited lifetime. If conduction of an electron through a DNA/DNA, DNA/PNA and PNA/PNA duplex occurs on a time scale longer than the return of the excited state to the ground state or consumption of the electron donor or acceptor, then the DNA/DNA, DNA/PNA and PNA/PNA duplex will act as if it is a “bioinsulator”, rather than a bioconductor.
The molecular and energetic features that control the rate and efficiency of electron transfer reactions have been extensively studied. The Marcus theory of electron transfer (Marcus, R. A. Ann. Rev. Phys. Chem. 1964, 15, 155. Marcus, R. A. J. Chem. Phys. 1965, 43, 679) is remarkably successful in its prediction of reaction rates. In its simplest formulation (eqs. 1 and 2), the theory identifies three factors that determine the rate constant for electron transfer (ket). These are the driving force (ΔGet) for the reaction, the reorganization energy (λ), and the maximum rate constant (kmax), which occurs when ΔG≠=0. In classical theory, kmax=kelνn, where kel is the electronic transmission coefficient and νn is the frequency of passage through the transition state.
Of most relevance to the present invention is the estimation of ket and its comparison with chemical or physical reactions that consume the electronically excited state, or other electron donor or electron acceptor species not having an essentially infinite lifetime. If the rate of electron transfer from donor to acceptor is greater than the rates of the competing chemical or physical reactions, then the electron transfer will be more efficient. Formation of a complex between the probe and the nucleic acid to be determined can change the magnitude of ket and/or the magnitude of the rates of the chemical or physical reactions that consume the electronically excited state, or other electron donor or electron acceptor species not having an essentially infinite lifetime. Formation of the complex between the probe molecule and the nucleic acid to be determined may modify ΔGet so that the rate of electron transfer increases or decreases upon complex formation. Alternatively, complex formation can cause changes in the environment of the electron donor or electron acceptor such that the rate of chemical or physical reactions that consume the electronically excited state, or other electron donor or electron acceptor species not having an essentially infinite lifetime, changes relative to the rate for the electron transfer reaction. Based on Marcus theory, the formation of the complex between the probe and the nucleic acid to be determined causes a change in the Marcus reorganization energy for the electron transfer or a change in the Marcus kmax. A change in either parameter may result in either an increase or decrease in the rate of electron transfer. In the case where formation of the complex between the probe and the nucleic acid to be determined results in an increase in the ket the resulting complex can be called a bioconductor, since it facilitates electron transfer. It is a surprising discovery that, depending on some specific details of structure, formation of the complex between the probe and the nucleic acid to be determined results in a decrease in the rate constant for electron transfer in the complex. Thus, in some cases, the complex formed by the probe molecule and the nucleic acid to be determined may be a “bioinsulator”.
FIG. 1 shows schematically the long range electron transfer (ET) mechanism of the present invention. The concept is illustrated by the example where the donor and acceptor are located on the same strand. The designations n and m refer to the electronic states of the donor and the acceptor.
FIG. 2 shows a first embodiment of the invention using only one probe molecule labeled with a distally positioned electron donor (Donor) and an internally positioned acceptor (acc.).
FIG. 3 shows an embodiment of the present invention wherein two labeled probes are used. Thus one probe contains the acceptor (distally located) and another probe contains the donor (distally located).
FIG. 4 shows a format using an immobilized probe containing an internally positioned acceptor and donor in the same probe.
FIG. 5 shows the proposed electron transfer mechanism via hole hopping.
FIG. 6 shows a format using a hairpin forming probe.
FIG. 7 shows an embodiment using two complementary probes.
FIG. 8 shows the flow scheme for the preparation of Q1, an electron accepting PNA monomer (linker length 1).
FIG. 9 shows the flow scheme for the preparation of Q2, an electron accepting PNA monomer (linker length 4).
FIG. 10 shows the flow scheme for the preparation of R1, an electron donating PNA monomer (linker length 4).
FIG. 11 shows the flow scheme for the preparation of R2, an electron donating PNA monomer (linker length 7).
FIG. 12 shows absorbency/temperature profiles for PNA/DNA hybrids with different quinone linker lengths for different wavelengths. AQ2 stands for PNA579 and AQ5 for PNA586.
FIG. 13 shows photoinduced cleavage of DNA within PNA/DNA hybrids.
FIG. 14 shows photoinduced cleavage of DNA with an 8-OxoG site.
FIG. 15 shows phosphorescence quenching in PNA/DNA hybrids due to electron transfer.
FIG. 16 shows absorbency/temperature profiles for PNA626 and 627 hairpins.
FIG. 17 shows absorbency/temperature profiles for DNA626A hybrids with PNA626 and PNA627.
FIG. 18 shows schematically the synthesis of a further PNA-electron donor monomer.
The term nucleic acid to be determined is understood in the present invention as an analyte nucleic acid or a nucleic acid derived therefrom. Analyte nucleic acids include nucleic acids of any origin, for example, nucleic acids of animal, human, viral, bacterial, or cellular origin. They may be present in solution, suspension, but also fixed to solid bodies or contained in cell-containing media, cell smears, fixed cells, tissues, or fixed organisms. Nucleic acids derived therefrom are nucleic acids prepared from analyte nucleic acids or parts thereof, for example as copies of the above-mentioned nucleic acids, or parts thereof. These copies include nucleic acids derived from that original analyte nucleic acids by amplification, including any replication or/and transcription/reverse transcription reactions, for example by the polymerase chain reaction.
Usually the nucleic acid to be determined will be pretreated to get into a condition ready for binding the probe molecule, if it is not already accessible. Such pretreatment may include denaturing double-stranded nucleic acids by changing the pH into the alkaline region, repeating extreme temperature changes (freezing/thawing), changing the physiological probe conditions (osmotic pressure), lysis of cell walls by detergents, chaotropic salts or enzymes (e.g. proteases, lipase). These steps may be used either alone or in combination in order to release the nucleic acids. In some instances, it may be advantageous to separate the nucleic acids from other components of the sample, like proteins, cells, cell fragments, but also nucleic acids which are not intended to be detected.
The probe molecule binding to the nucleic acid is defined by having a polymeric backbone different from the natural sugar phosphate backbone. Examples of such probe molecules are now well-known in the art. Those probes may be based upon monomeric subunits, which are linked in a repetitive way. It is preferable that the probe molecule contains at least five monomeric subunits, each monomeric subunit being connected to other monomeric subunits by peptide bonds. The peptide bond is understood to be the bond connecting a primary or secondary amine and a carboxylic acid residue. Other types of linkages within the monomers or connecting one or more monomers in the backbone are possible, for example, ether or amino bonds. Examples of such probe molecules are described in WO 92/20702 including probe molecules having ligands bound to aza-nitrogen atoms, WO 94/25477, and WO 96/20212. Probe molecules having mixed different linkages between the monomers are described in EP-A-0 672 677. The term probe molecule further contains molecules having the above stretch of the non-natural backbone and an additional stretch of the natural sugar phosphate backbone. Such chimera may possess some lower affinity to complementary nucleic acids, but are nevertheless useful in the present invention.
The term backbone in the present invention shall mean the polymeric moiety to which at different points of attachment heterocyclic base moieties are bound in a consecutive way.
The binding of the probe molecule to the nucleic acid is accomplished by hydrogen bonds mediated by said base moieties on said backbone. Such hydrogen bonding for example occurs between complementary nucleobases as in the base pairing in nucleic acids having the natural sugar phosphate backbone or artificial bases eliciting similar properties for hydrogen bonding to the natural bases. While the most prominent mode of binding is by duplex formation, some molecules bind to nucleic acids by triplex formation (see for example WO 95/01370). Artificial bases include diaminopurine, pseudouridine, thioguanine and 7-deazaguanine. The method of the invention requires a stretch of at least five consecutive monomeric units coupling the bases of the probe molecule. The probe molecule should have a length of at least 9 base-pairing monomeric units, preferably between 9 and 30 and most preferably between 10 and 25 base-pairing monomeric units.
Such probe molecules can be prepared by analogy to the methods described in the above-mentioned documents. Their preparation may include techniques usually employed in peptide chemistry. However, probe molecules especially preferred in the present method include new starting and intermediate compounds useful for introducing any electron donor or/and any electron acceptor moieties into the probe. Their preparation will be described later.
An electron acceptor or an electron donor or an electron acceptor and an electron donor are bound covalently to the probe molecule at defined, fixed positions, according to the invention. The generally possible positions for attaching moieties to polymeric backbones can be systematically divided into a first group wherein the moiety is attached at one or both ends of the polymeric backbone, a second group wherein at least one of the groups is bound to the polymeric backbone at a position on the backbone not located on the first and the last base bearing monomeric unit of said backbone, and a third group wherein the moiety is bound to a base position attached at any of the monomeric units. Surprisingly, the objects of the present invention are better met for the second group. Within this group, such probe molecules have proven to be the most successful in which the nucleobase moiety of one or more of the monomeric subunits is replaced totally by a group or groups containing the electron acceptor or an electron donor, preferably such that the rate or efficiency of electron transfer in the probe is modified by the nucleobases of the nucleic acid to be determined. It is possible not only to modify only one subunit by such moieties, but also more subunits, dependent upon the overall length of the probe molecule. However, as a general rule, it is preferred to not include more than 30% of modified monomeric units compared to the overall number of monomeric units in the probe molecule. If more than one modified monomeric subunit is contained within the probe molecule, these monomeric units should not all be located at the final monomeric subunits.
Preferred probe molecules are compounds of the general formula I
n is an integer of from at least 3,
x is an integer of from 2 to n−1,
each of L1-Ln is a ligand independently selected from the group consisting of hydrogen, hydroxyl, (C1-C4)alkanoyl, naturally occurring nucleobases, non-naturally occurring nucleobases, aromatic moieties, DNA intercalators, nucleobase-binding groups, heterocyclic moieties, reporter ligands and chelating moieties, wherein at least one of L1-Ln, preferably at least one of L2-Ln−1 is a non-nucleobase electron acceptor or a donor moiety and at least 2 of L1-Ln being a nucleobase binding group, or a naturally or non-naturally occurring nucleobase;
each of C1-Cn is (CR6R7)y (preferably CR6R7, CHR6CHR7 or CR6R7CH2) where R6 is hydrogen and R7 is selected from the group consisting of the side chains of naturally occurring alpha amino acids, or R6 and R7 are independently selected from the group consisting of hydrogen, (C1-C6)alkyl, aryl, aralkyl, heteroaryl, hydroxyl, (C1-C6)alkoxy, (C1-C6)alkylthio, NR3R4 and SR5, where R3 and R4 are as defined below, and R5 is hydrogen, (C1-C6)alkyl, hydroxyl, (C1-C6)alkoxy, or (C1-C6)alkylthio-substituted (C1-C6)alkyl or R6 and R7 taken together complete an alicyclic or heterocyclic system; or C1-Cn is CO, CS, CNR3;
each of D1-Dn is (CR6R7)z (preferably CR6R7, CHR6CHR7 or CH2CR6R7) where R6 and R7 are as defined above;
each of y and z is zero or an integer from 1 to 10, the sum y+z being at least 2, preferably greater than 2, but not more than 10;
each of G1-Gn−1 is —NR3CO—, —NR3CS—, —NR3SO— or —NR3SO2—, in either orientation, where R3 is as defined below;
each of A1-An and B1-Bn are selected such that:
(a) A1-An is a group of formula (I/A), (I/B), (I/C) or (I/D), and B1-Bn is N or R3N+; or
(b) A1-An is a group of formula (I/D) and B1-Bn is CH;
X is O, S, Se, NR3, CH2 or C(CH3)2;
Y is a single bond, O, S or NR4;
each of p and q is zero or an integer from 1 to 5, (the sum p+q being preferably not more than 5);
each of r and s is zero or an integer from 1 to 5, (the sum r+s being preferably not more than 5);
each R1 and R2 is independently selected from the group consisting of hydrogen, (C1-C4)alkyl which may be hydroxyl- or (C1-C4)alkoxy- or (C1-C4)alkylthio-substituted, hydroxyl, (C1-C4)alkoxy, (C1-C4)alkylthio, amino and halogen; and
each R3 and R4 is independently selected from the group consisting of hydrogen, (C1-C4)alkyl, hydroxyl- or alkoxy- or alkylthio-substituted (C1-C4)alkyl, hydroxyl, (C1-C6)-alkoxy, (C1-C6)-alkylthio and amino;
Q and I is independently selected from —CO2H, —CONR′R″, —SO3H or —SO2—NR′R″ or an activated derivative of —CO2H or —SO3H and —NR′R′″
where R′, R″ and R′″ are independently selected from the group consisting of hydrogen, alkyl, amino protecting groups, reporter ligands, intercalators, chelators, peptides, proteins, carbohydrates, lipids, steroids, nucleosides, nucleotides, nucleotide diphosphates, nucleotide triphosphates, oligonucleotides, including both oligoribonucleotides and oligodeoxyribo-nucleotides, oligonucleosides and soluble and non-soluble polymers and as well as nucleic acid binding moieties and
each of x1 and y1 is an integer offrom 0 to 10.
Alkoxy- and alkylthio groups contain preferably from 1 to 4 carbon atoms.
An example for a fused aromatic moiety is naphthol. A heterocyclic moiety is pyridine. Reporter groups are moieties that can be detected, like fluorescent compounds, for example fluorescein, or moieties that can be recognized by another molecular entity, like haptens which can be recognized by an antibody raised against this hapten.
In the above structures wherein Q or I is an oligonucleotide or oligonucleoside, such structures can be considered chimeric structures between PNA compounds and the oligonucleotide or oligonucleoside.
Linkers A1-An for binding acceptor moieties are generally preferred at a length of between 1 to 10 atoms, most preferred 2 to 6 atoms, for donor moieties at a length of 1 to 10, most preferred 2 to 8 atoms.
More preferable are compounds of subgroups Ia-Ib based on the general formula I wherein
(Ia): B1-Bn is N and A1-An is —CO—(CH2)6—
(Ib): B1-Bn is N and A1-An is —CO—NR3—(CH2)2—
(Ic): B1-Bn is CH and A1-An is —NR3—CO—(CH2)2—.
Preferred PNA-containing compounds useful to effect binding to RNA, ssDNA and dsDNA and to form triplexing structures are compounds of the formula IIa, IIb and IIc:
each L is independently selected from the definitions of L1-Ln in formula I;
each R7 is independently selected from the group consisting of hydrogen and the side chains of naturally occurring alpha amino acids;
n is an integer greater than 1,
each k, 1, and m is, independently, zero or an integer from 1 to 5;
each p is zero or 1;
Rh and Ri are as defined for R′, R″ and R′″.
Electron acceptor moieties and electron donor moieties are each referred to as electron transfer moieties. They are most preferably moieties not including nucleobases (non nucleobase moieties).
Electron acceptors according to the present invention can induce oxidation of other moieties by attracting an electron from their surroundings particularly after they have been stimulated with light or activated by some other means. Electron acceptors will typically be electron-deficient species. Preferable electron acceptors are organic molecules, preferably excluding organic molecules containing metal ions. Further, these organic molecules have a generally flat structure with a system of delocalized π-electrons. Therefore, organic molecules are especially preferred that contain aromatic hydrocarbon moieties containing functional groups, wherein the functional groups may have electron-withdrawing properties. It is possible that the electron acceptor will contain both electron-donating and electron-withdrawing functional groups. The substituents should not be so bulky that they disturb the regular base stacking between the probe and the nucleic acid to be determined. The electron deficiency of the electron acceptor will normally be increased by excitation with light or with stimulation by some other means. In the preferred case there is no measurable electron transfer to the acceptor without its stimulation in the assay medium. The acceptor moiety will typically have well-characterized redox and spectroscopic properties, although precise knowledge of these parameters is not required for their successful application in this invention. Electron acceptors useful in this invention typically undergo a change in chemical or optical properties when it accepts an electron. For example, such electron acceptors may be useful in this invention when, after conversion by electron attraction to become a radical anion, or any other one electron reduction product, it possesses a unique optical spectrum with strong absorption bands. However, the detection of the change in chemical or optical properties of the electron acceptor is not required for the successful application of this invention.
These acceptor moieties are typically composed of one to 10, preferably 1 to 4, fused cyclic hydrocarbon rings which may be substituted or unsubstituted. The cyclic hydrocarbons may independently have any ring size ranging from 5- to 10-membered rings, but preferably include 5- or 6-membered aromatic rings. The cyclic hydrocarbons may be fused in any position isomer.
Preferably, the acceptor or the donor moiety includes a group of general formula IIIa or IIIb
wherein X, Y, Z, Q, V and W are independently selected from the atoms C, N, S, and O, X, Y, Z, Q, V and W are connected by either single or double bonds. R1-R6 are independently selected from the group of R1-R6 are, independently from each other, selected from the group of —H, —O−, —OH, —OR′, —SH, —SR′, —NH2, NO2, —SO3 −, —SO2 −, —CN, —PO3 2−, —PO2 −, ——COOH, —CO—R′, —COOR′, —CS—R′, CSO—R′, —COO−, —N═N—, halogen (—F, —Cl, —Br, —I), —NHR′, N(R′R″), or derivatives hereof, or hydrocarbyls or heterocyclyl as defined below. R′ and R″ is chosen from the possible definitions of R1-R6. At least one of X, Y, Z, Q, V and W can together with the moiety R1-R6 bound thereto also mean —CO—, —SO— or —SO2—. At least one of R1-R6 can also be a single or a double bond.
Hydrocarbyl comprises groups such as alkyl, alkenyl, alkynyl, each having between 1 and 10 carbon (C)-atoms, aryl having 6-30 C-atoms, such as phenyl, naphthyl, biphenyl, tolyl, anthracenyl, etc., and combinations of these in different substitutions patterns. These hydrocarbyl groups may be straight chained or branched chained, symmetric or asymmetric, chiral or achiral, contain one or more hetero atoms selected among —N—, —NH—, —S—, —O—, and may also be fused. The hydrocarbyl groups may be unsubstituted or substituted by one or more of the above mentioned R1-R6.
Heterocyclyl are preferentially selected among cyclic aromatic or non-aromatic moieties containing hetero atoms selected among —N—, —NH—, —S— and —O—, preferably selected from the group consisting of pyridyl, imidazolyl, pyrimidinyl, pyradazinolyl, quinolyl, acridinolyl, pyrrolyl, furyl, thienyl, isoxazolyl, oxazolyl and thiazolyl. The heterocyclyl groups may optionally be substituted by one or more of the above mentioned R1-R6.
At least one of said moieties R1-R6 is modified such that it is capable of binding to the backbone of the probe. Preferably the electron acceptor is bound via a free sigma bond at one of said moieties to said backbone. It is understood according to the present invention that the acceptor moiety is defined as the moiety which is capable and responsible for accepting the electron. Any moiety between the backbone and the acceptor is defined to be a linker A.
While this is not the preferred case, the acceptor or/and donor moiety can include a complex of a transition metal chelated by one or more ligand moieties. Preferred transition metals are chosen from the group iron, copper, ruthenium, rhenium and osmium.
More preferably, the electron acceptor or donor may contain a group chosen from the class of imides. Imides may act as both acceptors and donors. These compounds will contain the essential —(C═O)—N(R)—(C═O)— unit with the carbonyls placed in a hydrocarbon ring structure which is bonded to an aromatic ring system via conjugation of the carbonyls. The imide is contained in a hydrocarbon ring which may be 5, 6, 7, 9 membered but preferentially a 5 or 6 membered ring. The acceptor/donor may contain from 1-10 imides linked via the aromatic system. An acceptor/donor may contain imides of the same ring size or containing imides of different ring sizes. The aromatic structure comprises of 1-10 fused cyclic aromatic hydrocarbons and optionally substituted with the substituents (R). The cyclic hydrocarbons may be bonded/fused in any position isomer. The cyclic hydrocarbons may independently of each other be any ring size (5, 6, 10, 14 membered rings) but preferentially 5, 6 membered aromatic rings which independently of each other may comprise one or several hetero atoms selected among —N—, —NH, —S— and —O—.
Preferred imides include groups of general formulae IVa-IVe.
wherein the definitions for R are selected from the possible definitions of R1-R6 of general formula IIIa and IIIb and wherein n and m is 0 or an integer from 1 to 10 and k, r and l is 0 or an integer from 1 to 4. It is clear that only so many substituents R are bound that the valency and charge of the atom of attachment of the substituent R is not changed.
A further preferred moiety is chosen from the group of n-alkyl-aza-aromatic compounds. N-alkyl-aza-aromatics may act as both acceptors and donors (formula Va-d, vide infra). In these compounds the nitrogen atoms in the aromatic rings are alkylated. This forms positively charged molecules. The aromatic rings where the aza atoms are located can be 5, 6, 10, 14 membered rings, but preferentially 5 and 6 membered rings. The acceptor/donor may contain 1 to 10 alkylated cyclic nitrogen atoms which are placed in an aromatic system of 1 to 10 fused aromatic rings.
The fused cyclic hydrocarbon system may be composed of 5, 6, 10, 14 membered aromatic rings, but preferentially 5 and 6 membered rings, which independently of each other may comprise one or several hetero atoms selected among —N—, —NH, —S— and —O—. The acceptor may be composed of alkylated cyclic nitrogen atoms of the same ring size and/or different ring size. The aromatic hydrocarbons carrying the alkylated cyclic nitrogen atoms and/or the fused aromatic ring not carrying the alkylated nitrogen atoms are preferentially substituted (R).
Within this group compounds of general formulae Va-Vd
wherein the definitions of the substituents of formula IV apply and wherein r is 0 or an integer of from 1 to 4.
Preferred as acceptors are compounds containing a quinoid structure. Such compounds are well-known to one skilled in the art. The quinones especially useful in this invention are composed of at least 2 conjugated carbonyl groups (optionally thiono or azo) placed in the same or in separate rings fused with 1-10 cyclic aromatic hydrocarbons and optionally substituted with the substituents (R1-R6) and having 2-20 conjugated carbonyl groups in pairs, provided that the number of conjugated carbonyl groups does not exceed twice the number of fused cyclic hydrocarbons. The cyclic hydrocarbons may be fused in any position isomer. The cyclic hydrocarbons may independently of each other be any ring size but preferentially 5, 6 carbon atom membered aromatic rings which independently of each other may comprise one or several hetero atoms selected among —N—, NH, —S— and —O—. The conjugated carbonyl groups may be located in any of these rings provided that the quinoid structure is maintained.
Especially preferred quinoid structures contain the general formulae VIa-VIc.
wherein the definitions of the substituents are chosen from the definitions of formulae IV and wherein o and p are 0 or an integer from 1 to 10 and t and q are 0 or an integer from 1 to 4. Preferred examples of this group are anthraquinone and phenanthraquinone.
As electron donors, generally all compounds are useful which can in the ground state or in the electronically excited state, or after stimulation by some other means, be oxidized by an electron acceptor by giving off an electron. Preferred donors are organic molecules, although organic molecules containing metal ions are not excluded. Preferably, the donor molecules are chosen such that the electron deficiency caused by the oxidation has the consequence of causing a change to one or more of its physical or chemical properties, whereby detection of the electron deficiency (oxidation event) is possible. Typical donors will be electron-rich fused aromatic systems carrying functional groups and will optimally be substituted. The donor will preferably have an aromatic structure substituted by electron-donating groups. Therefore, the donors are preferably chosen from the general formulae III-V, wherein the substitution with electron-donating groups supersedes.
Electron-withdrawing groups or electron-attracting groups are preferably the groups NO2, —SO3 −, —SO2 −, —CN, —PO3 2−, —PO2 −, —COOH, —CO—R′, —COOR′, —CS—R′, CSO—R′, —COO−, halogen (—F, —Cl, —Br, —I), while electron-releasing groups/electron-donating groups are preferably selected from the groups H, —O−, —OH, —OR′, —SH, —SR′, —NH2, —N═N—, —I, —NHR′, N(R′R″).
Compounds preferentially acting as donors are compounds containing an aza-aromatic structure. Examples of these compounds are aromatic ring structures containing nitrogen. The N-heterocyclic structure is bonded/fused with a cyclic hydrocarbon system composed of 5, 6, 10, 14 membered aromatic rings, but preferentially 5 and 6 membered rings, which independently of each other may comprise one or several hetero atoms selected among —N—, —NH, —S— and —O—. The donor may be composed of heterocyclic nitrogen containing rings of the same ring size and/or different ring size. The aromatic hydrocarbons carrying the heterocyclic structure and/or the bonded/fused aromatic ring system are preferentially substituted.
Preferred aza-aromatic structures are compounds of general formulae VIIa-VIIc
wherein the groups R, n, m, p, o, l, k, q, and r are chosen from the definition of formulae III to VI.
One possible donor moiety is the natural base guanine and derivatives thereof. It has been found in the present invention that guanine is an effective donor, forming a chemically reactive species.
FIG. 1 shows schematically one embodiment of the present invention. The probe strand contains a donor moiety D having n electrons and an electron acceptor A having m electrons covalently attached. Upon induction of electron transfer (ET) one electron is transferred from the donor moiety to the acceptor moiety, thereby changing the number of electrons in each moiety.
While applicants do not wish to be bound by this hypothesis, it appears that the use of nucleic acid analogues allows very selective control of the rate or efficiency of electron and hole transfer between a donor and an acceptor through modification of the distance or effectiveness of electron or hole transport through the π-electron stack of the DNA or the nucleic acid analogue when bound to a complementary nucleic acid strand. The orientation within such a complex is considered to be so rigid that the donor (acceptor) after excitation, or stimulation by some means, can donate (attract) an electron (hole) from a near-by acceptor (donor) or a nucleic acid base. The base where the electron or hole comes from is now oxidized or reduced and may bear a positive or negative charge. The acceptor as a consequence is negatively charged. When a chemical moiety (here a nucleobase) is missing an electron, an “electron hole” (or just hole) is created in the molecule. The electron hole in the first base can then be filled by transfer of an electron from another base (see FIG. 5); this process is called hole-hopping. By several such hole-hops, the excitation of the acceptor and subsequent electron transfers yield oxidation of a distal position in the binding product (for example, a hybrid between the nucleic acid analogue and the nucleic acid to be determined). Electron transfer may propagate in the strand where it is initiated but may as well propagate in the opposite strand. The rate or efficiency of electron or hole hopping will depend on its time scale and the time scale of chemical or physical processes that compete for consumption of the excited or stimulated electron donor or acceptor.
FIG. 5 shows schematically the process of hole-hopping initiated by irradiation at 350 nm for a peptide nucleic acid (lower strand) having an electron acceptor exemplified as Q within the sequence and replacing a base and a nucleic acid (upper strand) having bases B complementary to the bases of the peptide nucleic acid. Electron transfer creates a positively charged base at the nucleic acid (B−
The sample containing the nucleic acid to be determined is contacted with the probe molecule in order to create the complex between that probe molecule and the nucleic acid to be determined. This complex will form via base-mediated hydrogen bonding. Peptide nucleic acids as probe molecules have the advantage that they can strand invade into double-stranded nucleic acids. Therefore, it is not even necessary to separate double-stranded nucleic acids into single strands prior to adding the probe molecule. The binding of the probe molecule to the nucleic acid will be made according to conditions known to one skilled in the art. Such conditions are well described for peptide nucleic acids, for example in WO 92/20703, to which is made reference in this respect. Because in many cases, the amount of nucleic acids in the sample is not evident, it might be preferred to choose the probe molecule in the amount exceeding the highest expected amount of nucleic acids in the sample. However, in order to maintain a low level of background signal, it may be advisable to choose the amount of probe molecules to be in the order of magnitude of the nucleic acid to be expected. As described in WO 95/15971, there are several possibilities to determine the presence of any nucleic acid. The present invention, however, requires that at least one probe molecule is labeled by an electron donor moiety or an electron acceptor moiety or both. The ruling principle is that within the complex containing the nucleic acid to be determined and the probe molecule, there is at least one electron donor and at least one electron acceptor moiety. While one of the electron donor and electron acceptor moiety must be covalently bound to the probe molecule, at least one of the other species of electron transfer moieties can be located either within the same probe molecule, the nucleic acid to be determined or another probe molecule. This other probe molecule can either have a natural sugar phosphate backbone or a non-natural nucleic acid analogue backbone, while the nucleic acid to be determined preferably has the natural sugar phosphate backbone.
In a first embodiment, a complex is formed by a nucleic acid analogue probe molecule having an electron acceptor moiety bound at an internal backbone position, the nucleic acid to be determined or a nucleic acid being the product of an amplification process of a nucleic acid to be determined, and a further probe nucleic acid analogue probe molecule containing an electron donor moiety bound to the backbone at a non-terminal position, wherein the probe molecules are bound to different but adjacently located nucleotide sequences on the nucleic acid. It is important to avoid interrupting the base-stacking between the binding sites of the two probes. As a general rule, the electron donor and the electron acceptor of two different probe molecules should not be at termini facing each other when bound to the nucleic acid. It is obvious that groups at the termini facing each other would severely hinder the capability of the probes to bind to adjacent sequences on the nucleic acid to be determined.
In a second embodiment, the nucleic acid is labeled to contain an electron donor or electron acceptor moiety within the π stack. As described for the probe molecule, it is therefore preferred to insert a flat electron transfer moiety in place of a nucleobase. Such nucleic acids can for example be prepared by using a chemically synthesised oligonucleotide primer having the electron transfer moiety near the 3′-end. During amplification, if the original analyte nucleic acid is present, this primer is elongated by attachment of further mononucleotides. The probe molecule is then chosen to bind to a position containing both the part of the primer wherein the electron transfer moiety is located and a part of the adjacent extension product.
For the further determination of the complex, it is important that by this binding process a complex is formed having at least one electron acceptor moiety and at least one electron donor moiety covalently bound to either of the probe molecules or the nucleic acid to be determined. Thus, the complex contains at least one electron acceptor and at least one electron donor.
The formation of the complex is determined by inducing electron transfer from an electron donor within that said complex to an electron acceptor within said complex and determining the occurrence of this electronic transfer as a measure of said nucleic acid. This induction of electronic transfer of course depends upon the choice of the electron donor or/and the electron acceptor. There are at least two possibilities to start induction. In the first case, the electron donor is activated to give away an electron to the π-stack within the complex. In the second case, the electron acceptor is activated to absorb an electron from a moiety within the π-stack as described above. Generally, therefore the electron acceptor is changed in its electronic configuration, which will depend upon the π-systems involved. The activation process is described in the following using the example of compounds chosen from the general formula V. For example, anthraquinone derivatives have an nπ* excited state electronic configuration. Upon excitation by light, fast (≦10 ps) intersystem crossing to the triplet state will occur, precluding fluorescence emission. However, phosphorescence from the triplet state can be observed at low temperatures. This emission is efficiently quenched when the electron acceptor is intercalated into the π-stack of the complex formed due to rapid electron transfer from one of the bases adjacent to the intercalation site to the excited electron acceptor. When the electron acceptor is not located within the π-stack, electronic interaction with the bases is weak at best and significantly reduced quenching occurs.
In a first embodiment therefore the occurrence of electronic transfer is determined by determining the degree of the reduction of quenching compared to the case when no nucleic acid to be determined is present.
The preferred mode of inducing electron transfer in this aspect of the invention is therefore the inducement by irradiating the sample containing the complex with light at a wavelength wherein the activation of the electron acceptor is achieved. This will be for the electron acceptors mentioned above at wavelength of between 300 nm and 450 nm, most preferably between 330 nm and 360 nm. The vessel or containment wherein the sample is maintained preferably is transparent to the light used, such that the light yield is improved.
The generation of electron transfer (ET) is preferably done in an aqueous environment. Thus, the solvent used is preferentially water or water containing up to 40% (v/v) of an organic medium, preferentially up to 10% (v/v) of an organic solvent such as methanol. Higher content of organic solvents will not be used due to the instability of duplexes formed from nucleic acids and nucleic acid analogues, such as PNA, under these conditions.
The solvent will preferentially be buffered and the pH range will be 2-12. Buffers can be selected among the traditionally used buffers in biological assays, but will often be phosphate buffers at pH 7. The salt concentration in the buffer may vary but will preferentially be in the range of between 10 mM and 0.5 M.
It may be advantageous to mix the reaction mixture containing the complex during irradiation, such that homogeneous inducement is achieved.
Further possible ways for inducing electron transfer include stimulation of the electron acceptor at an electrode or by chemical means, or by irradiation with electrons, or by subjecting the samples to ionising electromagnetic radiation.
The time in which the stimulation conditions are maintained depends upon the strength of stimulus. It is apparent to one skilled in the art that if a strong stimulation source is used, inducement may be accomplished in a shorter time than when using a low strength source. In order to keep the time for the assay limited it is preferred to have irradiation times of less than 6 hours, more preferably less than 1 hour. A practical lower limit for the stimulation time may be less than 1 second. At high stimulation power there is a chance that damage to either the sample or the sample's environment will occur.
The electron transfer induced is determined as a measure of the nucleic acid to be determined. The occurrence of the electron transfer can be measured by several methods. They may include determining the electron transfer itself or may comprise the determination of any changes occurring within the complex or its components caused by the electron transfer. It is especially preferred to determine the changes within the electron acceptor either immediately after the electron transfer or at some time later when the process has reached its final state. The most obvious change of the electron acceptor is the absorption of the electron, changing the electronic configuration of the electron acceptor or donor. This change can be measured using either chemical or spectroscopic properties of the now electron-rich electron acceptor. Most preferred are determinations of the spectroscopic properties, such as light absorption or emission at characteristic wavelengths (or quenching thereof). The electron-rich electron acceptor may also be determined by chemical or physical means such as transferring its excess electron to a secondary acceptor or an electrode.
The electron donor has also changed its chemical and spectroscopic characteristics by losing an electron. For example, if guanine is the electron donor, by losing an electron a reactive species is created which renders the nucleic acid backbone susceptible to cleavage specifically at the site of oxidation by piperidine treatment. This chemical reaction is known to one skilled in the art. The occurrence of electronic transfer can then be determined by detecting nucleic acids or probe molecules being shorter in length than expected. This can be made either by direct gel-electrophoresis or after sequencing as well as by liquid chromatography methods.
In case of electron donors changing their spectroscopic characteristics upon losing an electron, the spectroscopic characteristics of the modified electron donor can be determined. Similar to the determination of any changes in the acceptor, it is possible to spectroscopically determine the presence of the created species using its light absorption or fluorescence at characteristic wavelength.
A further indication of the electron transfer is quenching of a possible luminescence of the electron donor or electron acceptor. It is well known to those skilled in the art that electronically excited states of many electron donors or electron acceptors emit light. Electron transfer reactions can effectively quench this emission. In one aspect of this invention, quenching of the luminescence of the excited electron donor or the excited electron acceptor is the means by which formation of the complex between the probe and the nucleic acid is determined. For example, luminescence from the electron donor or electron acceptor will be reduced by formation of the complex if the rate or efficiency of electron transfer is increased by complex formation. In the preferred application of this invention, the luminescence from the excited donor or acceptor is increased by complex formation because the rate or efficiency of electron transfer has decreased as a consequence of complex formation.
In addition to the dependence of the reaction rate on the free energy, reaction rate will also be dependent on distance. In particular, the reaction rate is expected to decrease exponentially with distance:
where r is the distance separating the donor and acceptor and β is a dampening factor with units of distance −1. The dampening factor β′ is a measure of the conductivity of the medium: a low β will allow electron transfer to occur over very long distances while a high β will permit electron transfer at short distances only.
There is currently some controversy regarding the β value for duplex DNA: Barton and coworkers have suggested that β could be very low (<0.2 Å−1) (Murphy, C. J.; Arkin, M. R.; Jenkins, Y.; Ghatlia, N. D.; Bossmann, S. H.; Turro, N. J.; Barton, J. K.; Science 1995, 262, 1025-1029), while Harriman and Brun have measured a β value of 0.88 Å−1 (Brun, A. M.; Harriman, A. J. Am. Chem. Soc. 1992, 114, 3656-3660). The Barton value would suggest that duplex DNA is an exceptional conductor while the Harriman/Brun value would suggest that duplex DNA is an ordinary conductor. The Barton value is not well-substantiated by experiment (i.e. the occurrence of electron transfer has not been demonstrated) so the Harriman/Brun value seems more reliable at this time. It should also be noted that the β values for PNA-DNA and DNA-DNA could be quite different since the electron which is transferred is passing through the π-stack and the structures of the π-stacks are quite different for the two types of duplexes.
If one assumes a “normal” β value of ca. 1.0 Å−1, then a change in distance from 10 to 20 Å would decrease the electron transfer rate by more than four orders of magnitude. Such a strong dependence of the electron transfer rate on distance is the foundation of our distinction between bioconductors and bioinsulators, particularly with respect to the severe time constraints imposed on the system by using photoinduction. The excited state species used as either the donor or acceptor will exist for a relatively short time. In a preferred embodiment, the donor absorbs a photon and has a relatively short lifetime, meaning it will relax to the ground state within a few nanoseconds of excitation. This relaxation can occur by several mechanisms but the most important pathway is fluorescence, in which photon of light is emitted. In the presence of a suitable electron acceptor, resulting in a decrease in the fluorescence. “Suitability” is determined by the acceptor reduction potential: ideally, the potential will be sufficiently low to yield a negative free energy for electron transfer. If the potential is too high, then the electron transfer will be too slow to compete effectively with fluorescence and little quenching will result. Therefore, the strong dependence of electron transfer on distance indicate that, with the proper donor/acceptor pair, one can convert a system from a bioconductor to a bioinsulator simply by increasing the donor-acceptor separation distance. The key is to increase the distance (decrease the reaction rate) until the electron transfer becomes too slow to compete with fluorescence.
In the PNA hairpin system we take advantage of the large increase in distance separating the donor and acceptor after hybridization in order to inhibit electron transfer. The fact that we observed significantly more fluorescence after hybridization than in the hairpin conformation indicates that β is not very small. The ideal fluorescer will have a short lifetime (the shorter the better), a high quantum yield, a low intersystem crossing rate, energetics so that energy transfer is fast, be flat so that it intercalates in the hairpin and the duplex without messing up base-pair recognition by hydrogen bonding, absorb where the anthraquinone does not, fluoresce in the “red” region of the spectrum, and it should also be easy to synthesize.
Such ideal fluorescor would give a substantially black background when no nucleic acid to be determined is present and bright fluorescence in the complex of probe and nucleic acid.
In a preferred mode of the method of the present invention, the rate or efficiency of the electron transfer is different in the probe and in the probe when bound to the nucleic acid. In this case the electron donor and acceptor are preferably bound covalently to the same probe molecule, most preferred at a defined, fixed first distance. Upon binding of the probe to the nucleic acid to be determined, the distance from the electron donor to the electron acceptor is altered, preferably increased, thus decreasing the reaction rate of the electron transfer. The increased distance results in a decreased rate of electron transfer and a concomminant increase in detectable signal. The detectable process will then serve as an indication or measure of the electron transfer and thus of the presence or absence of a nucleic acid to be detected.
A preferred method of the invention therefore is a method wherein the distance between acceptor and donor is a probe is altered upon binding to the nucleic acid, preferably by more than 5 base units in a stack.
In order to evaluate the results from determining of the occurrence of the electronic transfer, one skilled in the art will correlate the signals or results received from experiments wherein the amount or presence of a nucleic acid to be determined is defined. This can then be made by usual calibration experiments, for example, using standard samples containing different specified amounts of nucleic acids to be determined in subjecting the standards to the same conditions as the sample suspected to contain the nucleic acid to be determined. Qualitative and quantitative determination is possible.
An important feature of the formation of the probe-analyte complex is that the distance between the donor and acceptor is relatively fixed.
While these are the core steps of the method of the present invention, this method can be adapted to known convenient formats, for example, by introducing this concept into assays wherein the nucleic acid to be determined is determined in an immobilized state. A very advantageous format of the present invention is disclosed in FIG. 2. In this homogeneous (i.e. single solution) format, the sample containing the nucleic acid to be determined (comp. target DNA) and nucleic acids not to be determined (non-comp. target) in soluble form, the probe molecule containing the electron acceptor and an electron donor are mixed. Only nucleic acids to be determined will hybridize to the specific probe molecule. Nucleic acids not complementary to the probe molecule will not hybridize. Therefore, after inducement of electron transfer, there will be higher or lower light emission from the donor or acceptor if the nucleic acid to be determined was present.
In FIG. 3, the same homogeneous format is described wherein one probe containing an acceptor and another probe containing a donor is used. Only binding of both probes adjacent to each other, whereby the electron transfer is capable of propagating from donor to acceptor will yield a change in light emission.
In FIG. 4, a heterogeneous assay format is disclosed. The probe molecule containing both an electron acceptor (acc.) and an electron donor (Donor) is bound to a solid phase, for example, a polystyrene surface. This can be accomplished either by covalent bonding or using streptavidin-coated surfaces and biotin-labeled probes. Upon excitation of the electron acceptor, electron transfer is induced on the solid phase, and can be detected by identifying the modified donor only if the nucleic acid to be determined was bound to the solid phase near base-pairing via the immobilized probe molecule. The assay format using an immobilized probe is especially advantageous if the sample contains further ingredients disturbing the irradiation of a detection, for example, by absorbing light in the range of the irradiation or emission wavelength.
In an embodiment the nucleic acid to be determined is bound by nucleic acid analyte probe molecules capable of forming a hairpin structure. In this design, the electron donor (Do) and acceptor (Ac) are initially brought into relatively close proximity by the nature of the hairpin structure (FIG. 6). The close location of the electron transfer moieties will generally lead to high electron transfer rates and efficiencies between these two groups. Thus, spectroscopic changes induced by electron transfer will be pronounced. This could, for instance, be either luminescence initiation or luminescence quenching by electron transfer. A preferred application of this invention is the case when duplex DNA or the DNA-PNA complex of the probe and the nucleic acid to be determined has the characteristic of a bioinsulator. In this case when the hairpin probe is hybridised to a target nucleic acid (e.g. a PCR amplicon) electron transfer rates will be reduced compared with the rate in the probe itself and, therefore, less or no luminescence quenching will be observed from the complex than from the probe.
A hairpin structure is synthesized preferentially in a parallel mode. It consists of two hybridisable segments and a hinge segment (a segment linking the bybridisable segments). The two hybridisable segments are complementary to each other and to two non-overlapping segments of the nucleic acid to be determined. Due to the parallel synthesis of the whole hairpin, the two segments will form an antiparallel duplex. The two hybridisable segments are preferentially composed of 6-12 monomers each, each segment containing an electron transfer moiety capable or not capable of forming hydrogen bonds. The electron transfer moieties are preferentially placed in such a way that they are juxtaposed in the hairpin structure on that one of them is an electron donor and one of them is an electron acceptor. The close positioning of the electron transfer moieties makes electron transfer as optimal as possible. The preferred distance of the electron transfer moieties on different segments is between 1 and 10, more preferably 1 and 5 when the segments are hybridized to each other (a distance of 0 meaning base positions forming hydrogen bonds with each other, if bases were present at this position). The hinge segment is composed of 2-7 monomers, preferentially composed of 3-5 monomers.
Stem-loop (hairpin) RNA and DNA structures are known to form readily from appropriate nucleic acid sequences. We have now shown that PNA also form hairpin structures. Characteristically, the thermodynamic stability of the PNA structure, revealed by its melting temperature, is greater than that observed for DNA which contains an identical sequence. In particular, for cases where a change in spatial relationship provides the detected signal, PNA will provide greater spatial differentiation with fewer base pairs than the comparable DNA-based sequence. Exploitation of this advantage requires the ability to prepare specific PNA sequences covalently linked to “reporter” units that do not completely disrupt the hairpin structure. We have developed synthetic technology to incorporate reporter groups linked to internal PNA position.
The hairpin PNA contains an electron donor, for example an amnoacridine group in place of one of the bases on the PNA backbone. By design, the acridine group falls within the duplex region of the PNA hairpin. The melting behavior and, especially, circular dichroism spectroscopy indicate that incorporation of the acridine does not prohibit hairpin formation. Similarly, the evidence indicates that a PNA, which contains two non-nucleic acid groups (an acridine and an anthraquinone) bound in the duplex region of the hairpin, also maintains the ability to form stem-loop structures.
One of the key requirements to be satisfied by a nucleic acid biosensor is selectivity. The probe must recognize its complement selectively and initiate a unique, sensible event. The unrivaled ability of PNA to form duplex, and higher order, structures with complementary DNA and RNA suggested that their modification to include reporter groups would not destroy the recognition ability. The experiments described below reveal this to be the case. The melting behavior observed for mixtures of PNA and, its reaction with a dodecamer DNA complementary to only the loop and a portion of the stem region of the PNA. The melting behavior and the CD spectroscopy of the PNA/DNA structure indicates formation of a hybrid structure that has opened the PNA hairpin.
The preferred aminoacridine unit of non-nucleotide substituent is preferably linked to the PNA backbone through a carbon-nitrogen bond to its 9-amino group. While the properties of 9-aminoacridine has been extensively studied, there are only a few reports concerning the mono N-alkylated aminoacridines. We use 9-(N-methylamino)acridine as a model for the acridine group of Ar. In acetonitrile solution, the fluorescence quantum yield (Φfl) for 9-aminoacridine is 0.96 and its singlet lifetime is 15.8 ns (Kubota et al., J. Phys. Chem. 84, 2855-2861 (1980)). In contrast, under these conditions the (Φfl) for 9-(N-methylamino)acridine is only 0.011 and its singlet lifetime is correspondingly reduced to 2 ns. This dramatic change in photophysical properties is mirrored in a comparison of photoelectron spectra (Li et al., Zh. Obshch. Khim 61, 186-191, 1991). These effects are attributed to a change in excited state character caused by loss of overlap of the acridine π-electron system with the non-bonding electrons of the amino nitrogen atom due to steric effects when the methyl group of 9-(N-methylamino) derivative is co-planar with the acridine. The reduced lifetime of the methylaminoacridine makes it less susceptible to fluorescence quenching by guanine and adenine (Kubota, 1980). Consequently, we observe little difference in the aminoacridine fluorescence intensity between the hairpin and linear forms of PNA.
The aminoacridine fluorescence of the PNA is strongly quenched when it is in its hairpin form. We attribute this to electron transfer from the excited singlet state of the acridine to the nearby anthraquinone group. The oxidation potential of 9-(N-methylamino)acridine is reported to be 0.21 V vs SCE (Shen et al., Science in Clinc. (Ser. B) 35, 137-145 (1992)). The reduction potential of anthraquinone derivatives closely related to Q is −0.58 V vs SCE (Breslin et al., J. Am. Chem. Soc. 118, 2311-2319 (1996)). The energy of the singlet excited state of the aminoacridine in PNA calculated from its fluorescence spectrum is 3.45 eV. Application of the Weller equation (Rehm et al., Isr. J. Chem. 8, 259 (1970)) ignoring the Coulomb work term, which should be negligible, reveals that the free energy for electron transfer (ΔGet) is −2.66 e V for transfer of an electron from the singlet excited state of the aminoacridine to the anthraquinone. This reaction is sufficiently exothermic for it to occur at every encounter between the excited acridine and the quinone.
There is weak residual aminoacridine fluorescence from the hairpin form of the PNA. We attribute this to multiple conformations of the quinone and acridine groups as is suggested by its multi-phase melting behavior. If the excited state aminoacridine group is in contact with the quinone, then quenching should be nearly instantaneous and complete. The precise meaning of “contact” is not clearly defined by these experiments. We take it, provisionally, to mean that both the quinone and acridine are intercalated within the duplex region of the PNA hairpin. In this regard, if either the quinone or acridine is extrahelical when the acridine is excited, it seems unlikely that binding within the duplex will occur within the ca. 2 ns lifetime of the excited state. The data show that the aminoacridine fluorescence increase ca. 3.3 fold when the PNA is converted from its hairpin to linear form. Interpreted within the multi-conformational model, his finding indicates the ratio of PNA structures where there is quinone/acridine are in contact to those where one group is extrahelical is ca. 3.3:1.
The data show that the reaction of the hairpin PNA with either DNA or DNA results in a ca. 6.5 fold increase in aminoacridine fluorescence intensity. We attribute this to conversion of the hairpin form of the PNA to a linear hybrid duplex in which the acridine and quinone are not longer in contact. There has been considerable recent discussion concerning the ability of DNA to conduct holes (radical cations) and electrons (Beratan et al., Chemistry and Biology 4, 3-8 (1997)). Barton and co-workers (Arkin et al., Chem. & Biology 4, 389-400 (1997); Dandliker et al., Science 275, 1465-1468 (1997), Hall et al., Nature 382, 731-735 (1996)) have argued for very rapid transport. Meade and co-workers (Meade et al., Angew. Chem. Int. Ed. Engl. 34, 352-354 (1995)) report a slower rate. Our findings suggest that on the short time scale of the aminoacridine lifetime, the PNA/DNA hybrid duplex behaves as an insulator not a conductor. Thus, whether intercalated or not, the quinone does quench the aminoacridine fluorescence when the two groups are separated by twelve pairs in the hybrid duplex.
The use of PNA in the construction of probes to sense nucleic acids offers several advantages over using DNA as the sequence-selective recognition element. PNA/DNA hybrid duplexes provide exquisite single-base-mismatch selectivity (Wang et al., Anal. Chem. Acta 344, 111-118 (1997)) and offer greater stability than their DNA/DNA or DNA/RNA counterparts. This is especially advantageous in the hairpin format of the molecular beacons. The duplex region of the hairpin will generally not be fully complementary to the targeted nucleic acid, and this region can be shorter for a PNA probe than for a DNA probe. Also, since PNA is unnatural, there are no nuclease enzymes that destroy it. This offers a special advantage in homogeneous assays because fewer sample-preparation steps will be required. Finally, and perhaps of greatest significance, the fluorescer and quencher groups can be easily placed in internal positions in PNA. PNA is easily synthesized by standard peptide methods and the required monomeric derivatives are readily prepared by simple reactions. Internal positions for the fluorescer and quencher provide the prospect of high sensitivity since there will be no “fraying” that is likely to occur when the fluorescer and quencher are bound at opposite termini of the hairpin duplex.
In another embodiment the close electronic interaction between the donor and the acceptor is also used (see FIG. 7). In this embodiment the analyte specific probe (PNA2) contains the donor (or the acceptor). A second probe (PNA1) containing the other electron transfer moiety is complementary to a part of the first probe. When these are hybridised the interaction between the acceptor and the donor is as outlined above. When the analyte is introduced to the mixture, the short acceptor probe will be displaced by the analyte thus preventing the electron transfer interaction between the donor and the acceptor. Depending on the choice of donor/acceptor this could lead to either luminescence initiation or quenching.
Further subject matter of the invention are molecules of the general formulae I and II.
The compounds of the general formula I can be used advantageously in the above-disclosed method for determining a nucleic acid in a sample as the nucleic acid analogue probe molecule. These compounds can be prepared analogous to the synthesis described in WO 92/20702, WO 94/25477, WO 96/20212, EP-0 672 677 and EP-0 700 928, just replacing one or more monomers used in the oligomerisation with one ore more monomers containing non-nucleobase containing electron acceptors or non-nucleobase containing electron donors. A preferred way to produce the compounds of general formula I is the step-wise chemical synthesis according to WO 92/20702, incorporating as a monomer a compound of the general formula VIIIa-VIIIc
wherein the definitions of A, B, C and D are chosen from the definitions of A1-An, B1-Bn, C1-Cn and D1-Dn in formula I, respectively with the condition that any amino groups therein may be protected by amino protecting groups; E is COOH CSOH, SOOH, SO2OH or an activated derivative thereof; F is NHR3 or NPgaR3, where R3 is as defined above and Pga is an amino protecting group and L is a non-nucleobase electron donor or acceptor moiety.
Preferred monomers are amino acids having formula (IX)
wherein X′ is a carboxylic acid protection group or hydrogen, R7′ is independently selected from the group consisting of hydrogen and the side chains of naturally occurring alpha amino acids, or amino-protected and/or acid terminal activated derivatives thereof.
Therefore, a preferred method of preparing a molecule of general formula I is characterized in that a molecule of general formula VIIIa-VIIIc or IX wherein X′ is hydrogen is reacted with a compound of general formula X
K is a solid phase, like amino modified glass, or a solid phase bound or free compound of general formula I or II or a compound of formula I in protected form. These compounds can further be modified by attaching monomeric units not containing an electron transfer moiety.
Any reactive groups like primary amino groups or hydroxyl groups are preferably protected by removable or non-removable protecting groups, like benzyloxycarbonyl, Boc or Fmoc if they are not intended to react in the elongation reaction.
The following examples shall explain the invention in more detail using PNA as an example of a probe molecule. The PNA oligomers described above are synthesised by solid phase synthesis procedures. The preferred solid support is polystyrene but also other solid supports such as ®Tentagel and ®Controlled Pore Glass may be used. The synthesis proceeds from the C-terminal position of the PNA and proceeds via couplings of monomers and/or amino acids protected with acid labile protecting groups. Reactive groups on the bases/side chains are also protected. After deprotection of the moiety directly linked to the solid support the subsequent building blocks (monomers/amino acids) are then coupled in the desired sequence. The deprotection of the N-terminal position is performed prior to coupling of the following monomeric unit.
The coupling of the monomeric unit is performed via activation of the carboxylic residue. Such activation is well known to those skilled in this art and, as examples, are: HATU, HBTU, and carbodiimides. The coupling is performed in solvents normally used in peptide chemistry, that is NMP, DMF, acetonitrile, pyridine, dichloromethane or mixtures thereof Activation enhancers such as DMAP may be added. The activated monomeric derivative will be used in excess to the previously deprotected residue and typically in a 2 to 10 times excess. If a high coupling yield is detected (typically >99%) the coupling step will be followed by a capping step, which comprises treatment of the coupled resin bound oligomer with acetic anhydride. The capping mixture comprises 2-20% (v/v) acetic anhydride in NMP/pyridine. If unsatisfactory coupling is detected the coupling step will be repeated to obtain a high coupling yield.
Coupling of a conjugate containing a carboxylic acid can be done analogously to the above described procedure. Other ligands not containing a carboxylic acid can also be linked to the resin bound oligomers. Such other linkages can be performed via isocyanates, isothiocyanates (e.g. fluorescein isothiocyanate), carbonic acid active esters, and sulphonyl chlorides.
The described stepwise synthesis of PNA can be performed manually or automatically. In the latter case commercially available synthesisers (e.g. peptide synthesisers, multiple peptide synthesisers, DNA synthesisers) can be used, provided that the hardware in the instruments is compatible with the chemistry used during the PNA synthesis.
The synthesised oligomer is finally removed from the resin. This is preferentially done by treatment with strong acid, such as TFMSA or HF. The purification of the liberated oligomer is done by HPLC and/or ion exchange. The identity of the pure material is confirmed by mass spectrometry and HPLC.
The letter designation of amino acids and the orientation of PNA follows the traditionally used nomenclature.
Designates a monomeric unit composed by: 3,6-Diaza-(N3-2-
Designates a monomeric unit composed by: 3,6-Diaza(N3-boc-
Designates a monomeric unit composed by: 3,7-diaza-(N3-boc-
Designates a monomeric unit composed by: 3,10-diaza-(N3-boc-
Nuclear magnetic resonance
Peptide nucleic acid according to WO 92/20702
Oligomers are designated according to the PNA numbers assigned by PNA Diagnostics.
Anthraquinone-2-carboxylic acid (2 g, 7.9 mmol), DCC (1.7 g, 8.3 mmol), HOBT (1.08 g, 8.0 mmol) and methyl (N-(2-Boc-aminoethyl))glycinate (2 g, 8.6 mmol) were dissolved in DMF (25 mL) and stirred at room temperature overnight. The reaction mixture was filtered and the filtrate was washed with DCM (2×25 mL). The solution was extracted with diluted NaHCO3 (3×25 mL), 2 M NaHSO4 (2×25 mL) and brine. The organic phase was dried with MgSO4, filtered and evaporated to dryness under reduced pressure. The yellow foam was dissolved in THF (10 mL) and 1 M LiOH (30 mL) was added. The mixture was stirred for 2 h. THF was removed from the solution under reduced pressure and pH was adjusted to 2.8 with 2 M NaHSO4. The precipitate was extracted with DCM (2×25 mL), evaporated to dryness and then redissolved in ethyl acetate (3 mL). The ethyl acetate solution was poured into hexane (150 mL) whereby the target molecule precipitated. Yield 3.1 g (87%).
H1-NMR DMSO-d6 δ: 7.81-8.30 (m, 7 H, aromatics); 6.98 and 7.72 (m, 1 H, BocNH); 4.17 and 3.97 (s, 2 H, CH2O); 3.52 and 3.24 (m, 2 H, CH2); 3.21 and 3.02 (m, 2 H, CH2) 1.17 and 1.19 (s, 9 H, Boc).
Anthraquinone-2-carboxylic acid (3 g, 11.9 mmol), DIEA (3.1 ml, 24 mmol) and HATU (4.5 g, 11.9 mmol) in DCM (30 mL) was stirred for 10 min at room temperature. Methyl glycinate hydrochloride (1.5 g, 11.9 mmol) was added after which the product started to precipitate. The mixture was stirred for an additional 3 h. The precipitated material was collected by filtration. The volume of the mother liquor was reduced to ⅓ and an additional crop was collected. Yield 3.65 g (93%).
Methyl (2-anthraquinoyl)-3-aza-butanoate (3.65 g, 11.3 mmol) was stirred for 1 h at room temperature in 1 M LiOH (20 mL) and THF (3 mL) whereby the starting material dissolved. pH of the solution was adjusted to 2.8 with 2 M NaHSO4 and the target molecule precipitated. The organic solvents were removed from the suspension under reduced pressure and the precipitate was collected by filtration and dried. Yield 3.5 g (100%).
H1-NMR DMSO-d6 δ: 9.37 (t, 1 H, NH); 8.67 (d, 1 H, H-laq); 8.36 (dd, 1 H, H-3aq); 8.30 and 8.28 (1 H, H-4aq); 8.23 (m, 2 H, H-5 and H-8aq); 7.95 (m, 2 H, H-6 and H-7aq); 4.00 (d, 2 H, CH2).
3,6-Diaza(N3-Boc-aminoethyl)-4,7-dioxo-7-(2-anthraquinyl)-heptanoic acid (Q2)
4-(2-Anthraquinoyl)-4-oxo-3-aza-butanoic acid (3.5 g 11.5 mmol), DCC (2.6 g, 12.65 mmol), HOBT (1.7 g, 12.65 mmol) and methyl (N-(2-Boc-aminoethyl))glycinate (3.2 g, 13.8 mmol) was stirred in DMF (50 mL) at room temperature for 48 h. The reaction mixture was filtered and the residue was washed with DCM (2×30 mL). The organic phase was extracted with diluted NaHCO3 (3×30 mL), 2 M NaHSO4 (2×30 mL), brine, dried with MgSO4 and evaporated to dryness. Crystallization from ethyl acetate gave 4.0 g (67%) of methyl 3,6-Diaza(N3-Boc-aminoethyl)-4,7-dioxo-7-(2-anthraquinyl)-heptanoate. The methyl 3-aza-3-(N-2-Boc-aminoethyl)-4,7-dioxo-6-aza-7-(2-anthraquinoyl)-heptanoate was hydrolysed at room temperature in 1 M LiOH and 10% THF. The pH of the solution was adjusted to 2.8 with 2 M NaHSO4 and the target molecule precipitated as an oil. The oil was extracted with ethyl acetate and the organic phase was dried with MgSO4. The volume was reduced to 2 mL and hexane (250 mL) was added with stirring, whereby 3-aza-3-(N-2-Boc-aminoethyl)-4,7-dioxo-6-aza-7-(2-anthraquinoyl)-heptanoic acid precipitated. Yield 3.7 g (92%).
H1-NMR DMSO-d6 δ: 12.64 (s, 1 H, COOH); 9.14 (m, 1 H, NH); 8.69 (d, 1 H, H-laq); 8.67 and 8.35 (dd, 1 H, H-3aq); 8.32 and 8.30 (dd, 1 H, H-4aq); 8.25 (m, 2 H, H-5 and H-8aq); 7.69 (m, 2†H, H-6 and H-7aq); 6.90 and 6.75 (t, 1 H, BocNH); 4.24 (d, 2 H, CH2CO); 4.24 and 4.12 (dd, 2 H, CH2CO); 3.90 (s, 2 H, CH2CO); 3.44, 3.17 and 3.06 (m, 2 H, CH2); 1.40 and 1.38 (s, 9 H, Boc).
β-Alanine (1 g, 11.2 mmol) was added to a solution of 9-phenoxyacridine (2.7 g, 10 mmol) in phenol (15 g). The suspension was stirred at 120° C. for 2 h. The solution was cooled to room temperature and poured into ether whereby the product precipitated as a yellow-green solid. It was triturated with hot ethanol, filtered and then washed with ethanol, giving crude 9-(2-carboxyethyl)aminoacridine. Yield 1.9 g (71%).
H1-NMR DMSO-d6 δ: 8.27 (d, 2 H, acr. 1,8); 7.59 (m, 4 H, acr. 2,3,6,7); 7.26 (m, 2 H, acr. 4, 5); 4.05 (t, 2 H, CH2CO); 2.82 (m, 2 H, CH2N). C16H14N2O2: calc/found: 266/266.
3,7-diaza-(N3-Boc-aminoethyl)-(N7-9-acridinyl)-4-oxo-heptanoic acid (R1)
A mixture of 9-(2-carboxyethyl)aminoacridine (1.7 g 6.4 mmol), DIEA (1.8 g 13.8 mmol) and HATU 2.68 g, 7 mmol) was stirred in DMF (40 ml) at room temperature for 10 min. Methyl-Boc-aminoethylglycinate (1.8 g, 7.8 mmol) was added and the solution was stirred at 60° C. for 4 h. The solution was cooled and DCM (100 ml) was added. Subsequently the solution was extracted with 0.5 M NaHCO3 (3×40 ml), 2 M NaHSO4 (2×40 ml) and brine. The organic phase was dried with MgSO4 and evaporated to dryness. The residue was dissolved in THF/1 M LiOH (2/10) 60 ml The pH of the solution was adjusted to 2.8 with NaHSO4 whereby an oil precipitated. The water phase was decanted off and the oil was dissolved in ethanol (10 ml). The solution was purred into ether and 3,7-diaza-(N3-Boc-aminoethyl)-(N7-9-acridinyl)-4-oxo-heptanoic acid (R1) precipitated. Yield 2.3 g (77%).
H1-NMR DMSO-d6 δ: C16H30N4O5: 8.50 (dd, 2 H, acr. 1, 8); 7.91 (m, 2 H, acr. 2, 7); 7.84 (m, 3 H, acr. 3,6 and 9 NH); 7.49 (m, 2 H, acr. 3,6); 6.89 and 6.76 (m, 1 H, BocNH); 4.28 (m, 2 H, CH2CO); 4.05 and 3.92 (s, 2 H, CH2CO); 2.82 (m, 2 H, CH2N). C25H30N4O5: calc/found: 466/466
6-Aminohexanoic acid (1.47 g, 11.2 mmol) was added to a solution of 9-phenoxyacridine (2.7 g, 10 mmol) in phenol (15 g). The suspension was stirred at 120° C. for 2 h. The solution was cooled to room temperature and poured into ether whereby the product precipitated as a yellow-green solid. It was triturated with hot ethanol, filtered then washed with ethanol, giving crude 9-(5-carboxypentyl) Aminoacridine. Yield 2.4 g (78%).
H1-NMR DMSO-d6 δ: 8.27 (d, 2 H, acr. 1, 8); 7.61 (m, 4 H, acr. 2,3,6,7); 7.26 (m, 2 H acr. 4, 5); 3.79 (t, 2 H, CH2CO); 2.16 (t, 2 H, CH2N); 1.72, 1.50 and 1.37 (m, 2 H, CH2). C19H20N2O2: calc/found: 308/308.
3,10-diaza-(N3-Boc-aminoethyl)-(N10-9-acridinyl)-4-oxo-decanoic acid (R2)
A mixture of 9-(5-carboxypentyl)aminoacridine (1 g, 3.2 mmol), DIEA (0.9 g, 6.9 mmol) and HATU (1.34 g, 3.5 mmol) was stirred at room temperature for 10 min in DMF (30 mL). Methyl-Boc-aminoethylglycinate (0.9 g, 3.8 mmol) was added and the solution was stirred at 60° C. for 4†h. The solution was cooled and DCM (60 mL) was added. Subsequently the solution was extracted with 0.5 M NaHCO3 (3×30 mL), 2 M NaHSO4 (2×30 mL) and brine. The organic phase was dried with MgSO4 and evaporated to dryness. The residue was dissolved in THF/1M LiOH (2/10) (40 mL). The pH of the solution was adjusted to 2.8 with NaHSO4 whereby an oil precipitated. The water phase was decanted off and the oil was dissolved in ethanol (5 mL). The solution was then poured into ether and 3N-(Boc-arninoethyl)-10N-(9-acridinyl)-3,10-diaza-4-oxy-nonanicacid precipitated. Yield 1.2 g (76%).
H1-NMR DMSO-d6 δ: 8.51 (d, 2 H, acr. 1, 8); 7.90 (m, 2 H, acr. 2, 7); 7.82 (m, 2 H, acr. 3, 6); 7.71 (m, 1 H, NH); 6.83 and 6.45 (m, 1 H, BocNH); 7.51 (m, 2 H, acr. 4, 5); 4.01 (m, 2 H, CH2CO); 3.98 and 3.88 (s, 2 H, CH2CO); 3.23, 3.00, 2.30, 2.03 and 1.88 (m, 2 H, CH2); 2.16 (t, 2†H, CH2N); C27H36N4O5: calc/found: 508/508.
To a solution of methyl N-(Boc-aminoethyl)glycinate (2 g, 8.6 mmol) in DCM (30 mL) was added acetic anhydride (0.9 g, 8.8 mmol) and pyridine (1 mL). The mixture was stirred at room temperature for 2 h. The reaction was extracted with 0.5 M NaHCO3 (3×30 mL), 2 M NaHSO4 (2×30 mL) and brine. The organic phase was dried with MgSO4 and evaporated to dryness. The residue was dissolved in THF/LiOH(1M) 2/10 (40 mL) and stirred for 1 h after which the pH was adjusted to 2.8 with NaHSO4. The solution was extracted with ethyl acetate and the organic phase was dried with MgSO4; 3,6-diaza-(N3-acetyl, N6-Boc)-hexanoic acid was collected by evaporation to dryness. Yield 1.8 g (82%).
H1-NMR DMSO-d6 δ: 12.31 (s, 1 H, COOH); 6.83 and 6.69 (t, 1 H, BocNH); 4.07 and 3.89 (s, 2†H, CH2CO); 3.30 (m, 2 H, CH2); 3.09 (m, 2 H, CH2); 2.00 and 1.89 (s, 3 H, CH3); 1.37 (s,9 H, Boc).
The monomers described above were essentially incorporated by the oligomerisation procedure previously described and what has been disclosed in WO 92/20702. However, the PNA syntheses containing the donor moieties constituted a special case because the capping step was omitted in the part of the synthesis including and following the donor monomer. However, to obtain high coupling yields the monomeric moieties of this part of the synthesis was double coupled. Later experiments have shown that capping may be used during oligomerisation of donor containing molecules. All oligomers were cleaved from the resin by TFMSA and subsequently HPLC purified. The identity was confirmed by mass spectroscopy (MALDI-TOF).
PNA 554: calc./found: 5237/5239
PNA 555: calc./found: 5469/5467
PNA 579: calc./found: 5429/5426
PNA 586: calc./found: 5336/5333
PNA 626: calc./found: 6631/6636
PNA 627: calc./found: 6563/6564.
Concentrations of PNA stock solutions were estimated by UV absorbency at 260 nm. The extinction coefficients of the PNA oligomers were calculated using the nearest neighbor values for DNA, with Adenine substituted for Q or R. (Because of the length of the PNA oligomers, the error introduced by this substitution will be negligible.) In a typical measurement, PNA was dissolved in water and the absorbency at 260 nm was measured after incubation for 5 minutes at 75° C. in order to minimise secondary structure in the PNA. (Extinction coefficients determined from room temperature absorbency measurements tend to be 5-10% lower than those measured at the higher temperature). DNA oligomers were purchased from Midland Certified Reagent Company.
This example demonstrates the ability of PNAs containing an electron acceptor to hybridize with complementary DNA oligomers.
Samples were prepared consisting of equimolar concentrations of PNA and DNA oligomers (1.0 or 2.0 μM each) in 1.0 mL of 10 mM sodium phosphate buffer (pH=7.0). The PNA strand will often precipitate upon addition of the phosphate, presumably due to complexion of the lysine units by the phosphate. Mixing and addition of the DNA strand results in dissolution of the PNA. Samples were placed in cuvettes (1.5 mL capacity, 1.0 cm pathlength) and sealed with tape to prevent evaporation of water during heating/cooling cycles. The absorbency of the samples at 260 nm was monitored as a function of temperature for three consecutive runs: heating at 1.0° C./min, and cooling at 0.5° C./min and heating again at 0.5° C./min. In two experiments the absorbency was monitored at 330 nm, where the AQ unit absorbs. Due to the low extinction coefficient of the AQ, these experiments were performed at 20 μM each of PNA and DNA.
Data were analysed by exporting as an ASCII file and importing into a standard graphics software. The absorbency was plotted versus temperature for each sample. Melting temperatures (Tm) were determined as the maxima of plots of the first derivative of absorbency with respect to temperature, assuming a first order phase transition. Tm values given in the text and tables have error values of ±0.5° C.
An initial question to address regarded what could be tolerated on the DNA strand at the central site, i.e. directly across from the AQ in the hybrid. Five variations were considered: (1) An abasic site (X) was incorporated into the oligomer. This residue replaces the standard DNA base with a hydrogen atom at carbon-1 of the deoxyribose moiety and would be expected to provide the most room for accommodation of the AQ within the duplex. (2)-(5) Each of the four DNA bases: G, A, C and T were incorporated into the sequence. These DNA oligomers are identified as DNA579Z, where Z=X (Abasic), G, A, C, and T, respectively.
The PNA579/DNA579X hybrid melts with high cooperativity at Tm=61.4° C. (FIG. 12). There is no hysteresis in the transition indicative of rapid hybridisation kinetics as previously observed for full-length 1:1 PNA-DNA duplexes. The PNA586/DNA579X hybrid melts at 68.3° C., approximately seven degrees higher than for PNA579. The greater stabilization for PNA586 could be due to the increased flexibility which arises from the longer linker connecting the AQ to the PNA backbone. In both cases, the Tm is significantly higher than when the PNA bears only an acetyl group at the central position, indicating that the AQ moiety stabilizes the PNA/DNA hybrid (Table 1).
Monitoring the Tm at 330 nm, where only the AQ unit absorbs light, reveals monophasic transitions for both PNA/DNA hybrids (FIG. 12). The hyperchromicity observed for the PNA586/DNA579X hybrid is nearly twice as great as that for the PNA579/DNA579X hybrid, consistent with a structure in which the AQ unit is intercalated into the duplex and that insertion of the AQ is more favorable for the longer linker.
Effect of AQ Linker Length on PNA/DNA Hybrid Stability
Tm (° C.)
The PNAs form stable hybrids with the DNA579Z oligomers in which Z=G, A, C or T (Table 2). The resulting hybrids are only slightly less stable than the corresponding duplexes with Z=X.
Effect of Opposed Base on PNA/DNA Hybrid Stability (Tm given in ° C.)
Effect of Single Mismatches. PNA555 is analogous to PNA579 with the exception of replacing C with G at position 7. Hybridisation of this PNA with its abasic DNA complement (DNA555X) yields a duplex which melts at 63.1° C. The 2° C. stabilisation relative to PNA579/DNA579X could reflect greater base stacking in the region of the C-7 (PNA)/G-32 (DNA) base pair. Hibridization of PNA555 with DNA579 yields a duplex which possesses a G—G mismatch and melts at 53.4° C. Thus, a single mismatch depresses Tm by 8-10° C.
Effect of PNA Orientation on Hybrid Stability. The preferred orientation of PNA/DNA duplexes is antiparallel, where the PNA N-termini are aligned with the DNA 3′-termini. If the PNA579 sequence is inverted, hybridization with DNA579 produces a duplex which melts at 49.9° C. Thus, parallel orientation of the PNA and DNA strands results in reduction of the Tm by ca. 11° C.
These experiments demonstrate that the DNA recognition properties of the PNA(AQ) conjugates are completely analogous to those of unmodified PNA strands. While the stability of the resulting duplexes are most likely lower than would be observed for a 19 base pair PNA-DNA hybrid (due to the lack of base-pairing at the central AQ position), the presence of the AQ moiety stabilises the duplex by >4° C. relative to the case where both strands contain abasic residues at the central position, indicative of a strong interaction (stacking) between the AQ and the duplex.
This example describes the ability of the PNA containing an anthraquinone (AQ) to initiate photocleavage of the DNA strand within a hybrid duplex by photoinduced electron transfer. Photocleavage of the DNA strand within PNA-AQ/DNA hybrids was studied using radiolabelled DNA and polyacrylamide gel electrophoresis (PAGE). Synthetic DNA oligomers were labeled at the 5′-OH terminus using [γ-32P]-ATP and T4 polynucleotide kinase according to standard procedures. 5′-endlabelled oligomers were purified by 20% denaturing PAGE and precipitation. Samples were prepared for irradiation by mixing 5.0 μM each of PNA and unlabelled DNA as well as labeled DNA (2000-4000 cpm per sample) in 10 mM sodium phosphate buffer (pH=7.0). This ensures that there will be a slight excess of DNA over PNA, minimising contributions from unhybridised PNA. Samples were heated to 85° C. for 5 min then allowed to cool to room temperature. In cases where the only difference in samples would be the irradiation time, a single sample was prepared by this procedure and irradiated, with aliquots removed at the desired times.
Irradiation was performed using a Rayonet photoreactor equipped with 8 lamps (λmax=350 nm). Samples were placed in microcentrifuge tubes and suspended parallel to the lamps via a rotating sample holder. While most of the exciting light is either not incident on the tubes or scattered by them, the tubes are sufficiently transparent at 330 nm to permit photocleavage to proceed in reasonable times (less than one hour).
For experiments involving methylene blue, PNA-DNA hybrids were prepared as described above, then methylene blue was added to give 10 or 20 μM concentration. Samples were irradiated for 15 min using the filtered (λ>600 nm) output of a 150W Hg arc lamp.
For experiments in which D2O was substituted for water, all of the components (PNA, DNA, buffer, methylene blue where desired) were mixed, then the water was evaporated under vacuum. 20 μl D2O was added to the samples and then removed by evaporation. This procedure was repeated, then the samples were finally dissolved in the appropriate amount of D2O. An identical procedure was followed using H2O for control samples.
After irradiation, the salt concentration was increased by addition of sodium acetate (pH=5.2) to 0.3 M, and magnesium chloride to 10 mM in a total volume of 50 μL. 100 μL cold ethanol was added and after vortexing, samples were placed on dry ice for 30 min. followed by centrifugation at 12,000 g for 30 min. The supernatant was discarded and the pellet washed with 80% ethanol prior to drying. Samples were either then suspended in denaturing loading buffer or incubated with 100 μL piperidine (1 M) for 30 min at 90° C. After the piperidine was removed by vacuum evaporation, the DNA was dissolved in 20 μL water which was then evaporated. This procedure was performed once more and then the DNA was suspended in denaturing loading buffer. DNA fragments were ultimately separated on a 20% denaturing polyacrylamide gel and cleavage sites were visualised by autoradiography.
Irradiation of a PNA579/DNA579X duplex with 350 nm light for one hour led to very little spontaneous degradation of the DNA. However, after piperidine treatment, strong cleavage is observed at several positions (FIG. 13, Lane 8). Cleavage is not observed for a PNA554/DNA579X hybrid, which lacks an AQ moiety (Lane 4). The main cleavage sites are the three GG steps as well as the abasic site at the center of the duplex. Previous work has shown that irradiation of DNA-intercalated AQ leads to piperidine-dependent cleavage with high selectivity at the 5′-G of GG doublets. It has unambiguously been shown in the literature (Breslin, D. T. and Schuster, G. B. J. Am. Chem. Soc. 1996, 118, 2311-2319) that this GG-selective cleavage of DNA is initiated by photoinduced electron transfer from the DNA bases to the excited state AQ.
The similar selectivity observed in FIG. 13 suggests that the AQ reacts similarly with the PNA/DNA hybrid.
For the two GG sites on the 3′ end of the duplex, cleavage at the 5′ G is favored over that at the 3′ G. Thus, the two GG sites oriented in the 3′ direction relative to the AQ exhibit the same 5′-dependent cleavage as observed in duplex DNA. However, cleavage at the GG step on the 5′-side of the AQ is distributed evenly between the two sites. The 5′-νs 3′-distribution of cleavage at the GG step has been proposed to depend on the angle of rotation between the two guanines as they stack atop one another (Sugiyama, H. and Saito, I., J. Am. Chem. Soc. 1996, 118, 7063-7068). However, it is important to note that trapping requires not only a significantly lower oxidation potential at the GG site but also a chemical reaction to prevent the electron and hole from recombining. This step involves addition of either water or oxygen to the guanine radical cation, a process which will be influenced by the exposure of the base to solvent. It has been noted that the bases in PNA-DNA hybrids are closer to the exterior of the helix than in B-form DNA. NMR Erikkson, M.; Nielsen, P. E. Nature Struct. Biol. 1996, 3, 410-413) and x-ray diffraction data (13etts, L.; Josey, J. A.; Veal, J. M.; Jordan, S. R. Science 1995, 270, 1838-1841) indicate that the degree of inter- and intrastrand stacking within PNA-DNA hybrids is highly sequence dependent. Thus there is no reason to expect that the GG cleavage will exhibit the same 5′-preference as observed in duplex DNA. This is supported by the cleavage data on PNA/DNA hybrids disclosed in this invention.
Experiments were performed with PNA586/DNA579X which were analogous to those performed with PNA579. The results for the longer linker are similar to the shorter linker: After piperidine treatment, cleavage is observed at all three GG sites as well as at the abasic site (FIG. 13, Lane 12). The 5′-G is clearly favored for the two sites which are on the 3′-side of the abasic site while the two Gs are cleaved approximately equally at the other GG site. One notable observation is that the cleavage efficiency for the longer linker connecting the AQ to the PNA backbone is significantly lower than for the shorter linker.
It is clear that the cleavage is occurring at positions which are far from the AQ. In particular, the distal GG step in the 3′ direction is ca. 22 Å away from the AQ within the hybrid (based on a rise of 3.6 Å per base pair). Even if the AQ were able to adopt extrahelical conformations, the short linker connecting it with the backbone precludes direct reaction between the AQ and the remote G sites. We next investigated the possibility that the AQ was generating a freely diffusible species. If the PNA and DNA strands are annealed in the presence of a tenfold excess of unlabeled DNA579X, the cleavage is completely eliminated. However, if the annealing is performed in the presence of an excess of unlabeled, non-complementary DNA, no effect on the cleavage is observed. This rules out the possible intermediacy of a freely diffusing cleavage agent. Additionally, if the excess unlabeled DNA579X is added after annealing but prior to irradiation, minimal attenuation of the cleavage is observed. This indicates that the PNA/DNA hybrid is thermally stable throughout the irradiation time. These experiments demonstrate that the cleavage arises from excitation of AQ moiety and that the damage is localised at the same duplex wherein the photon is absorbed.
Selective cleavage at G residues by photonucleases in duplex DNA is typically the result of either electron transfer chemistry or singlet oxygen, which can be generated by triplet state molecules. We tested for the involvement of singlet oxygen by comparing the cleavage efficiencies for PNA/DNA hybrid in the presence of D2O with H2O. (The lifetime of singlet oxygen is increased nearly tenfold in the presence of D2O, leading to significantly enhanced cleavage by singlet oxygen generators). There is no enhancement of cleavage for the sample irradiated in D2O, arguing against a role for singlet oxygen in the mechanism. We also studied cleavage of the DNA strand in the PNA/DNA hybrid using a known singlet oxygen generator, methylene blue. Irradiation with visible light (λ>600 nm) results in selective cleavage at the guanine residues with a pattern which is similar to that of the AQ-sensitised cleavage. However, this chemistry is strongly enhanced when D2O is substituted for H2O.
At first consideration, the similar cleavage patterns observed for direct irradiation of the AQ and for singlet oxygen generation by methylene blue would suggest that the quinone-mediated cleavage also involves singlet oxygen. However, the lack of inhibition by non-specific DNA demonstrates that cleavage is not mediated by freely diffusing singlet oxygen. The only way that singlet oxygen could be responsible for the cleavage is if it is generated by the AQ at the central position of the duplex, then diffuses one-dimensionally along one of the grooves in either direction until it reacts with a guanine. The different intensities observed for various Gs would then reflect access of the base to the singlet oxygen molecule. Such a process should exhibit a distance dependence, however, since the first GG step in the 3′-direction would be expected to be a barrier to cleavage at the distal GG step. The equal cleavage observed at these two sites is clearly inconsistent with such a mechanism. Further evidence against a singlet oxygen dependent mechanism comes from the observation that anthraquinone-2-sulfonate, which is freely diffusing in solution, does not induce cleavage of the PNA-DNA hybrid. It is unlikely that the covalently linked AQ can generate singlet oxygen when the unbound AQ cannot, particularly in light of the strong phosphorescence quenching (see Example 10) within the duplex. A further argument against singlet oxygen derived cleavage comes from densitometric analysis of the ratio of cleavage at the 5′ and 3′ guanine residues in the distal GG site toward the 3′ end of the duplex. For the AQ-initiated cleavage, this ratio is 1.40, whereas for methylene blue it is 0.69. If the same intermediate were responsible for the DNA damage, then the ratio of 5′/3′ cleavage should be the same for AQ and methylene blue.
In light of the preceding discussion, the only explanation for the observed cleavage is electron transfer in the PNA/DNA duplex. The AQ triplet is quenched by electron transfer from an adjacent base, resulting in the injection of a hole within the duplex. This hole can subsequently migrate through the helix until it is trapped at a GG site (see FIG. 5). Incorporation into the duplex of a “trap” site, i.e. a site with an even lower oxidation potential than GG, should significantly alter the cleavage efficiency at the GG sites. The guanine oxidation product, 7,8-dihydro-8-oxoguanine (8-OxoG) was chosen as a potential trap site since its oxidation potential is estimated by Sheu and Foote to be ca. 0.4 V lower than that of guanine (Sheu, C.; Foote, C. S. J. Am. Chem. Soc. 1995, 117, 6439-6442). DNA579T(OxoG) is analogous to DNA579T except that G-13 is replaced by 8-OxoG. This DNA was radiolabeled on the 5′-terminus according to standard procedures, then hybridized with PNA579 and irradiated. After piperidine treatment, a significant enhancement is observed in the cleavage at the 8-OxoG position, while cleavage at the distal GG site is significantly suppressed relative to the case where a normal G is present at position 13. (See FIG. 14, compare lanes 8 and 4). This observation is consistent with a model in which the excited state AQ accepts an electron from an adjacent base at the intercalation site, injecting a hole into the PNA/DNA helix. This hole migrates along the helix by a series of discrete electron transfers until it reaches a GG site where it can be trapped by reaction with water or oxygen. The (8-OxoG)G site is a more effective trap than is a normal GG site, inhibiting migration of the hole to the distal GG site. Significantly, cleavage at the GG site in the other direction is unaffected by the 8-OxoG.
This example uses phosphorescence spectroscopy to study the reaction of the excited state AQ with the PNA/DNA hybrid.
Phosphorescence emission was measured as an indication of electron transfer from the bases of the hybrid to the photoexcited AQ, since electron transfer quenches this emission. (The quantum yields for phosphorescence are sufficiently low that the samples have to be prepared in a frozen glass matrix in order to be able to detect the phosphorescence. The matrix used was 30% ethylene glycol in 10 mM phosphate buffer, pH=7.0. Stable hybrids are formed even in the presence of the glycol as evidenced by a suppression of the Tm of less than 5° C.) Samples were prepared consisting of AQC(2) (a water soluble AQ), or PNA-AQ alone or with complementary DNA. In each case, the concentration of AQ (linked or unlinked to PNA) was 5.0 μM, as was the DNA concentration for the last sample. Samples were prepared in 10 mM sodium phosphate buffer (pH=7.0) and 30% ethylene glycol, which is required in order to form a low-temperature glass. For PNA/DNA hybrids, samples were heated to 85° C. for 5 min then allowed to cool to room temperature prior to addition of ethylene glycol. Samples (400 μl) were added to NMR tubes, shaken to position the liquid at the bottom of the tube, then frozen in liquid nitrogen. Phosphorescence spectra were recorded over the spectral region of 400-600 nm with excitation at 330 nm. Slits were set at 5.0 mm on both monochromators to maximise the signal intensity. Spectra were plotted after subtraction of the baseline recorded with a frozen glass lacking AQ and normalisation at 400 nm.
Emission spectra were recorded for three cases: (i) water soluble AQ in the absence of PNA and DNA, (ii) single-stranded PNA, and (iii) PNA-DNA hybrids. For PNA579, the phosphorescence is quenched by ca. 70% and >90% in the single-strand and hybrid forms, respectively, relative to the free AQ. Meanwhile, for PNA586, the phosphorescence is quenched by >90% in both the single-stranded and hybrid forms. (Data for PNA586 are plotted in FIG. 15.) These results indicate that there is efficient photoinduced electron transfer from the bases to the AQ in the hybrid. Moreover, electron transfer is also quite efficient in the single-stranded form, indicating the AQ moiety is capable of reacting with the PNA bases as well as the DNA bases.
This example describes the use of hybridization to inhibit electron transfer quenching, demonstrating the ability of PNA/DNA duplexes to act as “bioinsulators”.
In this example, sequence specific recognition of ssDNA by PNA hairpins 626 and 627 is demonstrated by thermal denaturation and fluorescence spectroscopy. A similar approach has been reported by Tyagi and Kramer (Tyagi, S.; Kramer, F. R. Nature Biotechnol. 1996, 14, 303-308). In that strategy, a DNA hairpin was labeled with an energy donor and acceptor. Hybridization led to decreased energy transfer between the donor and acceptor. The use of PNA in the present invention rather than DNA is preferred because of the superior hybridization characteristics of PNA, namely the much higher affinity and ionic strength independence of the PNA-DNA recognition. The choice of electron transfer rather than energy transfer in the present invention arises from the fact that fewer restrictions are placed on the donor and acceptor moieties for photoinduced electron transfer chemistry. In particular, either the donor or acceptor can be irradiated and there is no requirement that the donor absorb light of shorter wavelength than the acceptor.
PNA626 and PNA627 are shown below in their extended and folded conformations. PNA626 contains an acridine moiety (R2) which functions as a light absorber and electron donor plus an anthraquinone moiety (Q1) which functions as an electron acceptor. In the folded conformation, the acridine and quinone will be placed in close proximity to one another, particularly if both are stacked within the helix, leading to efficient photoinduced electron transfer, which is detected by quenching of the acridine fluorescence. PNA627 is analogous to PNA626 except it lacks the quinone acceptor. A thymine is included in place of the quinone, leading to a T—T mismatch in the folded conformation.
DNA Targets (single stranded, linear)
626A is perfectly complementary to the 12 base sequence separating the quinone from the acridine. 626B and 626C will hybridize with the PNAs but with single base mismatches. The mismatch for 626B will be located near the end of the resulting duplex, yielding a 10 base pair segment, whereas the mismatch for 626C will be located near the center of the resulting duplex, yielding 6 and 5 base pair segments. 626D will hybridize with the full length of the PNAs, placing thymines across from the acridine and quinone moieties.
Thermal Denaturation Experiments. Samples were prepared containing 2.5 μM PNA in 10 mM sodium phosphate buffer (pH=7.0) with and without 2.5 μM DNA626A. Melting curves were recorded by monitoring the absorbency at 260 nm over the temperature range 25-90° C.
FIG. 16 shows the curves for melting of the PNAs in the absence of DNA. The substantial hyperchromicity (>20%) is consistent with a folded hairpin structure rather than an extended linear conformation. The transition midpoint is approximately 55° C. for PNA627. Melting is more complex for PNA626 but the transition to the extended conformation occurs within the range 55-65° C.
FIG. 17 shows the curves for melting of the PNAs in the presence of DNA626A. The fact that the two curves in this figure are significantly different from those in FIG. 1 demonstrates that the PNA and DNA strands are interacting with one another. A clear transition is observed at ca. 76° C. for both PNAs. This transition is attributed to melting of the desired PNA/DNA hybrids.
Fluorescence Experiments. Samples were prepared containing 1.0 μM PNA in 10 mM sodium phosphate buffer (pH=7.0) with and without 1.0 μM DNA. Samples were heated to 90° C. for 5 min, then cooled to room temperature over a period of 90 minutes. Fluorescence emission spectra were recorded over the range 430-600 nm with excitation at 417 nm. 2.5 mm slits, 1.0 nm increment and 0.5 sec integration time were used for data collection. After collection, spectra were integrated to give the data shown below.
Effect of Hybridization on ET Quenching of Acridine Fluorescence
*Data for each PNA were normalized to the values for the samples lacking DNA (1 and 6).
Effect of Quinone in PNA Hairpin. Comparing samples 1 and 6, the fluorescence is 3.6 times lower when the quinone is present, consistent with quenching of the acridine fluorescence by electron transfer to the adjacent quinone. The lack of complete quenching is most likely due to the fact that the electron transfer reaction is not very favorable energetically and the lifetime of the acridine is probably fairly short (a few nanoseconds). An additional factor, indicated by the broad hairpin melting curves, is that the acridine and quinone likely have multiple conformations (e.g. intercalated or extrahelical). The incomplete quenching could arise from a population of hairpins in which only one of the two chromophores is actually intercalated at the time of excitation.
Effect of Hybridization. In the presence of complementary DNA, the hairpin is disrupted and a PNA/DNA hybrid is formed. In the hybrid, the acridine and anthraquinone are separated by 12 base pairs instead of being in contact in the hairpin. This results in a substantial increase in the fluorescence (compare samples 1 and 2). The increase is much greater for PNA626 than for 627, indicating that most of the enhanced fluorescence results from retarded electron transfer rather than from the change in the environment of the acridine fluorophore due to hybridization with the DNA strand. Note that the fluorescence is still greater for PNA627, suggesting that there is still some electron transfer quenching occurring over 12 base pairs within the hybrid.
Effect of Single Base Mismatches. A single mismatch near the end of the recognition site results in a 12% decrease in fluorescence enhancement while a mismatch near the center of the recognition site results in a 23% decrease in fluorescence enhancement. As expected, the central mismatch results in shorter (and, therefore, less stable) hybrid regions, leading to less fluorescence enhancement.
Effect of full-length Hybridization. The largest enhancements results from hybridization with a full length DNA complement (DNA626D, see Samples 5 and 10). This result emphasizes the sensitivity of the acridine fluorescence quantum yield to its environment: the emission is significantly greater when the acridine is part of a duplex as opposed to in a single strand. (The results for PNA627 also indicate that the acridine fluorescence quantum yield is greater for a PNA-DNA versus a PNA-PNA hybrid. The fluorescence increases by 2.67 in the presence of DNA.) The combination of this effect with the relieved electron transfer results in the large enhancement of fluorescence.
Assessment. The donor-acceptor PNA hairpins appear to work as designed and significant enhancements of fluorescence arise from hybridization with all four DNA oligomers. In each case, the fluorescence was enhanced 2.4-2.8 times more for PNA626 than for PNA627, demonstrating the importance of the electron acceptor in the system. The experiments were performed on samples containing 1 nanomole each of PNA and DNA target; this could easily be decreased by a factor of 10.
In this experiment, denomination is corresponding to example 11. N means the monomer containing an ANI residue. The N-monomer is shown in FIG. 18.
PNA 800: H-ATA-TQT-TGG-CTG-ATC-CAN-TAT-AT-LYS-Lys-NH2
PNA 801: H-ATA-TQX-TGG-CTG-ATC-CAN-XAT-AT-LYS-Lys-NH2
PNA 802: H-ATA-TTT-TGG-CTG-ATC-CAN-TAT-AT-LYS-Lys-NH2.
In PNA 800 T's are opposite Q and N. In PNA 801 a no-base X is opposite the Q and N supposed to give more room for the bulky chromophores. In 802 only ANI is placed.
DNAs: (single stranded, linear)
DNA A: 3′-ACC-GAC-TAG-GT-5′
DNA D: 3′-TAT-ATA-ACC-GAC-TAG-GTT-ATA-TA-5′.
Analogous Tm experiments as in the acridine/Q system was performed, and hairpin formation of individual PNA oligomers, and hybrid formation for each PNA oligomer to both DNA A and D was shown. Fluorescence experiments were then performed as described below and quantification was as follows:
1 μM PNA 800
3.6 × 106
1 μM PNA 800 + 1 μM DNA A
11.8 × 106
1 μM PNA 800 + 1 μM DNA D
20.3 × 106
1 μM PNA 801
2.4 × 106
1 μM PNA 801 + 1 μM DNA A
11.2 × 106
1 μM PNA 801 + 1 μM DNA D
12.9 × 106
1 μM PNA 802
13.6 × 106
1 μM PNA 802 + 1 μM DNA A
15.0 × 106
1 μM PNA 802 + 1 μM DNA D
14.7 × 106
PNA 802 has no Q monomer and there is therefore no difference in the fluorescence difference between the hairpin and the unfolded hairpin, PNA-DNA duplex. In both PNA 800 and 801 the quinone quenches the fluorescence when the hairpin is formed. This shown that excitation of ANI results in electron transfer to the quinone, thus no radiative event happens. When either the short or the long DNA is present the hairpin is unfolded and the quenching by the quinone is significantly reduced leading to fluorescence by the ANI chromophore. This experiment shows that electron transfer is not immediate in PNA/DNA and that the intervening bases pairs exerts this property.
To a suspension of 2.1 g (10 mmol) of 4-amino-1,8-napthalimide and potassium carbonate (4.14 g, 30 mmol) in DMF (50 ml) was stirred at 40° C. for 0.5 h . The suspension was cooled to room temperature and (12 mmol, 1.8 ml) benzyl bromoacetate was added in one portion. The suspension was stirred over night. To the reaction was added 10 g of celite and filtered through 1 cm of celite on a filter. The solvent was removed under reduced pressure and the residue was crystallised from ethylacetate.
Yield 3.1 g (89%); 1H NMR (DMSO-d6): δ 4.84 (s, 2 H, CH2CO); 5.19 (s, 2 H, CH2O); 6.87 (d, 1 H, H-3); 7.37 (m, 5 H, benzyl); 7 56 (s, 2 H, NH2); 7.68 (t, 1 H, H-6); 8.21 (d, 1 H, H-5); 8.46 (dd, 1 H, H-2); 8.67 (dd, 1 H, H-7).
To a solution of benzyl N-(4-amino-1,8-napthalimido)glycinate (2.5 g, 7.2 mmol) in THF (20 ml) was added lithium hydroxide (30 ml, 1 M) and stirred for 3 h. The THF was removed under reduced pressure and pH was adjusted to 2.8 and the precipitated free acid was collected by filtration and washed with water and dried.
Yield 1.6 g (86%). 1H NMR (DMSO-d6): δ 4.66 (s, 2 H, CH2CO); 6.86 (d, 1 H, H-3); 7 52 (s, 2 H, NH2); 7.67 (t, 1 H, H-6); 8.20 (d, 1 H, H-5); 8.45 (dd, 1 H, H-2); 8.66 (dd, 1 H, H-7).
N-(4-amino-1,8-napthalimido)glycine (1.5 g, 5.8 mmol), DCC (1.6 g, 6.2 mmol) and DhbtOH (1.0 g, 6.2 mmol) was mixed and stirred for 10 min in DMF (35 ml). Then methyl N-(-2-Boc-aminoethyl)glycinate (1.5 g, 6.5 mmol) was added and the reaction was stirred for additional 4 h. To the reaction mixture dichloromethane (70 ml) was added after which the mixture was cooled to 0° C. and filtered. The filtrate was extracted with sodium bicarbonate (3×35 ml, 0.5 M), sodium bisulphate (2×35 ml 2 M) and brine. The organic phase was dried with magnesium sulphate and evaporated to dryness. The residue was dissolved in tetrahydrofurane (20 ml) and lithium hydroxide (40 ml of 1 M) was added. The reaction mixture was stirred for 3 h. Tetrahydrofurane was removed under reduced pressure and the solution was filtrated and pH of the filtrate was adjusted to 2.8 which precipitated the title compound. The precipitate was recrystallised from ethylacetate.
Yield 2.1 g (72%); 1H NMR (DMSO-d6): δ 1.37 (d, 9H, Boc); 3.04 (m, 2 H, CH2); 3.27 (m, 2H, CH2); 4.02 (d, 2 H, CH2CO); 4.80 (d, 2H, CH2CO); 6.86 (m, 1 H, H-3); 6.95 (m, 1H, NH); 7 52 (s, 2 H, NH2); 7.63 (m, 1 H, H-6); 8.14 (m, 1 H, H-5); 8.36 (m, 1 H, H-2); 8.64 (m, 1 H, H-7).
b) PNA Synthesis
The three PNA oligomers were synthesised accordingly to the standard schemes. The oligomers were HPLC purified and the identity was checked by mass spectroscopy (MALDI-TOF).
c) Fluorescence Measurements
DNA oligomers were purchased from Midland Certified Reagent Company, purified by gel filtration and characterised by MALDI-TOF mass spectrometry. The extinction coefficients of DNA oligomers were calculated using the nearest-neighbor values: for DNA A, ε260=108,200 M−1cm−1; DNA D, ε260=238,200 M−1cm−1. Similarly, PNA oligomers extinction coefficients were determined using DNA values, with adenine substituted for 4-amino-1,8-naphthalimide or anthraquinone: PNA 800, ε260=232,900 M−1cm−1; PNA 801, ε260=214,700 M−1cm−1; PNA 802, ε260=228,900 M−1cm−1. The samples for all experiments were prepared in a 10 mM sodium phosphate buffer at pH=7.0. Fluorescence spectra were recorded with Cary 1E (UV-Vis) and SPEX 1681 FLUOROLOG. For PNA hairpins, samples were prepared consisting of 1.0 μM of PNA oligomer in 1.0 mL of 10 mM sodium phosphate (pH 7.0). In the case of PNA/DNA hybrids, equimolar concentration of PNA and DNA oligomers (1.0μ each) were used. Samples were placed in cuvettes (1.5 mL capacity, 1.0 cm pathlength) and sealed with Teflon tape to prevent evaporation of water during heating/cooling cycles. The same samples were heated to 90° C. for 5 min and then slowly cooled to RT to assure complete hybridisation. Samples were placed in a fluorescence cuvette (from Hellma: 1.5 mL capacity, 1.0 cm pathlength) and excited at 450 nm. The fluorescence emission was monitored from 460 to 700 nm at the rate of 1 nm/sec with the following slit sizes: ex=2.5 mm and em=0.5 mm.
|US7713519||29 Oct 2008||11 May 2010||Hallstar Innovations Corp.||Method of quenching electronic excitation of chromophore-containing organic molecules in photoactive compositions|
|US7754191||3 Nov 2008||13 Jul 2010||Hallstar Innovations Corp.||Method of quenching electronic excitation of chromophore-containing organic molecules photoactive compositions|
|US7776614 *||9 Aug 2007||17 Aug 2010||Hallstar Innovations Corp.||Test method for determining compounds capable of quenching electronic singlet state excitation of photoactive compounds|
|US7897345||13 Feb 2009||1 Mar 2011||Helicos Biosciences Corporation||Short cycle methods for sequencing polynucleotides|
|US7939670||23 Feb 2009||10 May 2011||Purdue Pharma L.P.||Nociceptin analogs|
|US7959834||14 Jun 2011||Hallstar Innovations Corp.||Alkoxycrylene/metal oxide photostabilized photoactive compositions and methods|
|US7964355 *||21 Jun 2011||Investigen, Inc.||Assays based on detection of photobleaching reaction products from dye catalytic complex|
|US7981604||19 Jul 2011||California Institute Of Technology||Methods and kits for analyzing polynucleotide sequences|
|US8070989||31 Jul 2009||6 Dec 2011||Hallstar Innovations Corp.||Photostabilization of retinoids with alkoxycrylene compounds|
|US8075808||13 Dec 2011||Hallstar Innovations Corp.||Method of quenching electronic excitation of chromophore-containing organic molecules in photoactive compositions|
|US8133477||4 Sep 2009||13 Mar 2012||Hallstar Innovations Corp.||Dispersions of inorganic particulates containing alkoxycrylene|
|US8257687||4 Sep 2012||Hallstar Innovations Corp.||Photostabilization of coenzyme Q compounds with alkoxycrylene compounds|
|US8263050||11 Sep 2012||Hallstar Innovations Corp.||Method of quenching electronic excitation of chromophore-containing organic molecules in photoactive compositions|
|US8263051||11 Sep 2012||Hallstar Innovations Corp.||Photostabilization of resveratrol with alkoxycrylene compounds|
|US8268294||22 Feb 2011||18 Sep 2012||Hallstar Innovations Corp.||Photostabilization of retinoids with alkoxycrylene compounds|
|US8278332||2 Oct 2012||Hallstar Innovations Corp.||Photostabilization of retinoids with alkoxycrylene compounds|
|US8329148||10 Aug 2012||11 Dec 2012||Hallstar Innovations Corp.||Photostabilization of coenzyme Q compounds with alkoxycrylene compounds|
|US8431112||5 Jan 2012||30 Apr 2013||Hallstar Innocations Corp.||Photostabilization of cholecalciferol with alkoxycrylene compounds|
|US8741936||20 Jun 2012||3 Jun 2014||Intermune, Inc.||Method of modulating stress-activated protein kinase system|
|US8916345||28 Mar 2011||23 Dec 2014||Integrated Dna Technologies, Inc.||Methods for enhancing nucleic acid hybridization|
|US8969347||15 Oct 2012||3 Mar 2015||Intermune, Inc.||Compounds and methods for treating inflammatory and fibrotic disorders|
|US9012144||18 Jan 2011||21 Apr 2015||Fluidigm Corporation||Short cycle methods for sequencing polynucleotides|
|US9096898||16 Dec 2010||4 Aug 2015||Life Technologies Corporation||Method of determining the nucleotide sequence of oligonucleotides and DNA molecules|
|US9212393||3 May 2011||15 Dec 2015||Life Technologies Corporation||Method of determining the nucleotide sequence of oligonucleotides and DNA molecules|
|US9290450||28 Apr 2014||22 Mar 2016||Intermune, Inc.||Compounds and methods for treating inflammatory and fibrotic disorders|
|US9359379||1 Oct 2013||7 Jun 2016||Intermune, Inc.||Anti-fibrotic pyridinones|
|US20020025529 *||18 Jul 2001||28 Feb 2002||Stephen Quake||Methods and apparatus for analyzing polynucleotide sequences|
|US20030069249 *||18 Apr 2002||10 Apr 2003||Qun Sun||Nociceptin analogs|
|US20030099999 *||31 Oct 2002||29 May 2003||Ai-Ping Wei||Assay procedure using fluorogenic tracers|
|US20030207306 *||7 Apr 2003||6 Nov 2003||California Institute Of Technology||Detectably and removably tagged nucleic acids|
|US20040101839 *||19 Sep 2001||27 May 2004||Hisafumi Ikeda||Novel functional peptide nucleic acid monomer and process for producing the same|
|US20050159452 *||11 Jan 2005||21 Jul 2005||Qun Sun||Nociceptin analogs|
|US20070218490 *||15 Mar 2007||20 Sep 2007||Integrated Dna Technologies, Inc.||Nucleic acid monomers with 2'-chemical moieties|
|US20080138817 *||7 Sep 2007||12 Jun 2008||Credia Japan Co., Ltd.||Novel Functional Peptide Nucleic Acid Monomer and Process for Producing the Same|
|US20090042312 *||9 Aug 2007||12 Feb 2009||Hallstar Innovations Corp.||Test method for determining compounds capable of quenching electronic singlet state excitation of photoactive compounds|
|US20090053821 *||16 Oct 2008||26 Feb 2009||Integrated Dna Technologies, Inc.||Fluorescence quenching azo dyes, their methods of preparation and use|
|US20090057627 *||3 Nov 2008||5 Mar 2009||Hallstar Innovations Corp.||Method of quenching electronic excitation of chromophore-containing organic molecules photoactive compositions|
|US20090074687 *||29 Oct 2008||19 Mar 2009||Hallstar Innovations Corp.|
|US20090208435 *||29 Apr 2009||20 Aug 2009||Hallstar Innovations Corp.|
|US20090246157 *||29 Apr 2009||1 Oct 2009||Hallstar Innovations Corp.|
|US20090291055 *||26 Nov 2009||Hallstar Innovations Corp.|
|US20090291141 *||26 Nov 2009||Hallstar Innovations Corp.|
|US20090324570 *||31 Dec 2009||Hallstar Innovations Corp.||Photostabilization of Retinoids with Alkoxycrylene Compounds|
|US20100021403 *||28 Jan 2010||Hallstar Innovations Corp.||Dispersions of inorganic particulates containing alkoxycrylene|
|US20100209909 *||17 Feb 2009||19 Aug 2010||Miaomiao Wang||Assays based on detection of photobleaching reaction products from dye catalytic complex|
|US20100283015 *||11 Nov 2010||Hallstar Innovations Corp.||Alkoxycrylene/Metal Oxide Photostabilized Photoactive Compositions and Methods|
|US20110142771 *||16 Jun 2011||Hallstar Innovations Corp.||Photostabilization of Retinoids with Alkoxycrylene Compounds|
|US20110152114 *||23 Jun 2011||Helicos Biosciences Corporation||Short cycle methods for sequencing polynucleotides|
|WO2003042398A2 *||8 Nov 2002||22 May 2003||Aclara Biosciences, Inc.||Multiplexed analysis by chromatographic separation of molecular tags|
|WO2003042398A3 *||8 Nov 2002||3 Jul 2003||Aclara Biosciences Inc||Multiplexed analysis by chromatographic separation of molecular tags|
|WO2003052134A2 *||18 Dec 2002||26 Jun 2003||Human Genetic Signatures Pty Ltd||Oligonucleotides comprising intercalator pseudonucleotide(s) for detection of nucleic acids and mutants hereof|
|WO2003052134A3 *||18 Dec 2002||25 Mar 2004||Ulf Bech Christensen||Oligonucleotides comprising intercalator pseudonucleotide(s) for detection of nucleic acids and mutants hereof|
|U.S. Classification||435/6.11, 435/810, 536/24.33, 536/25.3, 536/24.3, 536/23.1, 536/24.32, 536/24.31, 536/24.1, 436/501, 435/287.2|
|International Classification||C07H21/00, C12Q1/68, C07K14/00|
|Cooperative Classification||Y10S435/81, C07H21/00, C07K14/003, C12Q1/6818|
|European Classification||C07H21/00, C12Q1/68B2B, C07K14/00B1|
|16 Jun 1998||AS||Assignment|
Owner name: GEORGIA TECH RESEARCH CORPORATION, GEORGIA
Free format text: ASSIGNMENT OF ASSIGNORS INTEREST;ASSIGNORS:SCHUSTER, GARY B.;ARMITAGE, BRUCE A.;LY, DANITH;REEL/FRAME:009264/0406;SIGNING DATES FROM 19980526 TO 19980529
Owner name: BOEHRINGER MANNHEIM GMBH, GERMANY
Free format text: ASSIGNMENT OF ASSIGNORS INTEREST;ASSIGNORS:BATZ, HANS-GEORG;HANSEN, HENRIK FRYDENLUND;ORUM, HENRIK;AND OTHERS;REEL/FRAME:009264/0420;SIGNING DATES FROM 19980330 TO 19980406
|27 May 1999||AS||Assignment|
Owner name: ROCHE DIAGNOSTICS GMBH, GERMANY
Free format text: CHANGE OF NAME;ASSIGNOR:BOEHRINGER MANNHEIM GMBH;REEL/FRAME:009971/0915
Effective date: 19990104
Owner name: ROCHE DIAGNOSTICS GMBH,GERMANY
Free format text: CHANGE OF NAME;ASSIGNOR:BOEHRINGER MANNHEIM GMBH;REEL/FRAME:009971/0915
Effective date: 19990104
|1 Nov 2004||FPAY||Fee payment|
Year of fee payment: 4
|22 Jun 2005||AS||Assignment|
Owner name: PNA DIAGNOSTICS APS, SWEDEN
Free format text: ASSIGNMENT OF ASSIGNORS INTEREST;ASSIGNOR:ROCHE DIAGNOSTICS GMBH;REEL/FRAME:016386/0197
Effective date: 20050520
|10 Nov 2008||REMI||Maintenance fee reminder mailed|
|1 May 2009||LAPS||Lapse for failure to pay maintenance fees|
|23 Jun 2009||FP||Expired due to failure to pay maintenance fee|
Effective date: 20090501