# Patent

## Title
Invasion assays

## Abstract

The present invention relates to means for the detection and characterization of nucleic acid sequences, as well as variations in nucleic acid sequences. The present invention also relates to methods for forming a nucleic acid cleavage structure on a target sequence and cleaving the nucleic acid cleavage structure in a site-specific manner. The structure-specific nuclease activity of a variety of enzymes is used to cleave the target-dependent cleavage structure, thereby indicating the presence of specific nucleic acid sequences or specific variations thereof. The present invention further relates to methods and devices for the separation of nucleic acid molecules based on charge. The present invention also provides methods for the detection of non-target cleavage products via the formation of a complete and activated protein binding region. The invention further provides sensitive and specific methods for the detection of human cytomegalovirus nucleic acid in a sample.

## Background
This is a Continuation of U.S. patent application Ser. No. 09/350,597, filed Jul. 9, 1999, now U.S. Pat. No 6,458,535, which is a Continuation of U.S. application Ser. No. 08/823,516, filed Mar. 24, 1997, now U.S. Pat. No. 5,994,069, which is a Continuation-In-Part of U.S. application Ser. No. 08/759,038, filed Dec. 2, 1996, now U.S. Pat. No. 6,090,543, which is a Continuation-In-Part of U.S. application Ser. No. 08/756,386, filed Nov. 26, 1996, now U.S. Pat. No. 5,985,557, which is a Continuation-In-Part of U.S. application Ser. No. 08/682,853, filed Jul. 12, 1996, now U.S. Pat. No. 6,001,567, which is a Continuation-In-Part of U.S. application Ser. No. 08/599,491, filed Jan. 24, 1996, now U.S. Pat. No. 5,846,717.


FIELD OF THE INVENTION
The present invention relates to means for the detection and characterization of nucleic acid sequences and variations in nucleic acid sequences. The present invention relates to methods for forming a nucleic acid cleavage structure on a target sequence and cleaving the nucleic acid cleavage structure in a site-specific manner. The 5′ nuclease activity of a variety of enzymes is used to cleave the target-dependent cleavage structure, thereby indicating the presence of specific nucleic acid sequences or specific variations thereof. The present invention further provides novel methods and devices for the separation of nucleic acid molecules based by charge. The present invention further provides methods for the detection of non-target cleavage products via the formation of a complete and activated protein binding region.
BACKGROUND OF THE INVENTION
The detection and characterization of specific nucleic acid sequences and sequence variations has been utilized to detect the presence of viral or bacterial nucleic acid sequences indicative of an infection, the presence of variants or alleles of mammalian genes associated with disease and cancers and the identification of the source of nucleic acids found in forensic samples, as well as in paternity determinations.
Various methods are known to the art which may be used to detect and characterize specific nucleic acid sequences and sequence variants. Nonetheless, as nucleic acid sequence data of the human genome, as well as the genomes of pathogenic organisms accumulates, the demand for fast, reliable, cost-effective and user-friendly tests for the detection of specific nucleic acid sequences continues to grow. Importantly, these tests must be able to create a detectable signal from samples which contain very few copies of the sequence of interest. The following discussion examines two levels of nucleic acid detection assays currently in use: I. Signal Amplification Technology for detection of rare sequences; and II. Direct Detection Technology for quantitative detection of sequences.
I. Signal Amplification Technology Methods For Amplification
The “Polymerase Chain Reaction” (PCR) comprises the first generation of methods for nucleic acid amplification. However, several other methods have been developed that employ the same basis of specificity, but create signal by different amplification mechanisms. These methods include the “Ligase Chain Reaction” (LCR), “Self-Sustained Synthetic Reaction” (3SR/NASBA), and “Qβ-Replicase” (Qβ).
Polymerase Chain Reaction (PCR)
The polymerase chain reaction (PCR), as described in U.S. Pat. Nos. 4,683,195 and 4,683,202 to Mullis and Mullis et al. (the disclosures of which are hereby incorporated by reference), describe a method for increasing the concentration of a segment of target sequence in a mixture of genomic DNA without cloning or purification. This technology provides one approach to the problems of low target sequence concentration. PCR can be used to directly increase the concentration of the target to an easily detectable level. This process for amplifying the target sequence involves introducing a molar excess of two oligonucleotide primers which are complementary to their respective strands of the double-stranded target sequence to the DNA mixture containing the desired target sequence. The mixture is denatured and then allowed to hybridize. Following hybridization, the primers are extended with polymerase so as to form complementary strands. The steps of denaturation, hybridization, and polymerase extension can be repeated as often as needed, in order to obtain relatively high concentrations of a segment of the desired target sequence.
The length of the segment of the desired target sequence is determined by the relative positions of the primers with respect to each other, and, therefore, this length is a controllable parameter. Because the desired segments of the target sequence become the dominant sequences (in terms of concentration) in the mixture, they are said to be “PCR-amplified.”
Ligase Chain Reaction (LCR or LAR)
The ligase chain reaction (LCR; sometimes referred to as “Ligase Amplification Reaction” (LAR) described by Barany, Proc. Natl. Acad. Sci., 88:189 (1991); Barany, PCR Methods and Applic., 1:5 (1991); and Wu and Wallace, Genomics 4:560 (1989) has developed into a well-recognized alternative method for amplifying nucleic acids. In LCR, four oligonucleotides, two adjacent oligonucleotides which uniquely hybridize to one strand of target DNA, and a complementary set of adjacent oligonucleotides, which hybridize to the opposite strand are mixed and DNA ligase is added to the mixture. Provided that there is complete complementarity at the junction, ligase will covalently link each set of hybridized molecules. Importantly, in LCR, two probes are ligated together only when they base-pair with sequences in the target sample, without gaps or mismatches. Repeated cycles of denaturation, hybridization and ligation amplify a short segment of DNA. LCR has also been used in combination with PCR to achieve enhanced detection of single-base changes. Segev, PCT Public. No. WO9001069 A1 (1990). However, because the four oligonucleotides used in this assay can pair to form two short ligatable fragments, there is the potential for the generation of target-independent background signal. The use of LCR for mutant screening is limited to the examination of specific nucleic acid positions.
Self-Sustained Synthetic Reaction (3SR/NASBA)
The self-sustained sequence replication reaction (3SR) (Guatelli et al., Proc. Natl. Acad. Sci., 87:1874-1878 [1990], with an erratum at Proc. Natl. Acad. Sci., 87:7797 [1990]) is a transcription-based in vitro amplification system (Kwok et al., Proc. Natl. Acad. Sci., 86:1173-1177 [1989]) that can exponentially amplify RNA sequences at a uniform temperature. The amplified RNA can then be utilized for mutation detection (Fahy et al., PCR Meth. Appl., 1:25-33 [1991]). In this method, an oligonucleotide primer is used to add a phage RNA polymerase promoter to the 5′ end of the sequence of interest. In a cocktail of enzymes and substrates that includes a second primer, reverse transcriptase, RNase H, RNA polymerase and ribo-and deoxyribonucleoside triphosphates, the target sequence undergoes repeated rounds of transcription, cDNA synthesis and second-strand synthesis to amplify the area of interest. The use of 3SR to detect mutations is kinetically limited to screening small segments of DNA (e.g., 200-300 base pairs).
Q-Beta (Qβ) Replicase
In this method, a probe which recognizes the sequence of interest is attached to the replicatable RNA template for Qβ replicase. A previously identified major problem with false positives resulting from the replication of unhybridized probes has been addressed through use of a sequence-specific ligation step. However, available thermostable DNA ligases are not effective on this RNA substrate, so the ligation must be performed by T4 DNA ligase at low temperatures (37° C.). This prevents the use of high temperature as a means of achieving specificity as in the LCR, the ligation event can be used to detect a mutation at the junction site, but not elsewhere.
Table 1 below, lists some of the features desirable for systems useful in sensitive nucleic acid diagnostics, and summarizes the abilities of each of the major amplification methods (See also, Landgren, Trends in Genetics 9:199 [1993]).
A successful diagnostic method must be very specific. A straight-forward method of controlling the specificity of nucleic acid hybridization is by controlling the temperature of the reaction. While the 3SR/NASBA, and Qβ systems are all able to generate a large quantity of signal, one or more of the enzymes involved in each cannot be used at high temperature (i.e., >55° C.). Therefore the reaction temperatures cannot be raised to prevent non-specific hybridization of the probes. If probes are shortened in order to make them melt more easily at low temperatures, the likelihood of having more than one perfect match in a complex genome increases. For these reasons, PCR and LCR currently dominate the research field in detection technologies.









TABLE 1









Method















PCR &
3SR



Feature
PCR
LCR
LCR
NASBA
Qβ





Amplifies Target
+
+
+
+



Recognition of Independent
+
+
+
+
+


Sequences Required


Performed at High Temp.
+
+


Operates at Fixed Temp.



+
+


Exponential Amplification
+
+
+
+
+


Generic Signal Generation




+


Easily Automatable









The basis of the amplification procedure in the PCR and LCR is the fact that the products of one cycle become usable templates in all subsequent cycles, consequently doubling the population with each cycle. The final yield of any such doubling system can be expressed as: (1+X)n=y, where “X” is the mean efficiency (percent copied in each cycle), “n” is the number of cycles, and “y” is the overall efficiency, or yield of the reaction (Mullis, PCR Methods Applic., 1:1 [1991]). If every copy of a target DNA is utilized as a template in every cycle of a polymerase chain reaction, then the mean efficiency is 100%. If 20 cycles of PCR are performed, then the yield will be 220, or 1,048,576 copies of the starting material. If the reaction conditions reduce the mean efficiency to 85%, then the yield in those 20 cycles will be only 1.8520, or 220,513 copies of the starting material. In other words, a PCR running at 85% efficiency will yield only 21% as much final product, compared to a reaction running at 100% efficiency. A reaction that is reduced to 50% mean efficiency will yield less than 1% of the possible product.
In practice, routine polymerase chain reactions rarely achieve the theoretical maximum yield, and PCRs are usually run for more than 20 cycles to compensate for the lower yield. At 50% mean efficiency, it would take 34 cycles to achieve the million-fold amplification theoretically possible in 20, and at lower efficiencies, the number of cycles required becomes prohibitive. In addition, any background products that amplify with a better mean efficiency than the intended target will become the dominant products.
Also, many variables can influence the mean efficiency of PCR, including target DNA length and secondary structure, primer length and design, primer and dNTP concentrations, and buffer composition, to name but a few. Contamination of the reaction with exogenous DNA (e.g., DNA spilled onto lab surfaces) or cross-contamination is also a major consideration. Reaction conditions must be carefully optimized for each different primer pair and target sequence, and the process can take days, even for an experienced investigator. The laboriousness of this process, including numerous technical considerations and other factors, presents a significant drawback to using PCR in the clinical setting. Indeed, PCR has yet to penetrate the clinical market in a significant way. The same concerns arise with LCR, as LCR must also be optimized to use different oligonucleotide sequences for each target sequence. In addition, both methods require expensive equipment, capable of precise temperature cycling.
Many applications of nucleic acid detection technologies, such as in studies of allelic variation, involve not only detection of a specific sequence in a complex background, but also the discrimination between sequences with few, or single, nucleotide differences. One method for the detection of allele-specific variants by PCR is based upon the fact that it is difficult for Taq polymerase to synthesize a DNA strand when there is a mismatch between the template strand and the 3′ end of the primer. An allele-specific variant may be detected by the use of a primer that is perfectly matched with only one of the possible alleles; the mismatch to the other allele acts to prevent the extension of the primer, thereby preventing the amplification of that sequence. This method has a substantial limitation in that the base composition of the mismatch influences the ability to prevent extension across the mismatch, and certain mismatches do not prevent extension or have only a minimal effect (Kwok et al., Nucl. Acids Res., 18:999 [1990]).)
A similar 3′-mismatch strategy is used with greater effect to prevent ligation in the LCR (Barany, PCR Meth. Applic., 1:5 [1991]). Any mismatch effectively blocks the action of the thermostable ligase, but LCR still has the drawback of target-independent background ligation products initiating the amplification. Moreover, the combination of PCR with subsequent LCR to identify the nucleotides at individual positions is also a clearly cumbersome proposition for the clinical laboratory.
II. Direct Detection Technology
When a sufficient amount of a nucleic acid to be detected is available, there are advantages to detecting that sequence directly, instead of making more copies of that target, (e.g., as in PCR and LCR). Most notably, a method that does not amplify the signal exponentially is more amenable to quantitative analysis. Even if the signal is enhanced by attaching multiple dyes to a single oligonucleotide, the correlation between the final signal intensity and amount of target is direct. Such a system has an additional advantage that the products of the reaction will not themselves promote further reaction, so contamination of lab surfaces by the products is not as much of a concern. Traditional methods of direct detection including Northern and Southern blotting and RNase protection assays usually require the use of radioactivity and are not amenable to automation. Recently devised techniques have sought to eliminate the use of radioactivity and/or improve the sensitivity in automatable formats. Two examples are the “Cycling Probe Reaction” (CPR), and “Branched DNA” (bDNA)
The cycling probe reaction (CPR) (Duck et al., BioTech., 9:142 [1990]), uses a long chimeric oligonucleotide in which a central portion is made of RNA while the two termini are made of DNA. Hybridization of the probe to a target DNA and exposure to a thermostable RNase H causes the RNA portion to be digested. This destabilizes the remaining DNA portions of the duplex, releasing the remainder of the probe from the target DNA and allowing another probe molecule to repeat the process. The signal, in the form of cleaved probe molecules, accumulates at a linear rate. While the repeating process increases the signal, the RNA portion of the oligonucleotide is vulnerable to RNases that may be carried through sample preparation.
Branched DNA (bDNA), described by Urdea et al., Gene 61:253-264 (1987), involves oligonucleotides with branched structures that allow each individual oligonucleotide to carry 35 to 40 labels (e.g., alkaline phosphatase enzymes). While this enhances the signal from a hybridization event, signal from non-specific binding is similarly increased.
While both of these methods have the advantages of direct detection discussed above, neither the CPR or bDNA methods can make use of the specificity allowed by the requirement of independent recognition by two or more probe (oligonucleotide) sequences, as is common in the signal amplification methods described in section I. above. The requirement that two oligonucleotides must hybridize to a target nucleic acid in order for a detectable signal to be generated confers an extra measure of stringency on any detection assay. Requiring two oligonucleotides to bind to a target nucleic acid reduces the chance that false “positive” results will be produced due to the non-specific binding of a probe to the target. The further requirement that the two oligonucleotides must bind in a specific orientation relative to the target, as is required in PCR, where oligonucleotides must be oppositely but appropriately oriented such that the DNA polymerase can bridge the gap between the two oligonucleotides in both directions, further enhances specificity of the detection reaction. However, it is well known to those in the art that even though PCR utilizes two oligonucleotide probes (termed primers) “non-specific” amplification (i.e., amplification of sequences not directed by the two primers used) is a common artifact. This is in part because the DNA polymerase used in PCR can accommodate very large distances, measured in nucleotides, between the oligonucleotides and thus there is a large window in which non-specific binding of an oligonucleotide can lead to exponential amplification of inappropriate product. The LCR, in contrast, cannot proceed unless the oligonucleotides used are bound to the target adjacent to each other and so the full benefit of the dual oligonucleotide hybridization is realized.
An ideal direct detection method would combine the advantages of the direct detection assays (e.g., easy quantification and minimal risk of carry-over contamination) with the specificity provided by a dual oligonucleotide hybridization assay.
SUMMARY OF THE INVENTION
The present invention relates to means for cleaving a nucleic acid cleavage structure in a site-specific manner. In a preferred embodiment, the means for cleaving is a structure-specific nuclease. Particularly preferred structure-specific nucleases are thermostable structure-specific nucleases. In one embodiment, the structure-specific nuclease is an enzyme comprising 5′ nucleases derived from thermostable DNA polymerases. These polymerases form the basis of a novel method of detection of specific nucleic acid sequences. The present invention contemplates use of novel detection methods for various uses, including, but not limited to clinical diagnostic purposes.
In one embodiment, the present invention contemplates a DNA sequence encoding a DNA polymerase altered in sequence (i.e., a “mutant” DNA polymerase) relative to the native sequence, such that it exhibits altered DNA synthetic activity from that of the native (i.e., “wild type”) DNA polymerase. It is preferred that the encoded DNA polymerase is altered such that it exhibits reduced synthetic activity compared to that of the native DNA polymerase. In this manner, the enzymes of the invention are predominantly 5′ nucleases and are capable of cleaving nucleic acids in a structure-specific manner in the absence of interfering synthetic activity.
Importantly, the 5′ nucleases of the present invention are capable of cleaving linear duplex structures to create single discrete cleavage products. These linear structures are either 1) not cleaved by the wild type enzymes (to any significant degree), or 2) are cleaved by the wild type enzymes so as to create multiple products. This characteristic of the 5′ nucleases has been found to be a consistent property of enzymes derived in this manner from thermostable polymerases across eubacterial thermophilic species.
It is not intended that the invention be limited by the nature of the alteration necessary to render the polymerase synthesis-deficient. Nor is it intended that the invention be limited by the extent of the deficiency. The present invention contemplates various structures, including altered structures (primary, secondary, etc.), as well as native structures, that may be inhibited by synthesis inhibitors.
Where the polymerase structure is altered, it is not intended that the invention be limited by the means by which the structure is altered. In one embodiment, the alteration of the native DNA sequence comprises a change in a single nucleotide. In another embodiment, the alteration of the native DNA sequence comprises a deletion of one or more nucleotides. In yet another embodiment, the alteration of the native DNA sequence comprises an insertion of one or more nucleotides. It is contemplated that the change in DNA sequence may manifest itself as change in amino acid sequence.
The present invention contemplates structure-specific nucleases from a variety of sources, including mesophilic, psychrophilic, thermophilic, and hyperthermophilic organisms. The preferred structure-specific nucleases are thermostable. Thermostable structure-specific nucleases are contemplated as particularly useful in that they operate at temperatures where nucleic acid hybridization is extremely specific, allowing for allele-specific detection (including single-base mismatches). In one embodiment, the thermostable structure-specific are thermostable 5′ nucleases which are selected from the group consisting of altered polymerases derived from the native polymerases of Thermus species, including, but not limited to Thermus aquaticus, Thermus flavus, and Thermus thermophilus. However, the invention is not limited to the use of thermostable 5′ nucleases. Thermostable structure-specific nucleases from the FEN-1, RAD2 and XPG class of nucleases are also preferred.
Accordingly, the present invention provides improved enzymatic cleavage means. In one embodiment, the present invention provides a thermostable structure-specific nuclease having an amino acid sequence selected from the group consisting of SEQ ID NOS:61, 66, 69 and 72. In another embodiment, the nuclease is encoded by a DNA sequence selected from the group consisting of SEQ ID NO:60, 65, 68 and 70.
As noted above, the present invention contemplates the use of structure-specific nucleases in a detection method. In one embodiment, the present invention provides a method of detecting the presence of a target nucleic acid molecule comprising: a) providing: i) a cleavage means; ii) a source of a first target nucleic acid, the first target nucleic acid having a first region, a second region and a third region, wherein the first region is located adjacent to and downstream from the second region and wherein the second region is located adjacent to and downstream from the third region; iii) a first oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the first oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the first target nucleic acid and wherein the 3′ portion of the first oligonucleotide contains a sequence complementary to at least a portion of the third region of the first target nucleic acid; iv) a second oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the second oligonucleotide contains a sequence complementary to at least a portion of the first region of the first target nucleic acid and wherein the 3′ portion of the second oligonucleotide contains a sequence complementary to at least a portion of the second region of the first target nucleic acid; v) a source of a second target nucleic acid, the second target nucleic acid having a first region, a second region and a third region, wherein the first region is located adjacent to and downstream from the second region and wherein the second region is located adjacent to and downstream from the third region; vi) a third oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the third oligonucleotide contains a sequence complementary to at least a portion of the second region of the second target nucleic acid and wherein the 3′ portion of the third oligonucleotide contains a sequence complementary to at least a portion of the third region of the second target nucleic acid; b) generating a first cleavage structure wherein at least the 3′ portion of the first oligonucleotide is annealed to the first target nucleic acid and wherein at least the 5′ portion of the second oligonucleotide is annealed to the first target nucleic acid and wherein cleavage of the first cleavage structure occurs via the cleavage means thereby cleaving the first oligonucleotide to generate a fourth oligonucleotide, the fourth oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the fourth oligonucleotide contains a sequence complementary to at least a portion of the first region of the second target nucleic acid and wherein the 3′ portion of the fourth oligonucleotide contains a sequence complementary to at least a portion of the second region of the second target nucleic acid; c) generating a second cleavage structure under conditions wherein the at least the 3′ portion of the third oligonucleotide is annealed to the second target nucleic acid and wherein at least the 5′ portion of the fourth oligonucleotide is annealed to the second target nucleic acid oligonucleotide and wherein cleavage of the second cleavage structure occurs to generate a fifth oligonucleotide, the fifth oligonucleotide having a 3′-hydroxyl group; and d) detecting the fifth oligonucleotide.
It is contemplated that the first, second and third regions of the target nucleic acids be located adjacent to each other. However, the invention is not limited to the use of a target in which the three regions are contiguous with each other. Thus, the present invention contemplates the use of target nucleic acids wherein these three regions are contiguous with each other, as well as target acids wherein these three regions are not contiguous. It is further contemplated that gaps of approximately 2-10 nucleotides, representing regions of non-complementarity to the oligonucleotides (e.g., the first and/or second oligonucleotides), may be present between the three regions of the target nucleic acid.
The methods of the present invention are not limited by the size of the oligonucleotides employed. In a preferred embodiment, the first oligonucleotide has a length between eleven and fifteen nucleotides.
It is intended that the generation of the first and second cleavage structures and cleavage of these structures occurs under a variety of conditions. In a preferred format, the conditions of generating the cleavage structures comprises mixing together the target nucleic acids with the first, second and third oligonucleotides and the cleavage means in an aqueous solution in which a source of divalent cations is lacking. In this format, the cleavage reaction is initiated by the addition of a solution containing Mn2+ or Mg2+ ions. In another preferred format, the conditions of mixing comprises mixing together the target nucleic acid, and the first, second and third oligonucleotides in an aqueous solution containing Mn2+ or Mg2+ ions, and then adding the cleavage means to the reaction mixture.
It is contemplated that the oligonucleotides may be labelled. Thus, if the cleavage reaction employs a third oligonucleotide containing a label, detection of the cleavage product of the third oligonucleotide (i.e., the fifth oligonucleotide) may comprise detection of the label. The invention is not limited by the nature of the label chosen, including, but not limited to, labels which comprise a dye or a radionucleotide (e.g., 32P), fluorescein moiety, a biotin moiety, luminogenic, fluorogenic, phosphorescent, or fluors in combination with moieties that can suppress emission by fluorescence energy transfer (FET). Numerous methods are available for the detection of nucleic acids containing any of the above-listed labels. For example, biotin-labeled oligonucleotide(s) may be detected using non-isotopic detection methods which employ streptavidin-alkaline phosphatase conjugates. Fluorescein-labelled oligonucleotide(s) may be detected using a fluorescein-imager. Further the oligonucleotide and particularly the probe oligonucleotides may contain positively charged adducts (e.g., the Cy3 and Cy5 dyes, the dyes shown in FIG. 66, etc.) and/or positively charged amino acids and/or a phosphonate backbone to permit the detection of the fifth oligonucleotide [i.e., the non-target cleavage product generated by cleavage of the second (or terminal if more than two reactions are employed in the cascade) cleavage structure] by selective charge reversal as described herein (see section IV of the Description of the Invention). The oligonucleotides may be labelled with different labels (e.g., the first and the third oligonucleotides may each bear a different label).
It is also contemplated that labelled oligonucleotides (cleaved or uncleaved) may be separated by means other than electrophoresis. For example, biotin-labelled oligonucleotides may be separated from nucleic acid present in the reaction mixture using para-magnetic or magnetic beads, or particles which are coated with avidin (or streptavidin). In this manner, the biotinylated oligonucleotide/avidin-magnetic bead complex can be physically separated from the other components in the mixture by exposing the complexes to a magnetic field. Additionally, the signal from the cleaved oligonucleotides may be resolved from that of the uncleaved oligonucleotides without physical separation. For example, a change in size, and therefore rate of rotation in solution of fluorescent molecules can be detected by fluorescence polarization analysis.
In a preferred embodiment, the reaction conditions comprise a cleavage reaction temperature which is less than the melting temperature of the first oligonucleotide and greater than the melting temperature of the 3′ portion of the first oligonucleotide. In a particularly preferred embodiment, the reaction temperature is between approximately 40-75° C.; in another embodiment the reaction temperature is between approximately 40-60° C. It is contemplated that the reaction temperature at which the cleavage reaction occurs be selected with regard to the guidelines provided in the Description of the Invention.
The method of the present invention is not limited by the nature of the target nucleic acid. The target nucleic acid may comprise single-stranded or double-stranded DNA, RNA, and/or DNA/RNA hybrids. When a double-stranded target nucleic acid is employed, the reaction mixture may be treated such that the aid double-stranded DNA is rendered substantially single-stranded. A preferred method for rendering double-stranded DNA substantially single-stranded is by the use of increased temperature. When target nucleic acids comprising RNA are employed, the oligonucleotides may comprise DNA, RNA or an oligonucleotide comprising a mixture of RNA and DNA. It is not intended that the invention be limited by the nature of the oligonucleotides employed.
The oligonucleotides may comprise DNA, RNA or an oligonucleotide comprising a mixture of RNA and DNA. The invention also contemplates the use of a second oligonucleotide (i.e., the upstream oligonucleotide in the first cleavage structure) which comprises a functional group (e.g., a 5′ peptide region) which prevents the dissociation of the 5′ portion of the second oligonucleotide from the first region of the target nucleic acid. When such a functional group is present on the second oligonucleotide, the interaction between the 3′ portion of the second oligonucleotide and the first region of the target nucleic acid may be destabilized (i.e., designed to have a lower local melting temperature) through the use of A-T rich sequences, base analogs that form fewer hydrogen bonds (e.g., dG-DU pairs) or through the use of phosphorothioate backbones, in order to allow the 5′ region of the first oligonucleotide to compete successfully for hybridization.
The invention is not limited to use of oligonucleotides which are completely complementary to their cognate target sequences. In one embodiment, both the first and second oligonucleotides are completely complementary to the first target nucleic acid. In another embodiment, the first oligonucleotide is partially complementary to the first target nucleic acid. In yet another embodiment, the second oligonucleotide is partially complementary to the first target nucleic acid. In yet another embodiment, both the first and the second oligonucleotide are partially complementary to the first target nucleic acid. Likewise, the third and fourth oligonucleotides may be either completely or partially complementary to the second target nucleic acid.
The methods of the invention may employ a source of target nucleic acid which comprises a sample containing genomic DNA. In a preferred embodiment, the sample containing genomic DNA is selected from the group including, but not limited to blood, saliva, cerebral spinal fluid, pleural fluid, milk, lymph, sputum and semen.
In a preferred embodiment, the method employs reaction conditions which comprise providing a source of divalent cations. In a particularly preferred embodiment, the divalent cation is selected from the group comprising Mn2+ and Mg2+ ions.
The invention is not limited by the nature of the cleavage means. As discussed above, the invention contemplates that the cleavage means comprises a thermostable 5′ nuclease, although the invention is not limited to the use of a thermostable 5′ nuclease. When a thermostable 5′ nuclease is employed, a portion of the amino acid sequence of the nuclease may be homologous to a portion of the amino acid sequence of a thermostable DNA polymerase derived from a thermophilic organism. Particularly preferred cleavage means are structure-specific nucleases, with thermostable structure-specific nucleases being most preferred. In a preferred embodiment, the thermostable structure-specific nuclease is encoded by a DNA sequence selected from the group consisting of SEQ ID NOS:1-3, 9, 10, 12, 21, 25, 26, 60, 65, 68, 70, 74, and 78. In another preferred embodiment, the thermostable structure-specific nuclease is a nuclease from the FEN-1/RAD2/XPG class of nucleases. A preferred thermostable structure-specific nuclease is the Pyrococcus woesii FEN-1 endonuclease.
In another preferred embodiment, one or more of the first, second, and third oligonucleotides contain a dideoxynucleotide at the 3′ terminus. When dideoxynucleotide-containing oligonucleotides are employed, the detection of the fifth oligonucleotide preferably comprises: a) incubating the fifth oligonucleotide with a template-independent polymerase and at least one labelled nucleoside triphosphate under conditions such that at least one labelled nucleotide is added to the 3′-hydroxyl group of the fifth oligonucleotide to generate a labelled fifth oligonucleotide; and b) detecting the presence of the labelled fifth oligonucleotide. The invention is not limited by the nature of the template-independent polymerase employed; in one embodiment, the template-independent polymerase is selected from the group consisting of terminal deoxynucleotidyl transferase (TdT) and poly A polymerase. When TdT or polyA polymerase are employed in the detection step, the third oligonucleotide may contain a 5′ end label, the 5′ end label being a different label than the label present upon the labelled nucleoside triphosphate. The invention is not limited by the nature of the 5′ end label; a wide variety of suitable 5′ end labels are known to the art and include biotin, fluorescein, tetrachlorofluorescein, hexachlorofluorescein, Cy3, Cy5 and digoxigenin.
In another embodiment, detecting the fifth oligonucleotide comprises: a) incubating the fifth oligonucleotide with a template-independent polymerase and at least one nucleoside triphosphate under conditions such that at least one nucleotide is added to the 3′-hydroxyl group of the fifth oligonucleotide to generate a tailed fifth oligonucleotide; and b) detecting the presence of the tailed fifth oligonucleotide. The invention is not limited by the nature of the template-independent polymerase employed; in one embodiment, the template-independent polymerase is selected from the group consisting of terminal deoxynucleotidyl transferase (TdT) and poly A polymerase. When TdT or polyA polymerase are employed in the detection step, the second oligonucleotide may contain a 5′ end label. The invention is not limited by the nature of the 5′ end label; a wide variety of suitable 5′ end labels are known to the art and include biotin, fluorescein, tetrachlorofluorescein, hexachlorofluorescein, Cy3, Cy5 and digoxigenin.
The invention further provides a method of detecting the fifth oligonucleotide (i.e., the non-target cleavage product generated by cleavage of the second cleavage structure) comprising: a) providing: i) the fifth oligonucleotide; ii) a composition comprising two single-stranded nucleic acids annealed so as to define a single-stranded portion of a protein binding region; iii) a nucleic acid producing protein; b) exposing the fifth oligonucleotide to the single-stranded portion of the protein binding region under conditions such that the nucleic acid producing protein binds to the protein binding region and produces nucleic acid. In a preferred embodiment, the single-stranded portion of the protein binding region comprises: a) a first single continuous strand of nucleic acid comprising a sequence defining the template strand of an RNA polymerase binding region; and b) a second single continuous strand of nucleic acid having a 5′ and a 3′ end, the second nucleic acid comprising a region complementary to a portion of the first nucleic acid, wherein the second nucleic acid is annealed to the first nucleic acid so as to define the single-stranded portion of the protein binding region.
The invention is not limited by the nature of the protein binding region employed. In a preferred embodiment, the protein binding region is a template-dependent RNA polymerase binding region, more preferably a T7 RNA polymerase binding region.
The invention further provides a method of detecting the fifth oligonucleotide comprising: a) providing: i) the fifth oligonucleotide; ii) a single continuous strand of nucleic acid comprising a sequence defining a single strand of an RNA polymerase binding region; iii) a template-dependent DNA polymerase; iv) a template-dependent RNA polymerase; b) exposing the fifth oligonucleotide to the RNA polymerase binding region under conditions such that the fifth oligonucleotide binds to a portion of the single strand of the RNA polymerase binding region; c) exposing the bound fifth oligonucleotide to the template-dependent DNA polymerase under conditions such that a double-stranded RNA polymerase binding region is produced; and d) exposing the double-stranded RNA polymerase binding region to the template-dependent RNA polymerase under conditions such that RNA transcripts are produced. In a preferred embodiment, the method further comprises detecting the RNA transcripts.
The invention is not limited by the nature of the protein binding region employed. In a preferred embodiment, the protein binding region is a template-dependent RNA polymerase binding region, more preferably a T7 RNA polymerase binding region.
The present invention also provides a method of detecting the presence of a target nucleic acid molecule comprising: a) providing: i) a cleavage means, ii) a source of a first target nucleic acid, the first target nucleic acid having a first region, a second region, a third region and a fourth region, wherein the first region is located adjacent to and downstream from the second region, the second region is located adjacent to and downstream from the third region a n d the third region is located adjacent to and downstream from the fourth region; iii) a fir st oligonucleotide complementary to (at least a portion of) the fourth region of the first target nucleic acid; iv) a second oligonucleotide having a 5′ portion and a 3′ portion wherein the 5′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the first target nucleic acid and wherein the 3′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the third region of the first target nucleic acid; iv) a third oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the third oligonucleotide contains a sequence complementary to (at least a portion of) the first region of the first target nucleic acid and wherein the 3′ portion of the third oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the first target nucleic acid; v) a source of a second target nucleic acid, the second target nucleic acid having a first region, a second region and a third region, wherein the first region is located adjacent to and downstream from the second region and wherein the second region is located adjacent to and downstream from the third region; vi) a fourth oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the fourth oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the second target nucleic acid and wherein the 3′ portion of the fourth oligonucleotide contains a sequence complementary to (at least a portion of) the third region of the second target nucleic acid; b) generating a first cleavage structure wherein the first oligonucleotide is annealed to the fourth region of the first target nucleic acid and wherein at least the 3′ portion of the second oligonucleotide is annealed to the first target nucleic acid and wherein at least the 5′ portion of the third oligonucleotide is annealed to the first target nucleic acid and wherein cleavage of the first cleavage structure occurs thereby cleaving the second oligonucleotide to generate a fifth oligonucleotide, the fifth oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the fifth oligonucleotide contains a sequence complementary to (at least a portion of) the first region of the second target nucleic acid and wherein the 3′ portion of the fifth oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the second target nucleic acid; c) generating a second cleavage structure under conditions wherein the at least the 3′ portion of the fourth oligonucleotide is annealed to the second target nucleic acid and wherein at least the 5′ portion of the fifth oligonucleotide is annealed to the second target nucleic acid and wherein cleavage of the second cleavage structure occurs to generate a sixth oligonucleotide, the sixth oligonucleotide having a 3′-hydroxyl group; and d) detecting the sixth oligonucleotide.
The detection of the sixth oligonucleotide may be accomplished by a variety of methods, such as those described above for the method in which the fifth oligonucleotide is to be detected. As described above, the invention is not limited by the nature of the target nucleic acids, the nature of the cleavage means, the nature of the oligonucleotides, etc.
The invention also provides a method of detecting the presence of a target nucleic acid molecule comprising: a) providing: i) a cleavage means, ii) a source of a target nucleic acid, the target nucleic acid having a first region, a second region and a third region, wherein the first region is located adjacent to and downstream from the second region and wherein the second region is located adjacent to and downstream from the third region; iii) a first oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the first oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the target nucleic acid and wherein the 5′ portion of the first oligonucleotide contains a region of self-complementarity and wherein the 3′ portion of the first oligonucleotide contains a sequence complementary to (at least a portion of) the third region of the target nucleic acid; iv) a second oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the first region of the target nucleic acid and wherein the 3′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the target nucleic acid; v) a third oligonucleotide having a 5′ and a 3′ portion wherein the 3′ portion of the third oligonucleotide contains a sequence complementary to (at least a portion of) the 5′ portion of the first oligonucleotide; b) generating a first cleavage structure wherein at least the 3′ portion of the first oligonucleotide is annealed to the target nucleic acid and wherein at least the 5′ portion of the second oligonucleotide is annealed to the target nucleic acid and wherein cleavage of the first cleavage structure occurs thereby cleaving the first oligonucleotide to generate a fourth oligonucleotide, the fourth oligonucleotide having a first region, a second region and a third region, wherein the first region is located adjacent to and upstream of the second region and wherein the second region is located adjacent to and upstream of the third region and wherein the third region of the fourth oligonucleotide contains a region of self-complementarity; c) generating a second cleavage structure under conditions wherein the at least the 3′ portion of the third oligonucleotide is annealed to the first region of the fourth oligonucleotide and wherein at least the 5′ portion of the fourth oligonucleotide is annealed to the second region of the third oligonucleotide and wherein the third region of the fourth oligonucleotide forms a hairpin structure and wherein cleavage of the second cleavage structure occurs to generate a fifth oligonucleotide, the fifth oligonucleotide having a 3′-hydroxyl group; and d) detecting the fifth oligonucleotide.
The detection of the fifth oligonucleotide may be accomplished by a variety of methods, such as those described above. As described above, the invention is not limited by the nature of the target nucleic acids, the nature of the cleavage means, the nature of the oligonucleotides, etc.
The invention further provides a method of detecting the presence of human cytomegalovirus (HCMV) nucleic acid in a sample comprising: a) providing: i) a cleavage means, ii) a sample suspected of containing human cytomegalovirus target nucleic acid, the target nucleic acid having a first region, a second region and a third region, wherein the first region is located adjacent to and downstream from the second region and wherein the second region is located adjacent to and downstream from the third region; iii) a first oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the first oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the target nucleic acid and wherein the 3′ portion of the first oligonucleotide contains a sequence complementary to (at least a portion of) the third region of the target nucleic acid; iv) a second oligonucleotide having a 5′ and a 3′ portion wherein the 5′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the first region of the target nucleic acid and wherein the 3′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the target nucleic acid; b) generating a cleavage structure wherein at least the 3′ portion of the first oligonucleotide is annealed to the target nucleic acid and wherein at least the 5′ portion of the second oligonucleotide is annealed to the target nucleic acid and wherein cleavage of the cleavage structure occurs via the cleavage means to generate non-target cleavage products, each non-target cleavage product having a 3′ hydroxyl group; and c) detecting the non-target cleavage products and thereby detecting the presence of human cytomegalovirus nucleic acid in the sample. In one embodiment, the first oligonucleotide has a length between eleven and fifteen nucleotides.
The detection of the fifth oligonucleotide (indicative of the presence of HCMV nucleic acid in the sample) may be accomplished by a variety of methods, such as those described above. As described above, the invention is not limited by the nature of the target nucleic acids, the nature of the cleavage means, the nature of the oligonucleotides, etc.
The invention also provides a method of detecting the presence of human cytomegalovirus nucleic acid in a sample comprising: a) providing: i) a cleavage means, ii) a sample suspected of containing human cytomegalovirus target nucleic acid, the target nucleic acid having a first region, a second region, a third region and a fourth region, wherein the first region is located adjacent to and downstream from the second region, the second region is located adjacent to and downstream from the third region and the third region is located adjacent to and downstream from the fourth region; iii) a first oligonucleotide complementary to (at least a portion of) the fourth region of the target nucleic acid; iv) a second oligonucleotide having a 5′ portion and a 3′ portion wherein the 5′ portion of the second oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the target nucleic acid and wherein the 3′ portion of the first oligonucleotide contains a sequence complementary to (at least a portion of) the third region of the target nucleic acid; v) a third oligonucleotide having a 5′ portion and a 3′ portion wherein the 5′ portion of the third oligonucleotide contains a sequence complementary to (at least a portion of) the first region of the target nucleic acid and wherein the 3′ portion of the third oligonucleotide contains a sequence complementary to (at least a portion of) the second region of the target nucleic acid; b) generating a cleavage structure wherein the first oligonucleotide is annealed to the fourth region of the target nucleic acid and wherein at least the 3′ portion of the second oligonucleotide is annealed to the target nucleic acid and wherein at least the 5′ portion of the third oligonucleotide is annealed to the target nucleic acid and wherein cleavage of the cleavage structure occurs via the cleavage means to generate non-target cleavage products, each non-target cleavage product having a 3′ hydroxyl group; and c) detecting the non-target cleavage products and thereby detecting the presence of human cytomegalovirus nucleic acid in the sample. The detection of the non-target cleavage product (indicative of the presence of HCMV nucleic acid in the sample) may be accomplished by a variety of methods, such as those described above for the detection of the fifth oligonucleotide. As described above, the invention is not limited by the nature of the target nucleic acids, the nature of the cleavage means, the nature of the oligonucleotides, etc.



DESCRIPTION OF THE DRAWINGS
FIG. 1A to 1H is a comparison of the nucleotide structure of the DNAP genes isolated from Thermus aquaticus (SEQ ID NO:1), Thermus flavus (SEQ ID NO:2) and Thermus thermophilus (SEQ ID NO:3); the consensus sequence (SEQ ID NO:7) is shown at the top of each row.
FIG. 2A to 2C is a comparison of the amino acid sequence of the DNAP isolated from Thermus aquaticus (SEQ ID NO:4), Thermus flavus (SEQ ID NO:5), and Thermus thermophilus (SEQ ID NO:6); the consensus sequence (SEQ ID NO:8) is shown at the top of each row.
FIGS. 3A-G are a set of diagrams of wild-type and synthesis-deficient DNAPTaq genes.
FIG. 4A depicts the wild-type Thermus flavus polymerase gene.
FIG. 4B depicts a synthesis-deficient Thermus flavus polymerase gene.
FIG. 5 depicts a structure which cannot be amplified using DNAPTaq; this figure shows SEQ ID NO:17 (primer) and SEQ ID NO:15 (hairpin).
FIG. 6 is a ethidium bromide-stained gel demonstrating attempts to amplify a bifurcated duplex using either DNAPTaq or DNAPStf (i.e., the Stoffel fragment of DNAPTaq).
FIG. 7 is an autoradiogram of a gel analyzing the cleavage of a bifurcated duplex by DNAPTaq and lack of cleavage by DNAPStf.
FIGS. 8A-B are a set of autoradiograms of gels analyzing cleavage or lack of cleavage upon addition of different reaction components and change of incubation temperature during attempts to cleave a bifurcated duplex with DNAPTaq.
FIGS. 9A-B are an autoradiogram displaying timed cleavage reactions, with and without primer.
FIGS. 10A-B are a set of autoradiograms of gels demonstrating attempts to cleave a bifurcated duplex (with and without primer) with various DNAPs.
FIG. 11A shows the substrate and oligonucleotides [19-12 (SEQ ID NO:18) and 30-12 (SEQ ID NO:19)] used to test the specific cleavage of substrate DNAs targeted by pilot oligonucleotides.
FIG. 11B shows an autoradiogram of a gel showing the results of cleavage reactions using the substrates and oligonucleotides shown FIG. 12A.
FIG. 12A shows the substrate and oligonucleotide [30-0 (SEQ ID NO:20)] used to test the specific cleavage of a substrate RNA targeted by a pilot oligonucleotide.
FIG. 12B shows an autoradiogram of a gel showing the results of a cleavage reaction using the substrate and oligonucleotide shown in FIG. 13A.
FIG. 13 is a diagram of vector pTTQ18.
FIG. 14 is a diagram of vector pET-3c.
FIGS. 15A-E depicts a set of molecules which are suitable substrates for cleavage by the 5′ nuclease activity of DNAPs (SEQ ID NOS:15 and 17 are depicted in FIG. 15E).
FIG. 16 is an autoradiogram of a gel showing the results of a cleavage reaction run with synthesis-deficient DNAPs.
FIG. 17 is an autoradiogram of a PEI chromatogram resolving the products of an assay for synthetic activity in synthesis-deficient DNAPTaq clones.
FIG. 18A depicts the substrate molecule (SEQ ID NOS:15 and 17) used to test the ability of synthesis-deficient DNAPs to cleave short hairpin structures.
FIG. 18B shows an autoradiogram of a gel resolving the products of a cleavage reaction run using the substrate shown in FIG. 19A.
FIG. 19 provides the complete 206-mer duplex sequence (SEQ ID NO:27) employed as a substrate for the 5′ nucleases of the present invention
FIGS. 20A and B show the cleavage of linear nucleic acid substrates (based on the 206-mer of FIG. 21) by wild type DNAPs and 5′ nucleases isolated from Thermus aquaticus and Thermus flavus. 
FIG. 21A shows the “nibbling” phenomenon detected with the DNAPs of the present invention.
FIG. 21B shows that the “nibbling” of FIG. 25A is 5′ nucleolytic cleavage and not phosphatase cleavage.
FIG. 22 demonstrates that the “nibbling” phenomenon is duplex dependent.
FIG. 23 is a schematic showing how “nibbling” can be employed in a detection assay.
FIGS. 24A and B demonstrates that “nibbling” can be target directed.
FIG. 25 provides a schematic drawing of a target nucleic acid with an invader oligonucleotide and a probe oligonucleotide annealed to the target.
FIG. 26 provides a schematic showing the S-60 hairpin oligonucleotide (SEQ ID NO:29) with the annealed P-15 oligonucleotide (SEQ ID NO:30).
FIG. 27 is an autoradiogram of a gel showing the results of a cleavage reaction run using the S-60 hairpin in the presence or absence of the P-15 oligonucleotide.
FIG. 28 provides a schematic showing three different arrangements of target-specific oligonucleotides and their hybridization to a target nucleic acid which also has a probe oligonucleotide annealed thereto (SEQ ID NOS:31-35).
FIG. 29 is the image generated by a fluorescence imager showing that the presence of an invader oligonucleotide causes a shift in the site of cleavage in a probe/target duplex.
FIG. 30 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run using the three target-specific oligonucleotides diagrammed in FIG. 28.
FIG. 31 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run in the presence or absence of non-target nucleic acid molecules.
FIG. 32 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run in the presence of decreasing amounts of target nucleic acid.
FIG. 33 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run in the presence or absence of saliva extract using various thermostable 5′ nucleases or DNA polymerases.
FIG. 34 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run using various 5′ nucleases.
FIG. 35 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run using two target nucleic acids which differ by a single basepair at two different reaction temperatures.
FIG. 36A provides a schematic showing the effect of elevated temperature upon the annealing and cleavage of a probe oligonucleotide along a target nucleic acid wherein the probe contains a region of noncomplementarity with the target.
FIG. 36B provides a schematic showing the effect of adding an upstream oligonucleotide upon the annealing and cleavage of a probe oligonucleotide along a target nucleic acid wherein the probe contains a region of noncomplementarity with the target.
FIG. 37 provides a schematic showing an arrangement of a target-specific invader oligonucleotide (SEQ ID NO:39) and a target-specific probe oligonucleotide (SEQ ID NO:38) bearing a 5′ Cy3 label along a target nucleic acid (SEQ ID NO:31).
FIG. 38 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run in the presence of increasing concentrations of KCl.
FIG. 39 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run in the presence of increasing concentrations of MnCl2 or MgCl2.
FIG. 40 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run in the presence of increasing amounts of genomic DNA or tRNA.
FIG. 41 is the image generated by a fluorescence imager showing the products of invader-directed cleavage assays run use a HCV RNA target.
FIGS. 42A and B are images generated by a fluorescence imager showing the products of invaderTM-directed cleavage assays run using a HCV RNA target and demonstrate the stability of RNA targets under invaderTM-directed cleavage assay conditions.
FIG. 43 is the image generated by a fluorescence imager showing the sensitivity of detection and the stability of RNA in invader-directed cleavage assays run using a HCV RNA target.
FIG. 44 is the image generated by a fluorescence imager showing thermal degradation of oligonucleotides containing or lacking a 3′ phosphate group.
FIG. 45 depicts the structure of amino-modified oligonucleotides 70 and 74.
FIG. 46 depicts the structure of amino-modified oligonucleotide 75
FIG. 47 depicts the structure of amino-modified oligonucleotide 76.
FIG. 48 is the image generated by a fluorescence imager scan of an IEF gel showing the migration of substrates 70, 70 dp, 74, 74 dp, 75, 75 dp, 76 and 76 dp.
FIG. 49A provides a schematic showing an arrangement of a target-specific invader oligonucleotide (SEQ ID NO:50) and a target-specific probe oligonucleotide (SEQ ID NO:51) bearing a 5′ Cy3 label along a target nucleic acid (SEQ ID NO:52).
FIG. 49B is the image generated by a fluorescence imager showing the detection of specific cleavage products generated in an invasive cleavage assay using charge reversal (i.e., charge based separation of cleavage products).
FIG. 50 is the image generated by a fluorescence imager which depicts the sensitivity of detection of specific cleavage products generated in an invasive cleavage assay using charge reversal.
FIG. 51 depicts a first embodiment of a device for the charge-based separation of oligonucleotides.
FIG. 52 depicts a second embodiment of a device for the charge-based separation of oligonucleotides.
FIG. 53 shows an autoradiogram of a gel showing the results of cleavage reactions run in the presence or absence of a primer oligonucleotide; a sequencing ladder is shown as a size marker.
FIGS. 54A-D depict four pairs of oligonucleotides; in each pair shown, the upper arrangement of a probe annealed to a target nucleic acid lacks an upstream oligonucleotide and the lower arrangement contains an upstream oligonucleotide (SEQ ID NOS:32 and 54-58 are shown in FIGS. 54A-D).
FIG. 55 shows the chemical structure of several positively charged heterodimeric DNA-binding dyes.
FIG. 56 is a schematic showing alternative methods for the tailing and detection of specific cleavage products in the context of the InvaderTM-directed cleavage assay.
FIG. 57 provides a schematic drawing of a target nucleic acid with an InvaderTM oligonucleotide, a miniprobe, and a stacker oligonucleotide annealed to the target.
FIG. 58 provides a space-filling model of the 3-dimensional structure of the T5 5′-exonuclease.
FIG. 59 A-E provides an alignment of the amino acid sequences of several FEN-1 nucleases including the Methanococcus jannaschii FEN-1 protein (MJAFEN1.PRO), the Pyrococcus furiosus FEN-1 protein (PFUFEN1.PRO), the human FEN-1 protein (HUMFEN1.PRO), the mouse FEN-1 protein (MUSFEN1.PRO), the Saccharomyces cerevisiae YKL510 protein (YST510.PRO), the Saccharomyces cerevisiae RAD2 protein (YSTRAD2.PRO), the Shizosaccharomyces pombe RAD13 protein (SPORAD13.PRO), the human XPG protein (HUMXPG.PRO), the mouse XPG protein (MUSXPG.PRO), the Xenopus laevis XPG protein (XENXPG.PRO) and the C. elegans RAD2 protein (CELRAD2.PRO) (SEQ ID NOS:135-145, respectively); portions of the amino acid sequence of some of these proteins were not shown in order to maximize the alignment between proteins (specifically, amino acids 122 to 765 of the YSTRAD2 sequence were deleted; amino acids 122 to 746 of the SPORAD13 sequence were deleted; amino acids 122 to 757 of the HUMXPG sequence were deleted; amino acids 122 to 770 of the MUSXPG sequence were deleted; and amino acids 122 to 790 of the XENXPG sequence were deleted). The numbers to the left of each line of sequence refers to the amino acid residue number; dashes represent gaps introduced to maximize alignment.
FIG. 60 is a schematic showing the S-33 (SEQ ID NO:84) and 11-8-0 (SEQ ID NO:85) oligonucleotides in a folded configuration; the cleavage site is indicated by the arrowhead.
FIG. 61 shows a Coomassie stained SDS-PAGE gel showing the thrombin digestion of Cleavase® BN/thrombin.
FIG. 62 is the image generated by a fluorescence imager showing the products produced by the cleavage of the S-60 hairpin using Cleavase® BN/thrombin (before and after thrombin digestion).
FIG. 63 is the image generated by a fluorescence imager showing the products produced by the cleavage of circular M13 DNA using Cleavase® BN/thrombin.
FIG. 64 is an SDS-PAGE gel showing the migration of purified Cleavase® BN nuclease, Pfu FEN-1, Pwo FEN-1 and Mja FEN-1.
FIG. 65 is the image generated by a fluorescence imager showing the products produced by the cleavage of the S-33 and 11-8-0 oligonucleotides by Cleavase® BN and the Mja FEN-1 nucleases.
FIG. 66 is the image generated by a fluorescence imager showing the products produced by the incubation of an oligonucleotide either having or lacking a 3′-OH group with TdT.
FIG. 67 is the image generated by a fluorescence imager showing the products produced the incubation of cleavage products with TdT.
FIG. 68 is a photograph of a Universal GeneCombTM card showing the capture and detection of cleavage products on a nitrocellulose support.
FIG. 69 is the image generated by a fluorescence imager showing the products produced using the Cleavase® A/G and Pfu FEN-1 nucleases and a fluorescein-labeled probe.
FIG. 70 is the image generated by a fluorescence imager showing the products produced using the Cleavase® A/G and Pfu FEN-1 nucleases and a Cy3-labeled probe.
FIG. 71 is the image generated by a fluorescence imager showing the products produced using the Cleavase® A/G and Pfu FEN-1 nucleases and a TET-labeled probe.
FIGS. 72A and 72B are images generated by a fluorescence imager showing the products produced using the Cleavase® A/G and Pfu FEN-1 nucleases and probes having or lacking a 5′ positive charge; the gel shown in FIG. 83A was run in the standard direction and the gel shown in FIG. 84B was run in the reverse direction.
FIG. 73 shows the structure of 3-nitropyrrole and 5-nitroindole.
FIG. 74 shows the sequence of oligos 109, 61 and 67 (SEQ ID NOS:97, 50 and 51) annealed into a cleavage structure as well as the sequence of oligo 67 (SEQ ID NO:51) and a composite of SEQ ID NOS:98, 99, 101 and 102.
FIG. 75A-C show images generated by a fluorescence imager showing the products produced in an InvaderTM-directed cleavage assay performed at various temperatures using a miniprobe which is either completely complementary to the target or contains a single mismatch with the target.
FIG. 76 shows the sequence of oligos 166 (SEQ ID NO:103), 165 (SEQ ID NO:104), 161 (SEQ ID NO:106), 162 (SEQ ID NO:105) and 164 (SEQ ID NO:107) as well as a cleavage structure.
FIG. 77 shows the image generated by a fluorescence imager showing the products produced in an InvaderTM-directed cleavage assay performed using ras gene sequences as the target.
FIGS. 78A-C show the sequence of the S-60 hairpin (SEQ ID NO:29) (A), and the P-15 oligo (SEQ ID NO:30) (shown annealed to the S-60 hairpin in B) and the image generated by a fluorescence imager showing the products produced by cleavage of the S-60 hairpin in the presence of various InvaderTM oligos.
FIG. 79 shows the structure of various 3′ end substituents.
FIG. 80 is a composite graph showing the effect of probe concentration, temperature and a stacker oligonucleotide on the cleavage of miniprobes.
FIG. 81 shows the sequence of the IT-2 oligonucleotide (SEQ ID NO:115; shown in a folded configuration) as well as the sequence of the IT-1 (SEQ ID NO:116) and IT-A (SEQ ID NO:117) oligos.
FIG. 82 shows the image generated by a fluorescence imager showing the products produced by cleavage of the oligos shown in FIG. 92 by Cleavase® A/G nuclease.
FIG. 83 shows the image generated by a fluorescence imager which provides a comparison of the rates of cleavage by the Pfu FEN-1 and Mja FEN-1 nucleases.
FIG. 84 shows the image generated by a fluorescence imager which depicts the detection of RNA targets using a miniprobe and stacker oligonucleotides.
FIGS. 85A-C provide schematics showing particular embodiments of the present invention wherein a T7 promoter region and copy template annealed with either no oligo (A), a complete promoter oligo (B) or a complete promoter oligo with a 3′ tail (C); one strand of the T7 promoter region is indicated by the hatched line.
FIGS. 86A-D provide schematics showing particular embodiments of the present invention wherein a T7 promoter region and copy template annealed with either a cut probe(A), a partial promoter oligo (B), an uncut oligo (C) or both an uncut probe and a partial promoter oligo (D).
FIG. 87 provides a schematic illustrating one embodiment of the present invention wherein a template-dependent DNA polymerase is used to extend a cut probe to complete a T7 promoter region and thereby allow transcription.
FIGS. 88A and 88B provide a schematic illustrating that an uncut probe combined with a partial promoter oligo does not permit transcription while a cut probe combined with a partial promoter oligo generates a complete (but nicked) promoter which supports transcription.
FIG. 89 shows the image generated by a fluorescence imager which shows that primer extension can be used to complete a partial promoter formed by a cut probe (lanes 1-5) and that annealing a cut probe generated in an invasive cleavage assay can complete a partial T7 promoter to permit transcription (lanes 6-9).
FIGS. 90A-C provide schematics showing particular embodiments of the present invention which illustrate that the use of a partial promoter oligo with a paired 5′ tail can be used to block transcription from a composite promoter formed by the annealing of an uncut probe.
FIG. 91 shows the image generated by a fluorescence imager which shows that transcription from a “leaky” branched T7 composite promoter can be shut down by the use of a downstream partial promoter oligo having a paired 5′ tail.
FIG. 92 shows the image generated by a fluorescence imager which shows that the location of the nick site in a nicked composite T7 promoter can effect the efficiency of transcription.
FIG. 93 shows the image generated by a fluorescence imager which shows that the presence of an unpaired 3′ tail on a full-length promoter oligo decreases but does not abolish transcription. Beneath the image are schematics showing the nucleic acids tested in reactions 1-4; these schematics show SEQ ID NOS:123-125.
FIG. 94 is a schematic which illustrates one embodiment of the present invention where a composite T7 promoter region is created by the binding of the cut probe oligo downstream of the partial promoter oligo.
FIGS. 95A-D provide schematics showing particular embodiments of the present invention which show various ways in which a composite promoter can be formed wherein the nick is located in the template (or bottom) strand.
FIG. 96 is a schematic which illustrates one embodiment of the present invention where the cut probe from an initial invasive cleavage reaction is employed as the InvaderTM oligonucleotide in a second invasive cleavage reaction.
FIG. 97 is a schematic which illustrates one embodiment of the present invention where the cut probe from an initial invasive cleavage reaction is employed as an integrated InvaderTM-target complex in a second invasive cleavage reaction.
FIG. 98 shows the nucleotide sequence of the PR1 probe (SEQ ID NO:119), the IT3 Invader-Target oligonucleotide (SEQ ID NO:118), the IT3-8, IT3-6, IT3-4, IT3-3 and IT3-0 oligonucleotides (SEQ ID NOS:147-151, respectively).
FIG. 99 depicts structures that may be employed to determine the ability of an enzyme to cleave a probe in the presence and the absence of an upstream oligonucleotide. FIG. 99 displays the sequence of oligo 89-15-1 (SEQ ID NO:152), oligo 81-69-5 (SEQ ID NO:156), oligo 81-69-4 (SEQ ID NO155), oligo 81-69-3 (SEQ ID NO:154), oligo 81-69-2 (SEQ ID NO:153) and a portion of M13 mp18 (SEQ ID NO:163).
FIG. 100 shows the image generated by a fluorescence imager which shows the dependence of Pfu FEN-1 on the presence of an overlapping upstream oligonucleotide for specific cleavage of the probe.
FIG. 101a shows the image generated by a fluorescence imager which compares the amount of product generated in a standard (i.e., a non-sequential invasive cleavage reaction) and a sequential invasive cleavage reaction.
FIG. 101b is a graph comparing the amount of product generated in a standard or basic (i.e., a non-sequential invasive cleavage reaction) and a sequential invasive cleavage reaction (“invader sqrd”) (y axis=fluorescence units; x axix=attomoles of target).
FIG. 102 shows the image generated by a fluorescence imager which shows that the products of a completed sequential invasive cleavage reaction cannot cross contaminant a subsequent similar reaction.
FIG. 103 shows the sequence of the oligonucleotide employed in an invasive cleavage reaction for the detection of HCMV viral DNA; FIG. 103 shows the sequence of oligo 89-76 (SEQ ID NO:161), oligo 89-44 (SEQ ID NO:160) and nucleotides 3057-3110 of the HCMV genome (SEQ ID NO:162).
FIG. 104 shows the image generated by a fluorescence imager which shows the sensitive detection of HCMV viral DNA in samples containing human genomic DNA using an invasive cleavage reaction.



DEFINITIONS
As used herein, the terms “complementary” or “complementarity” are used in reference to polynucleotides (i.e., a sequence of nucleotides such as an oligonucleotide or a target nucleic acid) related by the base-pairing rules. For example, for the sequence “A-G-T,” is complementary to the sequence “T-C-A.” Complementarity may be “partial,” in which only some of the nucleic acids' bases are matched according to the base pairing rules. Or, there may be “complete” or “total” complementarity between the nucleic acids. The degree of complementarity between nucleic acid strands has significant effects on the efficiency and strength of hybridization between nucleic acid strands. This is of particular importance in amplification reactions, as well as detection methods which depend upon binding between nucleic acids.
The term “self-complementarity” when used in reference to a nucleic acid strand (e.g., an oligonucleotide) means that separate regions of that strand can base-pair. Because this term refers only to intramolecular base-pairing, any strand said to have a region of self-complementarity must have at least two regions capable of base-pairing with one another. As defined above, complementarity may be either “complete” or “partial”. As used in reference to the probe oligonucleotides of the present invention, regions are considered to have significant self-complementarity when they may form a duplex of at least 3 contiguous base pairs (i.e., three base pairs of complete complementarity), or when they may form a longer duplex that is partially complementary. The ability of an oligonucleotide having a region of self-complementarity to successfully serve both as a target strand for a probe, and as an upstream oligonucleotide that directs invasive cleavage of that probe as depicted in FIG. 97, is considered sufficient demonstration of self-complementarity as defined herein.
The term “homology” refers to a degree of identity. There may be partial homology or complete homology. A partially identical sequence is one that is less than 100% identical to another sequence.
As used herein, the term “hybridization” is used in reference to the pairing of complementary nucleic acids. Hybridization and the strength of hybridization (i.e., the strength of the association between the nucleic acids) is impacted by such factors as the degree of complementary between the nucleic acids, stringency of the conditions involved, the Tm of the formed hybrid, and the G:C ratio within the nucleic acids.
As used herein, the term “Tm” is used in reference to the “melting temperature.” The melting temperature is the temperature at which a population of double-stranded nucleic acid molecules becomes half dissociated into single strands. The equation for calculating the Tm of nucleic acids is well known in the art. As indicated by standard references, a simple estimate of the Tm value may be calculated by the equation: Tm=81.5+0.41(% G+C), when a nucleic acid is in aqueous solution at 1 M NaCl (see e.g., Anderson and Young, Quantitative Filter Hybridization, in Nucleic Acid Hybridization (1985). Other references include more sophisticated computations which take structural as well as sequence characteristics into account for the calculation of Tm.
As used herein the term “stringency” is used in reference to the conditions of temperature, ionic strength, and the presence of other compounds, under which nucleic acid hybridizations are conducted. With “high stringency” conditions, nucleic acid base pairing will occur only between nucleic acid fragments that have a high frequency of complementary base sequences. Thus, conditions of “weak” or “low” stringency are often required when it is desired that nucleic acids which are not completely complementary to one another be hybridized or annealed together.
The term “gene” refers to a DNA sequence that comprises control and coding sequences necessary for the production of a polypeptide or precursor. The polypeptide can be encoded by a full length coding sequence or by any portion of the coding sequence so long as the desired enzymatic activity is retained.
The term “wild-type” refers to a gene or gene product which has the characteristics of that gene or gene product when isolated from a naturally occurring source. A wild-type gene is that which is most frequently observed in a population and is thus arbitrarily designed the “normal” or “wild-type” form of the gene. In contrast, the term “modified” or “mutant” refers to a gene or gene product which displays modifications in sequence and or functional properties (i.e., altered characteristics) when compared to the wild-type gene or gene product. It is noted that naturally-occurring mutants can be isolated; these are identified by the fact that they have altered characteristics when compared to the wild-type gene or gene product.
The term “recombinant DNA vector” as used herein refers to DNA sequences containing a desired coding sequence and appropriate DNA sequences necessary for the expression of the operably linked coding sequence in a particular host organism. DNA sequences necessary for expression in procaryotes include a promoter, optionally an operator sequence, a ribosome binding site and possibly other sequences. Eukaryotic cells are known to utilize promoters, polyadenlyation signals and enhancers.
The term “LTR” as used herein refers to the long terminal repeat found at each end of a provirus (i.e., the integrated form of a retrovirus). The LTR contains numerous regulatory signals including transcriptional control elements, polyadenylation signals and sequences needed for replication and integration of the viral genome. The viral LTR is divided into three regions called U3, R and U5.
The U3 region contains the enhancer and promoter elements. The U5 region contains the polyadenylation signals. The R (repeat) region separates the U3 and U5 regions and transcribed sequences of the R region appear at both the 5′ and 3′ ends of the viral RNA.
The term “oligonucleotide” as used herein is defined as a molecule comprised of two or more deoxyribonucleotides or ribonucleotides, preferably at least 5 nucleotides, more preferably at least about 10-15 nucleotides and more preferably at least about 15 to 30 nucleotides. The exact size will depend on many factors, which in turn depends on the ultimate function or use of the oligonucleotide. The oligonucleotide may be generated in any manner, including chemical synthesis, DNA replication, reverse transcription, or a combination thereof.
Because mononucleotides are reacted to make oligonucleotides in a manner such that the 5′ phosphate of one mononucleotide pentose ring is attached to the 3′ oxygen of its neighbor in one direction via a phosphodiester linkage, an end of an oligonucleotide is referred to as the “5′ end” if its 5′ phosphate is not linked to the 3′ oxygen of a mononucleotide pentose ring and as the “3′ end” if its 3′ oxygen is not linked to a 5′ phosphate of a subsequent mononucleotide pentose ring. As used herein, a nucleic acid sequence, even if internal to a larger oligonucleotide, also may be said to have 5′ and 3′ ends. A first region along a nucleic acid strand is said to be upstream of another region if the 3′ end of the first region is before the 5′ end of the second region when moving along a strand of nucleic acid in a 5′ to 3′ direction.
When two different, non-overlapping oligonucleotides anneal to different regions of the same linear complementary nucleic acid sequence, and the 3′ end of one oligonucleotide points towards the 5′ end of the other, the former may be called the “upstream” oligonucleotide and the latter the “downstream” oligonucleotide.
The term “primer” refers to an oligonucleotide which is capable of acting as a point of initiation of synthesis when placed under conditions in which primer extension is initiated. An oligonucleotide “primer” may occur naturally, as in a purified restriction digest or may be produced synthetically.
A primer is selected to be “substantially” complementary to a strand of specific sequence of the template. A primer must be sufficiently complementary to hybridize with a template strand for primer elongation to occur. A primer sequence need not reflect the exact sequence of the template. For example, a non-complementary nucleotide fragment may be attached to the 5′ end of the primer, with the remainder of the primer sequence being substantially complementary to the strand. Non-complementary bases or longer sequences can be interspersed into the primer, provided that the primer sequence has sufficient complementarity with the sequence of the template to hybridize and thereby form a template primer complex for synthesis of the extension product of the primer.
“Hybridization” methods involve the annealing of a complementary sequence to the target nucleic acid (the sequence to be detected; the detection of this sequence may be by either direct or indirect means). The ability of two polymers of nucleic acid containing complementary sequences to find each other and anneal through base pairing interaction is a well-recognized phenomenon. The initial observations of the “hybridization” process by Marmur and Lane, Proc. Natl. Acad. Sci. USA 46:453 (1960) and Doty et al., Proc. Natl. Acad. Sci. USA 46:461 (1960) have been followed by the refinement of this process into an essential tool of modem biology.
With regard to complementarity, it is important for some diagnostic applications to determine whether the hybridization represents complete or partial complementarity. For example, where it is desired to detect simply the presence or absence of pathogen DNA (such as from a virus, bacterium, fungi, mycoplasma, protozoan) it is only important that the hybridization method ensures hybridization when the relevant sequence is present; conditions can be selected where both partially complementary probes and completely complementary probes will hybridize. Other diagnostic applications, however, may require that the hybridization method distinguish between partial and complete complementarity. It may be of interest to detect genetic polymorphisms. For example, human hemoglobin is composed, in part, of four polypeptide chains. Two of these chains are identical chains of 141 amino acids (alpha chains) and two of these chains are identical chains of 146 amino acids (beta chains). The gene encoding the beta chain is known to exhibit polymorphism. The normal allele encodes a beta chain having glutamic acid at the sixth position. The mutant allele encodes a beta chain having valine at the sixth position. This difference in amino acids has a profound (most profound when the individual is homozygous for the mutant allele) physiological impact known clinically as sickle cell anemia. It is well known that the genetic basis of the amino acid change involves a single base difference between the normal allele DNA sequence and the mutant allele DNA sequence.
The complement of a nucleic acid sequence as used herein refers to an oligonucleotide which, when aligned with the nucleic acid sequence such that the 5′ end of one sequence is paired with the 3′ end of the other, is in “antiparallel association.” Certain bases not commonly found in natural nucleic acids may be included in the nucleic acids of the present invention and include, for example, inosine and 7-deazaguanine. Complementarity need not be perfect; stable duplexes may contain mismatched base pairs or unmatched bases. Those skilled in the art of nucleic acid technology can determine duplex stability empirically considering a number of variables including, for example, the length of the oligonucleotide, base composition and sequence of the oligonucleotide, ionic strength and incidence of mismatched base pairs.
Stability of a nucleic acid duplex is measured by the melting temperature, or “Tm.” The Tm of a particular nucleic acid duplex under specified conditions is the temperature at which on average half of the base pairs have disassociated.
The term “label” as used herein refers to any atom or molecule which can be used to provide a detectable (preferably quantifiable) signal, and which can be attached to a nucleic acid or protein. Labels may provide signals detectable by fluorescence, radioactivity, colorimetry, gravimetry, X-ray diffraction or absorption, magnetism, enzymatic activity, and the like. A label may be a charged moeity (positive or negative charge) or alternatively, may be charge neutral.
The term “cleavage structure” as used herein, refers to a structure which is formed by the interaction of a probe oligonucleotide and a target nucleic acid to form a duplex, said resulting structure being cleavable by a cleavage means, including but not limited to an enzyme. The cleavage structure is a substrate for specific cleavage by said cleavage means in contrast to a nucleic acid molecule which is a substrate for non-specific cleavage by agents such as phosphodiesterases which cleave nucleic acid molecules without regard to secondary structure (i.e., no formation of a duplexed structure is required).
The term “cleavage means” as used herein refers to any means which is capable of cleaving a cleavage structure, including but not limited to enzymes. The cleavage means may include native DNAPs having 5′ nuclease activity (e.g., Taq DNA polymerase, E. coli DNA polymerase I) and, more specifically, modified DNAPs having 5′ nuclease but lacking synthetic activity. The ability of 5′ nucleases to cleave naturally occurring structures in nucleic acid templates (structure-specific cleavage) is useful to detect internal sequence differences in nucleic acids without prior knowledge of the specific sequence of the nucleic acid. In this manner, they are structure-specific enzymes. “Structure-specific nucleases” or “structure-specific enzymes” are enzymes which recognize specific secondary structures in a nucleic molecule and cleave these structures. The cleavage means of the invention cleave a nucleic acid molecule in response to the formation of cleavage structures; it is not necessary that the cleavage means cleave the cleavage structure at any particular location within the cleavage structure.
The cleavage means is not restricted to enzymes having solely 5′ nuclease activity. The cleavage means may include nuclease activity provided from a variety of sources including the Cleavase® enzymes, the FEN-1 endonucleases (including RAD2 and XPG proteins), Taq DNA polymerase and E. coli DNA polymerase I.
The term “thermostable” when used in reference to an enzyme, such as a 5′ nuclease, indicates that the enzyme is functional or active (i.e., can perform catalysis) at an elevated temperature, i.e., at about 55° C. or higher.
The term “cleavage products” as used herein, refers to products generated by the reaction of a cleavage means with a cleavage structure (i.e., the treatment of a cleavage structure with a cleavage means).
The term “target nucleic acid” refers to a nucleic acid molecule which contains a sequence which has at least partial complementarity with at least a probe oligonucleotide and may also have at least partial complementarity with an invader oligonucleotide. The target nucleic acid may comprise single- or double-stranded DNA or RNA.
The term “probe oligonucleotide” refers to an oligonucleotide which interacts with a target nucleic acid to form a cleavage structure in the presence or absence of an invader oligonucleotide. When annealed to the target nucleic acid, the probe oligonucleotide and target form a cleavage structure and cleavage occurs within the probe oligonucleotide. In the presence of an invader oligonucleotide upstream of the probe oligonucleotide along the target nucleic acid will shift the site of cleavage within the probe oligonucleotide (relative to the site of cleavage in the absence of the invader).
The term “non-target cleavage product” refers to a product of a cleavage reaction which is not derived from the target nucleic acid. As discussed above, in the methods of the present invention, cleavage of the cleavage structure occurs within the probe oligonucleotide. The fragments of the probe oligonucleotide generated by this target nucleic acid-dependent cleavage are “non-target cleavage products.”
The term “invader oligonucleotide” refers to an oligonucleotide which contains sequences at its 3′ end which are substantially the same as sequences located at the 5′ end of a probe oligonucleotide; these regions will compete for hybridization to the same segment along a complementary target nucleic acid.
The term “substantially single-stranded” when used in reference to a nucleic acid substrate means that the substrate molecule exists primarily as a single strand of nucleic acid in contrast to a double-stranded substrate which exists as two strands of nucleic acid which are held together by inter-strand base pairing interactions.
The term “sequence variation” as used herein refers to differences in nucleic acid sequence between two nucleic acids. For example, a wild-type structural gene and a mutant form of this wild-type structural gene may vary in sequence by the presence of single base substitutions and/or deletions or insertions of one or more nucleotides. These two forms of the structural gene are said to vary in sequence from one another. A second mutant form of the structural gene may exist. This second mutant form is said to vary in sequence from both the wild-type gene and the first mutant form of the gene.
The term “liberating” as used herein refers to the release of a nucleic acid fragment from a larger nucleic acid fragment, such as an oligonucleotide, by the action of a 5′ nuclease such that the released fragment is no longer covalently attached to the remainder of the oligonucleotide.
The term “Km” as used herein refers to the Michaelis-Menten constant for an enzyme and is defined as the concentration of the specific substrate at which a given enzyme yields one-half its maximum velocity in an enzyme catalyzed reaction.
The term “nucleotide analog” as used herein refers to modified or non-naturally occurring nucleotides such as 7-deaza purines (i.e., 7-deaza-dATP and 7-deaza-dGTP). Nucleotide analogs include base analogs and comprise modified forms of deoxyribonucleotides as well as ribonucleotides.
The term “polymorphic locus” is a locus present in a population which shows variation between members of the population (i.e., the most common allele has a frequency of less than 0.95). In contrast, a “monomorphic locus” is a genetic locus at little or no variations seen between members of the population (generally taken to be a locus at which the most common allele exceeds a frequency of 0.95 in the gene pool of the population).
The term “microorganism” as used herein means an organism too small to be observed with the unaided eye and includes, but is not limited to bacteria, virus, protozoans, fungi, and ciliates.
The term “microbial gene sequences” refers to gene sequences derived from a microorganism.
The term “bacteria” refers to any bacterial species including eubacterial and archaebacterial species.
The term “virus” refers to obligate, ultramicroscopic, intracellular parasites incapable of autonomous replication (i.e., replication requires the use of the host cell's machinery).
The term “multi-drug resistant” or “multiple-drug resistant” refers to a microorganism which is resistant to more than one of the antibiotics or antimicrobial agents used in the treatment of said microorganism.
The term “sample” in the present specification and claims is used in its broadest sense. On the one hand it is meant to include a specimen or culture (e.g., microbiological cultures). On the other hand, it is meant to include both biological and environmental samples.
Biological samples may be animal, including human, fluid, solid (e.g., stool) or tissue, as well as liquid and solid food and feed products and ingredients such as dairy items, vegetables, meat and meat by-products, and waste. Biological samples may be obtained from all of the various families of domestic animals, as well as feral or wild animals, including, but not limited to, such animals as ungulates, bear, fish, lagamorphs, rodents, etc.
Environmental samples include environmental material such as surface matter, soil, water and industrial samples, as well as samples obtained from food and dairy processing instruments, apparatus, equipment, utensils, disposable and non-disposable items. These examples are not to be construed as limiting the sample types applicable to the present invention.
The term “source of target nucleic acid” refers to any sample which contains nucleic acids (RNA or DNA). Particularly preferred sources of target nucleic acids are biological samples including, but not limited to blood, saliva, cerebral spinal fluid, pleural fluid, milk, lymph, sputum and semen.
An oligonucleotide is said to be present in “excess” relative to another oligonucleotide (or target nucleic acid sequence) if that oligonucleotide is present at a higher molar concentration that the other oligonucleotide (or target nucleic acid sequence). When an oligonucleotide such as a probe oligonucleotide is present in a cleavage reaction in excess relative to the concentration of the complementary target nucleic acid sequence, the reaction may be used to indicate the amount of the target nucleic acid present. Typically, when present in excess, the probe oligonucleotide will be present at least a 100-fold molar excess; typically at least 1 pmole of each probe oligonucleotide would be used when the target nucleic acid sequence was present at about 10 fmoles or less.
A sample “suspected of containing” a first and a second target nucleic acid may contain either, both or neither target nucleic acid molecule.
The term “charge-balanced” oligonucleotide refers to an oligonucleotide (the input oligonucleotide in a reaction) which has been modified such that the modified oligonucleotide bears a charge, such that when the modified oligonucleotide is either cleaved (i.e., shortened) or elongated, a resulting product bears a charge different from the input oligonucleotide (the “charge-unbalanced” oligonucleotide) thereby permitting separation of the input and reacted oligonucleotides on the basis of charge. The term “charge-balanced” does not imply that the modified or balanced oligonucleotide has a net neutral charge (although this can be the case). Charge-balancing refers to the design and modification of an oligonucleotide such that a specific reaction product generated from this input oligonucleotide can be separated on the basis of charge from the input oligonucleotide.
For example, in an invader-directed cleavage assay in which the probe oligonucleotide bears the sequence: 5′-TTCTTTTCACCAGCGAGACGGG-3′ (i.e., SEQ ID NO:50 without the modified bases) and cleavage of the probe occurs between the second and third residues, one possible charge-balanced version of this oligonucleotide would be: 5′-Cy3-Amino-T-Amino-TCTTTTCACCAGCGAGAC GGG-3′. This modified oligonucleotide bears a net negative charge. After cleavage, the following oligonucleotides are generated: 5′-Cy3-Amino-T-Amino-T-3′ and 5′-CTTTTCACCAGCGAGACGGG-3′ (residues 3-22 of SEQ ID NO:50). 5′-Cy3-Amino-T-Amino-T-3′ bears a detectable moeity (the positively-charged Cy3 dye) and two amino-modified bases. The amino-modified bases and the Cy3 dye contribute positive charges in excess of the negative charges contributed by the phosphate groups and thus the 5′-Cy3-Amino-T-Amino-T-3′ oligonucleotide has a net positive charge. The other, longer cleavage fragment, like the input probe, bears a net negative charge. Because the 5′-Cy3-Amino-T-Amino-T-3′ fragment is separable on the basis of charge from the input probe (the charge-balanced oligonucleotide), it is referred to as a charge-unbalanced oligonucleotide. The longer cleavage product cannot be separated on the basis of charge from the input oligonucleotide as both oligonucleotides bear a net negative charge; thus, the longer cleavage product is not a charge-unbalanced oligonucleotide.
The term “net neutral charge” when used in reference to an oligonucleotide, including modified oligonucleotides, indicates that the sum of the charges present (i.e, R-NH3+ groups on thymidines, the N3 nitrogen of cytosine, presence or absence or phosphate groups, etc.) under the desired reaction conditions is essentially zero. An oligonucleotide having a net neutral charge would not migrate in an electrical field.
The term “net positive charge” when used in reference to an oligonucleotide, including modified oligonucleotides, indicates that the sum of the charges present (i.e, R-NH3+ groups on thymidines, the N3 nitrogen of cytosine, presence or absence or phosphate groups, etc.) under the desired reaction conditions is +1 or greater. An oligonucleotide having a net positive charge would migrate toward the negative electrode in an electrical field.
The term “net negative charge” when used in reference to an oligonucleotide, including modified oligonucleotides, indicates that the sum of the charges present (i.e, R-NH3+ groups on thymidines, the N3 nitrogen of cytosine, presence or absence or phosphate groups, etc.) under the desired reaction conditions is −1 or lower. An oligonucleotide having a net negative charge would migrate toward the positive electrode in an electrical field.
The term “polymerization means” refers to any agent capable of facilitating the addition of nucleoside triphosphates to an oligonucleotide. Preferred polymerization means comprise DNA polymerases.
The term “ligation means” refers to any agent capable of facilitating the ligation (i.e., the formation of a phosphodiester bond between a 3′-OH and a 5′-P located at the termini of two strands of nucleic acid). Preferred ligation means comprise DNA ligases and RNA ligases.
The term “reactant” is used herein in its broadest sense. The reactant can comprise an enzymatic reactant, a chemical reactant or ultraviolet light (ultraviolet light, particularly short wavelength ultraviolet light is known to break oligonucleotide chains). Any agent capable of reacting with an oligonucleotide to either shorten (i.e., cleave) or elongate the oligonucleotide is encompassed within the term “reactant.”
The term “adduct” is used herein in its broadest sense to indicate any compound or element which can be added to an oligonucleotide. An adduct may be charged (positively or negatively) or may be charge neutral. An adduct may be added to the oligonucleotide via covalent or non-covalent linkages. Examples of adducts, include but are not limited to indodicarbocyanine dyes (e.g., Cy3 and Cy5), amino-substituted nucleotides, ethidium bromide, ethidium homodimer, (1,3-propanediamino)propidium, (diethylenetriamino)propidium, thiazole orange, (N—N′-tetramethyl-1,3-propanediamino)propyl thiazole orange, (N—N′-tetramethyl-1,2-ethanediamino)propyl thiazole orange, thiazole orange-thiazole orange homodimer (TOTO), thiazole orande-thiazole blue heterodimer (TOTAB), thiazole orange-ethidium heterodimer 1 (TOED1), thiazole orange-ethidium heterodimer 2 (TOED2) and florescien-ethidium heterodimer (FED), psoralens, biotin, streptavidin, avidin, etc.
Where a first oligonucleotide is complementary to a region of a target nucleic acid and a second oligonucleotide has complementary to the same region (or a portion of this region) a “region of overlap” exists along the target nucleic acid. The degree of overlap will vary depending upon the nature of the complementarity (see, e.g., region “X” in FIGS. 25 and 56 and the accompanying discussions).
As used herein, the term “purified” or “to purify” refers to the removal of contaminants from a sample. For example, recombinant Cleavase® nucleases are expressed in bacterial host cells and the nucleases are purified by the removal of host cell proteins; the percent of these recombinant nucleases is thereby increased in the sample.
The term “recombinant DNA molecule” as used herein refers to a DNA molecule which is comprised of segments of DNA joined together by means of molecular biological techniques.
The term “recombinant protein” or “recombinant polypeptide” as used herein refers to a protein molecule which is expressed from a recombinant DNA molecule.
As used herein the term “portion” when in reference to a protein (as in “a portion of a given protein”) refers to fragments of that protein. The fragments may range in size from four amino acid residues to the entire amino acid sequence minus one amino acid.
“Nucleic acid sequence” as used herein refers to an oligonucleotide, nucleotide or polynucleotide, and fragments or portions thereof, and to DNA or RNA of genomic or synthetic origin which may be single- or double-stranded, and represent the sense or antisense strand. Similarly, “amino acid sequence” as used herein refers to peptide or protein sequence.
“Peptide nucleic acid” (“PNA”) as used herein refers to a molecule which comprises an oligomer to which an amino acid residue, such as lysine, and an amino group have been added. These small molecules, also designated anti-gene agents, stop transcript elongation by binding to their complementary strand of nucleic acid [Nielsen PE et al. (1993) Anticancer Drug Des. 8:53-63].
As used herein, the terms “purified” or “substantially purified” refer to molecules, either nucleic or amino acid sequences, that are removed from their natural environment, isolated or separated, and are at least 60% free, preferably 75% free, and most preferably 90% free from other components with which they are naturally associated. An “isolated polynucleotide” or “isolated oligonucleotide” is therefore a substantially purified polynucleotide.
An isolated oligonucleotide (or polynucleotide) encoding a Pyrococcus woesei (Pwo) FEN-1 endonuclease having a region capable of hybridizing to SEQ ID NO:80 is an oligonucleotide containing sequences encoding at least the amino-terminal portion of Pwo FEN-1 endonuclease. An isolated oligonucleotide (or polynucleotide) encoding a Pwo FEN-1 endonuclease having a region capable of hybridizing to SEQ ID NO:81 is an oligonucleotide containing sequences encoding at least the carboxy-terminal portion of Pwo FEN-1 endonuclease. An isolated oligonucleotide (or polynucleotide) encoding a Pwo FEN-1 endonuclease having a region capable of hybridizing to SEQ ID NOS:82 and 83 is an oligonucleotide containing sequences encoding at least portions of Pwo FEN-1 endonuclease protein located internal to either the amino or carboxy-termini of the Pwo FEN-1 endonuclease protein.
As used herein, the term “fusion protein” refers to a chimeric protein containing the protein of interest (i.e., Cleavase® BN/thrombin nuclease and portions or fragments thereof) joined to an exogenous protein fragment (the fusion partner which consists of a non-Cleavase® BN/thrombin nuclease protein). The fusion partner may enhance solubility of recombinant chimeric protein (e.g., the Cleavase® BN/thrombin nuclease) as expressed in a host cell, may provide an affinity tag (e.g., a his-tag) to allow purification of the recombinant fusion protein from the host cell or culture supernatant, or both. If desired, the fusion protein may be removed from the protein of interest (e.g., Cleavase® BN/thrombin nuclease or fragments thereof) by a variety of enzymatic or chemical means known to the art.
The term “purified Pfu FEN-1 endonuclease having a molecular weight of about 38.7 kilodaltons” refers to a FEN-1 endonuclease isolated from Pyrococcus woesei which has a molecular weight on SDS-PAGE gels of about 38.7 kDa when the SDS-PAGE is conducted under the conditions described in Ex. 28. Those skilled in the art understand that the same protein preparation applied to separate gels of apparently the same composition can yield estimated molecular weights which vary somewhat from one another (approximately 5-15%).
The term “continuous strand of nucleic acid” as used herein is means a strand of nucleic acid that has a continuous, covalently linked, backbone structure, without nicks or other disruptions. The disposition of the base portion of each nucleotide, whether base-paired, single-stranded or mismatched, is not an element in the definition of a continuous strand. The backbone of the continuous strand is not limited to the ribose-phosphate or deoxyribose-phosphate compositions that are found in naturally occurring, unmodified nucleic acids. A nucleic acid of the present invention may comprise modifications in the structure of the backbone, including but not limited to phosphorothioate residues, phosphonate residues, 2′ substituted ribose residues (e.g., 2′-O-methyl ribose) and alternative sugar (e.g., arabinose) containing residues.
The term “continuous duplex” as used herein refers to a region of double stranded nucleic acid in which there is no disruption in the progression of basepairs within the duplex, i.e., the base pairs along the duplex are not distorted to accommodate a gap, bulge or mismatch with the confines of the region of continuous duplex. As used herein the term refers only to the arrangement of the basepairs within the duplex, without implication of continuity in the backbone portion of the nucleic acid strand. Duplex nucleic acids with uninterrupted basepairing, but with nicks in one or both strands are within the definition of a continuous duplex.
The term “duplex” refers to the state of nucleic acids in which the base portions of the nucleotides on one strand are bound through hydrogen bonding the their complementary bases arrayed on a second strand. The condition of being in a duplex form reflects on the state of the bases of a nucleic acid. By virtue of base pairing, the strands of nucleic acid also generally assume the tertiary structure of a double helix, having a major and a minor groove. The assumption of the helical form is implicit in the act of becoming duplexed.
The term “duplex dependent protein binding” refers to the binding of proteins to nucleic acid that is dependent on the nucleic acid being in a duplex,or helical form.
The term “duplex dependent protein binding sites or regions” as used herein refers to discrete regions or sequences within a nucleic acid that are bound with particular affinity by specific duplex-dependent nucleic acid binding proteins. This is in contrast to the generalized duplex-dependent binding of proteins that are not site-specific, such as the histone proteins that bind chromatin with little reference to specific sequences or sites.
The term “protein binding region” as used herein refers to a nucleic acid region identified by a sequence or structure as binding to a particular protein or class of proteins. It is within the scope of this definition to include those regions that contain sufficient genetic information to allow identifications of the region by comparison to known sequences, but which might not have the requisite structure for actual binding (e.g., a single strand of a duplex-depending nucleic acid binding protein site). As used herein “protein binding region” excludes restriction endonuclease binding regions.
The term “complete double stranded protein binding region” as used herein refers to the minimum region of continuous duplex required to allow binding or other activity of a duplex-dependent protein. This definition is intended to encompass the observation that some duplex dependent nucleic acid binding proteins can interact with full activity with regions of duplex that may be shorter than a canonical protein binding region as observed in one or the other of the two single strands. In other words, one or more nucleotides in the region may be allowed to remain unpaired without suppressing binding. As used here in, the term “complete double stranded binding region” refers to the minimum sequence that will accommodate the binding function. Because some such regions can tolerate non-duplex sequences in multiple places, although not necessarily simultaneously, a single protein binding region might have several shorter sub-regions that, when duplexed, will be fully competent for protein binding.
The term “template” refers to a strand of nucleic acid on which a complementary copy is built from nucleoside triphosphates through the activity of a template-dependent nucleic acid polymerase. Within a duplex the template strand is, by convention, depicted and described as the “bottom” strand. Similarly, the non-template strand is often depicted and described as the “top” strand.
The term “template-dependent RNA polymerase” refers to a nucleic acid polymerase that creates new RNA strands through the copying of a template strand as described above and which does not synthesize RNA in the absence of a template. This is in contrast to the activity of the template-independent nucleic acid polymerases that synthesize or extend nucleic acids without reference to a template, such as terminal deoxynucleotidyl transferase, or Poly A polymerase.
DESCRIPTION OF THE INVENTION
The present invention relates to methods and compositions for treating nucleic acid, and in particular, methods and compositions for detection and characterization of nucleic acid sequences and sequence changes.
The present invention relates to means for cleaving a nucleic acid cleavage structure in a site-specific manner. In particular, the present invention relates to a cleaving enzyme having 5′ nuclease activity without interfering nucleic acid synthetic ability.
This invention provides 5′ nucleases derived from thermostable DNA polymerases which exhibit altered DNA synthetic activity from that of native thermostable DNA polymerases. The 5′ nuclease activity of the polymerase is retained while the synthetic activity is reduced or absent. Such 5′ nucleases are capable of catalyzing the structure-specific cleavage of nucleic acids in the absence of interfering synthetic activity. The lack of synthetic activity during a cleavage reaction results in nucleic acid cleavage products of uniform size.
The novel properties of the nucleases of the invention form the basis of a method of detecting specific nucleic acid sequences. This method relies upon the amplification of the detection molecule rather than upon the amplification of the target sequence itself as do existing methods of detecting specific target sequences.
DNA polymerases (DNAPs), such as those isolated from E. coli or from thermophilic bacteria of the genus Thermus, are enzymes that synthesize new DNA strands. Several of the known DNAPs contain associated nuclease activities in addition to the synthetic activity of the enzyme.
Some DNAPs are known to remove nucleotides from the 5′ and 3′ ends of DNA chains [Komberg, DNA Replication, W. H. Freeman and Co., San Francisco, pp. 127-139 (1980)]. These nuclease activities are usually referred to as 5′ exonuclease and 3′ exonuclease activities, respectively. For example, the 5′ exonuclease activity located in the N-terminal domain of several DNAPs participates in the removal of RNA primers during lagging strand synthesis during DNA replication and the removal of damaged nucleotides during repair. Some DNAPs, such as the E. coli DNA polymerase (DNAPEc1), also have a 3′ exonuclease activity responsible for proof-reading during DNA synthesis (Komberg, supra).
A DNAP isolated from Thermus aquaticus, termed Taq DNA polymerase (DNAPTaq), has a 5′ exonuclease activity, but lacks a functional 3′ exonucleolytic domain [Tindall and Kunkell, Biochem. 27:6008 (1988)]. Derivatives of DNAPEc1 and DNAPTaq, respectively called the Klenow and Stoffel fragments, lack 5′ exonuclease domains as a result of enzymatic or genetic manipulations [Brutlag et al., Biochem. Biophys. Res. Commun. 37:982 (1969); Erlich et al., Science 252:1643 (1991); Setlow and Komberg, J. Biol. Chem. 247:232 (1972)].
The 5′ exonuclease activity of DNAPTaq was reported to require concurrent synthesis [Gelfand, PCR Technology—Principles and Applications for DNA Amplification (H. A. Erlich, Ed.), Stockton Press, New York, p. 19 (1989)]. Although mononucleotides predominate among the digestion products of the 5′ exonucleases of DNAPTaq and DNAPEc1, short oligonucleotides (≦12 nucleotides) can also be observed implying that these so-called 5′ exonucleases can function endonucleolytically [Setlow, supra; Holland et al., Proc. Natl. Acad. Sci. USA 88:7276 (1991)].
In WO 92/06200, Gelfand et al. show that the preferred substrate of the 5′ exonuclease activity of the thermostable DNA polymerases is displaced single-stranded DNA. Hydrolysis of the phosphodiester bond occurs between the displaced single-stranded DNA and the double-helical DNA with the preferred exonuclease cleavage site being a phosphodiester bond in the double helical region. Thus, the 5′ exonuclease activity usually associated with DNAPs is a structure-dependent single-stranded endonuclease and is more properly referred to as a 5′ nuclease. Exonucleases are enzymes which cleave nucleotide molecules from the ends of the nucleic acid molecule. Endonucleases, on the other hand, are enzymes which cleave the nucleic acid molecule at internal rather than terminal sites. The nuclease activity associated with some thermostable DNA polymerases cleaves endonucleolytically but this cleavage requires contact with the 5′ end of the molecule being cleaved. Therefore, these nucleases are referred to as 5′ nucleases.
When a 5′ nuclease activity is associated with a eubacterial Type A DNA polymerase, it is found in the one-third N-terminal region of the protein as an independent functional domain. The C-terminal two-thirds of the molecule constitute the polymerization domain which is responsible for the synthesis of DNA. Some Type A DNA polymerases also have a 3′ exonuclease activity associated with the two-third C-terminal region of the molecule.
The 5′ exonuclease activity and the polymerization activity of DNAPs have been separated by proteolytic cleavage or genetic manipulation of the polymerase molecule. To date thermostable DNAPs have been modified to remove or reduce the amount of 5′ nuclease activity while leaving the polymerase activity intact.
The Klenow or large proteolytic cleavage fragment of DNAPEc1 contains the polymerase and 3′ exonuclease activity but lacks the 5′ nuclease activity. The Stoffel fragment of DNAPTaq (DNAPStf) lacks the 5′ nuclease activity due to a genetic manipulation which deleted the N-terminal 289 amino acids of the polymerase molecule [Erlich et al., Science 252:1643 (1991)]. WO 92/06200 describes a thermostable DNAP with an altered level of 5′ to 3′ exonuclease. U.S. Pat. No. 5,108,892 describes a Thermus aquaticus DNAP without a 5′ to 3′ exonuclease. However, the art of molecular biology lacks a thermostable DNA polymerase with a lessened amount of synthetic activity.
The present invention provides 5′ nucleases derived from thermostable Type A DNA polymerases that retain 5′ nuclease activity but have reduced or absent synthetic activity. The ability to uncouple the synthetic activity of the enzyme from the 5′ nuclease activity proves that the 5′ nuclease activity does not require concurrent DNA synthesis as was previously reported (Gelfand, PCR Technology, supra).
The description of the invention is divided into: I. Generation of 5′ Nucleases Derived From Thermostable DNA Polymerases; II. Detection of Specific Nucleic Acid Sequences Using 5′ Nucleases in an InvaderTM-Directed Cleavage Assay; III. A Comparison Of Invasive Cleavage And Primer-Directed Cleavage; IV. Fractionation Of Specific Nucleic Acids By Selective Charge Reversal; V. InvaderTM-Directed Cleavage Using Miniprobes And Mid-Range Probes; VI. Signal Enhancement By Tailing Of Reaction Products In The InvaderTM-Directed Cleavage Assay; VII. Improved Enzymes For Use In InvaderTM-Directed Cleavage Reactions; VIII. Signal Enhancement By Completion Of An Activated Protein Binding Site; IX. Signal Enhancement By Incorporating The Products Of An Invasive Cleavage Reaction Into A Subsequent Invasive Cleavage Reaction; and X. Detection of Human Cytomegalovirus Viral DNA By Invasive Cleavage.
I. Generation of 5′ Nucleases from Thermostable DNA Polymerases
The genes encoding Type A DNA polymerases share about 85% homology to each other on the DNA sequence level. Preferred examples of thermostable polymerases include those isolated from Thermus aquaticus, Thermus flavus, and Thermus thermophilus. However, other thermostable Type A polymerases which have 5′ nuclease activity are also suitable. FIGS. 1 and 2 compare the nucleotide and amino acid sequences of the three above mentioned polymerases. In FIGS. 1 and 2, the consensus or majority sequence derived from a comparison of the nucleotide (FIG. 1) or amino acid (FIG. 2) sequence of the three thermostable DNA polymerases is shown on the top line. A dot appears in the sequences of each of these three polymerases whenever an amino acid residue in a given sequence is identical to that contained in the consensus amino acid sequence. Dashes are used to introduce gaps in order to maximize alignment between the displayed sequences. When no consensus nucleotide or amino acid is present at a given position, an “X” is placed in the consensus sequence. SEQ ID NOS:1-3 display the nucleotide sequences and SEQ ID NOS:4-6 display the amino acid sequences of the three wild-type polymerases. SEQ ID NO:1 corresponds to the nucleic acid sequence of the wild type Thermus aquaticus DNA polymerase gene isolated from the YT-1 strain [Lawyer et al., J. Biol. Chem. 264:6427 (1989)]. SEQ ID NO:2 corresponds to the nucleic acid sequence of the wild type Thermus flavus DNA polymerase gene [Akhmetzjanov and Vakhitov, Nucl. Acids Res. 20:5839 (1992)]. SEQ ID NO:3 corresponds to the nucleic acid sequence of the wild type Thermus thermophilus DNA polymerase gene [Gelfand et al., WO 91/09950 (1991)]. SEQ ID NOS:7-8 depict the consensus nucleotide and amino acid sequences, respectively for the above three DNAPs (also shown on the top row in FIGS. 2 and 3).
The 5′ nucleases of the invention derived from thermostable polymerases have reduced synthetic ability, but retain substantially the same 5′ exonuclease activity as the native DNA polymerase. The term “substantially the same 5′ nuclease activity” as used herein means that the 5′ nuclease activity of the modified enzyme retains the ability to function as a structure-dependent single-stranded endonuclease but not necessarily at the same rate of cleavage as compared to the unmodified enzyme. Type A DNA polymerases may also be modified so as to produce an enzyme which has increases 5′ nuclease activity while having a reduced level of synthetic activity. Modified enzymes having reduced synthetic activity and increased 5′ nuclease activity are also envisioned by the present invention.
By the term “reduced synthetic activity” as used herein it is meant that the modified enzyme has less than the level of synthetic activity found in the unmodified or “native” enzyme. The modified enzyme may have no synthetic activity remaining or may have that level of synthetic activity that will not interfere with the use of the modified enzyme in the detection assay described below. The 5′ nucleases of the present invention are advantageous in situations where the cleavage activity of the polymerase is desired, but the synthetic ability is not (such as in the detection assay of the invention).
As noted above, it is not intended that the invention be limited by the nature of the alteration necessary to render the polymerase synthesis deficient. The present invention contemplates a variety of methods, including but not limited to: 1) proteolysis; 2) recombinant constructs (including mutants); and 3) physical and/or chemical modification and/or inhibition.
1. Proteolysis
Thermostable DNA polymerases having a reduced level of synthetic activity are produced by physically cleaving the unmodified enzyme with proteolytic enzymes to produce fragments of the enzyme that are deficient in synthetic activity but retain 5′ nuclease activity. Following proteolytic digestion, the resulting fragments are separated by standard chromatographic techniques and assayed for the ability to synthesize DNA and to act as a 5′ nuclease. The assays to determine synthetic activity and 5′ nuclease activity are described below.
2. Recombinant Constructs
The examples below describe a preferred method for creating a construct encoding a 5′ nuclease derived from a thermostable DNA polymerase. As the Type A DNA polymerases are similar in DNA sequence, the cloning strategies employed for the Thermus aquaticus and flavus polymerases are applicable to other thermostable Type A polymerases. In general, a thermostable DNA polymerase is cloned by isolating genomic DNA using molecular biological methods from a bacteria containing a thermostable Type A DNA polymerase. This genomic DNA is exposed to primers which are capable of amplifying the polymerase gene by PCR.
This amplified polymerase sequence is then subjected to standard deletion processes to delete the polymerase portion of the gene. Suitable deletion processes are described below in the examples.
The example below discusses the strategy used to determine which portions of the DNAPTaq polymerase domain could be removed without eliminating the 5′ nuclease activity. Deletion of amino acids from the protein can be done either by deletion of the encoding genetic material, or by introduction of a translational stop codon by mutation or frame shift. In addition, proteolytic treatment of the protein molecule can be performed to remove segments of the protein.
In the examples below, specific alterations of the Taq gene were: a deletion between nucleotides 1601 and 2502 (the end of the coding region), a 4 nucleotide insertion at position 2043, and deletions between nucleotides 1614 and 1848 and between nucleotides 875 and 1778 (numbering is as in SEQ ID NO:1). These modified sequences are described below in the examples and at SEQ ID NOS:9-12.
Those skilled in the art understand that single base pair changes can be innocuous in terms of enzyme structure and function. Similarly, small additions and deletions can be present without substantially changing the exonuclease or polymerase function of these enzymes.
Other deletions are also suitable to create the 5′ nucleases of the present invention. It is preferable that the deletion decrease the polymerase activity of the 5′ nucleases to a level at which synthetic activity will not interfere with the use of the 5′ nuclease in the detection assay of the invention. Most preferably, the synthetic ability is absent. Modified polymerases are tested for the presence of synthetic and 5′ nuclease activity as in assays described below. Thoughtful consideration of these assays allows for the screening of candidate enzymes whose structure is heretofore as yet unknown. In other words, construct “X” can be evaluated according to the protocol described below to determine whether it is a member of the genus of 5′ nucleases of the present invention as defined functionally, rather than structurally.
In the example below, the PCR product of the amplified Thermus aquaticus genomic DNA did not have the identical nucleotide structure of the native genomic DNA and did not have the same synthetic ability of the original clone. Base pair changes which result due to the infidelity of DNAPTaq during PCR amplification of a polymerase gene are also a method by which the synthetic ability of a polymerase gene may be inactivated. The examples below and FIGS. 3A and 4A indicate regions in the native Thermus aquaticus and flavus DNA polymerases likely to be important for synthetic ability. There are other base pair changes and substitutions that will likely also inactivate the polymerase.
It is not necessary, however, that one start out the process of producing a 5′ nuclease from a DNA polymerase with such a mutated amplified product. This is the method by which the examples below were performed to generate the synthesis-deficient DNAPTaq mutants, but it is understood by those skilled in the art that a wild-type DNA polymerase sequence may be used as the starting material for the introduction of deletions, insertion and substitutions to produce a 5′ nuclease. For example, to generate the synthesis-deficient DNAPTf1 mutant, the primers listed in SEQ ID NOS:13-14 were used to amplify the wild type DNA polymerase gene from Thermus flavus strain AT-62. The amplified polymerase gene was then subjected to restriction enzyme digestion to delete a large portion of the domain encoding the synthetic activity.
The present invention contemplates that the nucleic acid construct of the present invention be capable of expression in a suitable host. Those in the art know methods for attaching various promoters and 3′ sequences to a gene structure to achieve efficient expression. The examples below disclose two suitable vectors and six suitable vector constructs. Of course, there are other promoter/vector combinations that would be suitable. It is not necessary that a host organism be used for the expression of the nucleic acid constructs of the invention. For example, expression of the protein encoded by a nucleic acid construct may be achieved through the use of a cell-free in vitro transcription/translation system. An example of such a cell-free system is the commercially available TnTTM Coupled Reticulocyte Lysate System (Promega Corporation, Madison, Wis.).
Once a suitable nucleic acid construct has been made, the 5′ nuclease may be produced from the construct. The examples below and standard molecular biological teachings enable one to manipulate the construct by different suitable methods.
Once the 5′ nuclease has been expressed, the polymerase is tested for both synthetic and nuclease activity as described below.
3. Physical And/Or Chemical Modification And/Or Inhibition
The synthetic activity of a thermostable DNA polymerase may be reduced by chemical and/or physical means. In one embodiment, the cleavage reaction catalyzed by the 5′ nuclease activity of the polymerase is run under conditions which preferentially inhibit the synthetic activity of the polymerase. The level of synthetic activity need only be reduced to that level of activity which does not interfere with cleavage reactions requiring no significant synthetic activity.
As shown in the examples below, concentrations of Mg++ greater than 5 mM inhibit the polymerization activity of the native DNAPTaq. The ability of the 5′ nuclease to function under conditions where synthetic activity is inhibited is tested by running the assays for synthetic and 5′ nuclease activity, described below, in the presence of a range of Mg+30  concentrations (5 to 10 mM). The effect of a given concentration of Mg++ is determined by quantitation of the amount of synthesis and cleavage in the test reaction as compared to the standard reaction for each assay.
The inhibitory effect of other ions, polyamines, denaturants, such as urea, formamide, dimethylsulfoxide, glycerol and non-ionic detergents (Triton X-100 and Tween-20), nucleic acid binding chemicals such as, actinomycin D, ethidium bromide and psoralens, are tested by their addition to the standard reaction buffers for the synthesis and 5′ nuclease assays. Those compounds having a preferential inhibitory effect on the synthetic activity of a thermostable polymerase are then used to create reaction conditions under which 5′ nuclease activity (cleavage) is retained while synthetic activity is reduced or eliminated.
Physical means may be used to preferentially inhibit the synthetic activity of a polymerase. For example, the synthetic activity of thermostable polymerases is destroyed by exposure of the polymerase to extreme heat (typically 96 to 100° C.) for extended periods of time (greater than or equal to 20 minutes). While these are minor differences with respect to the specific heat tolerance for each of the enzymes, these are readily determined. Polymerases are treated with heat for various periods of time and the effect of the heat treatment upon the synthetic and 5′ nuclease activities is determined.
II. Detection of Specific Nucleic Acid Sequences Using 5′ Nucleases in an Invader-Directed Cleavage Assay
The present invention provides means for forming a nucleic acid cleavage structure which is dependent upon the presence of a target nucleic acid and cleaving the nucleic acid cleavage structure so as to release distinctive cleavage products. 5′ nuclease activity is used to cleave the target-dependent cleavage structure and the resulting cleavage products are indicative of the presence of specific target nucleic acid sequences in the sample.
The present invention further provides assays in which the target nucleic acid is reused or recycled during multiple rounds of hybridization with oligonucleotide probes and cleavage without the need to use temperature cycling (i.e., for periodic denaturation of target nucleic acid strands) or nucleic acid synthesis (i.e., for the displacement of target nucleic acid strands). Through the interaction of the cleavage means (e.g., a 5′ nuclease) an upstream oligonucleotide, the cleavage means can be made to cleave a downstream oligonucleotide at an internal site in such a way that the resulting fragments of the downstream oligonucleotide dissociate from the target nucleic acid, thereby making that region of the target nucleic acid available for hybridization to another, uncleaved copy of the downstream oligonucleotide.
As illustrated in FIG. 25, the methods of the present invention employ at least a pair of oligonucleotides that interact with a target nucleic acid to form a cleavage structure for a structure-specific nuclease. More specifically, the cleavage structure comprises: i) a target nucleic acid that may be either single-stranded or double-stranded (when a double-stranded target nucleic acid is employed, it may be rendered single stranded, e.g., by heating); ii) a first oligonucleotide, termed the “probe,” which defines a first region of the target nucleic acid sequence by being the complement of that region (regions X and Z of the target as shown in FIG. 25); and iii) a second oligonucleotide, termed the “invader,” the 5′ part of which defines a second region of the same target nucleic acid sequence (regions Y and X in FIG. 25), adjacent to and downstream of the first target region (regions X and Z), and the second part of which overlaps into the region defined by the first oligonucleotide (region X depicts the region of overlap). The resulting structure is diagrammed in FIG. 25.
While not limiting the invention or the instant discussion to any particular mechanism of action, the diagram in FIG. 25 represents the effect on the site of cleavage caused by this type of arrangement of a pair of oligonucleotides. The design of such a pair of oligonucleotides is described below in detail. In FIG. 25, the 3′ ends of the nucleic acids (i.e., the target and the oligonucleotides) are indicated by the use of the arrowheads on the ends of the lines depicting the strands of the nucleic acids (and where space permits, these ends are also labelled “3′”). It is readily appreciated that the two oligonucleotides (the invader and the probe) are arranged in a parallel orientation relative to one another, while the target nucleic acid strand is arranged in an anti-parallel orientation relative to the two oligonucleotides. Further it is clear that the invader oligonucleotide is located upstream of the probe oligonucleotide and that with respect to the target nucleic acid strand, region Z is upstream of region X and region X is upstream of region Y (that is region Y is downstream of region X and region X is downstream of region Z). Regions of complementarity between the opposing strands are indicated by the short vertical lines. While not intended to indicate the precise location of the site(s) of cleavage, the area to which the site of cleavage within the probe oligonucleotide is shifted by the presence of the invader oligonucleotide is indicated by the solid vertical arrowhead. An alternative representation of the target/invader/probe cleavage structure is shown in FIG. 28C. Neither diagram (i.e., FIG. 25 or FIG. 28C) is intended to represent the actual mechanism of action or physical arrangement of the cleavage structure and further it is not intended that the method of the present invention be limited to any particular mechanism of action.
It can be considered that the binding of these oligonucleotides divides the target nucleic acid into three distinct regions: one region that has complementarity to only the probe (shown as “Z”); one region that has complementarity only to the invader (shown as “Y”); and one region that has complementarity to both oligonucleotides (shown as “X”).
Design of these oligonucleotides (i.e., the invader and the probe) is accomplished using practices which are standard in the art. For example, sequences that have self complementarity, such that the resulting oligonucleotides would either fold upon themselves, or hybridize to each other at the expense of binding to the target nucleic acid, are generally avoided.
One consideration in choosing a length for these oligonucleotides is the complexity of the sample containing the target nucleic acid. For example, the human genome is approximately 3×109 basepairs in length. Any 10 nucleotide sequence will appear with a frequency of 1:410, or 1:1048,576 in a random string of nucleotides, which would be approximately 2,861 times in 3 billion basepairs. Clearly an oligonucleotide of this length would have a poor chance of binding uniquely to a 10 nucleotide region within a target having a sequence the size of the human genome. If the target sequence were within a 3 kb plasmid, however, such an oligonucleotide might have a very reasonable chance of binding uniquely. By this same calculation it can be seen that an oligonucleotide of 16 nucleotides (i.e., a 16-mer) is the minimum length of a sequence which is mathematically likely to appear once in 3×109 basepairs.
A second consideration in choosing oligonucleotide length is the temperature range in which the oligonucleotides will be expected to function. A 16-mer of average base content (50% G-C basepairs) will have a calculated Tm (the temperature at which 50% of the sequence is dissociated) of about 41° C., depending on, among other things, the concentration of the oligonucleotide and its target, the salt content of the reaction and the precise order of the nucleotides. As a practical matter, longer oligonucleotides are usually chosen to enhance the specificity of hybridization. Oligonucleotides 20 to 25 nucleotides in length are often used as they are highly likely to be specific if used in reactions conducted at temperatures which are near their Tms (within about 5° of the Tm). In addition, with calculated Tms in the range of 50° to 70° C., such oligonucleotides (i.e, 20 to 25-mers) are appropriately used in reactions catalyzed by thermostable enzymes, which often display optimal activity near this temperature range.
The maximum length of the oligonucleotide chosen is also based on the desired specificity. One must avoid choosing sequences that are so long that they are either at a high risk of binding stably to partial complements, or that they cannot easily be dislodged when desired (e.g., failure to disassociate from the target once cleavage has occurred).
The first step of design and selection of the oligonucleotides for the invader-directed cleavage is in accordance with these sample general principles. Considered as sequence-specific probes individually, each oligonucleotide may be selected according to the guidelines listed above. That is to say, each oligonucleotide will generally be long enough to be reasonably expected to hybridize only to the intended target sequence within a complex sample, usually in the 20 to 40 nucleotide range. Alternatively, because the invader-directed cleavage assay depends upon the concerted action of these oligonucleotides, the composite length of the 2 oligonucleotides which span/bind to the X, Y, Z regions may be selected to fall within this range, with each of the individual oligonucleotides being in approximately the 13 to 17 nucleotide range. Such a design might be employed if a non-thermostable cleavage means were employed in the reaction, requiring the reactions to be conducted at a lower temperature than that used when thermostable cleavage means are employed. In some instances, it may be desirable to have these oligonucleotides bind multiple times within a target nucleic acid (e.g., which bind to multiple variants or multiple similar sequences within a target). It is not intended that the method of the present invention be limited to any particular size of the probe or invader oligonucleotide.
The second step of designing an oligonucleotide pair for this assay is to choose the degree to which the upstream “invader” oligonucleotide sequence will overlap into the downstream “probe” oligonucleotide sequence, and consequently, the sizes into which the probe will be cleaved. A key feature of this assay is that the probe oligonucleotide can be made to “turn over,” that is to say cleaved probe can be made to depart to allow the binding and cleavage of other copies of the probe molecule, without the requirements of thermal denaturation or displacement by polymerization. While in one embodiment of this assay probe turnover may be facilitated by an exonucleolytic digestion by the cleavage agent, it is central to the present invention that the turnover does not require this exonucleolytic activity.
Choosing The Amount Of Overlap (Length Of The X Region)
One way of accomplishing such turnover can be envisioned by considering the diagram in FIG. 25. It can be seen that the Tm of each oligonucleotide will be a function of the full length of that oligonucleotide: i.e., the Tm of the invader=Tm(Y+X), and the Tm of the probe=Tm(X+Y) for the probe. When the probe is cleaved the X region is released, leaving the Z section. If the Tm of Z is less than the reaction temperature, and the reaction temperature is less than the Tm(X+Z), then cleavage of the probe will lead to the departure of Z, thus allowing a new (X+Z) to hybridize. It can be seen from this example that the X region must be sufficiently long that the release of X will drop the Tm of the remaining probe section below the reaction temperature: a G-C rich X section may be much shorter than an A-T rich X section and still accomplish this stability shift.
Designing Oligonucleotides Which Interact With The Y And Z Regions
If the binding of the invader oligonucleotide to the target is more stable than the binding of the probe (e.g., if it is long, or is rich in G-C basepairs in the Y region), then the copy of X associated with the invader may be favored in the competition for binding to the X region of the target, and the probe may consequently hybridize inefficiently, and the assay may give low signal. Alternatively, if the probe binding is particularly strong in the Z region, the invader will still cause internal cleavage, because this is mediated by the enzyme, but portion of the probe oligonucleotide bound to the Z region may not dissociate at the reaction temperature, turnover may be poor, and the assay may again give low signal.
It is clearly beneficial for the portions of the oligonucleotide which interact with the Y and Z regions so be similar in stability, i.e., they must have similar melting temperatures. This is not to say that these regions must be the same length. As noted above, in addition to length, the melting temperature will also be affected by the base content and the specific sequence of those bases. The specific stability designed into the invader and probe sequences will depend on the temperature at which one desires to perform the reaction.
This discussion is intended to illustrate that (within the basic guidelines for oligonucleotide specificity discussed above) it is the balance achieved between the stabilities of the probe and invader sequences and their X and Y component sequences, rather than the absolute values of these stabilities, that is the chief consideration in the selection of the probe and invader sequences.
Design Of The Reaction Conditions
Target nucleic acids that may be analyzed using the methods of the present invention which employ a 5′ nuclease as the cleavage means include many types of both RNA and DNA. Such nucleic acids may be obtained using standard molecular biological techniques. For example, nucleic acids (RNA or DNA) may be isolated from a tissue sample (e.g, a biopsy specimen), tissue culture cells, samples containing bacteria and/or viruses (including cultures of bacteria and/or viruses), etc. The target nucleic acid may also be transcribed in vitro from a DNA template or may be chemically synthesized or generated in a PCR. Furthermore, nucleic acids may be isolated from an organism, either as genomic material or as a plasmid or similar extrachromosomal DNA, or they may be a fragment of such material generated by treatment with a restriction endonuclease or other cleavage agents or it may be synthetic.
Assembly of the target, probe, and invader nucleic acids into the cleavage reaction of the present invention uses principles commonly used in the design of oligonucleotide base enzymatic assays, such as dideoxynucleotide sequencing and polymerase chain reaction (PCR). As is done in these assays, the oligonucleotides are provided in sufficient excess that the rate of hybridization to the target nucleic acid is very rapid. These assays are commonly performed with 50 fmoles to 2 pmoles of each oligonucleotide per μl of reaction mixture. In the Examples described herein, amounts of oligonucleotides ranging from 250 fmoles to 5 pmoles per μl of reaction volume were used. These values were chosen for the purpose of case in demonstration and are not intended to limit the performance of the present invention to these concentrations. Other (e.g., lower) oligonucleotide concentrations commonly used in other molecular biological reactions are also contemplated.
It is desirable that an invader oligonucleotide be immediately available to direct the cleavage of each probe oligonucleotide that hybridizes to a target nucleic acid. For this reason, in the Examples described herein, the invader oligonucleotide is provided in excess over the probe oligonucleotide; often this excess is 10-fold. While this is an effective ratio, it is not intended that the practice of the present invention be limited to any particular ratio of invader-to-probe (a ratio of 2- to 100-fold is contemplated).
Buffer conditions must be chosen that will be compatible with both the oligonucleotide/target hybridization and with the activity of the cleavage agent. The optimal buffer conditions for nucleic acid modification enzymes, and particularly DNA modification enzymes, generally included enough mono- and di-valent salts to allow association of nucleic acid strands by base-pairing. If the method of the present invention is performed using an enzymatic cleavage agent other than those specifically described here, the reactions may generally be performed in any such buffer reported to be optimal for the nuclease function of the cleavage agent. In general, to test the utility of any cleavage agent in this method, test reactions are performed wherein the cleavage agent of interest is tested in the MOPS/MnCl2/KCl buffer or Mg-containing buffers described herein and in whatever buffer has been reported to be suitable for use with that agent, in a manufacturer's data sheet, a journal article, or in personal communication.
The products of the invader-directed cleavage reaction are fragments generated by structure-specific cleavage of the input oligonucleotides. The resulting cleaved and/or uncleaved oligonucleotides may be analyzed and resolved by a number of methods including electrophoresis (on a variety of supports including acrylamide or agarose gels, paper, etc.), chromatography, fluorescence polarization, mass spectrometry and chip hybridization. The invention is illustrated using electrophoretic separation for the analysis of the products of the cleavage reactions. However, it is noted that the resolution of the cleavage products is not limited to electrophoresis. Electrophoresis is chosen to illustrate the method of the invention because electrophoresis is widely practiced in the art and is easily accessible to the average practitioner.
The probe and invader oligonucleotides may contain a label to aid in their detection following the cleavage reaction. The label may be a radioisotope (e.g., a 32P or 35S-labelled nucleotide) placed at either the 5′ or 3′ end of the oligonucleotide or alternatively, the label may be distributed throughout the oligonucleotide (i.e., a uniformly labelled oligonucleotide). The label may be a nonisotopic detectable moiety, such as a fluorophore, which can be detected directly,or a reactive group which permits specific recognition by a secondary agent. For example, biotinylated oligonucleotides may be detected by probing with a streptavidin molecule which is coupled to an indicator (e.g., alkaline phosphatase or a fluorophore) or a hapten such as digoxigenin may be detected using a specific antibody coupled to a similar indicator.
Optimization Of Reaction Conditions
The invader-directed cleavage reaction is useful to detect the presence of specific nucleic acids. In addition to the considerations listed above for the selection and design of the invader and probe oligonucleotides, the conditions under which the reaction is to be performed may be optimized for detection of a specific target sequence.
One objective in optimizing the invader-directed cleavage assay is to allow specific detection of the fewest copies of a target nucleic acid. To achieve this end, it is desirable that the combined elements of the reaction interact with the maximum efficiency, so that the rate of the reaction (e.g., the number of cleavage events per minute) is maximized. Elements contributing to the overall efficiency of the reaction include the rate of hybridization, the rate of cleavage, and the efficiency of the release of the cleaved probe.
The rate of cleavage will be a function of the cleavage means chosen, and may be made optimal according to the manufacturer's instructions when using commercial preparations of enzymes or as described in the examples herein. The other elements (rate of hybridization, efficiency of release) depend upon the execution of the reaction, and optimization of these elements is discussed below.
Three elements of the cleavage reaction that significantly affect the rate of nucleic acid hybridization are the concentration of the nucleic acids, the temperature at which the cleavage reaction is performed and the concentration of salts and/or other charge-shielding ions in the reaction solution.
The concentrations at which oligonucleotide probes are used in assays of this type are well known in the art, and are discussed above. One example of a common approach to optimizing an oligonucleotide concentration is to choose a starting amount of oligonucleotide for pilot tests; 0.01 to 2 μM is a concentration range used in many oligonucleotide-based assays. When initial cleavage reactions are performed, the following questions may be asked of the data: Is the reaction performed in the absence of the target nucleic acid substantially free of the cleavage product?; Is the site of cleavage specifically shifted in accordance with the design of the invader oligonucleotide?; Is the specific cleavage product easily detected in the presence of the uncleaved probe (or is the amount of uncut material overwhelming the chosen visualization method)?
A negative answer to any of these questions would suggest that the probe concentration is too high, and that a set of reactions using serial dilutions of the probe should be performed until the appropriate amount is identified. Once identified for a given target nucleic acid in a give sample type (e.g., purified genomic DNA, body fluid extract, lysed bacterial extract), it should not need to be re-optimized. The sample type is important because the complexity of the material present may influence the probe optimum.
Conversely, if the chosen initial probe concentration is too low, the reaction may be slow, due to inefficient hybridization. Tests with increasing quantities of the probe will identify the point at which the concentration exceeds the optimum. Since the hybridization will be facilitated by excess of probe, it is desirable, but not required, that the reaction be performed using probe concentrations just below this point.
The concentration of invader oligonucleotide can be chosen based on the design considerations discussed above. In a preferred embodiment, the invader oligonucleotide is in excess of the probe oligonucleotide. In a particularly preferred embodiment, the invader is approximately 10-fold more abundant than the probe.
Temperature is also an important factor in the hybridization of oligonucleotides. The range of temperature tested will depend in large part, on the design of the oligonucleotides, as discussed above. In a preferred embodiment, the reactions are performed at temperatures slightly below the Tm of the least stable oligonucleotide in the reaction. Melting temperatures for the oligonucleotides and for their component regions (X, Y and Z, FIG. 25), can be estimated through the use of computer software or, for a more rough approximation, by assigning the value of 2° C. per A-T basepair, and 4° C. per G-C basepair, and taking the sum across an expanse of nucleic acid. The latter method may be used for oligonucleotides of approximately 10-30 nucleotides in length. Because even computer prediction of the Tm of a nucleic acid is only an approximation, the reaction temperatures chosen for initial tests should bracket the calculated Tm. While optimizations are not limited to this, 5° C. increments are convenient test intervals in these optimization assays.
When temperatures are tested, the results can be analyzed for specificity (the first two of the questions listed above) in the same way as for the oligonucleotide concentration determinations. Non-specific cleavage (i.e., cleavage of the probe at many or all positions along its length) would indicate non-specific interactions between the probe and the sample material, and would suggest that a higher temperature should be employed. Conversely, little or no cleavage would suggest that even the intended hybridization is being prevented, and would suggest the use of lower temperatures. By testing several temperatures, it is possible to identify an approximate temperature optimum, at which the rate of specific cleavage of the probe is highest. If the oligonucleotides have been designed as described above, the Tm of the Z-region of the probe oligonucleotide should be below this temperature, so that turnover is assured.
A third determinant of hybridization efficiency is the salt concentration of the reaction. In large part, the choice of solution conditions will depend on the requirements of the cleavage agent, and for reagents obtained commercially, the manufacturer's instructions are a resource for this information. When developing an assay utilizing any particular cleavage agent, the oligonucleotide and temperature optimizations described above should be performed in the buffer conditions best suited to that cleavage agent.
A “no enzyme” control allows the assessment of the stability of the labeled oligonucleotides under particular reaction conditions, or in the presence of the sample to be tested (i.e., in assessing the sample for contaminating nucleases). In this manner, the substrate and oligonucleotides are placed in a tube containing all reaction components, except the enzyme and treated the same as the enzyme-containing reactions. Other controls may also be included. For example, a reaction with all of the components except the target nucleic acid will serve to confirm the dependence of the cleavage on the presence of the target sequence.
Probing For Multiple Alleles
The invader-directed cleavage reaction is also useful in the detection and quantification of individual variants or alleles in a mixed sample population. By way of example, such a need exists in the analysis of tumor material for mutations in genes associated with cancers. Biopsy material from a tumor can have a significant complement of normal cells, so it is desirable to detect mutations even when present in fewer than 5% of the copies of the target nucleic acid in a sample. In this case, it is also desirable to measure what fraction of the population carries the mutation. Similar analyses may also be done to examine allelic variation in other gene systems, and it is not intended that the method of the present invention by limited to the analysis of tumors.
As demonstrated below, reactions can be performed under conditions that prevent the cleavage of probes bearing even a single-nucleotide difference mismatch within the region of the target nucleic acid termed “Z” in FIG. 25, but that permit cleavage of a similar probe that is completely complementary to the target in this region. Thus, the assay may be used to quantitate individual variants or alleles within a mixed sample.
The use of multiple, differently labelled probes in such an assay is also contemplated. To assess the representation of different variants or alleles in a sample, one would provide a mixture of probes such that each allele or variant to be detected would have a specific probe (i.e., perfectly matched to the Z region of the target sequence) with a unique label (e.g., no two variant probes with the same label would be used in a single reaction). These probes would be characterized in advance to ensure that under a single set of reaction conditions, they could be made to give the same rate of signal accumulation when mixed with their respective target nucleic acids. Assembly of a cleavage reaction comprising the mixed probe set, a corresponding invader oligonucleotide, the target nucleic acid sample, and the appropriate cleavage agent, along with performance of the cleavage reaction under conditions such that only the matched probes would cleave, would allow independent quantification of each of the species present, and would therefore indicate their relative representation in the target sample.
III. A Comparison of Invasive Cleavage and Primer-Directed Cleavage
As discussed herein, the terms “invasive” or “invader-directed” cleavage specifically denote the use of a first, upstream oligonucleotide, as defined below, to cause specific cleavage at a site within a second, downstream sequence. To effect such a direction of cleavage to a region within a duplex, it is required that the first and second oligonucleotides overlap in sequence. That is to say, a portion of the upstream oligonucleotide, termed the “invader”, has significant homology to a portion of the downstream “probe” oligonucleotide, so that these regions would tend to basepair with the same complementary region of the target nucleic acid to be detected. While not limiting the present invention to any particular mechanism, the overlapping regions would be expected to alternate in their occupation of the shared hybridization site. When the probe oligonucleotide fully anneals to the target nucleic acid, and thus forces the 3′ region of the invader to remain unpaired, the structure so formed is not a substrate for the 5′ nucleases of the present invention. By contrast, when the inverse is true, the structure so formed is substrate for these enzymes, allowing cleavage and release of the portion of the probe oligonucleotide that is displaced by the invader oligonucleotide. The shifting of the cleavage site to a region the probe oligonucleotide that would otherwise be basepaired to the target sequence is one hallmark of the invasive cleavage assay (i.e., the invader-directed cleavage assay) of the present invention.
It is beneficial at this point to contrast the invasive cleavage as described above with two other forms of probe cleavage that may lead to internal cleavage of a probe oligonucleotide, but which do not comprise invasive cleavage. In the first case, a hybridized probe may be subject to duplex-dependent 5′ to 3′ exonuclease “nibbling,” such that the oligonucleotide is shortened from the 5′ end until it cannot remain bound to the target (see, e.g., Examples 5-7 and FIGS. 26-28). The site at which such nibbling stops can appear to be discrete, and, depending on the difference between the melting temperature of the full-length probe and the temperature of the reaction, this stopping point may be 1 or several nucleotides into the probe oligonucleotide sequence. Such “nibbling” is often indicated by the presence of a “ladder” of longer products ascending size up to that of the full length of the probe, but this is not always the case. While any one of the products of such a nibbling reaction may be made to match in size and cleavage site the products of an invasive cleavage reaction, the creation of these nibbling products would be highly dependent on the temperature of the reaction and the nature of the cleavage agent, but would be independent of the action of an upstream oligonucleotide, and thus could not be construed to involve invasive cleavage.
A second cleavage structure that may be considered is one in which a probe oligonucleotide has several regions of complementarity with the target nucleic acid, interspersed with one or more regions or nucleotides of noncomplementarity. These noncomplementary regions may be thought of as “bubbles” within the nucleic acid duplex. As temperature is elevated, the regions of complementarity can be expected to “melt” in the order of their stability, lowest to highest. When a region of lower stability is near the end of a segment of duplex, and the next region of complementarity along the strand has a higher melting temperature, a temperature can be found that will cause the terminal region of duplex to melt first, opening the first bubble, and thereby creating a preferred substrate structure of the cleavage by the 5′ nucleases of the present invention (FIG. 36A). The site of such cleavage would be expected to be on the 5′ arm, within 2 nucleotides of the junction between the single and double-stranded regions (Lyamichev et al., supra. and U.S. Pat. No. 5,422,253)
An additional oligonucleotide could be introduced to basepair along the target nucleic acid would have a similar effect of opening this bubble for subsequent cleavage of the unpaired 5′ arm (FIG. 36B and FIG. 6). Note in this case, the 3′ terminal nucleotides of the upstream oligonucleotide anneals along the target nucleic acid sequence in such a manner that the 3′ end is located within the “bubble” region. Depending on the precise location of the 3′ end of this oligonucleotide, the cleavage site may be along the newly unpaired 5′ arm, or at the site expected for the thermally opened bubble structure as described above. In the former case the cleavage is not within a duplexed region, and is thus not invasive cleavage, while in the latter the oligonucleotide is merely an aide in inducing cleavage at a site that might otherwise be exposed through the use of temperature alone (i.e., in the absence of the additional oligonucleotide), and is thus not considered to be invasive cleavage.
In summary, any arrangement of oligonucleotides used for the cleavage-based detection of a target sequence can be analyzed to determine if the arrangement is an invasive cleavage structure as contemplated herein. An invasive cleavage structure supports cleavage of the probe in a region that, in the absence of an upstream oligonucleotide, would be expected to be basepaired to the target nucleic acid.
Example 26 below provides further guidance for the design and execution of a experiments which allow the determination of whether a given arrangement of a pair of upstream and downstream (i.e., the probe) oligonucleotides when annealed along a target nucleic acid would form an invasive cleavage structure.
IV. Fractionation of Specific Nucleic Acids by Selective Charge Reversal
Some nucleic acid-based detection assays involve the elongation and/or shortening of oligonucleotide probes. For example, as described herein, the primer-directed, primer-independent, and invader-directed cleavage assays, as well as the “nibbling” assay all involve the cleavage (i.e., shortening) of oligonucleotides as a means for detecting the presence of a target nucleic sequence. Examples of other detection assays which involve the shortening of an oligonucleotide probe include the “TaqMan” or nick-translation PCR assay described in U.S. Pat. No. 5,210,015 to Gelfand et al. (the disclosure of which is herein incorporated by reference), the assays described in U.S. Pat. Nos. 4,775,619 and 5,118,605 to Urdea (the disclosures of which are herein incorporated by reference), the catalytic hybridization amplification assay described in U.S. Pat. No. 5,403,711 to Walder and Walder (the disclosure of which is herein incorporated by reference), and the cycling probe assay described in U.S. Pat. Nos. 4,876,187 and 5,011,769 to Duck et al. (the disclosures of which are herein incorporated by reference). Examples of detection assays which involve the elongation of an oligonucleotide probe (or primer) include the polymerase chain reaction (PCR) described in U.S. Pat. Nos. 4,683,195 and 4,683,202 to Mullis and Mullis et al. (the disclosures of which are herein incorporated by reference) and the ligase chain reaction (LCR) described in U.S. Pat. Nos. 5,427,930 and 5,494,810 to Birkenmeyer et al. and Barany et al (the disclosures of which are herein incorporated by reference). The above examples are intended to be illustrative of nucleic acid-based detection assays that involve the elongation and/or shortening of oligonucleotide probes and do not provide an exhaustive list.
Typically, nucleic acid-based detection assays that involve the elongation and/or shortening of oligonucleotide probes require post-reaction analysis to detect the products of the reaction. It is common that, the specific reaction product(s) must be separated from the other reaction components, including the input or unreacted oligonucleotide probe. One detection technique involves the electrophoretic separation of the reacted and unreacted oligonucleotide probe. When the assay involves the cleavage or shortening of the probe, the unreacted product will be longer than the reacted or cleaved product. When the assay involves the elongation of the probe (or primer), the reaction products will be greater in length than the input. Gel-based electrophoresis of a sample containing nucleic acid molecules of different lengths separates these fragments primarily on the basis of size. This is due to the fact that in solutions having a neutral or alkaline pH, nucleic acids having widely different sizes (i.e., molecular weights) possess very similar charge-to-mass ratios and do not separate [Andrews, Electrophoresis, 2nd Edition, Oxford University Press (1986), pp. 153-154]. The gel matrix acts as a molecular sieve and allows nucleic acids to be separated on the basis of size and shape (e.g., linear, relaxed circular or covalently closed supercoiled circles).
Unmodified nucleic acids have a net negative charge due to the presence of negatively charged phosphate groups contained within the sugar-phosphate backbone of the nucleic acid. Typically, the sample is applied to gel near the negative pole and the nucleic acid fragments migrate into the gel toward the positive pole with the smallest fragments moving fastest through the gel.
The present invention provides a novel means for fractionating nucleic acid fragments on the basis of charge. This novel separation technique is related to the observation that positively charged adducts can affect the electrophoretic behavior of small oligonucleotides because the charge of the adduct is significant relative to charge of the whole complex. In addition, to the use of positively charged adducts (e.g., Cy3 and Cy5 fluorescent dyes, the positively charged heterodimeric DNA-binding dyes shown in FIG. 66, etc.), the oligonucleotide may contain amino acids (particularly useful amino acids are the charged amino acids: lysine, arginine, asparate, glutamate), modified bases, such as amino-modified bases, and/or a phosphonate backbone (at all or a subset of the positions). In addition as discussed further below, a neutral dye or detection moiety (e.g., biotin, streptavidin, etc.) may be employed in place of a positively charged adduct in conjunction with the use of amino-modified bases and/or a complete or partial phosphonate backbone.
This observed effect is of particular utility in assays based on the cleavage of DNA molecules. Using the assays described herein as an example, when an oligonucleotide is shortened through the action of a Cleavase® enzyme or other cleavage agent, the positive charge can be made to not only significantly reduce the net negative charge, but to actually override it, effectively “flipping” the net charge of the labeled entity. This reversal of charge allows the products of target-specific cleavage to be partitioned from uncleaved probe by extremely simple means. For example, the products of cleavage can be made to migrate towards a negative electrode placed at any point in a reaction vessel, for focused detection without gel-based electrophoresis; Example 24 provides examples of devices suitable for focused detection without gel-based electrophoresis. When a slab gel is used, sample wells can be positioned in the center of the gel, so that the cleaved and uncleaved probes can be observed to migrate in opposite directions. Alternatively, a traditional vertical gel can be used, but with the electrodes reversed relative to usual DNA gels (i.e., the positive electrode at the top and the negative electrode at the bottom) so that the cleaved molecules enter the gel, while the uncleaved disperse into the upper reservoir of electrophoresis buffer.
An important benefit of this type of readout is the absolute nature of the partition of products from substrates, i.e., the separation is virtually 100%. This means that an abundance of uncleaved probe can be supplied to drive the hybridization step of the probe-based assay, yet the unconsumed (i.e., unreacted) probe can, in essence, be subtracted from the result to reduce background by virtue of the fact that the unreacted probe will not migrate to the same pole as the specific reaction product.
Through the use of multiple positively charged adducts, synthetic molecules can be constructed with sufficient modification that the normally negatively charged strand is made nearly neutral. When so constructed, the presence or absence of a single phosphate group can mean the difference between a net negative or a net positive charge. This observation has particular utility when one objective is to discriminate between enzymatically generated fragments of DNA, which lack a 3 phosphate, and the products of thermal degradation, which retain a 3 phosphate (and thus two additional negative charges). Examples 22 and 23 demonstrate the ability to separate positively charged reaction products from a net negatively charged substrate oligonucleotide. As discussed in these examples, oligonucleotides may be transformed from net negative to net positively charged compounds. In Example 23, the positively charged dye, Cy3 was incorporated at the 5′ end of a 22-mer (SEQ ID NO:50) which also contained two amino-substituted residues at the 5′ end of the oligonucleotide; this oligonucleotide probe carries a net negative charge. After cleavage, which occurred 2 nucleotides into the probe, the following labelled oligonucleotide was released: 5′-Cy3-AminoT-AminoT-3′ (as well as the remaining 20 nucleotides of SEQ ID NO:50). This short fragment bears a net positive charge while the remainder of the cleaved oligonucleotide and the unreacted or input oligonucleotide bear net negative charges.
The present invention contemplates embodiments wherein the specific reaction product produced by any cleavage of any oligonucleotide can be designed to carry a net positive charge while the unreacted probe is charge neutral or carries a net negative charge. The present invention also contemplates embodiments where the released product may be designed to carry a net negative charge while the input nucleic acid carries a net positive charge. Depending on the length of the released product to be detected, positively charged dyes may be incorporated at the one end of the probe and modified bases may be placed along the oligonucleotide such that upon cleavage, the released fragment containing the positively charged dye carries a net positive charge. Amino-modified bases may be used to balance the charge of the released fragment in cases where the presence of the positively charged adduct (e.g., dye) alone is not sufficient to impart a net positive charge on the released fragment. In addition, the phosphate backbone may be replaced with a phosphonate backbone at a level sufficient to impart a net positive charge (this is particularly useful when the sequence of the oligonucleotide is not amenable to the use of amino-substituted bases); FIGS. 45 and 46 show the structure of short oligonucleotides containing a phosphonate group on the second T residue). An oligonucleotide containing a fully phosphonate-substituted backbone would be charge neutral (absent the presence of modified charged residues bearing a charge or the presence of a charged adduct) due to the absence of the negatively charged phosphate groups. Phosphonate-containing nucleotides (e.g., methylphosphonate-containing nucleotides are readily available and can be incorporated at any position of an oligonucleotide during synthesis using techniques which are well known in the art.
In essence, the invention contemplates the use of charge-based separation to permit the separation of specific reaction products from the input oligonucleotides in nucleic acid-based detection assays. The foundation of this novel separation technique is the design and use of oligonucleotide probes (typically termed “primers” in the case of PCR) which are “charge balanced” so that upon either cleavage or elongation of the probe it becomes “charge unbalanced,” and the specific reaction products may be separated from the input reactants on the basis of the net charge.
In the context of assays which involve the elongation of an oligonucleotide probe (i.e., a primer), such as is the case in PCR, the input primers are designed to carry a net positive charge. Elongation of the short oligonucleotide primer during polymerization will generate PCR products which now carry a net negative charge. The specific reaction products may then easily be separated and concentrated away from the input primers using the charge-based separation technique described herein (the electrodes will be reversed relative to the description in Example 23 as the product to be separated and concentrated after a PCR will carry a negative charge).
V. InvaderTM-Directed Cleavage Using Miniprobes and Mid-Range Probes
As discussed in section III above, the InvaderTM-directed cleavage assay may be performed using invader and probe oligonucleotides which have a length of about 13-25 nucleotides (typically 20-25 nucleotides). It is also contemplated that the oligonucleotides that span the X, Y and Z regions (see FIG. 25), the invader and probe oligonucleotides, may themselves be composed of shorter oligonucleotide sequences that align along a target strand but that are not covalently linked. This is to say that there is a nick in the sugar-phosphate backbone of the composite oligonucleotide, but that there is no disruption in the progression of base-paired nucleotides in the resulting duplex. When short strands of nucleic acid align contiguously along a longer strand the hybridization of each is stabilized by the hybridization of the neighboring fragments because the basepairs can stack along the helix as though the backbone was in fact uninterrupted. This cooperativity of binding can give each segment a stability of interaction in excess of what would be expected for the segment hybridizing to the longer nucleic acid alone. One application of this observation has been to assemble primers for DNA sequencing, typically about 18 nucleotides long, from sets of three hexamer oligonucleotides that are designed to hybridize in this way [Kotler, L. E., et al. (1993) Proc. Natl. Acad. Sci. USA 90:4241]. The resulting doubly-nicked primer can be extended enzymatically in reactions performed at temperatures that might be expected to disrupt the hybridization of hexamers, but not of 18-mers.
The use of composite or split oligonuceotides is applied with success in the InvaderTM-directed cleavage assay. The probe oligonucleotide may be split into two oligonucleotides which anneal in a contiguous and adjacent manner along a target oligonucleotide as diagrammed in FIG. 57. In this figure, the downstream oligonucleotide (analogous to the probe of FIG. 25) is assembled from two smaller pieces: a short segment of 6-10 nts (termed the “miniprobe”), that is to be cleaved in the course of the detection reaction, and an oligonucleotide that hybridizes immediately downstream of the miniprobe (termed the “stacker”), which serves to stabilize the hybridization of the probe. To form the cleavage structure, an upstream oligonucleotide (the “InvaderTM” oligo) is provided to direct the cleavage activity to the desired region of the miniprobe. Assembly of the probe from non-linked pieces of nucleic acid (i.e., the miniprobe and the stacker) allows regions of sequences to be changed without requiring the re-synthesis of the entire proven sequence, thus improving the cost and flexibility of the detection system. In addition, the use of unlinked composite oligonucleotides makes the system more stringent in its requirement of perfectly matched hybridization to achieve signal generation, allowing this to be used as a sensitive means of detecting mutations or changes in the target nucleic acid sequences.
As illustrated in FIG. 57, in one embodiment, the methods of the present invention employ at least three oligonucleotides that interact with a target nucleic acid to form a cleavage structure for a structure-specific nuclease. More specifically, the cleavage structure comprises i) a target nucleic acid that may be either single-stranded or double-stranded (when a double-stranded target nucleic acid is employed, it may be rendered single-stranded, e.g., by heating); ii) a first oligonucleotide, termed the “stacker,” which defines a first region of the target nucleic acid sequence by being the complement of that region (region W of the target as shown in FIG. 57); iii) a second oligonucleotide, termed the “miniprobe,” which defines a second region of the target nucleic acid sequence by being the complement of that region (regions X and Z of the target as shown in FIG. 57); iv) a third oligonucleotide, termed the “invader,” the 5′ part of which defines a third region of the same target nucleic acid sequence (regions Y and X in FIG. 57), adjacent to and downstream of the second target region (regions X and Z), and the second or 3′ part of which overlaps into the region defined by the second oligonucleotide (region X depicts the region of overlap). The resulting structure is diagrammed in FIG. 57.
While not limiting the invention or the instant discussion to any particular mechanism of action, the diagram in FIG. 57 represents the effect on the site of cleavage caused by this type of arrangement of three oligonucleotides. The design of these three oligonucleotides is described below in detail. In FIG. 57, the 3′ ends of the nucleic acids (i.e., the target and the oligonucleotides) are indicated by the use of the arrowheads on the ends of the lines depicting the strands of the nucleic acids (and where space permits, these ends are also labelled “3′”). It is readily appreciated that the three oligonucleotides (the invader, the miniprobe and the stacker) are arranged in a parallel orientation relative to one another, while the target nucleic acid strand is arranged in an anti-parallel orientation relative to the three oligonucleotides. Further it is clear that the invader oligonucleotide is located upstream of the miniprobe oligonucleotide and that the miniprobe olignuceotide is located upstream of the stacker oligonucleotide and that with respect to the target nucleic acid strand, region W is upstream of region Z, region Z is upstream of upstream of region X and region X is upstream of region Y (that is region Y is downstream of region X, region X is downstream of region Z and region Z is downstream of region W). Regions of complementarity between the opposing strands are indicated by the short vertical lines. While not intended to indicate the precise location of the site(s) of cleavage, the area to which the site of cleavage within the miniprobe oligonucleotide is shifted by the presence of the invader oligonucleotide is indicated by the solid vertical arrowhead. FIG. 57 is not intended to represent the actual mechanism of action or physical arrangement of the cleavage structure and further it is not intended that the method of the present invention be limited to any particular mechanism of action.
It can be considered that the binding of these oligonucleotides divides the target nucleic acid into four distinct regions: one region that has complementarity to only the stacker (shown as “W”); one region that has complementarity to only the miniprobe (shown as “Z”); one region that has complementarity only to the InvaderTM oligo (shown as “Y”); and one region that has complementarity to both the InvaderTM and miniprobe oligonucleotides (shown as “X”).
In addition to the benefits cited above, the use of a composite design for the oligonucleotides which form the cleavage structure allows more latitude in the design of the reaction conditions for performing the InvaderTM-directed cleavage assay. When a longer probe (e.g., 16-25 nt), as described in section III above, is used for detection in reactions that are performed at temperatures below the Tm of that probe, the cleavage of the probe may play a significant role in destabilizing the duplex of which it is a part, thus allowing turnover and reuse of the recognition site on the target nucleic acid. In contrast, with miniprobes, reaction temperatures that are at or above the Tm of the probe mean that the probe molecules are hybridizing and releasing from the target quite rapidly even without cleavage of the probe. When an upstream InvaderTM oligonucleotide and a cleavage means are provided the miniprobe will be specifically cleaved, but the cleavage will not be necessary to the turnover of the miniprobe. If a long probe (e.g., 16-25 nt) were to be used in this way the temperatures required to achieve this state would be quite high, around 65 to 70° C. for a 25-mer of average base composition. Requiring the use of such elevated temperatures limits the choice of cleavage agents to those that are very thermostable, and may contribute to background in the reactions, depending of the means of detection, through thermal degradation of the probe oligonucleotides. Thus, the shorter probes are preferable for use in this way.
The miniprobe of the present invention may vary in size depending on the desired application. In one embodiment, the probe may be relatively short compared to a standard probe (e.g., 16-25 nt), in the range of 6 to 10 nucleotides. When such a short probe is used reaction conditions can be chosen that prevent hybridization of the miniprobe in the absence of the stacker oligonucleotide. In this way a short probe can be made to assume the statistical specificity and selectivity of a longer sequence. In the event of a perturbation in the cooperative binding of the miniprobe and stacker nucleic acids, as might be caused by a mismatch within the short sequence (i.e., region “Z” which is the region of the miniprobe which does not overlap with the invader) or at the junction between the contiguous duplexes, this cooperativity can be lost, dramatically reducing the stability of the shorter oligonucleotide (i.e., the miniprobe), and thus reducing the level of cleaved product in the assay of the present invention.
It is also contemplated that probes of intermediate size may be used. Such probes, in the 11 to 15 nucleotide range, may blend some of the features associated with the longer probes as originally described, these features including the ability to hybridize and be cleaved absent the help of a stacker oligonucleotide. At temperatures below the expected Tm of such probes, the mechanisms of turnover may be as discussed above for probes in the 20 nt range, and be dependent on the removal of the sequence in the ‘X’ region for destabilization and cycling.
The mid-range probes may also be used at elevated temperatures, at or above their expected Tm, to allow melting rather than cleavage to promote probe turnover. In contrast to the longer probes described above, however, the temperatures required to allow the use of such a thermally driven turnover are much lower (about 40 to 60° C.), thus preserving both the cleavage means and the nucleic acids in the reaction from thermal degradation. In this way, the mid-range probes may perform in some instances like the miniprobes described above. In a further similarity to the miniprobes, the accumulation of cleavage signal from a mid-range probe may be helped under some reaction conditions by the presence of a stacker.
To summarize, a standard long probe usually does not benefit from the presence of a stacker oligonucleotide downstream (the exception being cases where such an oligonucleotide may also disrupt structures in the target nucleic acid that interfere with the probe binding), and it is usually used in conditions requiring several nucleotides to be removed to allow the oligonucleotide to release from the target efficiently.
The miniprobe is very short and performs optimally in the presence of a downstream stacker oligonucleotide. The miniprobes are well suited to reactions conditions that use the temperature of the reaction to drive rapid exchange of the probes on the target regardless of whether any bases have been cleaved. In reactions with sufficient amount of the cleavage means, the probes that do bind will be rapidly cleaved before they melt off.
The mid-range or midiprobe combines features of these probes and can be used in reactions like those designed long probes, with longer regions of overlap (“X” regions) to drive probe turnover at lower temperature. In a preferred embodiment, the midrange probes are used at temperatures sufficiently high that the probes are hybridizing to the target and releasing rapidly regardless of cleavage. This is known to be the behavior of oligonucleotides at or near their melting temperature. This mode of turnover is more similar to that used with miniprobe/stacker combinations than with long probes. The mid-range probe may have enhanced performance in the presence of a stacker under some circumstances. For example, with a probe in the lower end of the mid-range, e.g., 11 nt, or one with exceptional A/T content, in a reaction performed well in excess of the Tm of the probe (e.g., >10° C. above) the presence of a stacker would be likely to enhance the performance of the probe, while at a more moderate temperature the probe may be indifferent to a stacker.
The distinctions between the mini-, midi- (i.e., mid-range) and long probes are not contemplated to be inflexible and based only on length. The performance of any given probe may vary with its specific sequence, the choice of solution conditions, the choice of temperature and the selected cleavage means.
It is shown in Example 18 that the assemblage of oligonucleotides that comprises the cleavage structure of the present invention is sensitive to mismatches between the probe and the target. The site of the mismatch used in Ex. 18 provides one example and is not intended to be a limitation in location of a mismatch affecting cleavage. It is also contemplated that a mismatch between the InvaderTM oligonucleotide and the target may be used to distinguish related target sequences. In the 3-oligonucleotide system, comprising an InvaderTM, a probe and a stacker oligonucleotide, it is contemplated that mismatches may be located within any of the regions of duplex formed between these oligonucleotides and the target sequence. In a preferred embodiment, a mismatch to be detected is located in the probe. In a particularly preferred embodiment, the mismatch is in the probe, at the basepair immediately upstream (i.e., 5′) of the site that is cleaved when the probe is not mismatched to the target.
In another preferred embodiment, a mismatch to be detected is located within the region ‘Z’ defined by the hybridization of a miniprobe. In a particularly preferred embodiment, the mismatch is in the miniprobe, at the basepair immediately upstream (i.e., 5′) of the site that is cleaved when the miniprobe is not mismatched to the target.
It is also contemplated that different sequences may be detected in a single reaction. Probes specific for the different sequences may be differently labeled. For example, the probes may have different dyes or other detectable moieties, different lengths, or they may have differences in net charges of the products after cleavage. When differently labeled in one of these ways, the contribution of each specific target sequence to final product can be tallied. This has application in detecting the quantities of different versions of a gene within a mixture. Different genes in a mixture to be detected and quantified may be wild type and mutant genes, e.g., as may be found in a tumor sample (e.g., a biopsy). In this embodiment, one might design the probes to precisely the same site, but one to match the wild-type sequence and one to match the mutant. Quantitative detection of the products of cleavage from a reaction performed for a set amount of time will reveal the ratio of the two genes in the mixture. Such analysis may also be performed on unrelated genes in a mixture. This type of analysis is not intended to be limited to two genes. Many variants within a mixture may be similarly measured.
Alternatively, different sites on a single gene may be monitored and quantified to verify the measurement of that gene. In this embodiment, the signal from each probe would be expected to be the same.
It is also contemplated that multiple probes may be used that are not differently labeled, such that the aggregate signal is measured. This may be desirable when using many probes designed to detect a single gene to boost the signal from that gene. This configuration may also be used for detecting unrelated sequences within a mix. For example, in blood banking it is desirable to know if any one of a host of infectious agents is present in a sample of blood. Because the blood is discarded regardless of which agent is present, different signals on the probes would not be required in such an application of the present invention, and may actually be undesirable for reasons of confidentiality.
Just as described for the two-oligonucleotide system, above, the specificity of the detection reaction will be influenced by the aggregate length of the target nucleic acid sequences involved in the hybridization of the complete set of the detection oligonucleotides. For example, there may be applications in which it is desirable to detect a single region within a complex genome. In such a case the set of oligonucleotides may be chosen to require accurate recognition by hybridization of a longer segment of a target nucleic acid, often in the range of 20 to 40 nucleotides. In other instances it may be desirable to have the set of oligonucleotides interact with multiple sites within a target sample. In these cases one approach would be to use a set of oligonucleotides that recognize a smaller, and thus statistically more common, segment of target nucleic acid sequence.
In one preferred embodiment, the invader and stacker oligonucleotides may be designed to be maximally stable, so that they will remain bound to the target sequence for extended periods during the reaction. This may be accomplished through any one of a number of measures well known to those skilled in the art, such as adding extra hybridizing sequences to the length of the oligonucleotide (up to about 50 nts in total length), or by using residues with reduced negative charge, such as phosphorothioates or peptide-nucleic acid residues, so that the complementary strands do not repel each other to degree that natural strands do. Such modifications may also serve to make these flanking oligonucleotides resistant to contaminating nucleases, thus further ensuring their continued presence on the target strand during the course of the reaction. In addition, the InvaderTM and stacker oligonucleotides may be covalently attached to the target (e.g., through the use of psoralen cross-linking).
The use of the reaction temperatures at or near the Tm of the probe oligonucleotide, rather than that used for cleavage, to drive the turnover of the probe oligonucleotide in these detection reactions means that the amount of the probe oligonucleotide cleaved off may be substantially reduced without adversely affecting the turnover rate. It has been determined that the relationship between the 3′ end of the upstream oligonucleotide and the desired site of cleavage on the probe must be carefully designed. It is known that the preferred site of cleavage for the types of structure specific endonucleases employed herein is one basepair into a duplex (Lyamichev et al., supra). It was previously believed that the presence of an upstream oligonucleotide or primer allowed the cleavage site to be shifted away from this preferred site, into the single stranded region of the 5′ arm (Lyamichev et al., supra and U.S. Pat. No. 5,422,253). In contrast to this previously proposed mechanism, and while not limiting the present invention to any particular mechanism, it is believed that the nucleotide immediately 5′, or upstream of the cleavage site on the probe (including miniprobe and mid-range probes) must be able to basepair with the target for efficient cleavage to occur. In the case of the present invention, this would be the nucleotide in the probe sequence immediately upstream of the intended cleavage site. In addition, as described herein, it has been observed that in order to direct cleavage to that same site in the probe, the upstream oligonucleotide must have its 3′ base (i.e., nt) immediately upstream of the intended cleavage site of the probe. This places the 3′ terminal nucleotide of the upstream oligonucleotide and the base of the probe oligonucleotide 5′ of the cleavage site in competition for pairing with the corresponding nucleotide of the target strand.
To examine the outcome of this competition, i.e. which base is paired during a successful cleavage event, substitutions were made in the probe and invader oligonucleotides such that either the probe or the InvaderTM oligonucleotide were mismatched with the target sequence at this position. The effects of both arrangements on the rates of cleavage were examined. When the InvaderTM oligonucleotide is unpaired at the 3′ end, the rate of cleavage was not reduced. If this base was removed, however, the cleavage site was shifted upstream of the intended site. In contrast, if the probe oligonucleotide was not base-paired to the target just upstream of the site to which the InvaderTM oligonucleotide was directing cleavage, the rate of cleavage was dramatically reduced, suggesting that when a competition exists, the probe oligonucleotide was the molecule to be base-paired in this position.
It appears that the 3′ end of the upstream invader oligonucleotide is unpaired during cleavage, and yet is required for accurate positioning of the cleavage. To examine which part(s) of the 3′ terminal nucleotide are required for the positioning of cleavage, InvaderTM oligonucleotides were designed that terminated on this end with nucleotides that were altered in a variety of ways. Sugars examined included 2′ deoxyribose with a 3′ phosphate group, a dideoxyribose, 3′ deoxyribose, 2′ O-methyl ribose, arabinose and arabinose with a 3′ phosphate. Abasic ribose, with and without 3′ phosphate were tested. Synthetic “universal” bases such at 3-nitropyrrole and 5-3nitroindole on ribose sugars were tested. Finally, a base-like aromatic ring structure, acridine, linked to the 3′ end the previous nucleotide without a sugar group was tested. The results obtained support the conclusion that the aromatic ring of the base (at the 3′ end of the invader oligonuceotide) is the required moiety for accomplishing the direction of cleavage to the desired site within the downstream probe.
VI. Signal Enhancement by Tailing of Reaction Products in the InvaderTM-Directed Cleavage Assay
It has been determined that when oligonucleotide probes are used in cleavage detection assays at elevated temperature, some fraction of the truncated probes will have been shortened by nonspecific thermal degradation, and that such breakage products can make the analysis of the target-specific cleavage data more difficult. The thermal degradation that creates a background ladder of bands when the probes of the present invention are treated at high temperature for more than a few minutes occurs as a two step process. In the first step the N-glycosyl bond breaks, leaving an abasic site in the DNA strand. At the abasic site the DNA chain is weakened and undergoes spontaneous cleavage through a beta-elimination process. It has been determined that purine bases are about 20 times more prone to breakage than pyrimidine bases [Lindahl (1993) Nature 362:709]. This suggests that one way of reducing background in methods using oligonucleotides at elevated temperatures is to select target sequences that allow the use of pyrimidine-rich probes. It is preferable, where possible, to use oligonucleotides that are entirely composed of pyrimidine residues. If only one or a few purines are used, the background breakage will appear primarily at the corresponding sites, and these bands (due to thermal breakdown) may be mistaken for the intended cleavage products if care is not taken in the data analysis (i.e., proper controls must be run).
Background cleavage due to thermal breakdown of probe oligonucleotides can, when not resolved from specific cleavage products, reduce the accuracy of quantitation of target nucleic acids based on the amount of accumulated product in a set timeframe. One means of distinguishing the specific from the nonspecific products is disclosed above, and is based on partitioning the products of these reactions by differences in the net charges carried by the different molecular species in the reaction. As was noted in that discussion, the thermal breakage products usually retain 3′ phosphates after breakage, while the enzyme-cleaved products do not. The two negative charges on the phosphate facilitate charge-based partition of the products.
The absence of a 3′ phosphate on the desired subset of the probe fragments may be used to advantage in enzymatic assays as well. Nucleic acid polymerases, both non-templated (e.g., terminal deoxynucleotidyl transferase, polyA polymerase) and template-dependent (e.g., Pol I-type DNA polymerases), require an available 3′ hydroxyl by which to attach further nucleotides. This enzymatic selection of 3′ end structure may be used as an effective means of partitioning specific from non-specific products.
In addition to the benefits of the partitioning described above, the addition of nucleotides to the end of the specific product of an invader-specific cleavage offers an opportunity to either add label to the products, to add capturable tails to facilitate solid-support based readout systems, or to do both of these things at the same time. Some possible embodiments of this concept are illustrated in FIG. 56.
In FIG. 56, an InvaderTM cleavage struture comprising an InvaderTM oligonuclotide containing a blocked or non-extendible 3′ end (e.g., a 3′ dideoxynucleotide) and a probe oligonucleotide containing a blocked or non-extendable 3′ end (the open circle at the 3′ end of the oligonucleotides represents a non-extendible nucleotide) and a target nucleic acid is shown; the probe oligonucleotide may contain a 5′ end label such as a biotin or a fluorescein (indicated by the stars) label (cleavage structures which employ a 5′ biotin-labeled probe or a 5′ fluorescein-labeled probe are shown below the large diagram of the cleavage structure to the left and the right, respectively). Following, cleavage of the probe (the site of cleavage is indicated by the large arrowhead), the cleaved biotin-labeled probe is extended using a template-independent polymerase (e.g., TdT) and fluoresceinated nucleotide triphosphates. The fluorescein tailed cleaved probe molecule is then captured by binding via its 5′ biotin label to streptavidin and the fluroescence is then measured. Alternatively, following, cleavage of a 5′-fluoresceinated probe, the cleaved probe is extended using a template-independent polymerase (e.g., TdT) and dATP. The polyadenylated (A-tailed) cleaved probe molecule is then captured by binding via the polyA tail to oligo dT attached to a solid support.
The examples described in FIG. 56 are based on the use of TdT to tail the specific products of InvaderTM-directed cleavage. The description of the use of this particular enzyme is presented by way of example and is not intended as a limitation (indeed, when probe oligos comprising RNA are employed, cleaved RNA probes may be extended using polyA polymerase). It is contemplated that an assay of this type could be configured to use a template-dependent polymerase, as described above. While this would require the presence of a suitable copy template distinct from the target nucleic acid, on which the truncated oligonucleotide could prime synthesis, it can be envisaged that a probe which before cleavage would be unextendible, due to either mismatch or modification of the 3′ end, could be activated as a primer when cleaved by an invader directed cleavage. A template directed tailing reaction also has the advantage of allowing greater selection and control of the nucleotides incorporated.
The use of nontemplated tailing does not require the presence of any additional nucleic acids in the detection reaction, avoiding one step of assay development and troubleshooting. In addition, the use of non templated synthesis eliminated the step of hybridization, potentially speeding up the assay. Furthermore, the TdT enzyme is fast, able to add at least >700 nucleotides to substrate oligonucleotides in a 15 minute reaction.
As mentioned above, the tails added can be used in a number of ways. It can be used as a straight-forward way of adding labeled moieties to the cleavage product to increase signal from each cleavage event. Such a reaction is depicted in the left side of FIG. 66. The labeled moieties may be anything that can, when attached to a nucleotide, be added by the tailing enzyme, such as dye molecules, haptens such as digoxigenin, or other binding groups such as biotin.
In a preferred embodiment the assay includes a means of specifically capturing or partitioning the tailed invader-directed cleavage products in the mixture. It can be seen that target nucleic acids in the mixture may be tailed during the reaction. If a label is added, it is desirable to partition the tailed invader-directed cleavage products from these other labeled molecules to avoid background in the results. This is easily done if only the cleavage product is capable of being captured. For example, consider a cleavage assay of the present invention in which the probe used has a biotin on the 5′ end and is blocked from extension on the 3′ end, and in which a dye is added during tailing. Consider further that the products are to be captured onto a support via the biotin moiety, and the captured dye measured to assess the presence of the target nucleic acid. When the label is added by tailing, only the specifically cleaved probes will be labeled. The residual uncut probes can still bind in the final capture step, but they will not contribute to the signal. In the same reaction, nicks and cuts in the target nucleic acid may be tailed by the enzyme, and thus become dye labeled. In the final capture these labeled targets will not bind to the support and thus, though labeled, they will not contribute to the signal. If the final specific product is considered to consist of two portions, the probe-derived portion and the tail portion, can be seen from this discussion that it is particularly preferred that when the probe-derived portion is used for specific capture, whether by hybridization, biotin/streptavidin, or other method, that the label be associated with the tail portion. Conversely, if a label is attached to the probe-derived portion, then the tail portion may be made suitable for capture, as depicted on the right side of FIG. 66. Tails may be captured in a number of ways, including hybridization, biotin incorporation with streptavidin capture, or by virtue if the fact that the longer molecules bind more predictably and efficiently to a number of nucleic acid minding matrices, such as nitrocellulose, nylon, or glass, in membrane, paper, resin, or other form. While not required for this assay, this separation of functions allows effective exclusion from signal of both unreacted probe and tailed target nucleic acid.
In addition to the supports decribed above, the tailed products may be captured onto any support that contains a suitable capture moiety. For example, biotinylated products are generally captured with avidin-treated surfaces. These avidin surfaces may be in microtitre plate wells, on beads, on dipsticks, to name just a few of the possibilities. Such surfaces can also be modified to contain specific oligonucleotides, allowing capture of product by hybridization. Capture surfaces as described here are generally known to those skilled in the art and include nitrocellulose dipsticks (e.g., GeneCombTM, BioRad, Hercules, Calif.).
VII. Improved Enzymes for Use in InvaderTM-Directed Cleavage Reactions
A cleavage structure is defined herein as a structure which is formed by the interaction of a probe oligonucleotide and a target nucleic acid to form a duplex, the resulting structure being cleavable by a cleavage means, including but not limited to an enzyme. The cleavage structure is further defined as a substrate for specific cleavage by the cleavage means in contrast to a nucleic acid molecule which is a substrate for nonspecific cleavage by agents such as phosphodiesterases. Examples of some possible cleavage structures are shown in FIG. 15. In considering improvements to enzymatic cleavage means, one may consider the action of said enzymes on any of these structures, and on any other structures that fall within the definition of a cleavage structure. The cleavage sites indicated on the structures in FIG. 15 are presented by way of example. Specific cleavage at any site within such a structure is contemplated.
Improvements in an enzyme may be an increased or decreased rate of cleavage of one or more types of structures. Improvements may also result in more or fewer sites of cleavage on one or more of said cleavage structures. In developing a library of new structure-specific nucleases for use in nucleic acid cleavage assays, improvements may have many different embodiments, each related to the specific substrate structure used in a particular assay.
As an example, one embodiment of the InvaderTM-directed cleavage assay of the present invention may be considered. In the InvaderTM directed cleavage assay, the accumulation of cleaved material is influenced by several features of the enzyme behavior. Not surprisingly, the turnover rate, or the number of structures that can be cleaved by a single enzyme molecule in a set amount of time, is very important in determining the amount of material processed during the course of an assay reaction. If an enzyme takes a long time to recognize a substrate (e.g., if it is presented with a less-than-optimal structure), or if it takes a long time to execute cleavage, the rate of product accumulation is lower than if these steps proceeded quickly. If these steps are quick, yet the enzyme “holds on” to the cleaved structure, and does not immediately proceed to another uncut structure, the rate will be negatively affected.
Enzyme turnover is not the only way in which enzyme behavior can negatively affect the rate of accumulation of product. When the means used to visualize or measure product is specific for a precisely defined product, products that deviate from that definition may escape detection, and thus the rate of product accumulation may appear to be lower than it is. For example, if one had a sensitive detector for trinucleotides that could not see di- or tetranucleotides, or any sized oligonucleotide other that 3 residues, in the InvaderTM-directed cleavage assay of the present invention any errant cleavage would reduce the detectable signal proportionally. It can be seen from the cleavage data presented here that, while there is usually one site within a probe that is favored for cleavage, there are often products that arise from cleavage one or more nucleotides away from the primary cleavage site. These are products that are target dependent, and are thus not non-specific background. Nevertheless, if a subsequent visualization system can detect only the primary product, these represent a loss of signal. One example of such a selective visualization system is the charge reversal readout presented herein, in which the balance of positive and negative charges determines the behavior of the products. In such a system the presence of an extra nucleotide or the absence of an expected nucleotide can excluded a legitimate cleavage product from ultimate detection by leaving that product with the wrong balance of charge. It can be easily seen that any assay that can sensitively distinguish the nucleotide content of an oligonucleotide, such as standard stringent hybridization, suffers in sensitivity when some fraction of the legitimate product is not eligible for successful detection by that assay.
These discussions suggest two highly desirable traits in any enzyme to be used in the method of the present invention. First, the more rapidly the enzyme executes an entire cleavage reaction, including recognition, cleavage and release, the more signal it may potentially created in the invader-directed cleavage assay. Second, the more successful an enzyme is at focusing on a single cleavage site within a structure, the more of the cleavage product can be successfully detected in a selective read-out.
The rationale cited above for making improvements in enzymes to be used in the InvaderTM-directed cleavage assay are meant to serve as an example of one direction in which improvements might be sought, but not as a limit on either the nature or the applications of improved enzyme activities. As another direction of activity change that would be appropriately considered improvement, the DNAP-associated 5′ nucleases may be used as an example. In creating some of the polymerase-deficient 5′ nucleases described herein it was found that the those that were created by deletion of substantial portions of the polymerase domain, as depicted in FIG. 4, assumed activities that were weak or absent in the parent proteins. These activities included the ability to cleave the non-forked structure shown in FIG. 15D, a greatly enhanced ability to exonucleolytically remove nucleotides from the 5′ ends of duplexed strands, and a nascent ability to cleave circular molecules without benefit of a free 5′ end.
In addition to the 5′ nucleases derived from DNA polymerases, the present invention also contemplates the use of structure-specific nucleases that are not derived from DNA polymerases. For example, a class of eukaryotic and archaebacterial endonucleases have been identified which have a similar substrate specificity to 5′ nucleases of Pol I-type DNA polymerases. These are the FEN1 (Flap EndoNuclease), RAD2, and XPG (Xeroderma Pigmentosa-complementation group G) proteins. These proteins are involved in DNA repair, and have been shown to favor the cleavage of structures that resemble a 5′ arm that has been displaced by an extending primer during polymerization, similar to the model depicted in FIG. 15B. Similar DNA repair enzymes have been isolated from single cell and higher eukaryotes and from archaea, and there are related DNA repair proteins in eubacteria. Similar 5′ nucleases have also be associated with bacteriophage such as T5 and T7.
Recently, the 3-dimensional structures of DNAPTaq and T5 phage 5′-exonuclease (FIG. 58) were determined by X-ray diffraction [Kim et al. (1995) Nature 376:612 and Ceska et al. (1995) Nature 382:90). The two enzymes have very similar 3-dimensional structures despite limited amino acid sequence similarity. The most striking feature of the T5 5′-exonuclease structure is the existence of a triangular hole formed by the active site of the protein and two alpha helices (FIG. 58). This same region of DNAPTaq is disordered in the crystal structure, indicating that this region is flexible, and thus is not shown in the published 3-dimensional structure. However, the 5′ nuclease domain of DNAPTaq is likely to have the same structure, based its overall 3-dimensional similarity to T5 5′-exonuclease, and that the amino acids in the disordered region of the DNAPTaq protein are those associated with alpha helix formation. The existence of such a hole or groove in the 5′ nuclease domain of DNAPTaq was predicted based on its substrate specificity [Lyamichev et al., supra].
It has been suggested that the 5′ arm of a cleavage structure must thread through the helical arch described above to position said structure correctly for cleavage (Ceska et al., supra). One of the modifications of 5′ nucleases described herein opened up the helical arch portion of the protein to allow improved cleavage of structures that cut poorly or not at all (e.g., structures on circular DNA targets that would preclude such threading of a 5′ arm). The gene construct that was chosen as a model to test this approach was the one called Cleavase® BN, which was derived from DNAPTaq but does not contain the polymerase domainn (Ex. 2). It comprises the entire 5′ nuclease domain of DNAP Taq, and thus should be very close in structure to the T5 5′ exonuclease. This 5′ nuclease was chosen to demonstrate the principle of such a physical modification on proteins of this type. The arch-opening modification of the present invention is not intended to be limited to the 5′ nuclease domains of DNA polymerases, and is contemplated for use on any structure-specific nuclease which includes such an aperture as a limitation on cleavage activity. The present invention contemplates the insertion of a thrombin cleavage site into the helical arch of DNAPs derived from the genus Thermus as well as 5′ nucleases derived from DNAPs derived from the genus Thermus. The specific example shown herein using the Cleavase® BN/thrombin nuclease merely illustrates the concept of opening the helical arch located within a nuclease domain. As the amino acid sequence of DNAPs derived from the genus Thermus are highly conserved, the teachings of the present invention enable the insertion of a thrombin site into the helical arch present in these DNAPs and 5′ nucleases derived from these DNAPs.
The opening of the helical arch was accomplished by insertion of a protease site in the arch. This allowed post-translational digestion of the expressed protein with the appropriate protease to open the arch at its apex. Proteases of this type recognize short stretches of specific amino acid sequence. Such proteases include thrombin and factor Xa. Cleavage of a protein with such a protease depends on both the presence of that site in the amino acid sequence of the protein and the accessibility of that site on the folded intact protein. Even with a crystal structure it can be difficult to predict the susceptibility of any particular region of a protein to protease cleavage. Absent a crystal structure it must be determined empirically.
In selecting a protease for a site-specific cleavage of a protein that has been modified to contain a protease cleavage site, a first step is to test the unmodified protein for cleavage at alternative sites. For example, DNAPTaq and Cleavase® BN nuclease were both incubated under protease cleavage conditions with factor Xa and thrombin proteases. Both nuclease proteins were cut with factor Xa within the 5′ nuclease domain, but neither nuclease was digested with large amounts of thrombin. Thus, thrombin was chosen for initial tests on opening the arch of the Cleavase® BN enzyme.
In the protease/Cleavase® modifications described herein the factor Xa protease cleaved strongly in an unacceptable position in the unmodified nuclease protein, in a region likely to compromise the activity of the end product. Other unmodified nucleases contemplated herein may not be sensitive to the factor Xa, but may be sensitive to thrombin or other such proteases. Alternatively, they may be sensitive to these or other such proteases at sites that are immaterial to the function of the nuclease sought to be modified. In approaching any protein for modification by addition of a protease cleavage site, the unmodified protein should be tested with the proteases under consideration to determine which proteases give acceptable levels of cleavage in other regions.
Working with the cloned segment of DNAPTaq from which the Cleavase® BN protein is expressed, nucleotides encoding a thrombin cleavage site were introduced in-frame near the sequence encoding amino acid 90 of the nuclease gene. This position was determined to be at or near the apex of the helical arch by reference to both the 3-dimensional structure of DNAPTaq, and the structure of T5 5′ exonuclease. The encoded amino acid sequence, LVPRGS, was inserted into the apex of the helical arch by site-directed mutagenesis of the nuclease gene. The proline (P) in the thrombin cleavage site was positioned to replace a proline normally in this position in Cleavase® BN because proline is an alpha helix-breaking amino acid, and may be important for the 3-dimensional structure of this arch. This construct was expressed, purified and then digested with thrombin. The digested enzyme was tested for its ability to cleave a target nucleic acid, bacteriophage M13 genomic DNA, that does not provide free 5′ ends to facilitate cleavage by the threading model.
While the helical arch in this nuclease was opened by protease cleavage, it is contemplated that a number of other techniques could be used to achieve the same end. For example, the nucleotide sequence could be rearranged such that, upon expression, the resulting protein would be configured so that the top of the helical arch (amino acid 90) would be at the amino terminus of the protein, the natural carboxyl and amino termini of the protein sequence would be joined, and the new carboxyl terminus would lie at natural amino acid 89. This approach has the benefit that no foreign sequences are introduced and the enzyme is a single amino acid chain, and thus may be more stable that the cleaved 5′ nuclease. In the crystal structure of DNAPTaq, the amino and carboxyl termini of the 5′-exonuclease domain lie in close proximity to each other, which suggests that the ends may be directly joined without the use of a flexible linker peptide sequence as is sometimes necessary. Such a rearrangement of the gene, with subsequent cloning and expression could be accomplished by standard PCR recombination and cloning techniques known to those skilled in the art.
The present invention also contemplates the use of nucleases isolated from a organisms that grow under a variety of conditions. The genes for the FEN-1/XPG class of enzymes are found in organisms ranging from bacteriophage to humans to the extreme thermophiles of Kingdom Archaea. For assays in which high temperature is to be used, it is contemplated that enzymes isolated from extreme thermophiles may exhibit the thermostability required of such an assay. For assays in which it might be desirable to have peak enzyme activity at moderate temperature or in which it might be desirable to destroy the enzyme with elevated temperature, those enzymes from organisms that favor moderate temperatures for growth may be of particular value.
An alignment of a collection of FEN-1 proteins sequenced by others is shown in FIGS. 59A-E (SEQ ID NOS:135-145). It can be seen from this alignment that there are some regions of conservation in this class of proteins, suggesting that they are related in function, and possibly in structure. Regions of similarity at the amino acid sequence level can be used to design primers for in vitro amplification (PCR) by a process of back translating the amino acid sequence to the possible nucleic acid sequences, then choosing primers with the fewest possible variations within the sequences. These can be used in low stringency PCR to search for related DNA sequences. This approach permits the amplification of DNA encoding a FEN-1 nuclease without advance knowledge of the actual DNA sequence.
It can also be seen from this alignment that there are regions in the sequences that are not completely conserved. The degree of difference observed suggests that the proteins may have subtle or distinct differences is substrate specificity. In other words, they may have different levels of cleavage activity on the cleavage structures of the present invention. When a particular structure is cleaved at a higher rate than the others, this is referred to a preferred substrate, while a structure that is cleaved slowly is considered a less preferred substrate. The designation of preferred or less preferred substrates in this context is not intended to be a limitation of the present invention. It is contemplated that some embodiments the present invention will make use of the interactions of an enzyme with a less preferred substrate. Candidate enzymes are tested for suitability in the cleavage assays of the present invention using the assays described below.
1. Structure Specific Nuclease Assay
Testing candidate nucleases for structure-specific activities in these assays is done in much the same way as described for testing modified DNA polymerases in Example 2, but with the use of a different library of model structures. In addition to assessing the enzyme performance in primer-independent and primer-directed cleavage, a set of synthetic hairpins are used to examine the length of duplex downstream of the cleavage site preferred by the enzyme.
The FEN-1 and XPG 5′ nucleases used in the present invention must be tested for activity in the assays in which they are intended to be used, including but not limited to the InvaderTM-directed cleavage detection assay of the present invention and the CFLP® method of characterizing nucleic acids (the CFLP® method is described in co-pending application Ser. Nos. 08/337,164, 08/402,601, 08/484,956 and 08/520,946; the disclosures of these applications are incorporated herein by reference). The InvaderTM assay uses a mode of cleavage that has been termed “primer directed” of “primer dependent” to reflect the influence of the an oligonucleotide hybridized to the target nucleic acid upstream of the cleavage site. In contrast, the CFLP® reaction is based on the cleavage of folded structure, or hairpins, within the target nucleic acid, in the absence of any hybridized oligonucleotide. The tests described herein are not intended to be limited to the analysis of nucleases with any particular site of cleavage or mode of recognition of substrate structures. It is contemplated that enzymes may be described as 3′ nucleases, utilizing the 3′ end as a reference point to recognize structures, or may have a yet a different mode of recognition. Further, the use of the term 5′ nucleases is not intended to limit consideration to enzymes that cleave the cleavage structures at any particular site. It refers to a general class of enzymes that require some reference or access to a 5′ end to effect cleavage of a structure.
A set of model cleavage structures have been created to allow the cleavage ability of unknown enzymes on such structures to be assessed. Each of the model structures is constructed of one or more synthetic oligonucleotides made by standard DNA synthesis chemistry. Examples of such synthetic model substrate structures are shown in FIGS. 26 and 60. These are intended only to represent the general folded configuration desirable is such test structures. While a sequence that would assume such a structure is indicated in the figures, there are numerous other sequence arrangements of nucleotides that would be expected to fold in such ways. The essential features to be designed into a set of oligonucleotides to perform the tests described herein are the presence or absence of a sufficiently long 3′ arm to allow hybridization of an additional nucleic acid to test cleavage in a “primer-directed” mode, and the length of the duplex region. In the set depicted in FIG. 60, the duplex lengths of the S-33 and the 11-8-0 structures are 12 and 8 basepairs, respectively. This difference in length in the test molecules facilitates detection of discrimination by the candidate nuclease between longer and shorter duplexes. Additions to this series expanding the range of duplex molecules presented to the enzymes, both shorter and longer, may be used. The use of a stabilizing DNA tetraloop [Antao et al. (1991) Nucl. Acids Res. 19:5901] or triloop [Hiraro et al. (1994) Nuc. Acids Res. 22:576] at the closed end of the duplex helps ensure formation of the expected structure by the oligonucleotide.
The model substrate for testing primer directed cleavage, the “S-60 hairpin” (SEQ ID NO:40) is described in Example 11. In the absence of a primer this hairpin is usually cleaved to release 5′ arm fragments of 18 and 19 nucleotides length. An oligonucleotide, termed P-14 (5′-CGAGAGACCACGCT-3′; SEQ ID NO:108), that extends to the base of the duplex when hybridized to the 3′ arm of the S-60 hairpin gives cleavage products of the same size, but at a higher rate of cleavage.
To test invasive cleavage a different primer is used, termed P-15 (5′-CGAGAGACCACGCTG-3′; SEQ ID NO:30). In a successful invasive cleavage the presence of this primer shifts the site of cleavage of S-60 into the duplex region, usually releasing products of 21 and 22 nucleotides length.
The S-60 hairpin may also be used to test the effects of modifications of the cleavage structure on either primer-directed or invasive cleavage. Such modifications include, but are not limited to, use of mismatches or base analogs in the hairpin duplex at one, a few or all positions, similar disruptions or modifications in the duplex between the primer and the 3′ arm of the S-60, chemical or other modifications to one or both ends of the primer sequence, or attachment of moieties to, or other modifications of the 5′ arm of the structure. In all of the analyses using the S-60 or a similar hairpin described herein, activity with and without a primer may be compared using the same hairpin structure.
The assembly of these test reactions, including appropriate amounts of hairpin, primer and candidate nuclease are described in Example 2. As cited therein, the presence of cleavage products is indicated by the presence of molecules which migrate at a lower molecular weight than does the uncleaved test structure. When the reversal of charge of a label is used the products will carry a different net charge than the uncleaved material. Any of these cleavage products indicate that the candidate nuclease has the desired structure-specific nuclease activity. By “desired structure-specific nuclease activity” it is meant only that the candidate nuclease cleaves one or more test molecules. It is not necessary that the candidate nuclease cleave at any particular rate or site of cleavage to be considered successful cleavage.
VIII. Signal Enhancement by Completion of an Activated Protein Binding Site
In addition to the DNA polymerase tailing reaction described above, the present invention also contemplates the use of the products of the invasive cleavage reaction to form activated protein binding sites, such as RNA polymerase promoter duplexes, thereby allowing the interaction of the completed site to be used as an indicator of the presence of the nucleic acid that is the target of the invasive cleavage reaction. By way of example, when an RNA polymerase promoter duplex is activated by being made complete (i.e., double-stranded over that portion of the promoter region required for polymerase binding) through the hybridization of the oligonucleotide product of the invasive cleavage reaction, the synthesis of RNA can be used as such an indicator.
It is not intended that the transcription reaction of the present invention be limited to the use of any particular RNA polymerase or RNA polymerase promoter region. Promoter sequences are well characterized for several bacteriophage, including bacteriophage SP6, T7 and T3. In addition, promoter sequences have been well characterized for a number of both eukaryotic and prokaryotic RNA polymerases. In a preferred embodiment, the promoter used enables transcription from one of the bacteriophage RNA polymerases. In a particularly preferred embodiment, the promoter used enables transcription by T7 RNA polymerase. Means of performing transcription in vitro are well known in the art and commercial kits are available for performing transcription with eukaryotic, prokaryotic or bacteriophage RNA polymerases (e.g., from Promega).
The protein binding regions of the present invention are not limited to the bacteriophage RNA polymerase promoters described above. Other promoter sequences that are contemplated are those of prokaryotes and eukaryotes. For example, many strains of bacteria and fungi are used for the expression of heterologous proteins. The minimal promoters required for transcription by the RNA polymerases of organisms such as yeast and other fungi, eubacteria, nematodes, and cultured mammalian cells are well described in the literature and in the catalogs of commercial suppliers of DNA vectors for the expression of foreign proteins in these organisms.
The binding sites for other types of nucleic acid (e.g., DNA) binding proteins are contemplated for use in the present invention. For example, proteins involved in the regulation of genes exert their effects by binding to the DNA in the vicinity of the promoter from which the RNA from that gene is transcribed. The lac operator of E. coli is one example of a particularly well characterized and commonly used gene regulation system in which the lac repressor protein binds to specific sequences that overlap, and thus block, the promoter for the genes under the repressor's control [Jacob and Monod (1961) Cold Spring Harbor Symposium on Quantitative Biol. XXVI: 193-211]. Many similar systems have been described in bacteria, including the trp and AraC regulatory systems. Given the large amount of information available about bacterial promoters, the steps described below for the design of suitable partial promoters for the bacteriophage RNA polymerases can be readily adapted to the design of detection systems based on these other promoters.
As noted above, many of the bacterial promoters are under the control of a repressor or other regulatory protein. It is considered to be within the scope of the present invention to include the creation of composite binding sites for these regulatory proteins through the provision of a nucleic acid fragment (e.g., a non-target cleavage product generated in an invasive cleavage reaction). The binding of the regulatory protein to the completed protein binding region (e.g., the composite binding region) can be assessed by any one of a number of means, including slowed electrophoretic migration of either the protein or the DNA fragment, or by a conformational change in the protein or DNA upon binding. In addition, transcription from a downstream promoter can be monitored for up- or down-regulation as a result of the binding of the regulatory protein to the completed protein binding region.
In addition to the bacterial systems described above, many genes in eukaryotic systems have also been found to be under the control of specific proteins that bind to specific regions of duplex DNA. Examples include, but are not limited to, the OCT-1, OCT-2 and AP-4 proteins in mammals and the GAL4 and GCN4 proteins in yeast. Such regulatory proteins usually have a structural motif associated with duplex nucleic acid binding, such as a helix-turn-helix, a zinc finger or a leucine zipper [for review, see, Molecular and Cellular Biology (1993) S. L. Wolfe, Ed., Wadsworth Publishing Co., Belmont, Calif., pp. 694-715].
For simplicity the test reaction described here will refer to T7 RNA polymerase, and its promoter. This is not intended to limit the invention to the use of this RNA polymerase, and those skilled in the art of molecular biology would be able to readily adapt this described test to the examination of any of the DNA binding proteins, RNA polymerases and their binding or promoter sites discussed above.
It is known in the art that active T7 promoters can be formed by the hybridization of two oligonucleotides, each comprising either the top or bottom strand of the promoter sequence, such that a complete un-nicked duplex promoter is formed [J. F. Milligan, D. R. Groebe, G. W. Witherall, O. C. Uhlenbeck, Nucl. Acids Res. 15:21, 8783-8798 (1987)]. We show here that one way of making the initiation of transcription dependent on the products of an invasive cleavage reaction is to design the probe for the cleavage reaction such that a portion of an RNA polymerase promoter is released as product. The remaining DNA piece or pieces required to assemble a promoter duplex may either be provided as elements in the reaction mixture, or they may be produced by other invasive cleavage events. If the oligonucleotide pieces are designed to comprise appropriate regions of complementarity they may base pair to form a complete promoter duplex composed of three or more nucleic acid fragments, as depicted in FIG. 88B. A promoter assembled in this way will have nicks in the backbone of one or both strands. In one embodiment, these nicks may be covalently closed through the use of a DNA ligase enzyme. In a preferred embodiment, the nicks are positioned such that transcription can proceed without ligation. In selecting the site of a nick created by the assembly of the partial promoter fragment, at least one nick should be within the recognized promoter region for the RNA polymerase to be used. When a bacteriophage promoter is used, a nick should be between nucleotides −17 and −1, measured from the site of transcription initiation at +1. In a preferred embodiment, a nick will be between nucleotides −13 and −8. In a particularly preferred embodiment, a nick will be between nucleotides −12 and −10 on the non-template strand of the bacteriophage promoter.
When nicks are to be left unrepaired (i.e., not covalently closed with a DNA ligase) it is important to assess the effect of the nick location on the level of transcription from the assembled promoter. A simple test is to combine the oligonucleotides that comprise the separate portions of the promoter with an oligonucleotide that comprises one entire strand of the promoter to be assembled, thereby forming a duplex promoter with a nick in one strand. If the nick is in the top, or non-template strand of the promoter, then the oligonucleotide that comprises the complete promoter is made to include additional non-promoter sequence on its 5′ end to serve as a template to be copied in the transcription. This arrangement is depicted in FIG. 88B. Alternatively, if the nick is to be in the bottom, or template strand of the promoter, then the partial promoter oligonucleotide that covers the +1 position, the transcription start site, will include the additional template sequence. This arrangement is depicted in FIGS. 95A-D (this figure shows several different embodiments in which a cut probe or non-target cleavage product is used to form a composite promoter which contains one or more nicks on the template strand). In either case, the separate oligonucleotides are combined to form the complete promoter, and the assembly is used in a transcription reaction to create RNA.
To measure the effect of the nick, a substantially identical promoter fragment is created by hybridization of two oligonucleotides that each comprise one strand of the full-length promoter to create an un-nicked version of the same promoter. These two molecular assemblies are tested in parallel transcription reactions and the amount of the expected RNA that is produced in each reaction is measured for both size and yield. A preferred method of assessing the size of the RNA is by electrophoresis with subsequent visualization. If a labeled nucleotide (e.g., 32P-GTP, or fluorescein-UTP) is used in the transcription, the RNA can be detected and quantitated by autoradiography, fluorescence imaging or by transfer to support membrane with subsequent detection, e.g., by antibody or hybridization probing. Alternatively, if unlabeled RNA is produced the amounts may be determined by other methods known in the art, such as by spectrophotometry or by electrophoresis with subsequent staining and comparison to known standards.
If the size of the RNA is as predicted by the template sequence, or if it matches that produced from the control promoter, it can be presumed to have initiated transcription at the same site in the complex, and to have produced essentially the same RNA product. If the product is much shorter then transcription is either initiating at an internal site or is terminating prematurely [E. T. Schenborn and R. C. Mierendorf, Jr., Nucl. Acids Res. 13:17, 6223 (1985); Milligan et al., supra]. While this does not indicate that the assembly tested is completely unsuitable for the assay, the partial transcripts will reduce the gross amount of RNA created, perhaps compromising the signal from the assay, and such products would require further characterization (e.g., finger printing or sequencing) to identify the nucleotide content of the product. It is preferred that the size of the RNA produced matches that of the RNA produced in the control reaction.
The yield of the reaction is also examined. It is not necessary that the level of transcription matches that of the control reaction. In some instances (see Ex. 41, below) the nicked promoter may have an enhanced rate of transcription, while in other arrangements transcription may be reduced (relative to the rate from the un-nicked promoter assembly). It is only required that the amount of product be within the detection limits of the method to be used with the test promoter.
It is reported that transcription from a bacteriophage promoter can produce 200 to 1000 copies of each transcription template (template plus active promoter) in a reaction. These levels of transcription are not required by the present invention. Reactions in which one RNA is produced for each template are also contemplated.
The test described above will allow a promoter with a nick in any position to be assessed for utility in this assay. It is an objective of this invention to provide one or more of the oligos which comprise a partial promoter region through invasive cleavage event(s). In this embodiment, the partial promoter sequences are attached to the probe oligonucleotide in the invasive cleavage assay, and are released by cleavage at specific site, as directed by the InvaderTM oligonucleotide. It is also intended that transcription be very poor or nonexistent in the absence of the correctly cleaved probe. To assess the success of any oligonucleotide design at meeting these objectives, several transcription reaction tests can be performed.
For a promoter assembly that will have a nick on the non-template strand, several partial assemblies that should be tested are shown in FIGS. 86 A-D. By way of example, but not by way of limitation, this figure depicts the tests for a nicked promoter in which the upstream, or 5′ portion of the non-template strand is to be provided by the invasive cleavage assay. This fragment is seen in FIG. 86A labeled as “cut probe”. Transcription reactions incubated in the presence of the duplex shown in FIG. 86A will test the ability of the upstream partial promoter to allow initiation of transcription when hybridized to a bottom strand, termed a “copy template.” Similarly, a reaction performed in the presence of the duplex depicted in FIG. 86B will test the ability of the partial promoter fragment nearest the initiation site (the +1 site, as indicated in FIG. 85B) to support transcription of the copy template. It is an important feature of the present invention that neither of these partial promoter duplexes be able to support transcription at the same level as would by seen in transcription from an intact promoter as depicted in FIG. 85B. It is preferred that neither of these partial promoters be sufficient to initiate detectable transcription in the time course of an average transcription reaction, i.e., within about an hour of incubation.
FIGS. 86C and 86D depict two other duplex arrangements designed to test the effect of uncut probe within the transcription reaction. FIG. 86C depicts the duplex formed between only the uncut probe and the copy template, while FIG. 86D includes the other portion of the promoter. The 3′ region of the probe is not complementary to the promoter sequence and therefore produces an unpaired branch in the middle of the promoter. It is an important feature of the present invention that neither of these branched promoter duplexes be able to support transcription at the same level as would by seen in transcription from an intact promoter as depicted in FIG. 85B. It is preferred that neither of these branched promoters be sufficient to initiate detectable transcription in the time course of an average transcription reaction, i.e., within about an hour of incubation.
In one embodiment of the transcription system of the present invention, the initiation of transcription from the copy template in the absence of a complete promoter, or in the presence of a branched promoter, is prevented by the judicious placement of the nick or nicks in the composite promoter. For example, as shown in the examples below, placement of a nick between the −12 and −11 nucleotides of the non-template strand of the bacteriophage T7 promoter allows transcription to take place only when the probe has been successfully cut, as in an invasive cleavage reaction. However, in some instances where the invasive cleavage reaction is to provide the upstream portion of the non-template strand of the promoter (e.g., as depicted in FIG. 88B) it may be necessary or desirable to place the nick on that strand in a particular position for reasons other than providing an optimal composite promoter (i.e., one that is inactive in the absence of any one of the promoter pieces). It may be necessary or desirable to place the nick in such a way that the creation of a branched complete promoter (FIG. 86D) has an undesirable level of transcription, reducing dependence of RNA production on the success of the invasive cleavage step. It is shown in the examples below that transcription from such a branched promoter can be suppressed by a modification of the downstream non-template promoter piece, shown as the “Partial Promoter Oligonucleotide” in FIGS. 86, 88, 90 and 95D. As depicted in FIG. 90, the partial promoter oligonucleotide can be provided with a 5′ “tail” of nucleotides that are not complementary to the template strand of the promoter, but which are complementary to the 3′ portion of the probe oligonucleotide that would be removed in the invasive cleavage reaction. When uncut probe hybridizes to the copy template with the bound 5′ tailed partial promoter oligonucleotide, the 5′ tail can basepair to the 3′ region of the probe, forming a three-way junction as depicted in FIG. 90A. This can effectively shut off transcription, as shown below. When a cut probe hybridizes, as shown in FIG. 90B, a promoter with a small branch is formed, and it is shown herein that such a branched promoter can initiate transcription. Furthermore, if care is taken in selecting the sequence of the 5′ tail (i.e., if the first unpaired base is the same nucleotide at the 3′ nucleotide of the cut probe, so that they compete for hybridization to the same template strand base), the resulting branched structure may also be cleaved by one of the structure specific nucleases of the present invention, creating the un-branched promoter depicted in FIG. 90C, in some instances enhancing transcription over that seen with the FIG. 90B promoter.
The promoter duplex that is intended to be created, in this embodiment, by the successful execution of the InvaderTM directed cleavage assay will include both the “cut probe” and the partial promoter oligonucleotide depicted in FIGS. 86A and B, aligned on a single copy template nucleic acid. The testing of the efficiency of transcription of such a nicked promoter segment in comparison to the intact promoter is described above. All of the oligonucleotides described for these test molecules may be created using standard synthesis chemistries.
The set of test molecules depicted in FIG. 86 is designed to assess the transcription capabilities of the variety of structures that may be present in reactions in which the 5′ portion of the non-template strand of the promoter is to be supplied by the invader directed cleavage. It is also envisioned that a different portion of partial promoter may be supplied by the invasive cleavage reaction, e.g., the downstream segment of the non-template strand of the promoter, as is shown in FIG. 94. Portions of the template strand of the promoter may also be provided by the cut probe, as shown in FIGS. 95A-D. An analogous set of test molecules, including “cut” and uncut versions of the probe to be used in the invasive cleavage assay may be created to test any alternative design, whether the nick is to be located on the template or non template strand of the promoter.
The transcription-based visualization methods of the present invention may also be used in a multiplex fashion. Reactions can be constructed such that the presence of one particular target leads to transcription from one type of promoter, while the presence of a different target sequence (e.g., a mutant or variant) or another target suspected of being present, may lead to transcription from a different (i.e., a second) type of promoter. In such an embodiment, the identity of the promoter from which transcription was initiated could be deduced from the type or size of the RNA produced.
By way of example, but not by way of limitation, the bacteriophage promoters can be compared with such an application in view. The promoters for the phage T7, T3 and SP6 are quite similar, each being about 15 to 20 basepairs long, and sharing about 45% identity between −17 and −1 nucleotides, relative to the start of transcription. Despite these similarities, the RNA polymerases from these phage are highly specific for their cognate promoters, such that the other promoters may be present in a reaction, but will not be transcribed [Chamberlin and Ryan (1982) The Enzymes XV:87-108]. Because these promoters are similar in size and in the way in which they are recognized by their polymerases [Li et al. (1996) Biochem. 35:3722] similar nicked versions of the promoters may be designed for use in the methods of the present invention by analogy to the examples described herein which employ the T7 promoter. Because of the high degree of specificity of the RNA polymerases, these nicked promoters may be used together to detect multiple targets in a single reaction. There are many instances in which it would be highly desirable to detect multiple nucleic acid targets in a single sample, including cases in which multiple infectious agents may be present, or in which variants of a single type of target may need to be identified. Alternatively, it is often desirable to use a combination of probes to detect both a target sequence and an internal control sequence, to gauge the effects of sample contaminants on the output of the assay. The use of multiple promoters allows the reaction to be assessed for both the efficiency of the invasive cleavage and the robustness of the transcription.
As stated above, the phage promoters were described in detail as an example of suitable protein binding regions (e.g., which can be used to generate a composite promoter) for use in the methods of the present invention. The invention is not limited to the use of phage RNA polymerase promoter regions, in particular, and RNA polymerase promoter regions, in general. Suitably specific, well characterized promoters are also found in both prokaryotic and eukaryotic systems.
The RNA that is produced in a manner that is dependent of the successful detection of the target nucleic acid in the invasive cleavage reaction may be detected in any of several ways. If a labeled nucleotide is incorporated into the RNA during transcription, the RNA may be detected directly after fractionation, e.g., by electrophoresis or chromatography. The labeled RNA may also be captured onto a solid support, such as a microtitre plate, a bead or a dipstick, e.g., by hybridization, antibody capture, or through an affinity interaction such as that between biotin and avidin. Capture may facilitate the measuring of incorporated label, or it may be an intermediate step before probe hybridization or similar detection means. If the maximum amount of label is desired to be incorporated into each transcript, it is preferred that the copy template be very long, around 3 to 10 kilobases, so that each RNA molecule will carry many labels. Alternatively, it may be desired that a single site or a limited number of sites within the transcript be specifically labeled. In this case, it may be desirable to have a short copy template with only one or a few residues that would allow incorporation of the labeled nucleotide.
The copy template may also be selected to produce RNAs that perform specified functions. In a simple case, if an duplex-dependent intercalating fluorophore is to be used to detect the RNA product, it may be desirable to transcribe an RNA that is known to form duplexed secondary structures, such as a ribosomal RNA or a tRNA. In another embodiment, the RNA may be designed to interact specifically, or with particular affinity, with a different substance. It has been shown that a process of alternating steps of selection (e.g., by binding to a target substance) and in vitro amplification (e.g., by PCR) can be used to identify nucleic acid ligands with novel and useful properties [Tuerk and Gold (1990) Science 249:505]. This system has been used to identify RNAs, termed ligands or aptamers, that bind tightly and specifically to proteins and to other types of molecules, such as antibiotics [Wang et al. (1996) Biochem. 35:12338] and hormones. RNAs can even be selected to bind to other RNAs through non-Watson-Crick interactions [Schmidt et al. (1996) Ann. N.Y. Acad. Sci. 782:526]. A ligand RNA may be used to either inactivate or enhance the activity of a molecule to which it binds. Any RNA segment identified through such a process may also be produced by the methods of the present invention, so that the observation of the activity of the RNA ligand may be used as a specific sign of the presence of the target material in the invasive cleavage reaction. The ligand binding to its specific partner may also be used as another way of capturing a readout signal to a solid support.
The product RNA might also be designed to have a catalytic function, e.g., to act as a ribozyme, allowing cleavage another molecule to be indicative of the success of the primary invasive cleavage reaction [Uhlenbeck (1987) Nature 328:596]. In yet another embodiment, the RNA may be made to encode a peptide sequence. When coupled to an in vitro translation system (e.g., the S-30 system derived from E. coli [Lesley (1985) Methods Mol. Biol. 37:265] or a rabbit reticulocyte lysate system [Dasso and Jackson (1989) Nucleic Acids Res. 17:3129], available from Promega), the production of the appropriate protein may be detected. In a preferred embodiment, the proteins produced include those that allow either calorimetric or luminescent detection, such as beta-galactosidase (lac-Z) or luciferase, respectively.
The above discussion focused on the use of the present transcription visualization methods in the context of the InvaderTM-directed cleavage assay (i.e., the non-target cleavage products produced in the InvaderTM assay were used to complete and activate a protein binding region, such as a promoter region). However, the transcription visualization methods are not limited to this context. Any assay which produces an oligonucleotide product having relatively discrete ends can be used in conjunction with the present transcription visualization methods. For example, the homogenous assay described in U.S. Pat. No. 5,210,015, particularly when conducted under conditions where polymerization cannot occur, produces short oligonucleotide fragments as the result of cleavage of a probe. If this assay is conducted under conditions where polymerase occurs, the site of cleavage of the probe may be focused through the use of nucleotide analogs that have uncleavable linkages at particular positions within the probe. These short oligonucleotides can be employed in a manner analogous to the cut probe or non-target cleavage products produced in the invasive cleavage reactions of the present invention. Additional assays which generate suitable oligonucleotide products are known to the art. For example, the non-target cleavage products produced in assays such as the “Cycling Probe Reaction” (Duck et al., BioTech., 9:142 [1990] and U.S. Pat. Nos. 4,876,187 and 5,011,769), in which shorter oligonucleotides are released from longer oligonucleotides after hybridization to a target sequence would be suitable, as would short restriction fragments released in assays where a probe is designed to be cleaved when successfully hybridized to an appropriate restriction recognition sequence (U.S. Pat. No. 4,683,194).
Assays which generate short oligonucleotides having “ragged” (i.e., not discrete) 3′ ends can also be employed with success in the transcription reactions of the present invention when the oligonucleotide provided by this non-transcription reaction are used to provide a portion of the promoter region located downstream of the other oligonucleotide(s) which are required to complete the promoter region (that is a 3′ tail or unpaired extension can be tolerated when the oligo is being used as the “Cut Probe” is in FIGS. 94 and 95A).
IX. Signal Enhancement by Incorporating the Products of an Invasive Cleavage Reaction into a Subsequent Invasive Cleavage Reaction
As noted above, the oligonucleotide product released by the invasive cleavage can be used subsequently in any reaction or read-out method that uses oligonucleotides in the size range of a cleavage product. In addition to the reactions involving primer extension and transcription, described above, another enzymatic reaction that makes use of oligonucleotides is the invasive cleavage reaction. The present invention provide means of using the oligonucleotide released in a primary invasive cleavage reaction as a component to complete a cleavage structure to enable a secondary invasive cleavage reaction. One possible configuration of a primary cleavage reaction supplying a component for a secondary cleavage structure is diagrammed in FIG. 96. Is not intended that the sequential use of the invasive cleavage product be limited to a single additional step. It is contemplated that many distinct invasive cleavage reactions may be performed in sequence.
The polymerase chain reaction uses a DNA replication method to create copies of a targeted segment of nucleic acid at a logarithmic rate of accumulation. This is made possible by the fact that when the strands of DNA are separated, each individual strand contains sufficient information to allow assembly of a new complementary strand. When the new strands are synthesized the number of identical molecules has doubled. Within 20 iterations of this process, the original may be copied 1 million-fold, making very rare sequences easily detectable. The mathematical power of a doubling reaction has been incorporated into a number of amplification assays, several of which are cited in Table 1.
By performing multiple, sequential invasive cleavage reactions the method of the present invention captures an exponential mathematical advantage without producing additional copies of the target analyte. In a simple invasive cleavage reaction the yield, Y, is simply the turnover rate, K, multiplied by the time of the reaction, t [i.e., Y=(K)(t)]. If Y is used to represent the yield of a simple reaction, then the yield of a compound (i.e., a multiple, sequential reaction), assuming that each of the individual invasive cleavage steps has the same turnover rate, can be simply represented as Yn, where n is the number of invasive cleavage reactions that have been performed in the series. If the yields of each step differ the ultimate yield can be represented as the product of the multiplication of the yields of each individual reaction in the series. For example, if a primary invasive cleavage reaction can produce one thousand products in 30 minutes, and each of those products can in turn participate in 1000 additional reactions, there will be 10002 copies (1000×1000) of the ultimate product in a second reaction. If a third reaction is added to the series, then the theoretical yield will be 10003 (1000×1000×1000). In the methods of the present invention the exponent comes from the number of invasive cleavage reactions in the cascade. This can be contrasted to the amplification methods described above (e.g., PCR) in which Y is limited to 2 by the number of strands in duplex DNA, and the exponent n is the number of cycles performed, so that many iterations are necessary to accumulate large amounts of product.
To distinguish the exponential amplifications described above from those of the present invention, the former can be consider reciprocating reactions because the products the reaction feed back into the same reaction, e.g., event one leads to some number of events 2, and each event 2 leads back to some number of events 1. In contrast, the events of the present invention are sequential, e.g., event 1 leads to some number of events 2; each event 2 leads to some number of events 3, etc., and no event can contribute to an event earlier in the chain.
The sensitivity of the reciprocating methods is also one of the greatest weaknesses when these assays are used to determine if a target nucleic acid sequence is present or absent in a sample. Because the product of these reactions is detectable copy of the starting material, contamination of a new reaction with the products of an earlier reaction can lead to false positive results, i.e., the apparent detection of the target nucleic acid in samples that do not actually contain any of that target analyte. Furthermore, because the concentration of the product in each positive reaction is so high, amounts of DNA sufficient to create a strong false positive signal can be communicated to new reactions very easily either by contact with contaminated instruments or by aerosol. In contrast to the reciprocating methods, the most concentrated product of the sequential reaction, i.e., the product released in the ultimate invasive cleavage event, is not capable of initiating a like reaction or cascade if carried over to a fresh test sample. This is a marked advantage over the exponential amplification methods described above because the reactions of the present invention may be performed without the costly containment arrangements (e.g. either by specialized instruments or by separate laboratory space) required by any reciprocating reaction. While the products of a penultimate event may be inadvertently transferred to produce a background of the ultimate product in the absence of the a target analyte, the contamination would need to be of much greater volume to give an equivalent risk of a false positive result.
When the term sequential is used it is not intended to limit the invention to configurations in which that one invasive cleavage reaction or assay must be completed before the initiation of a subsequent reaction for invasive cleavage of a different probe. Rather, the term refers to the order of events as would occur if only single copies of each of the oligonucleotide species were used in an assay. The primary invasive cleavage reaction refers to that which occurs first, in response to the formation of the cleavage structure on the target nucleic acid. Subsequent reactions may be referred to as secondary, tertiary and so forth, and may involve artificial “target” strands that serve only to support assembly of a cleavage structure, and which are unrelated to the nucleic acid analyte of interest. While the complete assay may, if desired, be configured with each step of invasive cleavage separated either in space (e.g., in different reaction vessels) or in time (e.g., using a shift in reaction conditions, such as temperature, enzyme identity or solution condition, to enable the later cleavage events), it is also contemplated that all of the reaction components may be mixed so that secondary reactions may be initiated as soon as product from a primary cleavage becomes available. In such a format, primary, secondary and subsequent cleavage events involving different copies of the cleavage structures may take place simultaneously.
Several levels of this sort of linear amplification can be envisioned, in which each successive round of cleavage produces an oligonucleotide that can participate in the cleavage of a different probe in subsequent rounds. The primary reaction would be specific for the analyte of interest with secondary (and tertiary, etc.) reactions being used to generate signal while still being dependent on the primary reaction for initiation.
The released product may perform in several capacities in the subsequent reactions. One of the possible variations are shown in FIG. 96, in which the product of one invasive cleavage reaction becomes the InvaderTM oligonucleotide to direct the specific cleavage of another probe in a second reaction. In FIG. 96, the first invasive cleavage structure is formed by the annealing of the InvaderTM oligonucleotide (“Invader”) and the probe oligonucleotide (“Probe 1”) to the first target nucleic acid (“Target 1”). The target nucleic acid is divided into three regions based upon which portions of the InvaderTM and probe oligonucleotides are capable of hybridizing to the target (as discussed above and as shown in FIG. 25). Region 1 (region Y in FIG. 25) of the target has complementarity to only the InvaderTM oligonucleotide; region 3 (region Z in FIG. 25) of the target has complementarity to only the probe; and region 2 (region X in FIG. 25) of the target has complementarity to both the InvaderTM and probe oligonucleotides. It is noted that the sequential invasive cleavage reaction diagrammed in FIG. 96 employs an InvaderTM and a probe oligonucleotide; the sequential cleavage reaction is not limited to the use of such a first cleavage structure. The first cleavage structure in the sequential reaction may also employ an InvaderTM oligonucleotide, a mini probe and a stacker oligonucleotide as discussed in section V above.
In FIG. 96, cleavage of Probe 1 releases the “Cut Probe 1” (indicated by the hatched line in both the cleaved and uncleaved Probe 1 in FIG. 96). The released Probe 1 is then used as the InvaderTM oligonucleotide in second cleavage. The second cleavage structure is formed by the annealing of the Cut Probe 1, a second probe oligonucleotide (“Probe 2”) and a second target nucleic acid (“Target 2”). Probe 2 may be labelled (indicated by the star in FIG. 96) and detection of cleavage of the second cleavage structure may be accomplished by detecting the labelled cut Probe 2; the label may a radioisotope (e.g., 32P, 35S), a fluorophore (e.g., fluorescein), a reactive group capable of detection by a secondary agent (e.g., biotin/streptavidin), a positively charged adduct which permits detection by selective charge reversal (as discussed in section IV above), etc. Alternatively, the cut Probe 2 may used in a tailing reaction (as discussed in section VI above) or may used to complete or activate a protein binding site (as discussed in section VIII above).
Another possible configuration for performing a sequential invasive cleavage reaction is diagrammed in FIG. 97. In this case, probe oligonucleotides that are cleaved in the primary reaction can be designed to fold back on themselves (i.e., they contain a region of self-complementarity) to create a molecule that can serve as both the target and InvaderTM oligonucleotide (termed here an “IT” complex). The IT complex then enables cleavage of a different probe present in the secondary reaction. Inclusion of an excess of the secondary probe molecule (“Probe 2”), allows each IT molecule to serve as the platform for the generation of multiple copies of cleaved secondary probe. In FIG. 97, the regions of self-complementarity contained within the 5′ portion of the InvaderTM oligonucleotide is indicated by the hatched ovals; the arrow between these two ovals indicates that these two regions can self-pair (as shown in the “Cut Probe 1”). The target nucleic acid is divided into three regions based upon which portions of the InvaderTM and probe oligonucleotides are capable of hybridizing to the target (as discussed above and it is noted that the target may be divided into four regions if a stacker oligonucleotide is employed). The second cleavage structure is formed by the annealing of the second probe (“Probe 2”) to the fragment of Probe 1 (“Cut Probe 1”) that was released by cleavage of the first cleavage structure. The Cut Probe 1 forms a hairpin or stem/loop structure near its 3′ terminus by virtue of the annealing of the regions of self-complementarity contained within Cut Probe 1 (this self-annealed Cut Probe 1 forms the IT complex). The IT complex (Cut Probe 1) is divided into three regions. Region 1 of the IT complex has complementarity to the 3′ portion of Probe 2; region 2 has complementarity to both the 3′ end of Cut Probe 1 and to the 5′ portion of Probe 2 (analogous to the region of overlap “X” shown in FIG. 25); and region 3 contains the region of self-complementarity (i.e., region 3 is complementary to the 3′ portion of the Cut Probe 1). Note that with regard to the IT complex (i.e., Cut Probe 1), region 1 is located upstream of region 2 and region 2 is located upstream of region 3.
The cleavage products of the secondary invasive cleavage reaction (i.e., Cut Probe 2) can either be detected, or can in turn be designed to constitute yet another integrated InvaderTM-target complex to be used with a third probe molecule, again unrelated to the preceding targets.
The present invention is not limited to the configurations diagrammed in FIGS. 96 and 97. It is envisioned that the oligonucleotide product of a primary cleavage reaction may fill the role of any of the oligonucleotides described herein, e.g., it may serve as a target strand without an attached InvaderTM oligonucleotide-like sequence, or it may serve as a stacker oligonucleotide, as described above, to enhance the turnover rate seen in the secondary reaction by stabilizing the probe hybridization through coaxial stacking.
In a preferred embodiment, each subsequent reaction is enabled by (i.e., is dependent upon) the product of the previous cleavage, so that the presence of the ultimate product may serve as an indicator of the presence of the target analyte. However, cleavage in the second reaction need not be dependent upon the presence of the product of the primary cleavage reaction; the product of the primary cleavage reaction may merely measurably enhance the rate of the second cleavage reaction.
In summary, the InvaderTM assay cascade (i.e., sequential invasive cleavage reactions) of the present invention is a combination of two or more linear assays that allow the accumulation of the ultimate product at an exponential rate, but without significant risk of carryover contamination.
The sequential invasive cleavage amplification of the present invention can be used as an intermediate boost to any of the detection methods, e.g., gel based analysis (standard or by charge reversal), polymerase tailing, and incorporation into a protein binding region, described herein. When used is such combinations the increased production of a specific cleavage product in the invasive cleavage assay reduces the burdens of sensitivity and specificity on the read-out systems, thus facilitating their use.
In addition to enabling a variety of detection platforms, the cascade strategy is suitable for multiplex analysis of individual analytes (i.e., individual target nucleic acids) in a single reaction. The multiplex format can be categorized into two types. In one case, it is desirable to know the identity (and amount) of each of the analytes that can be present in a clinical sample, or the identity of each of the analytes as well as an internal control. To identify the presence of multiple individual analytes in a single sample, several distinct secondary amplification systems may be included. Each probe cleaved in response to the presence of a particular target sequence (or internal control) can be designed to trigger a different cascade coupled to different detectable moieties, such as different sequences to be extended by DNA polymerase or different dyes in an FET format. The contribution of each specific target sequence to final product can thereby be tallied, allowing quantitative detection of different genes or alleles in a sample containing a mixture of genes/alleles.
In the second configuration, it is desirable to determine if any of several analytes are present in a sample, but the exact identity of each does not need to be known. For example, in blood banking it is desirable to know if any one of a host of infectious agents is present in a sample of blood. Because the blood is discarded regardless of which agent is present, different signals on the probes would not be required in such an application of the present invention, and may actually be undesirable for reasons of confidentiality. In this case, the 5′ arms (i.e., the 5′ portion which will be released upon cleavage) of the different analyte-specific probes would be identical and would therefore trigger the same secondary signal cascade. A similar configuration would permit multiple probes complementary to a single gene to be used to boost the signal from that gene or to ensure inclusivity when there are numerous alleles of a gene to be detected.
In the primary InvaderTM reaction, there are two potential sources of background. The first is from InvaderTM-independent cleavage of probe annealed to the target, to itself, or to one of the other oligonucleotides present in the reaction. It can be seen by consideration of FIGS. 96 and 97 that the probes of the primary cleavage reactions depicted are designed to have regions of complementarity to the other oligonucleotides involved in the subsequent reactions, and, as depicted in FIG. 97, to other regions of the same molecule. The use of an enzyme that cannot efficiently cleave a structure that lacks a primer (e.g., that cannot cleave the structures diagrammed in FIG. 16A or 16D) is preferred for this reason. As shown in FIGS. 99 and 100, the enzyme Pfu FEN-1 gives no detectable cleavage in the absence of the upstream oligonucleotide or even in the presence of an upstream oligonucleotide that fails to invade the probe-target complex. This indicates that the Pfu FEN-1 endonuclease is a suitable enzyme for use in the methods of the present invention.
Other structure-specific nucleases may be suitable as a well. As discussed in the first example, some 5′ nucleases can be used in conditions that significantly reduce this primer-independent cleavage. For example, it has been shown that when the 5′ nuclease of DNAPTaq is used to cleave hairpins the primer-independent cleavage is markedly reduced by the inclusion of a monovalent salt in the reaction [Lyamichev, et al. (1993), supra].
Test For InvaderTM Oligonucleotide-Independent Cleavage
A simple test can be performed for any enzyme in combination with any reaction buffer to gauge the amount of InvaderTM oligonucleotide-independent cleavage to be expected from that combination. A small hairpin-like test molecule that can be used with or without a primer hybridized to a 3′ arm, the S-60 molecule, is depicted in FIG. 30. The S-60 and the oligonucleotide P15 are a convenient set of molecules for testing the suitability of an enzyme for application in the present invention and conditions for using these molecules are described in Example 11. Other similar hairpins may be used, of a cleavage structure may be assembled from separate oligonucleotides as diagrammed in FIGS. 99a-e. Reactions using these structures to examine the activity of the Pfu FEN-1 enzyme in the presence or absence of an upstream overlapping oligonucleotide are described in Example 45 and the results are displayed in FIG. 100. To test any particular combination of enzyme and cleavage conditions, similar reactions can be assembled. Outside of the variables of reaction conditions to be tested for any particular enzyme (e.g., salt sensitivities, divalent cation requirements) the test reactions should accommodate any known limitations of the test enzyme. For example, the test reactions should be performed at a temperature that is within the operating temperature range of the candidate enzyme, if known.
It is not necessary that multiple lengths of overlap be demonstrated for each candidate enzyme, but the activity of the enzyme in the absence of an upstream oligonucleotide (as shown in FIG. 99a) and in the presence of an oligonucleotide that does not overlap (FIG. 99b) should be assessed. It is preferable that structures lacking an upstream oligonucleotide be cleaved at less than one half of the rate seen in the presence of an upstream overlapping oligonucleotide. It is more preferable that these structures be cleaved at less than about on tenth the rate of the invasive cleavage structure. It is most preferred that cleavage of these structures occur at less than one percent the rate of the invasive cleavage structure.
If the cleaved product is to serve as an upstream oligonucleotide in a subsequent cleavage reaction, as diagrammed in FIG. 96, the most rapid reaction will be achieved if the other components of the second cleavage structure (i.e., Target 2 and Probe 2 in FIG. 96) are provided in excess so that cleavage may proceed immediately after the upstream oligonucleotide (i.e, Cut Probe 1 in FIG. 96) is made available. To provide an abundance of the second target strand (Target 2 in FIG. 96) one may use an isolated natural nucleic acid, such as bacteriophage M13 DNA, or one may use a synthetic oligonucleotide. If a synthetic oligonucleotide is chosen as the second target sequence, the sequence employed must be examined for regions of self-complementarity (similar considerations apply to short isolated natural nucleic acids such as restriction enzyme fragments or PCR products; natural nucleic acid targets whose 3′ end is located≧100 nucleotides downstream of the probe binding site on the target strand are sufficiently long enough to obviate the design considerations discussed below). Specifically, it must be determined that the 3′ end of the synthetic oligonucleotide may not hybridize to the target strand (i.e., intra-strand hybridization) upstream of the probe, triggering unintended cleavage. Simple examination of the sequence of the synthetic oligonucleotide should reveal if the 3′ end has sufficient complementarity to the region of the target upstream of the probe binding site to pose a problem (i.e, it would reveal whether the synthetic oligonucleotide can form a hairpin at its 3′ end which could act as an invading oligonucleotide to cause cleavage of the probe in the absence of the hybridization of the intended InvaderTM oligonucleotide, i.e., the cleavage product from the first invasive cleavage reaction). If 3 or more of the last 4 to 7 nucleotides (the 3′ terminal region) of the synthetic target can basepair upstream of the probe such that there is an invasion into the probe-target duplex, or such that the duplexes formed by the synthetic target strand with its own 3′ terminal region and with the probe abut without a gap and the 3′ terminal region has an additional 1 or 2 nucleotides unpaired at the extreme 3′ end of the synthetic target, then the sequence of the synthetic target oligonucleotide should be modified. The sequence may be changed to disrupt the interaction of the 3′ terminal region or to increase the distance between the probe binding site and the regions to which the 3′ terminus is binding. Alternatively, the 3′ end may be modified to reduce its ability to direct cleavage, e.g., by adding a 3′ phosphate during synthesis (see Ex. 35, Table 3) or by adding several additional nucleotides that will not basepair in a self-complementary manner (i.e., they will not participate in the formation of a hairpin structure).
When the product of a first invasive cleavage reaction is designed to form a target which can fold on itself to direct cleavage of a second probe, the IT complex as diagrammed in FIG. 97, the design of the sequence used to form the stem/loop of the IT complex must be considered. To be factored into the design of such a probe are 1) the length of the region of self-complementarity, 2) the length of the region of overlap (region “X” in FIG. 25) and 3) the stability of the hairpin or stem/loop structure as predicted by both Watson-Crick base pairing and by the presence or absence of a particularly stable loop sequence [e.g., a tetraloop (Tinoco et al., supra) or a triloop (Hirao et al., supra)]. It is desirable that this sequence have nucleotides that can base pair (intrastrand), so that the second round of invasive cleavage may occur, but that the structure not be so strong that its presence will prevent the cleavage of the probe in the primary reaction (i.e., Probe 1 in FIG. 96). As shown herein, the presence of a secondary structure in the 5′ arm of a cleavage structure cleaved by a structure-specific nuclease may inhibit cleavage by some structure-specific nucleases (Ex. 1).
The length of the region of self-complementarity within Probe 1 determines the length of the region of the duplex upstream of Probe 2 in the second cleavage structure (see FIG. 97). Different enzymes have different length requirements for this duplex to effect invasive cleavage efficiently. For example, the Pfu FEN-1 and Mja FEN-1 enzymes have been tested for the effect of this duplex length using the set of target/invader oligonucleotide molecules depicted in FIG. 98 (i.e., SEQ ID NOS:118, 119, 147-151). The invasive cleavage reactions were performed as described in Example 38, using 1 pM IT3 (SEQ ID NO:118), 2 μM probe PR1 (SEQ ID NO:119) for 5 min, and the rates of cleavage are shown in Table 2.









TABLE 2





Length of
Pfu FEN-1 Turnover,
Mja FEN-1 Turnover,


Duplex
per min.
per min.







0
 0
 0


3
 1
29


4
10
57


6
44
51


8
45
46









The data shown in Table 2 demonstrate that the Pfu FEN-1 enzyme can be used with stems of 3 or 4 bases, but that the rate of cleavage is maximized when the stem is greater than 4 basepairs in length. Table 2 shows that the Mja FEN-1 enzyme can cleave efficiently using shorter stems; however, as this enzyme can also cleave a probe in the absence of an upstream oligonucleotide, Mja FEN-1 is not preferred for use in the methods of the present invention.
A similar test can be performed using any candidate enzyme to determine how much self-complementarity may be designed into the Probe 1. The use of a shorter stem means that the overall probe may be shorter. This is beneficial because shorter probes are less costly to synthesize, and because shorter probes will have fewer sequences that might form unintended intrastrand structures. In assessing the activity of a candidate enzyme on the structures such as those shown in FIG. 98 it is not required that the stem length chosen allow the maximum rate of cleavage to occur. For example, in considering the case of Pfu FEN-1, the advantages of using a 4 basepair stem (e.g., cost or sequence limitations), with a cleavage rate of 10 cleavages per minute, may outweigh the rate advantage of using a longer 6 basepair stem (44 cleavages/min.), in the context of a particular experiment. It is within the scope of the present invention that some elements chosen for use in the assay be sub-optimal for performance of that particular element, if the use of a sub-optimal design benefits the objectives of that particular experiment as a whole.
In designing oligonucleotides to be employed as a probe that once cleaved forms a stem-loop structure as diagrammed in FIG. 97 (i.e., Probe 1 in FIG. 97), it has been found that the stability of the loop is not a factor in the efficiency of cleavage of either Probe 1 or Probe 2. Loops tested have included stable triloops, loops of 3 and 4 nucleotides that were not predicted to be particularly stable (i.e., the stability is determined by the duplex sequence and not by additional stabilizing interactions within the loop), and large loops of up to about 25 nucleotides.
X. Detection of Human Cytomegalovirus Viral DNA by Invasive Cleavage
Human cytomegalovirus (HCMV) causes, or is associated with, a wide variety of diseases in humans (Table 3). More than 90% of bone marrow or kidney transplant recipients (immunocompromised hosts) develop HCMV infections, most of which are due to reactivation of latent virus by immunosuppressive drugs, as well as transmission of virus by latently infected donor tissue or blood [Ackerman et al. (1988) Transplant. Proc. 20(S1):468 and Peterson et al. (1980) Medicine 59:283].







TABLE 3





Diseases Caused By Human Cytomegalovirus







cytomegalic inclusion


heterophil-negative disease in neonates


mononucleosis


interstitial pneumonia


pneumonitis


retinitis


hepatitis


pancreatitis


meningoencephalitis


gastrointestinal disease


disseminated infection









There are instances in which rapid, sensitive, and specific diagnosis of HCMV disease is imperative. In recent years, the number of patients undergoing organ and tissue transplantations has increased markedly. HCMV is the most frequent cause of death in immunocompromised transplant recipients, thereby confirming the need for rapid and reliable laboratory diagnosis. Lymphocytes, monocytes, and possibly arterial endothelial or smooth muscle cells, are sites of HCMV latency. Therefore, prevention of HCMV infections in immunocompromised individuals (e.g., transplant recipients) includes use of HCMV-negative blood products and organs. Additionally, HCMV can be spread transplacentally, and to newborns by contact with infected cervical secretions during birth. Thus, a rapid, sensitive, and specific assay for detecting HCMV in body fluids or secretions may be desirable as a means to monitor infection, and consequently, determine the necessity of cesarean section.
Diagnosis of HCMV infection may be performed by conventional cell culture using human fibroblasts; shell vial centrifugation culture utilizing monoclonal antibodies and immunofluorescent staining techniques; serological methods; the HCMV antigenemia assay which employs a monoclonal antibody to detect HCMV antigen in peripheral blood leukocytes (PBLs); or by nucleic acid hybridization assays. These various methods have their advantages and limitations. Conventional cell culture is sensitive but slow, as cytopathic effect (CPE) may take 30 or more days to develop. Shell vial centrifugation is more rapid but still requires 24-48 hours for initial results. Both culture methods are affected by antiviral therapy. In immunocompromised patients, the ability to mount IgG and/or IgM antibody responses to HCMV infection are impaired, and serological methods are thus not reliable in this setting. Alternatively, IgM antibodies may be persistent for months after infection is resolved, and thus their presence may not be indicative of active infection. The HCMV antigenemia assay is labor intensive and is not applicable to specimens other than PBLs.
Recent advances in molecular biology have spurred the use of DNA probes in attempts to provide a more rapid, sensitive and specific assay for detecting HCMV in clinical specimens. For example, radiolabeled DNA probes have been used to hybridize to tissue cultures infected with or by HCMV, or in clinical samples suspected of containing HCMV (“hybridization assays”). However, probing of tissue cultures requires at least 18-24 hours for growth to amplify the antigen (HCMV) to be detected, if present, and additional time for development of autoradiographic detection systems. Using hybridization assays for assaying clinical specimens for HCMV may lack sensitivity, depending upon the titer of virus and the clinical sample assayed. Detection of HCMV in clinical samples has been reported using the polymerase chain reaction (PCR) to enzymatically amplify HCMV DNA. Methods using PCR compare favorably with virus isolation, in situ hybridization assays, and Southern blotting [See, e.g., Bamborschke et al. (1992) J. Neurol. 239:205; Drouet et al. (1993) J. Virol. Methods 45:259; Einsele et al. (1991) Blood 77:1104-1110; Einsele et al. (1991) Lancet 338:1170; Lee et al. (1992) Aust. NZ J. Med. 22:249; Miller et al. (1994) J. Clin. Microbiol. 32:5; Rowley et al. (1991) Transplant. 51:1028; Spector et al. (1992) J. Clin. Microbiol. 30:2359; and Stanier et al. (1992) Molec. Cell. Probes 8:51]. Others, comparing the HCMV antigenemia assay with PCR methods, have found PCR methods as efficient or slightly more efficient in the detection of HCMV (van Dorp et al. (1992) Transplant. 54:661; Gerna et al. (1991) J. Infect. Dis. 164:488; Vleiger et al. (1992) Bone Marrow Transplant. 9:247; Zipeto et al. (1992) J. Clin. Microbiol. 30:527]. In addition, PCR methods have exhibited great sensitivity when specimens other than PBLs are assayed [Natori et al. (1993) Kansenshogaku Zasshi 67:1011; Peterson et al. (1980) Medicine 59:283; Prosch et al. (1992) J. Med. Virol. 38:246; Ratnamohan et al. (1992) J. Med. Virol. 38:252]. However, because of the dangers of false positive reactions, these PCR-based procedures require rigid controls to prevent contamination and carry over [Ehrlich et al. (1994) in PCR-Based Diagnostics in Infectious Diseases, G D Ehrlich and S J Greenberg (eds), Blackwell Scientific Publications, pp.3-18]. Therefore, there exists a need for a rapid, sensitive, and specific assay for HCMV that has a reduced risk of false positive result due to contamination by reaction product carried over from other samples.
As shown herein, the InvaderTM-directed cleavage assay is rapid, sensitive and specific. Because the accumulated products do not contribute to the further accumulation of signal, reaction products carried over from one standard (i.e., non-sequential) InvaderTM-directed cleavage assay to another cannot promote false positive results. The use of multiple sequential InvaderTM-directed cleavage assays will further boost the sensitivity of HCMV detection without sacrifice of these advantages.
EXPERIMENTAL
The following examples serve to illustrate certain preferred embodiments and aspects of the present invention and are not to be construed as limiting the scope thereof.
In the disclosure which follows, the following abbreviations apply:°C. (degrees Centigrade); fig (figure); g (gravitational field); hr (hour); min (minute); oligo (oligonucleotide); rxn (reaction); vol (volume); w/v (weight to volume); v/v (volume to volume); BSA (bovine serum albumin); CTAB (cetyltrimethylammonium bromide); HPLC (high pressure liquid chromatography); DNA (deoxyribonucleic acid); p (plasmid); μl (microliters); ml (milliliters); μg (micrograms); pmoles (picomoles); mg (milligrams); M (molar); mM (milliMolar); μM (microMolar); nm (nanometers); kdal (kilodaltons); OD (optical density); EDTA (ethylene diamine tetra-acetic acid); FITC (fluorescein isothiocyanate); SDS (sodium dodecyl sulfate); NaPO4 (sodium phosphate); NP-40 (Nonidet P-40); Tris (tris(hydroxymethyl)-aminomethane); PMSF (phenylmethylsulfonylfluoride); TBE (Tris-Borate-EDTA, i.e., Tris buffer titrated with boric acid rather than HCl and containing EDTA); PBS (phosphate buffered saline); PPBS (phosphate buffered saline containing 1 mM PMSF); PAGE (polyacrylamide gel electrophoresis); Tween (polyoxyethylene-sorbitan); Ambion (Ambion, Inc., Austin, Tex.); Boehringer (Boehringer Mannheim Biochemical, Indianapolis, Ind.); Dynal (Dynal A. S., Oslo, Norway); Epicentre (Epicentre Technologies, Madison, Wis.); MJ Research (MJ Research, Watertown, Mass.); National Biosciences (Plymouth, Minn.); New England Biolabs (Beverly, Mass.); Novagen (Novagen, Inc., Madison, Wis.); Perkin Elmer (Norwalk, Conn.); Promega (Promega Corp., Madison, Wis.); Stratagene (Stratagene Cloning Systems, La Jolla, Calif.); USB (U.S. Biochemical, Cleveland, Ohio).
EXAMPLE 1 
Characteristics of Native Thermostable DNA Polymerases
A. 5′ Nuclease Activity Of DNAPTaq
During the polymerase chain reaction (PCR) [Saiki et al., Science 239:487 (1988); Mullis and Faloona, Methods in Enzymology 155:335 (1987)], DNAPTaq is able to amplify many, but not all, DNA sequences. One sequence that cannot be amplified using DNAPTaq is shown in FIG. 5 (Hairpin structure is SEQ ID NO:15, FIG. 5 also shows a primer: SEQ ID NO:17.) This DNA sequence has the distinguishing characteristic of being able to fold on itself to form a hairpin with two single-stranded arms, which correspond to the primers used in PCR.
To test whether this failure to amplify is due to the 5′ nuclease activity of the enzyme, we compared the abilities of DNAPTaq and DNAPStf to amplify this DNA sequence during 30 cycles of PCR. Synthetic oligonucleotides were obtained from The Biotechnology Center at the University of Wisconsin-Madison. The DNAPTaq and DNAPStf were from Perkin Elmer (i.e., AmplitaqTM DNA polymerase and the Stoffel fragment of AmplitaqTM DNA polymerase). The substrate DNA comprised the hairpin structure shown in FIG. 6 cloned in a double-stranded form into pUC19. The primers used in the amplification are listed as SEQ ID NOS:16-17. Primer SEQ ID NO:17 is shown annealed to the 3′ arm of the hairpin structure in FIG. 5. Primer SEQ ID NO:16 is shown as the first 20 nucleotides in bold on the 5′ arm of the hairpin in FIG. 5.
Polymerase chain reactions comprised 1 ng of supercoiled plasmid target DNA, 5 pmoles of each primer, 40 μM each dNTP, and 2.5 units of DNAPTaq or DNAPStf, in a 50 μl solution of 10 mM Tris.Cl pH 8.3. The DNAPTaq reactions included 50 mM KCl and 1.5 mM MgCl2. The temperature profile was 95° C. for 30 sec., 55° C. for 1 min. and 72° C. for 1 min., through 30 cycles. Ten percent of each reaction was analyzed by gel electrophoresis through 6% polyacrylamide (cross-linked 29:1) in a buffer of 45 mM Tris.Borate, pH 8.3, 1.4 mM EDTA.
The results are shown in FIG. 6. The expected product was made by DNAPStf (indicated simply as “S”) but not by DNAPTaq (indicated as “T”). We conclude that the 5′ nuclease activity of DNAPTaq is responsible for the lack of amplification of this DNA sequence.
To test whether the 5′ unpaired nucleotides in the substrate region of this structured DNA are removed by DNAPTaq, the fate of the end-labeled 5′ arm during four cycles of PCR was compared using the same two polymerases (FIG. 7). The hairpin templates, such as the one described in FIG. 5, were made using DNAPStf and a 32P-5′-end-labeled primer. The 5′-end of the DNA was released as a few large fragments by DNAPTaq but not by DNAPStf. The sizes of these fragments (based on their mobilities) show that they contain most or all of the unpaired 5′ arm of the DNA. Thus, cleavage occurs at or near the base of the bifurcated duplex. These released fragments terminate with 3′ OH groups, as evidenced by direct sequence analysis, and the abilities of the fragments to be extended by terminal deoxynucleotidyl transferase.
FIGS. 8-10 show the results of experiments designed to characterize the cleavage reaction catalyzed by DNAPTaq. Unless otherwise specified, the cleavage reactions comprised 0.01 pmoles of heat-denatured, end-labeled hairpin DNA (with the unlabeled complementary strand also present), 1 pmole primer (complementary to the 3′ arm) and 0.5 units of DNAPTaq (estimated to be 0.026 pmoles) in a total volume of 10 μl of 10 mM Tris-Cl, ph 8.5, 50 mM KCl and 1.5 mM MgCl2. As indicated, some reactions had different concentrations of KCl, and the precise times and temperatures used in each experiment are indicated in the individual figures. The reactions that included a primer used the one shown in FIG. 5 (SEQ ID NO:17). In some instances, the primer was extended to the junction site by providing polymerase and selected nucleotides.
Reactions were initiated at the final reaction temperature by the addition of either the MgCl2 or enzyme. Reactions were stopped at their incubation temperatures by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. The Tm calculations listed were made using the OligoTM primer analysis software from National Biosciences, Inc. These were determined using 0.25 μM as the DNA concentration, at either 15 or 65 mM total salt (the 1.5 mM MgCl2 in all reactions was given the value of 15 mM salt for these calculations).
FIG. 8 is an autoradiogram containing the results of a set of experiments and conditions on the cleavage site. FIG. 8A is a determination of reaction components that enable cleavage. Incubation of 5′-end-labeled hairpin DNA was for 30 minutes at 55° C., with the indicated components. The products were resolved by denaturing polyacrylamide gel electrophoresis and the lengths of the products, in nucleotides, are indicated. FIG. 8B describes the effect of temperature on the site of cleavage in the absence of added primer. Reactions were incubated in the absence of KCl for 10 minutes at the indicated temperatures. The lengths of the products, in nucleotides, are indicated.
Surprisingly, cleavage by DNAPTaq requires neither a primer nor dNTPs (see FIG. 8A). Thus, the 5′ nuclease activity can be uncoupled from polymerization. Nuclease activity requires magnesium ions, though manganese ions can be substituted, albeit with potential changes in specificity and activity. Neither zinc nor calcium ions support the cleavage reaction. The reaction occurs over a broad temperature range, from 25° C. to 85° C., with the rate of cleavage increasing at higher temperatures.
Still referring to FIG. 8, the primer is not elongated in the absence of added dNTPs. However, the primer influences both the site and the rate of cleavage of the hairpin. The change in the site of cleavage (FIG. 8A) apparently results from disruption of a short duplex formed between the arms of the DNA substrate. In the absence of primer, the sequences indicated by underlining in FIG. 5 could pair, forming an extended duplex. Cleavage at the end of the extended duplex would release the 11 nucleotide fragment seen on the FIG. 8A lanes with no added primer. Addition of excess primer (FIG. 8A, lanes 3 and 4) or incubation at an elevated temperature (FIG. 8B) disrupts the short extension of the duplex and results in a longer 5′ arm and, hence, longer cleavage products.
The location of the 3′ end of the primer can influence the precise site of cleavage. Electrophoretic analysis revealed that in the absence of primer (FIG. 8B), cleavage occurs at the end of the substrate duplex (either the extended or shortened form, depending on the temperature) between the first and second base pairs. When the primer extends up to the base of the duplex, cleavage also occurs one nucleotide into the duplex. However, when a gap of four or six nucleotides exists between the 3′ end of the primer and the substrate duplex, the cleavage site is shifted four to six nucleotides in the 5′ direction.
FIG. 9 describes the kinetics of cleavage in the presence (FIG. 9A) or absence (FIG. 9B) of a primer oligonucleotide. The reactions were run at 55° C. with either 50 mM KCl (FIG. 9A) or 20 mM KCl (FIG. 9B). The reaction products were resolved by denaturing polyacrylamide gel electrophoresis and the lengths of the products, in nucleotides, are indicated. “M”, indicating a marker, is a 5′ end-labeled 19-nt oligonucleotide. Under these salt conditions, FIGS. 9A and 9B indicate that the reaction appears to be about twenty times faster in the presence of primer than in the absence of primer. This effect on the efficiency may be attributable to proper alignment and stabilization of the enzyme on the substrate.
The relative influence of primer on cleavage rates becomes much greater when both reactions are run in 50 mM KCl. In the presence of primer, the rate of cleavage increases with KCl concentration, up to about 50 mM. However, inhibition of this reaction in the presence of primer is apparent at 100 mM and is complete at 150 mM KCl. In contrast, in the absence of primer the rate is enhanced by concentration of KCl up to 20 mM, but it is reduced at concentrations above 30 mM. At 50 mM KCl, the reaction is almost completely inhibited. The inhibition of cleavage by KCl in the absence of primer is affected by temperature, being more pronounced at lower temperatures.
Recognition of the 5′ end of the arm to be cut appears to be an important feature of substrate recognition. Substrates that lack a free 5′ end, such as circular M13 DNA, cannot be cleaved under any conditions tested. Even with substrates having defined 5′ arms, the rate of cleavage by DNAPTaq is influenced by the length of the arm. In the presence of primer and 50 mM KCl, cleavage of a 5′ extension that is 27 nucleotides long is essentially complete within 2 minutes at 55° C. In contrast, cleavages of molecules with 5′ arms of 84 and 188 nucleotides are only about 90% and 40% complete after 20 minutes. Incubation at higher temperatures reduces the inhibitory effects of long extensions indicating that secondary structure in the 5′ arm or a heat-labile structure in the enzyme may inhibit the reaction. A mixing experiment, run under conditions of substrate excess, shows that the molecules with long arms do not preferentially tie up the available enzyme in non-productive complexes. These results may indicate that the 5′ nuclease domain gains access to the cleavage site at the end of the bifurcated duplex by moving down the 5′ arm from one end to the other. Longer 5′ arms would be expected to have more adventitious secondary structures (particularly when KCl concentrations are high), which would be likely to impede this movement.
Cleavage does not appear to be inhibited by long 3′ arms of either the substrate strand target molecule or pilot nucleic acid, at least up to 2 kilobases. At the other extreme, 3′ arms of the pilot nucleic acid as short as one nucleotide can support cleavage in a primer-independent reaction, albeit inefficiently. Fully paired oligonucleotides do not elicit cleavage of DNA templates during primer extension.
The ability of DNAPTaq to cleave molecules even when the complementary strand contains only one unpaired 3′ nucleotide may be useful in optimizing allele-specific PCR. PCR primers that have unpaired 3′ ends could act as pilot oligonucleotides to direct selective cleavage of unwanted templates during preincubation of potential template-primer complexes with DNAPTaq in the absence of nucleoside triphosphates.
B. 5′ Nuclease Activities Of Other DNAPs
To determine whether other 5′ nucleases in other DNAPs would be suitable for the present invention, an array of enzymes, several of which were reported in the literature to be free of apparent 5′ nuclease activity, were examined. The ability of these other enzymes to cleave nucleic acids in a structure-specific manner was tested using the hairpin substrate shown in FIG. 5 under conditions reported to be optimal for synthesis by each enzyme.
DNAPEcl and DNAP Klenow were obtained from Promega Corporation; the DNAP of Pyrococcus furious [“Pfu”, Bargseid et al., Strategies 4:34 (1991)] was from Stratagene; the DNAP of Thermococcus litoralis [“Tli”, VentTM (exo-), Perler et al., Proc. Natl. Acad. Sci. USA 89:5577 (1992)] was from New England Biolabs; the DNAP of Thermus flavus [“Tfl”, Kaledin et al., Biokhimiya 46:1576 (1981)] was from Epicentre Technologies; and the DNAP of Thermus thermophilus [“Tth”, Carballeira et al., Biotechniques 9:276 (1990); Myers et al., Biochem. 30:7661 (1991)] was from U.S. Biochemicals.
0.5 units of each DNA polymerase was assayed in a 20 μl reaction, using either the buffers supplied by the manufacturers for the primer-dependent reactions, or 10 mM Tris.Cl, pH 8.5, 1.5 mM MgCl2, and 20 mM KCl. Reaction mixtures were at held 72° C. before the addition of enzyme.
FIG. 10 is an autoradiogram recording the results of these tests. FIG. 10A demonstrates reactions of endonucleases of DNAPs of several thermophilic bacteria. The reactions were incubated at 55° C. for 10 minutes in the presence of primer or at 72° C. for 30 minutes in the absence of primer, and the products were resolved by denaturing polyacrylamide gel electrophoresis. The lengths of the products, in nucleotides, are indicated. FIG. 10B demonstrates endonucleolytic cleavage by the 5′ nuclease of DNAPEcl. The DNAPEcl and DNAP Klenow reactions were incubated for 5 minutes at 37° C. Note the light band of cleavage products of 25 and 11 nucleotides in the DNAPEcl lanes (made in the presence and absence of primer, respectively). FIG. 8A also demonstrates DNAPTaq reactions in the presence (+) or absence (−) of primer. These reactions were run in 50 mM and 20 mM KCl, respectively, and were incubated at 55° C. for 10 minutes.
Referring to FIG. 10A, DNAPs from the eubacteria Thermus thermophilus and Thermus flavus cleave the substrate at the same place as DNAPTaq, both in the presence and absence of primer. In contrast, DNAPs from the archaebacteria Pyrococcus furiosus and Thermococcus litoralis are unable to cleave the substrates endonucleolytically. The DNAPs from Pyrococcus furious and Thermococcus litoralis share little sequence homology with eubacterial enzymes (Ito et al., Nucl. Acids Res. 19:4045 (1991); Mathur et al., Nucl. Acids. Res. 19:6952 (1991); see also Perler et al.). Referring to FIG. 10B, DNAPEcl also cleaves the substrate, but the resulting cleavage products are difficult to detect unless the 3′ exonuclease is inhibited. The amino acid sequences of the 5′ nuclease domains of DNAPEcl and DNAPTaq are about 38% homologous (Gelfand, supra).
The 5′ nuclease domain of DNAPTaq also shares about 19% homology with the 5′ exonuclease encoded by gene 6 of bacteriophage T7 [Dunn et al., J. Mol. Biol. 166:477 (1983)]. This nuclease, which is not covalently attached to a DNAP polymerization domain, is also able to cleave DNA endonucleolytically, at a site similar or identical to the site that is cut by the 5′ nucleases described above, in the absence of added primers.
C. Transcleavage
The ability of a 5′ nuclease to be directed to cleave efficiently at any specific sequence was demonstrated in the following experiment. A partially complementary oligonucleotide termed a “pilot oligonucleotide” was hybridized to sequences at the desired point of cleavage. The non-complementary part of the pilot oligonucleotide provided a structure analogous to the 3′ arm of the template (see FIG. 5), whereas the 5′ region of the substrate strand became the 5′ arm. A primer was provided by designing the 3′ region of the pilot so that it would fold on itself creating a short hairpin with a stabilizing tetra-loop [Antao et al., Nucl. Acids Res. 19:5901 (1991)]. Two pilot oligonucleotides are shown in FIG. 11A. Oligonucleotides 19-12 (SEQ ID NO:18), 30-12 (SEQ ID NO:19) and 30-0 (SEQ ID NO:20) are 31, 42 or 30 nucleotides long, respectively. However, oligonucleotides 19-12 (SEQ ID NO:18) and 34-19 (SEQ ID NO:19) have only 19 and 30 nucleotides, respectively, that are complementary to different sequences in the substrate strand. The pilot oligonucleotides are calculated to melt off their complements at about 50° C. (19-12) and about 75° C. (30-12). Both pilots have 12 nucleotides at their 3′ ends, which act as 3′ arms with base-paired primers attached.
To demonstrate that cleavage could be directed by a pilot oligonucleotide, we incubated a single-stranded target DNA with DNAPTaq in the presence of two potential pilot oligonucleotides. The transcleavage reactions, where the target and pilot nucleic acids are not covalently linked, includes 0.01 pmoles of single end-labeled substrate DNA, 1 unit of DNAPTaq and 5 pmoles of pilot oligonucleotide in a volume of 20 μl of the same buffers. These components were combined during a one minute incubation at 95° C., to denature the PCR-generated double-stranded substrate DNA, and the temperatures of the reactions were then reduced to their final incubation temperatures. Oligonucleotides 30-12 and 19-12 can hybridize to regions of the substrate DNAs that are 85 and 27 nucleotides from the 5′ end of the targeted strand.
FIG. 19 shows the complete 206-mer sequence (SEQ ID NO:27). The 206-mer was generated by PCR. The M13/pUC 24-mer reverse sequencing (−48) primer and the M13/pUC sequencing (−47) primer from New England Biolabs (catalogue nos. 1233 and 1224 respectively) were used (50 pmoles each) with the pGEM3z(f+) plasmid vector (Promega Corp.) as template (10 ng) containing the target sequences. The conditions for PCR were as follows: 50 μM of each dNTP and 2.5 units of Taq DNA polymerase in 100 μl of 20 mM Tris-Cl, pH 8.3, 1.5 mM MgCl2, 50 mM KCl with 0.05% Tween-20 and 0.05% NP-40. Reactions were cycled 35 times through 95° C. for 45 seconds, 63° C. for 45 seconds, then 72° C. for 75 seconds. After cycling, reactions were finished off with an incubation at 72° C. for 5 minutes. The resulting fragment was purified by electrophoresis through a 6% polyacrylamide gel (29:1 cross link) in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA, visualized by ethidium bromide staining or autoradiography, excised from the gel, eluted by passive diffusion, and concentrated by ethanol precipitation.
Cleavage of the substrate DNA occurred in the presence of the pilot oligonucleotide 19-12 at 50° C. (FIG. 11B, lanes 1 and 7) but not at 75° C. (lanes 4 and 10). In the presence of oligonucleotide 30-12 cleavage was observed at both temperatures. Cleavage did not occur in the absence of added oligonucleotides (lanes 3, 6 and 12) or at about 80° C. even though at 50° C. adventitious structures in the substrate allowed primer-independent cleavage in the absence of KCl (FIG. 11B, lane 9). A non-specific oligonucleotide with no complementarity to the substrate DNA did not direct cleavage at 50° C., either in the absence or presence of 50 mM KCl (lanes 13 and 14). Thus, the specificity of the cleavage reactions can be controlled by the extent of complementarity to the substrate and by the conditions of incubation.
D. Cleavage Of RNA
A shortened RNA version of the sequence used in the transcleavage experiments discussed above was tested for its ability to serve as a substrate in the reaction. The RNA is cleaved at the expected place, in a reaction that is dependent upon the presence of the pilot oligonucleotide. The RNA substrate, made by T7 RNA polymerase in the presence of [α-32P]UTP, corresponds to a truncated version of the DNA substrate used in FIG. 11B. Reaction conditions were similar to those in used for the DNA substrates described above, with 50 mM KCl; incubation was for 40 minutes at 55° C. The pilot oligonucleotide used is termed 30-0 (SEQ ID NO:20) and is shown in FIG. 12A.
The results of the cleavage reaction is shown in FIG. 13B. The reaction was run either in the presence or absence of DNAPTaq or pilot oligonucleotide as indicated in FIG. 12B.
Strikingly, in the case of RNA cleavage, a 3′ arm is not required for the pilot oligonucleotide. It is very unlikely that this cleavage is due to previously described RNaseH, which would be expected to cut the RNA in several places along the 30 base-pair long RNA-DNA duplex. The 5′ nuclease of DNAPTaq is a structure-specific RNaseH that cleaves the RNA at a single site near the 5′ end of the heteroduplexed region.
It is surprising that an oligonucleotide lacking a 3′ arm is able to act as a pilot in directing efficient cleavage of an RNA target because such oligonucleotides are unable to direct efficient cleavage of DNA targets using native DNAPs. However, some 5′ nucleases of the present invention (for example, clones E, F and G of FIG. 4) can cleave DNA in the absence of a 3′ arm. In other words, a non-extendable cleavage structure is not required for specific cleavage with some 5′ nucleases of the present invention derived from thermostable DNA polymerases.
We tested whether cleavage of an RNA template by DNAPTaq in the presence of a fully complementary primer could help explain why DNAPTaq is unable to extend a DNA oligonucleotide on an RNA template, in a reaction resembling that of reverse transcriptase. Another thermophilic DNAP, DNAPTth, is able to use RNA as a template, but only in the presence of Mn++, so we predicted that this enzyme would not cleave RNA in the presence of this cation. Accordingly, we incubated an RNA molecule with an appropriate pilot oligonucleotide in the presence of DNAPTaq or DNAPTth, in buffer containing either Mg++ or Mn++. As expected, both enzymes cleaved the RNA in the presence of Mg++. However, DNAPTaq, but not DNAPTth, degraded the RNA in the presence of Mn++. We conclude that the 5′ nuclease activities of many DNAPs may contribute to their inability to use RNA as templates.
EXAMPLE 2 
Generation of 5′ Nucleases from Thermostable DNA Polymerases
Thermostable DNA polymerases were generated which have reduced synthetic activity, an activity that is an undesirable side-reaction during DNA cleavage in the detection assay of the invention, yet have maintained thermostable nuclease activity. The result is a thermostable polymerase which cleaves nucleic acids DNA with extreme specificity.
Type A DNA polymerases from eubacteria of the genus Thermus share extensive protein sequence identity (90% in the polymerization domain, using the Lipman-Pearson method in the DNA analysis software from DNAStar, Wis.) and behave similarly in both polymerization and nuclease assays. Therefore, we have used the genes for the DNA polymerase of Thermus aquaticus (DNAPTaq) and Thermus flavus (DNAPTfl) as representatives of this class. Polymerase genes from other eubacterial organisms, such as Thermus thermophilus, Thermus sp., Thermotoga maritima, Thermosipho africanus and Bacillus stearothermophilus are equally suitable. The DNA polymerases from these thermophilic organisms are capable of surviving and performing at elevated temperatures, and can thus be used in reactions in which temperature is used as a selection against non-specific hybridization of nucleic acid strands.
The restriction sites used for deletion mutagenesis, described below, were chosen for convenience. Different sites situated with similar convenience are available in the Thermus thermophilus gene and can be used to make similar constructs with other Type A polymerase genes from related organisms.
A. Creation Of 5′ Nuclease Constructs
1. Modified DNAPTaq Genes
The first step was to place a modified gene for the Taq DNA polymerase on a plasmid under control of an inducible promoter. The modified Taq polymerase gene was isolated as follows: The Taq DNA polymerase gene was amplified by polymerase chain reaction from genomic DNA from Thermus aquaticus, strain YT-1 (Lawyer et al., supra), using as primers the oligonucleotides described in SEQ ID NOS:13-14. The resulting fragment of DNA has a recognition sequence for the restriction endonuclease EcoRI at the 5′ end of the coding sequence and a BglII sequence at the 3′ end. Cleavage with BgllI leaves a 5′ overhang or “sticky end” that is compatible with the end generated by BamHI. The PCR-amplified DNA was digested with EcoRI and BamHI. The 2512 bp fragment containing the coding region for the polymerase gene was gel purified and then ligated into a plasmid which contains an inducible promoter.
In one embodiment of the invention, the pTTQ18 vector, which contains the hybrid trp-lac (tac) promoter, was used [M. J. R. Stark, Gene 5:255 (1987)] and shown in FIG. 13. The tac promoter is under the control of the E. coli lac repressor. Repression allows the synthesis of the gene product to be suppressed until the desired level of bacterial growth has been achieved, at which point repression is removed by addition of a specific inducer, isopropyl-β-D-thiogalactopyranoside (IPTG). Such a system allows the expression of foreign proteins that may slow or prevent growth of transformants.
Bacterial promoters, such as tac, may not be adequately suppressed when they are present on a multiple copy plasmid. If a highly toxic protein is placed under control of such a promoter, the small amount of expression leaking through can be harmful to the bacteria. In another embodiment of the invention, another option for repressing synthesis of a cloned gene product was used. The non-bacterial promoter, from bacteriophage T7, found in the plasmid vector series pET-3 was used to express the cloned mutant Taq polymerase genes [FIG. 15; Studier and Moffatt, J. Mol. Biol. 189:113 (1986)]. This promoter initiates transcription only by T7 RNA polymerase. In a suitable strain, such as BL21(DE3)pLYS, the gene for this RNA polymerase is carried on the bacterial genome under control of the lac operator. This arrangement has the advantage that expression of the multiple copy gene (on the plasmid) is completely dependent on the expression of T7 RNA polymerase, which is easily suppressed because it is present in a single copy.
For ligation into the pTTQ18 vector (FIG. 13), the PCR product DNA containing the Taq polymerase coding region (mutTaq, clone 4B, SEQ ID NO:21) was digested with EcoRI and BglII and this fragment was ligated under standard “sticky end” conditions [Sambrook et al. Molecular Cloning, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, pp. 1.63-1.69 (1989)] into the EcoRI and BamHI sites of the plasmid vector pTTQ18. Expression of this construct yields a translational fusion product in which the first two residues of the native protein (Met-Arg) are replaced by three from the vector (Met-Asn-Ser), but the remainder of the natural protein would not change. The construct was transformed into the JM109 strain of E. coli and the transformants were plated under incompletely repressing conditions that do not permit growth of bacteria expressing the native protein. These plating conditions allow the isolation of genes containing pre-existing mutations, such as those that result from the infidelity of Taq polymerase during the amplification process.
Using this amplification/selection protocol, we isolated a clone (depicted in FIG. 3B) containing a mutated Taq polymerase gene (mutTaq, clone 3B). The mutant was first detected by its phenotype, in which temperature-stable 5′ nuclease activity in a crude cell extract was normal, but polymerization activity was almost absent (approximately less than 1% of wild type Taq polymerase activity).
DNA sequence analysis of the recombinant gene showed that it had changes in the polymerase domain resulting in two amino acid substitutions: an A to G change at nucleotide position 1394 causes a Glu to Gly change at amino acid position 465 (numbered according to the natural nucleic and amino acid sequences, SEQ ID NOS:1 and 4) and another A to G change at nucleotide position 2260 causes a Gln to Arg change at amino acid position 754. Because the Gln to Gly mutation is at a nonconserved position and because the Glu to Arg mutation alters an amino acid that is conserved in virtually all of the known Type A polymerases, this latter mutation is most likely the one responsible for curtailing the synthesis activity of this protein. The nucleotide sequence for the FIG. 3B construct is given in SEQ ID NO:21. The enzyme encoded by this sequence is referred to as Cleavase® A/G.
Subsequent derivatives of DNAPTaq constructs were made from the mutTaq gene, thus, they all bear these amino acid substitutions in addition to their other alterations, unless these particular regions were deleted. These mutated sites are indicated by black boxes at these locations in the diagrams in FIG. 3. In FIG. 3, the designation “3′ Exo” is used to indicate the location of the 3′ exonuclease activity associated with Type A polymerases which is not present in DNAPTaq. All constructs except the genes shown in FIGS. 3E, F and G were made in the pTTQ18 vector.
The cloning vector used for the genes in FIGS. 3E and F was from the commercially available pET-3 series, described above. Though this vector series has only a BamHI site for cloning downstream of the T7 promoter, the series contains variants that allow cloning into any of the three reading frames. For cloning of the PCR product described above, the variant called pET-3c was used (FIG. 14). The vector was digested with BamHI, dephosphorylated with calf intestinal phosphatase, and the sticky ends were filled in using the Klenow fragment of DNAPEcl and dNTPs. The gene for the mutant Taq DNAP shown in FIG. 3B (mutTaq, clone 3B) was released from pTTQ18 by digestion with EcoRI and SalI, and the “sticky ends” were filled in as was done with the vector. The fragment was ligated to the vector under standard blunt-end conditions (Sambrook et al., Molecular Cloning, supra), the construct was transformed into the BL21(DE3)pLYS strain of E. coli, and isolates were screened to identify those that were ligated with the gene in the proper orientation relative to the promoter. This construction yields another translational fusion product, in which the first two amino acids of DNAPTaq (Met-Arg) are replaced by 13 from the vector plus two from the PCR primer (Met-Ala-Ser-Met-Thr-Gly-Gly-Gln-Gln-Met-Gly-Arg-Ile-Asn-Ser) (SEQ ID NO:24).
Our goal was to generate enzymes that lacked the ability to synthesize DNA, but retained the ability to cleave nucleic acids with a 5′ nuclease activity. The act of primed, templated synthesis of DNA is actually a coordinated series of events, so it is possible to disable DNA synthesis by disrupting one event while not affecting the others. These steps include, but are not limited to, primer recognition and binding, dNTP binding and catalysis of the inter-nucleotide phosphodiester bond. Some of the amino acids in the polymerization domain of DNAPEcI have been linked to these functions, but the precise mechanisms are as yet poorly defined.
One way of destroying the polymerizing ability of a DNA polymerase is to delete all or part of the gene segment that encodes that domain for the protein, or to otherwise render the gene incapable of making a complete polymerization domain. Individual mutant enzymes may differ from each other in stability and solubility both inside and outside cells. For instance, in contrast to the 5′ nuclease domain of DNAPEcI, which can be released in an active form from the polymerization domain by gentle proteolysis [Setlow and Kornberg, J. Biol. Chem. 247:232 (1972)], the Thermus nuclease domain, when treated similarly, becomes less soluble and the cleavage activity is often lost.
Using the mutant gene shown in FIG. 3B as starting material, several deletion constructs were created. All cloning technologies were standard (Sambrook et al, supra) and are summarized briefly, as follows:
FIG. 3C: The mutTaq construct was digested with PstI, which cuts once within the polymerase coding region, as indicated, and cuts immediately downstream of the gene in the multiple cloning site of the vector. After release of the fragment between these two sites, the vector was re-ligated, creating an 894-nucleotide deletion, and bringing into frame a stop codon 40 nucleotides downstream of the junction. The nucleotide sequence of this 5′ nuclease (clone 4C) is given in SEQ ID NO:9.
FIG. 3D: The mutTaq construct was digested with NheI, which cuts once in the gene at position 2047. The resulting four-nucleotide 5′ overhanging ends were filled in, as described above, and the blunt ends were re-ligated. The resulting four-nucleotide insertion changes the reading frame and causes termination of translation ten amino acids downstream of the mutation. The nucleotide sequence of this 5′ nuclease (clone 3D) is given in SEQ ID NO:10.
FIG. 3E: The entire mutTaq gene was cut from pTTQ18 using EcoRI and SalI and cloned into pET-3c, as described above. This clone was digested with BstXI and XcmI, at unique sites that are situated as shown in FIG. 3E. The DNA was treated with the Klenow fragment of DNAPEcl and dNTPs, which resulted in the 3′ overhangs of both sites being trimmed to blunt ends. These blunt ends were ligated together, resulting in an out-of-frame deletion of 1540 nucleotides. An in-frame termination codon occurs 18 triplets past the junction site. The nucleotide sequence of this 5′ nuclease (clone 3E) is given in SEQ ID NO:11, with the appropriate leader sequence given in SEQ ID NO:25. It is also referred to as Cleavase® BX.
FIG. 3F: The entire mutTaq gene was cut from pTTQ18 using EcoRI and SalI and cloned into pET-3c, as described above. This clone was digested with BstXI and BamHI, at unique sites that are situated as shown in the diagram. The DNA was treated with the Klenow fragment of DNAPEcl and dNTPs, which resulted in the 3′ overhang of the BstXI site being trimmed to a blunt end, while the 5′ overhang of the BamHI site was filled in to make a blunt end. These ends were ligated together, resulting in an in-frame deletion of 903 nucleotides. The nucleotide sequence of the 5′ nuclease (clone 3F) is given in SEQ ID NO:12. It is also referred to as Cleavase® BB.
FIG. 3G: This polymerase is a variant of that shown in FIG. 4E. It was cloned in the plasmid vector pET-21 (Novagen). The non-bacterial promoter from bacteriophage T7, found in this vector, initiates transcription only by T7 RNA polymerase. See Studier and Moffatt, supra. In a suitable strain, such as (DES)pLYS, the gene for this RNA polymerase is carried on the bacterial genome under control of the lac operator. This arrangement has the advantage that expression of the multiple copy gene (on the plasmid) is completely dependent on the expression of T7 RNA polymerase, which is easily suppressed because it is present in a single copy. Because the expression of these mutant genes is under this tightly controlled promoter, potential problems of toxicity of the expressed proteins to the host cells are less of a concern.
The pET-21 vector also features a “His*Tag”, a stretch of six consecutive histidine residues that are added on the carboxy terminus of the expressed proteins. The resulting proteins can then be purified in a single step by metal chelation chromatography, using a commercially available (Novagen) column resin with immobilized Ni++ ions. The 2.5 ml columns are reusable, and can bind up to 20 mg of the target protein under native or denaturing (guanidine*HCl or urea) conditions.
E. coli (DES)pLYS cells are transformed with the constructs described above using standard transformation techniques, and used to inoculate a standard growth medium (e.g., Luria-Bertani broth). Production of T7 RNA polymerase is induced during log phase growth by addition of IPTG and incubated for a further 12 to 17 hours. Aliquots of culture are removed both before and after induction and the proteins are examined by SDS-PAGE. Staining with Coomassie Blue allows visualization of the foreign proteins if they account for about 3-5% of the cellular protein and do not co-migrate with any of the major protein bands. Proteins that co-migrate with major host protein must be expressed as more than 10% of the total protein to be seen at this stage of analysis.
Some mutant proteins are sequestered by the cells into inclusion bodies. These are granules that form in the cytoplasm when bacteria are made to express high levels of a foreign protein, and they can be purified from a crude lysate, and analyzed by SDS-PAGE to determine their protein content. If the cloned protein is found in the inclusion bodies, it must be released to assay the cleavage and polymerase activities. Different methods of solubilization may be appropriate for different proteins, and a variety of methods are known. See e.g., Builder & Ogez, U.S. Pat. No. 4,511,502 (1985); Olson, U.S. Pat. No. 4,518,526 (1985); Olson & Pai, U.S. Pat. No. 4,511,503 (1985); Jones et al., U.S. Pat. No. 4,512,922 (1985), all of which are hereby incorporated by reference.
The solubilized protein is then purified on the Ni++ column as described above, following the manufacturers instructions (Novagen). The washed proteins are eluted from the column by a combination of imidazole competitor (1 M) and high salt (0.5 M NaCl), and dialyzed to exchange the buffer and to allow denature proteins to refold. Typical recoveries result in approximately 20 μg of specific protein per ml of starting culture. The DNAP mutant is referred to as the Cleavase® BN nuclease and the sequence is given in SEQ ID NO:26 (the amino acid sequence of the Cleavase® BN nuclease is obtained by translating the DNA sequence of SEQ ID NO:26).
2. Modified DNAPTfl Gene
The DNA polymerase gene of Thermus flavus was isolated from the “T. flavus” AT-62 strain obtained from the American Type Tissue Collection (ATCC 33923). This strain has a different restriction map then does the T. flavus strain used to generate the sequence published by Akhmetzjanov and Vakhitov, supra. The published sequence is listed as SEQ ID NO:2. No sequence data has been published for the DNA polymerase gene from the AT-62 strain of T. flavus. 
Genomic DNA from T. flavus was amplified using the same primers used to amplify the T. aquaticus DNA polymerase gene (SEQ ID NOS:13-14). The approximately 2500 base pair PCR fragment was digested with EcoRI and BamHI. The over-hanging ends were made blunt with the Klenow fragment of DNAPEcl and dNTPs. The resulting approximately 1800 base pair fragment containing the coding region for the N-terminus was ligated into pET-3c, as described above. This construct, clone 4B, is depicted in FIG. 4B. The wild type T. flavus DNA polymerase gene is depicted in FIG. 4A. The 4B clone has the same leader amino acids as do the DNAPTaq clones 4E and F which were cloned into pET-3c; it is not known precisely where translation termination occurs, but the vector has a strong transcription termination signal immediately downstream of the cloning site.
B. Growth And Induction Of Transformed Cells
Bacterial cells were transformed with the constructs described above using standard transformation techniques and used to inoculate 2 mls of a standard growth medium (e.g., Luria-Bertani broth). The resulting cultures were incubated as appropriate for the particular strain used, and induced if required for a particular expression system. For all of the constructs depicted in FIGS. 3 and 4, the cultures were grown to an optical density (at 600 nm wavelength) of 0.5 OD.
To induce expression of the cloned genes, the cultures were brought to a final concentration of 0.4 mM IPTG and the incubations were continued for 12 to 17 hours. 50 μl aliquots of each culture were removed both before and after induction and were combined with 20 μl of a standard gel loading buffer for sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Subsequent staining with Coomassie Blue (Sambrook et al., supra) allows visualization of the foreign proteins if they account for about 3-5% of the cellular protein and do not co-migrate with any of the major E. coli protein bands. Proteins that do co-migrate with a major host protein must be expressed as more than 10% of the total protein to be seen at this stage of analysis.
C. Heat Lysis And Fractionation
Expressed thermostable proteins, i.e., the 5′ nucleases, were isolated by heating crude bacterial cell extracts to cause denaturation and precipitation of the less stable E. coli proteins. The precipitated E. coli proteins were then, along with other cell debris, removed by centrifugation. 1.7 mls of the culture were pelleted by microcentrifugation at 12,000 to 14,000 rpm for 30 to 60 seconds. After removal of the supernatant, the cells were resuspended in 400 μl of buffer A (50 mM Tris-HCl, pH 7.9, 50 mM dextrose, 1 mM EDTA), re-centrifuged, then resuspended in 80 μl of buffer A with 4mg/ml lysozyme. The cells were incubated at room temperature for 15 minutes, then combined with 80 μl of buffer B (10 mM Tris-HCl, pH 7.9, 50 mM KCl, 1 mM EDTA, I mM PMSF, 0.5% Tween-20, 0.5% Nonidet-P40).
This mixture was incubated at 75° C. for 1 hour to denature and precipitate the host proteins. This cell extract was centrifuged at 14,000 rpm for 15 minutes at 4° C., and the supernatant was transferred to a fresh tube. An aliquot of 0.5 to 1 μl of this supernatant was used directly in each test reaction, and the protein content of the extract was determined by subjecting 7 μl to electrophoretic analysis, as above. The native recombinant Taq DNA polymerase [Englke, Anal. Biochem 191:396 (1990)], and the double point mutation protein shown in FIG. 3B are both soluble and active at this point.
The foreign protein may not be detected after the heat treatments due to sequestration of the foreign protein by the cells into inclusion bodies. These are granules that form in the cytoplasm when bacteria are made to express high levels of a foreign protein, and they can be purified from a crude lysate, and analyzed SDS PAGE to determine their protein content. Many methods have been described in the literature, and one approach is described below.
D. Isolation And Solubilization Of Inclusion Bodies
A small culture was grown and induced as described above. A 1.7 ml aliquot was pelleted by brief centrifugation, and the bacterial cells were resuspended in 100 μl of Lysis buffer (50 mM Tris-HCl, pH 8.0, 1 mM EDTA, 100 mM NaCl). 2.5 μl of 20 mM PMSF were added for a final concentration of 0.5 mM, and lysozyme was added to a concentration of 1.0 mg/ml. The cells were incubated at room temperature for 20 minutes, deoxycholic acid was added to 1 mg/ml (1 μl of 100 mg/ml solution), and the mixture was further incubated at 37° C. for about 15 minutes or until viscous. DNAse I was added to 10 μg/ml and the mixture was incubated at room temperature for about 30 minutes or until it was no longer viscous.
From this mixture the inclusion bodies were collected by centrifugation at 14,000 rpm for 15 minutes at 4° C., and the supematant was discarded. The pellet was resuspended in 100 μl of lysis buffer with 10 mM EDTA (pH 8.0) and 0.5% Triton X-100. After 5 minutes at room temperature, the inclusion bodies were pelleted as before, and the supernatant was saved for later analysis. The inclusion bodies were resuspended in 50 μl of distilled water, and 5 μl was combined with SDS gel loading buffer (which dissolves the inclusion bodies) and analyzed electrophoretically, along with an aliquot of the supernatant.
If the cloned protein is found in the inclusion bodies, it may be released to assay the cleavage and polymerase activities and the method of solubilization must be compatible with the particular activity. Different methods of solubilization may be appropriate for different proteins, and a variety of methods are discussed in Molecular Cloning (Sambrook et al., supra). The following is an adaptation we have used for several of our isolates.
20 μl of the inclusion body-water suspension were pelleted by centrifugation at 14,000 rpm for 4 minutes at room temperature, and the supematant was discarded. To further wash the inclusion bodies, the pellet was resuspended in 20 μl of lysis buffer with 2M urea, and incubated at room temperature for one hour. The washed inclusion bodies were then resuspended in 2 μl of lysis buffer with 8M urea; the solution clarified visibly as the inclusion bodies dissolved. Undissolved debris was removed by centrifugation at 14,000 rpm for 4 minutes at room temperature, and the extract supernatant was transferred to a fresh tube.
To reduce the urea concentration, the extract was diluted into KH2PO4. A fresh tube was prepared containing 180 μl of 50 mM KH2PO4, pH 9.5, 1 mM EDTA and 50 mM NaCl. A 2 μl aliquot of the extract was added and vortexed briefly to mix. This step was repeated until all of the extract had been added for a total of 10 additions. The mixture was allowed to sit at room temperature for 15 minutes, during which time some precipitate often forms. Precipitates were removed by centrifugation at 14,000 rpm, for 15 minutes at room temperature, and the supernatant was transferred to a fresh tube. To the 200 μl of protein in the KH2PO4 solution, 140-200 μl of saturated (NH4)2SO4 were added, so that the resulting mixture was about 41% to 50% saturated (NH4)2SO4. The mixture was chilled on ice for 30 minutes to allow the protein to precipitate, and the protein was then collected by centrifugation at 14,000 rpm, for 4 minutes at room temperature. The supernatant was discarded, and the pellet was dissolved in 20 μl Buffer C (20 mM HEPES, pH 7.9, 1 mM EDTA, 0.5% PMSF, 25 mM KCl and 0.5% each of Tween-20 and Nonidet P 40). The protein solution was centrifuged again for 4 minutes to pellet insoluble materials, and the supernatant was removed to a fresh tube. The protein contents of extracts prepared in this manner were visualized by resolving 1-4 μl by SDS-PAGE; 0.5 to 1 μl of extract was tested in the cleavage and polymerization assays as described.
E. Protein Analysis For Presence Of Nuclease And Synthetic Activity
The 5′ nucleases described above and shown in FIGS. 3 and 4 were analyzed by the following methods.
1. Structure Specific Nuclease Assay
A candidate modified polymerase is tested for 5′ nuclease activity by examining its ability to catalyze structure-specific cleavages. By the term “cleavage structure” as used herein, is meant a nucleic acid structure which is a substrate for cleavage by the 5′ nuclease activity of a DNAP.
The polymerase is exposed to test complexes that have the structures shown in FIG. 15. Testing for 5′ nuclease activity involves three reactions: 1) a primer-directed cleavage (FIG. 15B) is performed because it is relatively insensitive to variations in the salt concentration of the reaction and can, therefore, be performed in whatever solute conditions the modified enzyme requires for activity; this is generally the same conditions preferred by unmodified polymerases; 2) a similar primer-directed cleavage is performed in a buffer which permits primer-independent cleavage, i.e., a low salt buffer, to demonstrate that the enzyme is viable under these conditions; and 3) a primer-independent cleavage (FIG. 15A) is performed in the same low salt buffer.
The bifurcated duplex is formed between a substrate strand and a template strand as shown in FIG. 15. By the term “substrate strand” as used herein, is meant that strand of nucleic acid in which the cleavage mediated by the 5′ nuclease activity occurs. The substrate strand is always depicted as the top strand in the bifurcated complex which serves as a substrate for 5′ nuclease cleavage (FIG. 15). By the term “template strand” as used herein, is meant the strand of nucleic acid which is at least partially complementary to the substrate strand and which anneals to the substrate strand to form the cleavage structure. The template strand is always depicted as the bottom strand of the bifurcated cleavage structure (FIG. 15). If a primer (a short oligonucleotide of 19 to 30 nucleotides in length) is added to the complex, as when primer-dependent cleavage is to be tested, it is designed to anneal to the 3′ arm of the template strand (FIG. 15B). Such a primer would be extended along the template strand if the polymerase used in the reaction has synthetic activity.
The cleavage structure may be made as a single hairpin molecule, with the 3+ end of the target and the 5′ end of the pilot joined as a loop as shown in FIG. 15E. A primer oligonucleotide complementary to the 3′ arm is also required for these tests so that the enzyme's sensitivity to the presence of a primer may be tested.
Nucleic acids to be used to form test cleavage structures can be chemically synthesized, or can be generated by standard recombinant DNA techniques. By the latter method, the hairpin portion of the molecule can be created by inserting into a cloning vector duplicate copies of a short DNA segment, adjacent to each other but in opposing orientation. The double-stranded fragment encompassing this inverted repeat, and including enough flanking sequence to give short (about 20 nucleotides) unpaired 5′ and 3′ arms, can then be released from the vector by restriction enzyme digestion, or by PCR performed with an enzyme lacking a 5′ exonuclease (e.g., the Stoffel fragment of AmplitaqTM DNA polymerase, VentTM DNA polymerase).
The test DNA can be labeled on either end, or internally, with either a radioisotope, or with a non-isotopic tag. Whether the hairpin DNA is a synthetic single strand or a cloned double strand, the DNA is heated prior to use to melt all duplexes. When cooled on ice, the structure depicted in FIG. 16E is formed, and is stable for sufficient time to perform these assays.
To test for primer-directed cleavage (Reaction 1), a detectable quantity of the test molecule (typically 1-100 fmol of 32P-labeled hairpin molecule) and a 10 to 100-fold molar excess of primer are placed in a buffer known to be compatible with the test enzyme. For Reaction 2, where primer-directed cleavage is performed under condition which allow primer-independent cleavage, the same quantities of molecules are placed in a solution that is the same as the buffer used in Reaction 1 regarding pH, enzyme stabilizers (e.g., bovine serum albumin, nonionic detergents, gelatin) and reducing agents (e.g., dithiothreitol, 2-mercaptoethanol) but that replaces any monovalent cation salt with 20 mM KCl; 20 mM KCl is the demonstrated optimum for primer-independent cleavage. Buffers for enzymes, such as DNAPEc1, that usually operate in the absence of salt are not supplemented to achieve this concentration. To test for primer-independent cleavage (Reaction 3) the same quantity of the test molecule, but no primer, are combined under the same buffer conditions used for Reaction 2.
All three test reactions are then exposed to enough of the enzyme that the molar ratio of enzyme to test complex is approximately 1:1. The reactions are incubated at a range of temperatures up to, but not exceeding, the temperature allowed by either the enzyme stability or the complex stability, whichever is lower, up to 80° C. for enzymes from thermophiles, for a time sufficient to allow cleavage (10 to 60 minutes). The products of Reactions 1, 2 and 3 are resolved by denaturing polyacrylamide gel electrophoresis, and visualized by autoradiography or by a comparable method appropriate to the labeling system used. Additional labeling systems include chemiluminescence detection, silver or other stains, blotting and probing and the like. The presence of cleavage products is indicated by the presence of molecules which migrate at a lower molecular weight than does the uncleaved test structure. These cleavage products indicate that the candidate polymerase has structure-specific 5′ nuclease activity.
To determine whether a modified DNA polymerase has substantially the same 5′ nuclease activity as that of the native DNA polymerase, the results of the above-described tests are compared with the results obtained from these tests performed with the native DNA polymerase. By “substantially the same 5′ nuclease activity” we mean that the modified polymerase and the native polymerase will both cleave test molecules in the same manner. It is not necessary that the modified polymerase cleave at the same rate as the native DNA polymerase.
Some enzymes or enzyme preparations may have other associated or contaminating activities that may be functional under the cleavage conditions described above and that may interfere with 5′ nuclease detection. Reaction conditions can be modified in consideration of these other activities, to avoid destruction of the substrate, or other masking of the 5′ nuclease cleavage and its products. For example, the DNA polymerase I of E. Coli (Pol I), in addition to its polymerase and 5′ nuclease activities, has a 3′ exonuclease that can degrade DNA in a 3′ to 5′ direction. Consequently, when the molecule in FIG. 15E is exposed to this polymerase under the conditions described above, the 3′ exonuclease quickly removes the unpaired 3′ arm, destroying the bifurcated structure required of a substrate for the 5′ exonuclease cleavage and no cleavage is detected. The true ability of Pol I to cleave the structure can be revealed if the 3′ exonuclease is inhibited by a change of conditions (e.g., pH), mutation, or by addition of a competitor for the activity. Addition of 500 pmoles of a single-stranded competitor oligonucleotide, unrelated to the FIG. 15E structure, to the cleavage reaction with Pol I effectively inhibits the digestion of the 3′ arm of the FIG. 15E structure without interfering with the 5′ exonuclease release of the 5′ arm. The concentration of the competitor is not critical, but should be high enough to occupy the 3′ exonuclease for the duration of the reaction.
Similar destruction of the test molecule may be caused by contaminants in the candidate polymerase preparation. Several sets of the structure specific nuclease reactions may be performed to determine the purity of the candidate nuclease and to find the window between under and over exposure of the test molecule to the polymerase preparation being investigated.
The above described modified polymerases were tested for 5′ nuclease activity as follows: Reaction 1 was performed in a buffer of 10 mM Tris-Cl, pH 8.5 at 20° C., 1.5 mM MgCl2 and 50 mM KCl and in Reaction 2 the KCl concentration was reduced to 20 mM. In Reactions 1 and 2, 10 fmoles of the test substrate molecule shown in FIG. 15E were combined with 1 pmole of the indicated primer and 0.5 to 1.0 μl of extract containing the modified polymerase (prepared as described above). This mixture was then incubated for 10 minutes at 55° C. For all of the mutant polymerases tested these conditions were sufficient to give complete cleavage. When the molecule shown in FIG. 15E was labeled at the 5′end, the released 5′ fragment, 25 nucleotides long, was conveniently resolved on a 20% polyacrylamide gel (19:1 cross-linked) with 7 M urea in a buffer containing 45 mM Tris-borate pH 8.3, 1.4 mM EDTA. Clones 3C-F and 4B exhibited structure-specific cleavage comparable to that of the unmodified DNA polymerase. Additionally, clones 3E, 3F and 3G have the added ability to cleave DNA in the absence of a 3′ arm as discussed above. Representative cleavage reactions are shown in FIG. 16.
For the reactions shown in FIG. 16, the mutant polymerase clones 3E (Taq mutant) and 4B (Tfl mutant) were examined for their ability to cleave the hairpin substrate molecule shown in FIG. 15E. The substrate molecule was labeled at the 5′ terminus with 32P. Ten fmoles of heat-denatured, end-labeled substrate DNA and 0.5 units of DNAPTaq (lane 1) or 0.5 μl of 3E or 4B extract (FIG. 16, lanes 2-7, extract was prepared as described above) were mixed together in a buffer containing 10 mM Tris-Cl, pH 8.5, 50 mM KCl and 1.5 mM MgCl2. The final reaction volume was 10 μl. Reactions shown in lanes 4 and 7 contain in addition 50 μM of each dNTP. Reactions shown in lanes 3, 4, 6 and 7 contain 0.2 μM of the primer oligonucleotide (complementary to the 3′ arm of the substrate and shown in FIG. 15E). Reactions were incubated at 55° C. for 4 minutes. Reactions were stopped by the addition of 8 μl of 95% formamide containing 20 mM EDTA and 0.05% marker dyes per 10 μl reaction volume. Samples were then applied to 12% denaturing acrylamide gels. Following electrophoresis, the gels were autoradiographed. FIG. 16 shows that clones 3E and 4B exhibit cleavage activity similar to that of the native DNAPTaq. Note that some cleavage occurs in these reactions in the absence of the primer. When long hairpin structure, such as the one used here (FIG. 15E), are used in cleavage reactions performed in buffers containing 50 mM KCl a low level of primer-independent cleavage is seen. Higher concentrations of KCl suppress, but do not eliminate, this primer-independent cleavage under these conditions.
2. Assay for Synthetic Activity
The ability of the modified enzyme or proteolytic fragments is assayed by adding the modified enzyme to an assay system in which a primer is annealed to a template and DNA synthesis is catalyzed by the added enzyme. Many standard laboratory techniques employ such an assay. For example, nick translation and enzymatic sequencing involve extension of a primer along a DNA template by a polymerase molecule.
In a preferred assay for determining the synthetic activity of a modified enzyme an oligonucleotide primer is annealed to a single-stranded DNA template, e.g., bacteriophage M13 DNA, and the primer/template duplex is incubated in the presence of the modified polymerase in question, deoxynucleoside triphosphates (dNTPs) and the buffer and salts known to be appropriate for the unmodified or native enzyme. Detection of either primer extension (by denaturing gel electrophoresis) or dNTP incorporation (by acid precipitation or chromatography) is indicative of an active polymerase. A label, either isotopic or non-isotopic, is preferably included on either the primer or as a dNTP to facilitate detection of polymerization products. Synthetic activity is quantified as the amount of free nucleotide incorporated into the growing DNA chain and is expressed as amount incorporated per unit of time under specific reaction conditions.
Representative results of an assay for synthetic activity is shown in FIG. 17. The synthetic activity of the mutant DNAPTaq clones 3B-F was tested as follows: A master mixture of the following buffer was made: 1.2×PCR buffer (I×PCR buffer contains 50 mM KCl, 1.5 mM MgCl2, 10 mM Tris-Cl, ph 8.5 and 0.05% each Tween 20 and Nonidet P40), 50 μM each of dGTP, DATP and dTTP, 5 μM dCTP and 0.125 μM α-32P-dCTP at 600 Ci/mmol. Before adjusting this mixture to its final volume, it was divided into two equal aliquots. One received distilled water up to a volume of 50 μl to give the concentrations above. The other received 5 μg of single-stranded M13mp18 DNA (approximately 2.5 pmol or 0.05 μM final concentration) and 250 pmol of M13 sequencing primer (5 μM final concentration) and distilled water to a final volume of 50 μl. Each cocktail was warmed to 75° C. for 5 minutes and then cooled to room temperature. This allowed the primers to anneal to the DNA in the DNA-containing mixtures.
For each assay, 4 μl of the cocktail with the DNA was combined with 1 μl of the mutant polymerase, prepared as described, or 1 unit of DNAPTaq (Perkin Elmer) in 1 μl of dH2O. A “no DNA” control was done in the presence of the DNAPTaq (FIG. 17, lane 1), and a “no enzyme” control was done using water in place of the enzyme (lane 2). Each reaction was mixed, then incubated at room temperature (approx. 22° C.) for 5 minutes, then at 55° C. for 2 minutes, then at 72° C. for 2 minutes. This step incubation was done to detect polymerization in any mutants that might have optimal temperatures lower than 72° C. After the final incubation, the tubes were spun briefly to collect any condensation and were placed on ice. One μl of each reaction was spotted at an origin 1.5 cm from the bottom edge of a polyethyleneimine (PEI) cellulose thin layer chromatography plate and allowed to dry. The chromatography plate was run in 0.75 M NaH2PO4, pH 3.5, until the buffer front had run approximately 9 cm from the origin. The plate was dried, wrapped in plastic wrap, marked with luminescent ink, and exposed to X-ray film. Incorporation was detected as counts that stuck where originally spotted, while the unincorporated nucleotides were carried by the salt solution from the origin.
Comparison of the locations of the counts with the two control lanes confirmed the lack of polymerization activity in the mutant preparations. Among the modified DNAPTaq clones, only clone 3B retains any residual synthetic activity as shown in FIG. 17.
EXAMPLE 3
5′ Nucleases Derived from Thermostable DNA Polymerases can Cleave Short Hairpin Structures with Specificity
The ability of the 5′ nucleases to cleave hairpin structures to generate a cleaved hairpin structure suitable as a detection molecule was examined. The structure and sequence of the hairpin test molecule is shown in FIG. 18A (SEQ ID NO:15). The oligonucleotide (labeled “primer” in FIG. 18A, SEQ ID NO:22) is shown annealed to its complementary sequence on the 3′ arm of the hairpin test molecule. The hairpin test molecule was single-end labeled with 32P using a labeled T7 promoter primer in a polymerase chain reaction. The label is present on the 5′ arm of the hairpin test molecule and is represented by the star in FIG. 18A.
The cleavage reaction was performed by adding 10 fmoles of heat-denatured, end-labeled hairpin test molecule, 0.2 μM of the primer oligonucleotide (complementary to the 3′ arm of the hairpin), 50 μM of each dNTP and 0.5 units of DNAPTaq (Perkin Elmer) or 0.5μ of extract containing a 5μ nuclease (prepared as described above) in a total volume of 10 μl in a buffer containing 10 mM Tris-Cl, pH 8.5, 50 mM KCl and 1.5 mM MgCl2. Reactions shown in lanes 3, 5 and 7 were run in the absence of dNTPs.
Reactions were incubated at 55° C. for 4 minutes. Reactions were stopped at 55° C. by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes per 10 μl reaction volume. Samples were not heated before loading onto denaturing polyacrylamide gels (10% polyacrylamide, 19:1 crosslinking, 7 M urea, 89 mM Tris-borate, pH 8.3, 2.8 mM EDTA). The samples were not heated to allow for the resolution of single-stranded and re-duplexed uncleaved hairpin molecules.
FIG. 18B shows that altered polymerases lacking any detectable synthetic activity cleave a hairpin structure when an oligonucleotide is annealed to the single-stranded 3′ arm of the hairpin to yield a single species of cleaved product (FIG. 18B, lanes 3 and 4). 5′ nucleases, such as clone 3D, shown in lanes 3 and 4, produce a single cleaved product even in the presence of dNTPs. 5′ nucleases which retain a residual amount of synthetic activity (less than 1% of wild type activity) produce multiple cleavage products as the polymerase can extend the oligonucleotide annealed to the 3′ arm of the hairpin thereby moving the site of cleavage (clone 3B, lanes 5 and 6). Native DNATaq produces even more species of cleavage products than do mutant polymerases retaining residual synthetic activity and additionally converts the hairpin structure to a double-stranded form in the presence of dNTPs due to the high level of synthetic activity in the native polymerase (FIG. 18B, lane 8).
EXAMPLE 4 
Cleavage of Linear Nucleic Acid Substrates
From the above, it should be clear that native (i.e., “wild type”) thermostable DNA polymerases are capable of cleaving hairpin structures in a specific manner and that this discovery can be applied with success to a detection assay. In this example, the mutant DNAPs of the present invention are tested against three different cleavage structures shown in FIG. 20A. Structure 1 in FIG. 20A is simply single stranded 206-mer (the preparation and sequence information for which was discussed in Example 1C). Structures 2 and 3 are duplexes; structure 2 is the same hairpin structure as shown in FIG. 11A (bottom), while structure 3 has the hairpin portion of structure 2 removed.
The cleavage reactions comprised 0.01 pmoles of the resulting substrate DNA, and 1 pmole of pilot oligonucleotide in a total volume of 10 μl of 10 mM Tris-Cl, pH 8.3, 100 mM KCl, 1 mM MgCl2. Reactions were incubated for 30 minutes at 55° C., and stopped by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 10% polyacrylamide gel (19:1 cross link), with 7M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA.
The results were visualized by autoradiography and are shown in FIG. 20B with the enzymes indicated as follows: I is native Taq DNAP; II is native Tfl DNAP; III is Cleavase® BX shown in FIG. 3E; IV is Cleavase® BB shown in FIG. 3F; V is the mutant shown in FIG. 4B; and VI is Cleavase® BN shown in FIG. 3G.
Structure 2 was used to “normalize” the comparison. For example, it was found that it took 50 ng of Taq DNAP and 300 ng of Cleavase® BN to give similar amounts of cleavage of Structure 2 in thirty (30) minutes. Under these conditions native Taq DNAP is unable to cleave Structure 3 to any significant degree. Native Tfl DNAP cleaves Structure 3 in a manner that creates multiple products.
By contrast, all of the mutants tested cleave the linear duplex of Structure 3. This finding indicates that this characteristic of the mutant DNA polymerases is consistent of thermostable polymerases across thermophilic species.
EXAMPLE 5
5′ Exonucleolytic Cleavage (“Nibbling”) by Thermostable DNAPs
It has been found that thermostable DNAPs, including those of the present invention, have a true 5′ exonuclease capable of nibbling the 5′ end of a linear duplex nucleic acid structures. In this example, the 206 base pair DNA duplex substrate is again employed (see Example 1C). In this case, it was produced by the use of one 32P-labeled primer and one unlabeled primer in a polymerase chain reaction. The cleavage reactions comprised 0.01 pmoles of heat-denatured, end-labeled substrate DNA (with the unlabeled strand also present), 5 pmoles of pilot oligonucleotide (see pilot oligos in FIG. 11A) and 0.5 units of DNAPTaq or 0.5μ of Cleavase® BB in the E. coli extract (see above), in a total volume of 10 μl of 10 mM Tris.Cl, pH 8.5, 50 mM KCl, 1.5 mM MgCl2.
Reactions were initiated at 65° C. by the addition of pre-warmed enzyme, then shifted to the final incubation temperature for 30 minutes. The results are shown in FIG. 21A. Samples in lanes 1-4 are the results with native Taq DNAP, while lanes 5-8 shown the results with Cleavase® BB. The reactions for lanes 1, 2, 5, and 6 were performed at 65° C. and reactions for lanes 3, 4, 7, and 8 were performed at 50° C. and all were stopped at temperature by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 10% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris.Borate, pH 8.3, 1.4 mM EDTA. The expected product in reactions 1, 2, 5, and 6 is 85 nucleotides long; in reactions 3 and 7, the expected product is 27 nucleotides long. Reactions 4 and 8 were performed without pilot, and should remain at 206 nucleotides. The faint band seen at 24 nucleotides is residual end-labeled primer from the PCR.
The surprising result is that Cleavase® BB under these conditions causes all of the label to appear in a very small species, suggesting the possibility that the enzyme completely hydrolyzed the substrate. To determine the composition of the fastest-migrating band seen in lanes 5-8 (reactions performed with the deletion mutant), samples of the 206 base pair duplex were treated with either T7 gene 6 exonuclease (USB) or with calf intestine alkaline phosphatase (Promega), according to manufacturers' instructions, to produce either labeled mononucleotide (lane a of FIG. 21B) or free 32P-labeled inorganic phosphate (lane b of FIG. 21B), respectively. These products, along with the products seen in lane 7 of panel A were resolved by brief electrophoresis through a 20% acrylamide gel (19:1 cross-link), with 7 M urea, in a buffer of 45 mM Tris.Borate, pH 8.3, 1.4 mM EDTA. Cleavase® BB is thus capable of converting the substrate to mononucleotides.
EXAMPLE 6 
Nibbling is Duplex Dependent
The nibbling by Cleavase® BB is duplex dependent. In this example, internally labeled, single strands of the 206-mer were produced by 15 cycles of primer extension incorporating α-32P labeled dCTP combined with all four unlabeled dNTPs, using an unlabeled 206-bp fragment as a template. Single and double stranded products were resolved by electrophoresis through a non-denaturing 6% polyacrylamide gel (29:1 cross-link) in a buffer of 45 mM Tris.Borate, pH 8.3, 1.4 mM EDTA, visualized by autoradiography, excised from the gel, eluted by passive diffusion, and concentrated by ethanol precipitation.
The cleavage reactions comprised 0.04 pmoles of substrate DNA, and 2 μl of Cleavase® BB (in an E. coli extract as described above) in a total volume of 40 μl of 10 mM Tris.Cl, pH 8.5, 50 mM KCl, 1.5 mM MgCl2. Reactions were initiated by the addition of pre-warmed enzyme; 10 μl aliquots were removed at 5, 10, 20, and 30 minutes, and transferred to prepared tubes containing 8 μl of 95% formamide with 30 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 10% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris.Borate, pH 8.3, 1.4 mM EDTA. Results were visualized by autoradiography as shown in FIG. 22. Clearly, the cleavage by Cleavase® BB depends on a duplex structure; no cleavage of the single strand structure is detected whereas cleavage of the 206-mer duplex is complete.
EXAMPLE 7
Nibbling can be Target Directed
The nibbling activity of the DNAPs of the present invention can be employed with success in a detection assay. One embodiment of such an assay is shown in FIG. 23. In this assay, a labelled oligo is employed that is specific for a target sequence. The oligo is in excess of the target so that hybridization is rapid. In this embodiment, the oligo contains two fluorescein labels whose proximity on the oligo causes their emission to be quenched. When the DNAP is permitted to nibble the oligo the labels separate and are detectable. The shortened duplex is destabilized and disassociates. Importantly, the target is now free to react with an intact labelled oligo. The reaction can continue until the desired level of detection is achieved. An analogous, although different, type of cycling assay has been described employing lambda exonuclease. See C. G. Copley and C. Boot, BioTechniques 13:888 (1992).
The success of such an assay depends on specificity. In other words, the oligo must hybridize to the specific target. It is also preferred that the assay be sensitive; the oligo ideally should be able to detect small amounts of target. FIG. 24A shows a 5′-end 32P-labelled primer bound to a plasmid target sequence. In this case, the plasmid was pUC19 (commercially available) which was heat denatured by boiling two (2) minutes and then quick chilling. The primer is a 21-mer (SEQ ID NO:28). The enzyme employed was Cleavase® BX (a dilution equivalent to 5×10−1 μl extract) in 100 mM KCl, 10 mM Tris-Cl, pH 8.3, 2 mM MnCl2. The reaction was performed at 55° C. for sixteen (16) hours with or without genomic background DNA (from chicken blood). The reaction was stopped by the addition of 8 μl of 95% formamide with 20 mM EDTA and marker dyes.
The products of the reaction were resolved by PAGE (10% polyacrylamide, 19:1 cross link, 1×TBE) as seen in FIG. 24B. Lane “M” contains the labelled 21-mer. Lanes 1-3 contain no specific target, although Lanes 2 and 3 contain 100 ng and 200 ng of genomic DNA, respectively. Lanes 4, 5 and 6 all contain specific target with either 0 ng, 100 ng or 200 ng of genomic DNA, respectively. It is clear that conversion to mononucleotides occurs in Lanes 4, 5 and 6 regardless of the presence or amount of background DNA. Thus, the nibbling can be target directed and specific.
EXAMPLE 8 
Cleavase Purification
As noted above, expressed thermostable proteins, i.e., the 5′ nucleases, were isolated by crude bacterial cell extracts. The precipitated E. coli proteins were then, along with other cell debris, removed by centrifugation. In this example, cells expressing the BN clone were cultured and collected (500 grams). For each gram (wet weight) of E. coli, is buffer (50 mM Tris-HCl, pH 8.0, 1 mM EDTA, 100 μM NaCl) was added. The cells were lysed with 200 μg/ml lysozyme at room temperature for 20 minutes. Thereafter deoxycholic acid was added to make a 0.2% final concentration and the mixture was incubated 15 minutes at room temperature.
The lysate was sonicated for approximately 6-8 minutes at 0° C. The precipitate was removed by centrifugation (39,000 g for 20 minutes). Polyethyleneimine was added (0.5%) to the supematant and the mixture was incubated on ice for 15 minutes. The mixture was centrifuged (5,000 g for 15 minutes) and the supernatant was retained. This was heated for 30 minutes at 60° C. and then centrifuged again (5,000 g for 15 minutes) and the supernatant was again retained.
The supernatant was precipitated with 35% ammonium sulfate at 4° C. for 15 minutes. The mixture was then centrifuged (5,000 g for 15 minutes) and the supernatant was removed. The precipitate was then dissolved in 0.25M KCl, 20 Tris pH 7.6, 0.2% Tween and 0.1 EDTA) and then dialyzed against Binding Buffer (8×Binding Buffer comprises: 40 mM imidazole, 4M NaCl, 160 mM Tris-HCl, pH 7.9).
The solubilized protein is then purified on the Ni++ column (Novagen). The Binding Buffer is allows to drain to the top of the column bed and load the column with the prepared extract. A flow rate of about 10 column volumes per hour is optimal for efficient purification. If the flow rate is too fast, more impurities will contaminate the eluted fraction.
The column is washed with 25 ml (10 volumes) of 1×Binding Buffer and then washed with 15 ml (6 volumes) of 1×Wash Buffer (8×Wash Buffer comprises: 480 mM imidazole, 4M NaCl, 160 mM Tris-HCl, pH 7.9). The bound protein was eluted with 15 ml (6 volumes) of 1×Elute Buffer (4×Elute Buffer comprises: 4 mM imidazole, 2M NaCl, 8mM Tris-HCl, pH 7.9). Protein is then reprecipitated with 35% Ammonium Sulfate as above. The precipitate was then dissolved and dialyzed against: 2 mM Tris, 100 mM KCl, 1 mM EDTA). The solution was brought up to 0.1% each of Tween 20 and NP-40 and stored at 4° C.
EXAMPLE 9 
The Use of Various Divalent Cations in the Cleavage Reaction Influences the Nature of the Resulting Cleavage Products
In comparing the 5′ nucleases generated by the modification and/or deletion of the C-terminal polymerization domain of Thermus aquaticus DNA polymerase (DNAPTaq), as diagrammed in FIG. 3B-G, significant differences in the strength of the interactions of these proteins with the 3′ end of primers located upstream of the cleavage site (as depicted in FIG. 5) were noted. In describing the cleavage of these structures by Pol I-type DNA polymerases [Example 1 and Lyamichev et al. (1993) Science 260:778], it was observed that in the absence of a primer, the location of the junction between the double-stranded region and the single-stranded 5′ and 3′ arms determined the site of cleavage, but in the presence of a primer, the location of the 3′ end of the primer became the determining factor for the site of cleavage. It was postulated that this affinity for the 3′ end was in accord with the synthesizing function of the DNA polymerase.
Structure 2, shown in FIG. 20A, was used to test the effects of a 3′ end proximal to the cleavage site in cleavage reactions comprising several different solutions [e.g., solutions containing different salts (KCl or NaCl), different divalent cations (Mn2+ or Mg2+), etc.] as well as the use of different temperatures for the cleavage reaction. When the reaction conditions were such that the binding of the enzyme (e.g., a DNAP comprising a 5′ nuclease, a modified DNAP or a 5′ nuclease) to the 3′ end (of the pilot oligonucleotide) near the cleavage site was strong, the structure shown is cleaved at the site indicated in FIG. 20A. This cleavage releases the unpaired 5′ arm and leaves a nick between the remaining portion of the target nucleic acid and the folded 3′ end of the pilot oligonucleotide. In contrast, when the reaction conditions are such that the binding of the DNAP (comprising a 5′ nuclease) to the 3′ end was weak, the initial cleavage was as described above, but after the release of the 5′ arm, the remaining duplex is digested by the exonuclease function of the DNAP.
One way of weakening the binding of the DNAP to the 3′ end is to remove all or part of the domain to which at least some of this function has been attributed. Some of 5′ nucleases created by deletion of the polymerization domain of DNAPTaq have enhanced true exonuclease function, as demonstrated in Example 5.
The affinity of these types of enzymes (i.e., 5′ nucleases associated with or derived from DNAPs) for recessed 3′ ends may also be affected by the identity of the divalent cation present in the cleavage reaction. It was demonstrated by Longley et al. [Nucl. Acids Res. 18:7317 (1990)] that the use of MnCl2 in a reaction with DNAPTaq enabled the polymerase to remove nucleotides from the 5′ end of a primer annealed to a template, albeit inefficiently. Similarly, by examination of the cleavage products generated using Structure 2 from FIG. 20A, as described above, in a reaction containing either DNAPTaq or the Cleavase® BB nuclease, it was observed that the substitution of MnCl2 for MgCl2 in the cleavage reaction resulted in the exonucleolytic “nibbling” of the duplex downstream of the initial cleavage site. While not limiting the invention to any particular mechanism, it is thought that the substitution of MnCl2 for MgCl2 in the cleavage reaction lessens the affinity of these enzymes for recessed 3′ ends.
In all cases, the use of MnCl2 enhances the 5′ nuclease function, and in the case of the Cleavase® BB nuclease, a 50- to 100-fold stimulation of the 5′ nuclease function is seen. Thus, while the exonuclease activity of these enzymes was demonstrated above in the presence of MgCl2, the assays described below show a comparable amount of exonuclease activity using 50 to 100-fold less enzyme when MnCl2 is used in place of MgCl2. When these reduced amounts of enzyme are used in a reaction mixture containing MgCl2, the nibbling or exonuclease activity is much less apparent than that seen in Examples 5-7.
Similar effects are observed in the performance of the nucleic acid detection assay described in Examples 10-39 below when reactions performed in the presence of either MgCl2 or MnCl2 are compared. In the presence of either divalent cation, the presence of the invader oligonucleotide (described below) forces the site of cleavage into the probe duplex, but in the presence of MnCl2 the probe duplex can be further nibbled producing a ladder of products that are visible when a 3′ end label is present on the probe oligonucleotide. When the invader oligonucleotide is omitted from a reaction containing Mn2+, the probe is nibbled from the 5′ end. Mg2+-based reactions display minimal nibbling of the probe oligonucleotide. In any of these cases, the digestion of the probe is dependent upon the presence of the target nucleic acid. In the examples below, the ladder produced by the enhanced nibbling activity observed in the presence of Mn2+ is used as a positive indicator that the probe oligonucleotide has hybridized to the target sequence.
EXAMPLE 10 
Invasive 5′ Endonucleolytic Cleavage by Thermostable 5′ Nucleases in the Absence of Polymerization
As described in the examples above, 5′ nucleases cleave near the junction between single-stranded and base-paired regions in a bifurcated duplex, usually about one base pair into the base-paired region. In this example, it is shown that thermostable 5′ nucleases, including those of the present invention (e.g., Cleavase® BN nuclease, Cleavase® A/G nuclease), have the ability to cleave a greater distance into the base paired region when provided with an upstream oligonucleotide bearing a 3′ region that is homologous to a 5′ region of the subject duplex, as shown in FIG. 26.
FIG. 26 shows a synthetic oligonucleotide which was designed to fold upon itself which consists of the following sequence: 5′-GTTCTCTGCTCTCTGGTCGCTGTCTCGCTTGTGAAACAAGCGAGACAGCGTGGTCTCTCG-3′ (SEQ ID NO:29). This oligonucleotide is referred to as the “S-60 Hairpin.” The 15 basepair hairpin formed by this oligonucleotide is further stabilized by a “tri-loop” sequence in the loop end (i.e., three nucleotides form the loop portion of the hairpin) [Hiraro, I. et al. (1994) Nucleic Acids Res. 22(4):576]. FIG. 26 also show the sequence of the P-15 oligonucleotide and the location of the region of complementarity shared by the P-15 and S-60 hairpin oligonucleotides. The sequence of the P-15 oligonucleotide is 5′-CGAGAGACCACGCTG-3′ (SEQ ID NO:30). As discussed in detail below, the solid black arrowheads shown in FIG. 26 indicate the sites of cleavage of the S-60 hairpin in the absence of the P-15 oligonucleotide and the hollow arrow heads indicate the sites of cleavage in the presence of the P-15 oligonucleotide. The size of the arrow head indicates the relative utilization of a particular site.
The S-60 hairpin molecule was labeled on its 5′ end with biotin for subsequent detection. The S-60 hairpin was incubated in the presence of a thermnostable 5′ nuclease in the presence or the absence of the P-15 oligonucleotide. The presence of the full duplex which can be formed by the S-60 hairpin is demonstrated by cleavage with the Cleavase® BN 5′ nuclease, in a primer-independent fashion (i.e., in the absence of the P-15 oligonucleotide). The release of 18 and 19-nucleotide fragments from the 5′ end of the S-60 hairpin molecule showed that the cleavage occurred near the junction between the single and double stranded regions when nothing is hybridized to the 3′ arm of the S-60 hairpin (FIG. 27, lane 2).
The reactions shown in FIG. 27 were conducted as follows. Twenty fmole of the 5° biotin-labeled hairpin DNA (SEQ ID NO:29) was combined with 0.1 ng of Cleavase® BN enzyme and 1 μl of 100 mM MOPS (pH 7.5) containing 0.5% each of Tween-20 and NP-40 in a total volume of 9 μl. In the reaction shown in lane 1, the enzyme was omitted and the volume was made up by addition of distilled water (this served as the uncut or no enzyme control). The reaction shown in lane 3 of FIG. 27 also included 0.5 pmole of the P15 oligonucleotide (SEQ ID NO:30), which can hybridize to the unpaired 3′ arm of the S-60 hairpin (SEQ ID NO:29), as diagrammed in FIG. 26.
The reactions were overlaid with a drop of mineral oil, heated to 95° C. for 15 seconds, then cooled to 37° C., and the reaction was started by the addition of 1 μl of 10 mM MnCl2 to each tube. After 5 minutes, the reactions were stopped by the addition of 6 μl of 95% formamide containing 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 15% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA.
After electrophoresis, the gel plates were separated allowing the gel to remain flat on one plate. A 0.2 mm-pore positively-charged nylon membrane (NYTRAN, Schleicher and Schuell, Keene, N.H.), pre-wetted in H2O, was laid on top of the exposed gel. All air bubbles were removed. Two pieces of 3MM filter paper (Whatman) were then placed on top of the membrane, the other glass plate was replaced, and the sandwich was clamped with binder clips. Transfer was allowed to proceed overnight. After transfer, the membrane was carefully peeled from the gel and allowed to air dry. After complete drying, the membrane was washed in 1.2×Sequenase Images Blocking Buffer (United States Biochemical) using 0.3 ml of buffer/cm2 of membrane. The wash was performed for 30 minutes at room temperature. A streptavidin-alkaline phosphatase conjugate (SAAP, United States Biochemical) was added to a 1:4000 dilution directly to the blocking solution, and agitated for 15 minutes. The membrane was rinsed briefly with H2O and then washed three times for 5 minutes per wash using 0.5 ml/cm2 of 1×X SAAP buffer (100 mM Tris-HCl, pH 10, 50 mM NaCl) with 0.1% sodium dodecyl sulfate (SDS). The membrane was rinsed briefly with H2O between each wash. The membrane was then washed once in 1×SAAP buffer containing 1 mM MgCl2 without SDS, drained thoroughly and placed in a plastic heat-sealable bag. Using a sterile pipet, 5 mls of CDP-StarTM (Tropix, Bedford, Mass.) chemiluminescent substrate for alkaline phosphatase were added to the bag and distributed over the entire membrane for 2-3 minutes. The CDP-Star®-treated membrane was exposed to XRP X-ray film (Kodak) for an initial exposure of 10 minutes.
The resulting autoradiograph is shown in FIG. 27. In FIG. 27, the lane labelled “M” contains the biotinylated P-15 oligonucleotide which served as a marker. The sizes (in nucleotides) of the uncleaved S-60 hairpin (60 nuc; lane 1), the marker (15 nuc; lane “M”) and the cleavage products generated by cleavage of the S-60 hairpin in the presence (lane 3) or absence (lane 2) of the P-15 oligonucleotide are indicated.
Because the complementary regions of the S-60 hairpin are located on the same molecule, essentially no lag time should be needed to allow hybridization (i.e., to form the duplex region of the hairpin). This hairpin structure would be expected to form long before the enzyme could locate and cleave the molecule. As expected, cleavage in the absence of the primer oligonucleotide was at or near the junction between the duplex and single-stranded regions, releasing the unpaired 5′ arm (FIG. 27, lane 2). The resulting cleavage products were 18 and 19 nucleotides in length.
It was expected that stability of the S-60 hairpin with the tri-loop would prevent the P-15 oligonucleotide from promoting cleavage in the “primer-directed” manner described in Example 1 above, because the 3′ end of the “primer” would remain unpaired. Surprisingly, it was found that the enzyme seemed to mediate an “invasion” by the P-15 primer into the duplex region of the S-60 hairpin, as evidenced by the shifting of the cleavage site 3 to 4 basepairs further into the duplex region, releasing the larger products (22 and 21 nuc.) observed in lane 3 of FIG. 27.
The precise sites of cleavage of the S-60 hairpin are diagrammed on the structure in FIG. 26, with the solid black arrowheads indicating the sites of cleavage in the absence of the P-15 oligonucleotide and the hollow arrow heads indicating the sites of cleavage in the presence of P-15.
These data show that the presence on the 3° arm of an oligonucleotide having some sequence homology with the first several bases of the similarly oriented strand of the downstream duplex can be a dominant factor in determining the site of cleavage by 5° nucleases. Because the oligonucleotide which shares some sequence homology with the first several bases of the similarly oriented strand of the downstream duplex appears to invade the duplex region of the hairpin, it is referred to as an“invader” oligonucleotide. As shown in the examples below, an invader oligonucleotide appears to invade (or displace) a region of duplexed nucleic acid regardless of whether the duplex region is present on the same molecule (i.e., a hairpin) or whether the duplex is formed between two separate nucleic acid strands.
EXAMPLE 11 
The Invader Oligonucleotide Shifts the Site of Cleavage in a Pre-Formed Probe/Target Duplex
In Example 10 it was demonstrated that an invader oligonucleotide could shift the site at which a 5° nuclease cleaves a duplex region present on a hairpin molecule. In this example, the ability of an invader oligonucleotide to shift the site of cleavage within a duplex region formed between two separate strands of nucleic acid molecules was examined.
A single-stranded target DNA comprising the single-stranded circular M13mp19 molecule and a labeled (fluorescein) probe oligonucleotide were mixed in the presence of the reaction buffer containing salt (KCl) and divalent cations (Mg2+ or Mn2+) to promote duplex formation. The probe oligonucleotide refers to a labelled oligonucleotide which is complementary to a region along the target molecule (e.g., M13mp19). A second oligonucleotide (unlabelled) was added to the reaction after the probe and target had been allowed to anneal. The second oligonucleotide binds to a region of the target which is located downstream of the region to which the probe oligonucleotide binds. This second oligonucleotide contains sequences which are complementary to a second region of the target molecule. If the second oligonucleotide contains a region which is complementary to a portion of the sequences along the target to which the probe oligonucleotide also binds, this second oligonucleotide is referred to as an invader oligonucleotide (see FIG. 28c).
FIG. 32 depicts the annealing of two oligonucleotides to regions along the M13mp19 target molecule (bottom strand in all three structures shown). In FIG. 28 only a 52 nucleotide portion of the M13mp19 molecule is shown; this 52 nucleotide sequence is listed in SEQ ID NO:31. The probe oligonucleotide contains a fluorescein label at the 3′ end; the sequence of the probe is 5′-AGAAAGGAAGGGAAGAAAGCGAAAGG-3′ (SEQ ID NO:32). In FIG. 28, sequences comprising the second oligonucleotide, including the invader oligonucleotide are underlined. In FIG. 28a, the second oligonucleotide, which has the sequence 5′-GACGGGGAAAGCCGGCGAACG-3′ (SEQ ID NO:33), is complementary to a different and downstream region of the target molecule than is the probe oligonucleotide (labeled with fluorescein or “Fluor”); there is a gap between the second, upstream oligonucleotide and the probe for the structure shown in FIG. 28a. In FIG. 28b, the second, upstream oligonucleotide, which has the sequence 5′-GAAAGCCGGCGAACGTGGCG-3′ (SEQ ID NO:34), is complementary to a different region of the target molecule than is the probe oligonucleotide, but in this case, the second oligonucleotide and the probe oligonucleotide abut one another (that is the 3′ end of the second, upstream oligonucleotide is immediately adjacent to the 5′ end of the probe such that no gap exists between these two oligonucleotides). In FIG. 28c, the second, upstream oligonucleotide [5′-GGCGAACGTGGCGAGAAAGGA-3′ (SEQ ID NO:35)] and the probe oligonucleotide share a region of complementarity with the target molecule. Thus, the upstream oligonucleotide has a 3′ arm which has a sequence identical to the first several bases of the downstream probe. In this situation, the upstream oligonucleotide is referred to as an “invader” oligonucleotide.
The effect of the presence of an invader oligonucleotide upon the pattern of cleavage in a probe/target duplex formed prior to the addition of the invader was examined. The invader oligonucleotide and the enzyme were added after the probe was allowed to anneal to the target and the position and extent of cleavage of the probe were examined to determine a) if the invader was able to shift the cleavage site to a specific internal region of the probe, and b), if the reaction could accumulate specific cleavage products over time, even in the absence of thermal cycling, polymerization, or exonuclease removal of the probe sequence.
The reactions were carried out as follows. Twenty μl each of two enzyme mixtures were prepared, containing 2 μl of Cleavase® A/G nuclease extract (prepared as described in Example 2), with or without 50 pmole of the invader oligonucleotide (SEQ ID NO:35), as indicated, per 4 μl of the mixture. For each of the eight reactions shown in FIG. 29, 150 fmole of M13mp19 single-stranded DNA (available from Life Technologies, Inc.) was combined with 5 pmoles of fluorescein labeled probe (SEQ ID NO:32), to create the structure shown in FIG. 28c, but without the invader oligonucleotide present (the probe/target mixture). One half (4 tubes) of the probe/target mixtures were combined with 1 μl of 100 mM MOPS, pH 7.5 with 0.5% each of Tween-20 and NP-40, 0.5 μl of 1 M KCl and 0.25 μl of 80 mM MnCl2, and distilled water to a volume of 6 μl. The second set of probe/target mixtures were combined with 1 μl of 100 mM MOPS, pH 7.5 with 0.5% each of Tween-20 and NP-40, 0.5 μl of 1 M KCl and 0.25 μl of 80 mM MgCl2. The second set of mixtures therefore contained MgCl2 in place of the MnCl2 present in the first set of mixtures.
The mixtures (containing the probe/target with buffer, KCl and divalent cation) were covered with a drop of ChillOut® evaporation barrier and were brought to 60° C. for 5 minutes to allow annealing. Four μl of the above enzyme mixtures without the invader oligonucleotide was added to reactions whose products are shown in lanes 1, 3, 5 and 7 of FIG. 29. Reactions whose products are shown lanes 2, 4, 6, and 8 of FIG. 29 received the same amount of enzyme mixed with the invader oligonucleotide (SEQ ID NO:35). Reactions 1, 2, 5 and 6 were incubated for 5 minutes at 60° C. and reactions 3, 4, 7 and 8 were incubated for 15 minutes at 60° C.
All reactions were stopped by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 90° C. for 1 minute immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), containing 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. Following electrophoresis, the reaction products and were visualized by the use of an Hitachi FMBIO fluorescence imager, the output of which is seen in FIG. 29. The very low molecular weight fluorescent material seen in all lanes at or near the salt front in FIG. 29 and other fluoro-imager figures is observed when fluorescently-labeled oligonucleotides are electrophoresed and imaged on a fluoro-imager. This material is not a product of the cleavage reaction.
The use of MnCl2 in these reactions (lanes 1-4) stimulates the true exonuclease or “nibbling” activity of the Cleavase® enzyme, as described in Example 6, as is clearly seen in lanes 1 and 3 of FIG. 29. This nibbling of the probe oligonucleotide (SEQ ID NO:32) in the absence of invader oligonucleotide (SEQ ID NO:35) confirms that the probe oligonucleotide is forming a duplex with the target sequence. The ladder-like products produced by this nibbling reaction may be difficult to differentiate from degradation of the probe by nucleases that might be present in a clinical specimen. In contrast, introduction of the invader oligonucleotide (SEQ ID NO:35) caused a distinctive shift in the cleavage of the probe, pushing the site of cleavage 6 to 7 bases into the probe, confirming the annealing of both oligonucleotides. In presence of MnCl2, the exonuclease “nibbling” may occur after the invader-directed cleavage event, until the residual duplex is destabilized and falls apart.
In a magnesium based cleavage reaction (lanes 5-8), the nibbling or true exonuclease function of the Cleavase® A/G is enzyme suppressed (but the endonucleolytic function of the enzyme is essentially unaltered), so the probe oligonucleotide is not degraded in the absence of the invader (FIG. 29, lanes 5 and 7). When the invader is added, it is clear that the invader oligonucleotide can promote a shift in the site of the endonucleolytic cleavage of the annealed probe. Comparison of the products of the 5 and 15 minute reactions with invader (lanes 6 and 8 in FIG. 29) shows that additional probe hybridizes to the target and is cleaved. The calculated melting temperature (Tm) of the portion of probe that is not invaded (i.e., nucleotides 9-26 of SEQ ID NO:32) is 56° C., so the observed turnover (as evidenced by the accumulation of cleavage products with increasing reaction time) suggests that the full length of the probe molecule, with a calculated Tm of 76° C., is must be involved in the subsequent probe annealing events in this 60° C. reaction.
EXAMPLE 12 
The Overlap of the 3μ Invader Oligonucleotide Sequence with the 5′ Region of the Probe Causes a Shift in the Site of Cleavage
In Example 11, the ability of an invader oligonucleotide to cause a shift in the site of cleavage of a probe annealed to a target molecule was demonstrated. In this example, experiments were conducted to examine whether the presence of an oligonucleotide upstream from the probe was sufficient to cause a shift in the cleavage site(s) along the probe or whether the presence of nucleotides on the 3′ end of the invader oligonucleotide which have the same sequence as the first several nucleotides at the 5′ end of the probe oligonucleotide were required to promote the shift in cleavage.
To examine this point, the products of cleavage obtained from three different arrangements of target-specific oligonucleotides are compared. A diagram of these oligonucleotides and the way in which they hybridize to a test nucleic acid, M13mp19, is shown in FIG. 28. In FIG. 28a, the 3′ end of the upstream oligonucleotide (SEQ ID NO:33) is located upstream of the 5′ end of the downstream “probe” oligonucleotide (SEQ ID NO:32) such that a region of the M13 target which is not paired to either oligonucleotide is present. In FIG. 28b, the sequence of the upstream oligonucleotide (SEQ ID NO:34) is immediately upstream of the probe (SEQ ID NO:32), having neither a gap nor an overlap between the sequences. FIG. 28c diagrams the arrangement of the substrates used in the assay of the present invention, showing that the upstream “invader” oligonucleotide (SEQ ID NO:35) has the same sequence on a portion of its 3′ region as that present in the 5′ region of the downstream probe (SEQ ID NO:32). That is to say, these regions will compete to hybridize to the same segment of the M13 target nucleic acid.
In these experiments, four enzyme mixtures were prepared as follows (planning 5 μl per digest): Mixture 1 contained 2.251 μl of Cleavase® A/G nuclease extract (prepared as described in Example 2) per 5 μl of mixture, in 20 mM MOPS, pH 7.5 with 0.1% each of Tween 20 and NP-40, 4 mM MnCl2 and 100 mM KCl. Mixture 2 contained 11.25 units of Taq DNA polymerase (Promega) per 5 μof mixture in 20 mM MOPS, pH 7.5 with 0.1% each of Tween 20 and NP-40, 4 mM MnCl2 and 100 mM KCl. Mixture 3 contained 2.25 μl of Cleavase® A/G nuclease extract per 5 μl of mixture in 20 mM Tris-HCl, pH 8.5, 4 mM MgCl2 and 100 mM KCl. Mixture 4 contained 11.25 units of Taq DNA polymerase per 5 μl of mixture in 20 mM Tris-HCl, pH 8.5, 4 mM MgCl2 and 100 mM KCl.
For each reaction, 50 fmole of M13mp19 single-stranded DNA (the target nucleic acid) was combined with 5 pmole of the probe oligonucleotide (SEQ ID NO:32 which contained a fluorescein label at the 3′μend) and 50 pmole of one of the three upstream oligonucleotides diagrammed in FIG. 28 (i.e., one of SEQ ID NOS:33-35), in a total volume of 5 μl of distilled water. The reactions were overlaid with a drop of ChillOutTM evaporation barrier and warmed to 62° C. The cleavage reactions were started by the addition of 5 μl of an enzyme mixture to each tube, and the reactions were incubated at 62° C. for 30 min. The reactions shown in lanes 1-3 of FIG. 30 received Mixture 1; reactions 4-6 received Mixture 2; reactions 7-9 received Mixture 3 and reactions 10-12 received Mixture 4.
After 30 minutes at 62° C., the reactions were stopped by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA.
Following electrophoresis, the products of the reactions were visualized by the use of an Hitachi FMBIO fluorescence imager, the output of which is seen in FIG. 30. The reaction products shown in lanes 1, 4, 7 and 10 of FIG. 30 were from reactions which contained SEQ ID NO:33 as the upstream oligonucleotide (see FIG. 28a). The reaction products shown in lanes 2, 5, 8 and 11 of FIG. 30 were from reactions which contained SEQ ID NO:34 as the upstream oligonucleotide (see FIG. 28b). The reaction products shown in lanes 3, 6, 9 and 12 of FIG. 30 were from reactions which contained SEQ ID NO:35, the invader oligonucleotide, as the upstream oligonucleotide (see FIG. 28c).
Examination of the Mn2+ based reactions using either Cleavase® A/G nuclease or DNAPTaq as the cleavage agent (lanes 1 through 3 and 4 through 6, respectively) shows that both enzymes have active exonuclease function in these buffer conditions. The use of a 3′ label on the probe oligonucleotide allows the products of the nibbling activity to remain labeled, and therefore visible in this assay. The ladders seen in lanes 1, 2, 4 and 5 confirm that the probe hybridize to the target DNA as intended. These lanes also show that the location of the non-invasive oligonucleotides have little effect on the products generated. The uniform ladder created by these digests would be difficult to distinguish from a ladder causes by a contaminating nuclease, as one might find in a clinical specimen. In contrast, the products displayed in lanes 3 and 6, where an invader oligonucleotide was provided to direct the cleavage, show a very distinctive shift, so that the primary cleavage product is smaller than those seen in the non-invasive cleavage. This product is then subject to further nibbling in these conditions, as indicated by the shorter products in these lanes. These invader-directed cleavage products would be easily distinguished from a background of non-specific degradation of the probe oligonucleotide.
When Mg2+ is used as the divalent cation the results are even more distinctive. In lanes 7, 8, 10 and 11 of FIG. 30, where the upstream oligonucleotides were not invasive, minimal nibbling is observed. The products in the DNAPTaq reactions show some accumulation of probe that has been shortened on the 5′ end by one or two nucleotides consistent with previous examination of the action of this enzyme on nicked substrates (Longley et al., supra). When the upstream oligonucleotide is invasive, however, the appearance of the distinctively shifted probe band is seen. These data clearly indicated that it is the invasive 3′ portion of the upstream oligonucleotide that is responsible for fixing the site of cleavage of the downstream probe.
Thus, the above results demonstrate that it is the presence of the free or initially non-annealed nucleotides at the 3′ end of the invader oligonucleotide which mediate the shift in the cleavage site, not just the presence of an oligonucleotide annealed upstream of the probe. Nucleic acid detection assays which employ the use of an invader oligonucleotide are termed “invader-directed cleavage” assays.
EXAMPLE 13
Invader-Directed Cleavage Recognizes Single and Double Stranded Target Molecules in a Background of Non-Target DNA Molecules
For a nucleic acid detection method to be broadly useful, it must be able to detect a specific target in a sample that may contain large amounts of other DNA, e.g., bacterial or human chromosomal DNA. The ability of the invader directed cleavage assay to recognize and cleave either single- or double-stranded target molecules in the presence of large amounts of non-target DNA was examined. In these experiments a model target nucleic acid, M13, in either single or double stranded form (single-stranded M13mp18 is available from Life Technologies, Inc and double-stranded M13mp19 is available from New England Biolabs), was combined with human genomic DNA (Novagen, Madison, Wis.) and then utilized in invader-directed cleavage reactions. Before the start of the cleavage reaction, the DNAs were heated to 95° C. for 15 minutes to completely denature the samples, as is standard practice in assays, such as polymerase chain reaction or enzymatic DNA sequencing, which involve solution hybridization of oligonucleotides to double-stranded target molecules.
For each of the reactions shown in lanes 2-5 of FIG. 31, the target DNA (25 fmole of the ss DNA or 1 pmole of the ds DNA) was combined with 50 pmole of the invader oligonucleotide (SEQ ID NO:35); for the reaction shown in lane 1 the target DNA was omitted. Reactions 1, 3 and 5 also contained 470 ng of human genomic DNA. These mixtures were brought to a volume of 10 μl with distilled water, overlaid with a drop of ChillOutTM evaporation barrier, and brought to 95° C. for 15 minutes. After this incubation period, and still at 95° C., each tube received 10 μl of a mixture comprising 2.25 μof Cleavase® A/G nuclease extract (prepared as described in Example 2) and 5 pmole of the probe oligonucleotide (SEQ ID NO:32), in 20 mM MOPS, pH 7.5 with 0.1% each of Tween 20 and NP-40, 4 mM MnCl2 and 100 mM KCl. The reactions were brought to 62° C. for 15 minutes and stopped by the addition of 12 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. The products of the reactions were visualized by the use of an Hitachi FMBIO fluorescence imager. The results are displayed in FIG. 31.
In FIG. 31, lane 1 contains the products of the reaction containing the probe (SEQ ID NO:32), the invader oligonucleotide (SEQ ID NO:35) and human genomic DNA. Examination of lane 1 shows that the probe and invader oligonucleotides are specific for the target sequence, and that the presence of genomic DNA does not cause any significant background cleavage.
In FIG. 31, lanes 2 and 3 contain reaction products from reactions containing the single-stranded target DNA (M13mp18), the probe (SEQ ID NO:32) and the invader oligonucleotide (SEQ ID NO:35) in the absence or presence of human genomic DNA, respectively. Examination of lanes 2 and 3 demonstrate that the invader detection assay may be used to detect the presence of a specific sequence on a single-stranded target molecule in the presence or absence of a large excess of competitor DNA (human genomic DNA).
In FIG. 31, lanes 4 and 5 contain reaction products from reactions containing the double-stranded target DNA (M13mp18), the probe (SEQ ID NO:32) and the invader oligonucleotide (SEQ ID NO:35) in the absence or presence of human genomic DNA, respectively. Examination of lanes 4 and 5 show that double stranded target molecules are eminently suitable for invader-directed detection reactions. The success of this reaction using a short duplexed molecule, M13mp19, as the target in a background of a large excess of genomic DNA is especially noteworthy as it would be anticipated that the shorter and less complex M13 DNA strands would be expected to find their complementary strand more easily than would the strands of the more complex human genomic DNA. If the M13 DNA reannealed before the probe and/or invader oligonucleotides could bind to the target sequences along the M13 DNA, the cleavage reaction would be prevented. In addition, because the denatured genomic DNA would potentially contain regions complementary to the probe and/or invader oligonucleotides it was possible that the presence of the genomic DNA would inhibit the reaction by binding these oligonucleotides thereby preventing their hybridization to the M13 target. The above results demonstrate that these theoretical concerns are not a problem under the reaction conditions employed above.
In addition to demonstrating that the invader detection assay may be used to detect sequences present in a double-stranded target, these data also show that the presence of a large amount of non-target DNA (470 ng/20 μl reaction) does not lessen the specificity of the cleavage. While this amount of DNA does show some impact on the rate of product accumulation, probably by binding a portion of the enzyme, the nature of the target sequence, whether single- or double-stranded nucleic acid, does not limit the application of this assay.
EXAMPLE 14 
Signal Accumulation in the Invader-Directed Cleavage Assay as a Function of Target Concentration
To investigate whether the invader-directed cleavage assay could be used to indicate the amount of target nucleic acid in a sample, the following experiment was performed. Cleavage reactions were assembled which contained an invader oligonucleotide (SEQ ID NO:35), a labelled probe (SEQ ID NO:32) and a target nucleic acid, M13mp19. A series of reactions, which contained smaller and smaller amounts of the M13 target DNA, was employed in order to examine whether the cleavage products would accumulate in a manner that reflected the amount of target DNA present in the reaction.
The reactions were conducted as follows. A master mix containing enzyme and buffer was assembled. Each 5 μl of the master mixture contained 25 ng of Cleavase® BN nuclease in 20 mM MOPS (pH 7.5) with 0.1% each of Tween 20 and NP-40, 4 mM MnCl2 and 100 mM KCl. For each of the cleavage reactions shown in lanes 4-13 of FIG. 32, a DNA mixture was generated which contained 5 pmoles of the fluorescein-labelled probe oligonucleotide (SEQ ID NO:32), 50 pmoles of the invader oligonucleotide (SEQ ID NO:35) and 100, 50, 10, 5, 1, 0.5, 0.1, 0.05, 0.01 or 0.005 fmoles of single-stranded M13mp19, respectively, for every 5 μl of the DNA mixture. The DNA solutions were covered with a drop of ChillOut® evaporation barrier and brought to 61° C. The cleavage reactions were started by the addition of 5 μl of the enzyme mixture to each of tubes (final reaction volume was 10 μl). After 30 minutes at 61° C., the reactions were terminated by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 90° C. for 1 minutes immediately before electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked) with 7 M urea, in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA. To provide reference (i.e., standards), 1.0, 0.1 and 0.01 pmole aliquots of fluorescein-labelled probe oligonucleotide (SEQ ID NO:32) were diluted with the above formamide solution to a final volume of 18 μl. These reference markers were loaded into lanes 1-3, respectively of the gel. The products of the cleavage reactions (as well as the reference standards) were visualized following electrophoresis by the use of a Hitachi FMBIO fluorescence imager. The results are displayed in FIG. 32.
In FIG. 32, boxes appear around fluorescein-containing nucleic acid (i.e., the cleaved and uncleaved probe molecules) and the amount of fluorescein contained within each box is indicated under the box. The background fluorescence of the gel (see box labelled “background”) was subtracted by the fluoro-imager to generate each value displayed under a box containing cleaved or uncleaved probe products (the boxes are numbered 1-14 at top left with a V followed by a number below the box). The lane marked “M” contains fluoresceinated oligonucleotides which served as markers.
The results shown in FIG. 32, demonstrate that the accumulation of cleaved probe molecules in a fixed-length incubation period reflects the amount of target DNA present in the reaction. The results also demonstrate that the cleaved probe products accumulate in excess of the copy number of the target. This is clearly demonstrated by comparing the results shown in lane 3, in which 10 fmole (0.01 pmole) of uncut probe are displayed with the results shown in 5, where the products which accumulated in response to the presence of 10 fmole of target DNA are displayed. These results show that the reaction can cleave hundreds of probe oligonucleotide molecules for each target molecule present, dramatically amplifying the target-specific signal generated in the invader-directed cleavage reaction.
EXAMPLE 15 
Effect of Saliva Extract on the Invader-Directed Cleavage Assay
For a nucleic acid detection method to be useful in a medical (i.e., a diagnostic) setting, it must not be inhibited by materials and contaminants likely to be found in a typical clinical specimen. To test the susceptibility of the invader-directed cleavage assay to various materials, including but not limited to nucleic acids, glycoproteins and carbohydrates, likely to be found in a clinical sample, a sample of human saliva was prepared in a manner consistent with practices in the clinical laboratory and the resulting saliva extract was added to the invader-directed cleavage assay. The effect of the saliva extract upon the inhibition of cleavage and upon the specificity of the cleavage reaction was examined.
One and one-half milliliters of human saliva were collected and extracted once with an equal volume of a mixture containing phenol:chloroform:isoamyl alcohol (25:24:1). The resulting mixture was centrifuged in a microcentrifuge to separate the aqueous and organic phases. The upper, aqueous phase was transferred to a fresh tube. One-tenth volumes of 3 M NaOAc were added and the contents of the tube were mixed. Two volumes of 100% ethyl alcohol were added to the mixture and the sample was mixed and incubated at room temperature for 15 minutes to allow a precipitate to form. The sample was centrifuged in a microcentrifuge at 13,000 rpm for 5 minutes and the supernatant was removed and discarded. A milky pellet was easily visible. The pellet was rinsed once with 70% ethanol, dried under vacuum and dissolved in 200 μl of 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA (this constitutes the saliva extract). Each μl of the saliva extract was equivalent to 7.5 μl of saliva. Analysis of the saliva extract by scanning ultraviolet spectrophotometry showed a peak absorbance at about 260 nm and indicated the presence of approximately 45 ng of total nucleic acid per μl of extract.
The effect of the presence of saliva extract upon the following enzymes was examined: Cleavase® BN nuclease, Cleavase® A/G nuclease and three different lots of DNAPTaq: AmpliTaq® (Perkin Elmer; a recombinant form of DNAPTaq), AmpliTaq® LD (Perkin-Elmer; a recombinant DNAPTaq preparation containing very low levels of DNA) and Taq DNA polymerase (Fischer). For each enzyme tested, an enzyme/probe mixture was made comprising the chosen amount of enzyme with 5 pmole of the probe oligonucleotide (SEQ ID NO:32) in 10 μl of 20 mM MOPS (pH 7.5) containing 0.1% each of Tween 20 and NP-40, 4 mM MnCl2, 100 mM KCl and 100 μg/ml BSA. The following amounts of enzyme were used: 25 ng of Cleavase® BN prepared as described in Example 8; 2 μl of Cleavase® A/G nuclease extract prepared as described in Example 2; 2.25 μl (11.25 polymerase units) the following DNA polymerases: AmpliTaq® DNA polymerase (Perkin Elmer); AmpliTaq® DNA polymerase LD (low DNA; from Perkin Elmer); Taq DNA polymerase (Fisher Scientific).
For each of the reactions shown in FIG. 33, except for that shown in lane 1, the target DNA (50 fmoles of single-stranded M13mp19 DNA) was combined with 50 pmole of the invader oligonucleotide (SEQ ID NO:35) and 5 pmole of the probe oligonucleotide (SEQ ID NO:32); target DNA was omitted in reaction 1 (lane 1). Reactions 1, 3, 5, 7, 9 and 11 included 1.5 μl of saliva extract. These mixtures were brought to a volume of 5 μl with distilled water, overlaid with a drop of ChillOut® evaporation barrier and brought to 95° C. for 10 minutes. The cleavage reactions were then started by the addition of 5 μl of the desired enzyme/probe mixture; reactions 1, 4 and 5 received Cleavase® A/G nuclease. Reactions 2 and 3 received Cleavase® BN; reactions 6 and 7 received AmpliTaq®; reactions 8 and 9 received AmpliTaq® LD; and reactions 10 and 11 received Taq DNA Polymerase from Fisher Scientific.
The reactions were incubated at 63° C. for 30 minutes and were stopped by the addition of 6 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. The products of the reactions were visualized by the use of an Hitachi FMBIO fluorescence imager, and the results are displayed in FIG. 33.
A pairwise comparison of the lanes shown in FIG. 33 without and with the saliva extract, treated with each of the enzymes, shows that the saliva extract has different effects on each of the enzymes. While the Cleavase® BN nuclease and the AmpliTaq® are significantly inhibited from cleaving in these conditions, the Cleavase® A/G nuclease and AmpliTaq® LD display little difference in the yield of cleaved probe. The preparation of Taq DNA polymerase from Fisher Scientific shows an intermediate response, with a partial reduction in the yield of cleaved product. From the standpoint of polymerization, the three DNAPTaq variants should be equivalent; these should be the same protein with the same amount of synthetic activity. It is possible that the differences observed could be due to variations in the amount of nuclease activity present in each preparation caused by different handling during purification, or by different purification protocols. In any case, quality control assays designed to assess polymerization activity in commercial DNAP preparations would be unlikely to reveal variation in the amount of nuclease activity present. If preparations of DNAPTaq were screened for full 5° nuclease activity (i.e., f the 5° nuclease activity was specifically quantitated), it is likely that the preparations would display sensitivities (to saliva extract) more in line with that observed using Cleavase® A/G nuclease, from which DNAPTaq differs by a very few amino acids.
It is worthy of note that even in the slowed reactions of Cleavase® BN and the DNAPTaq variants there is no noticeable increase in non-specific cleavage of the probe oligonucleotide due to inappropriate hybridization or saliva-borne nucleases.
EXAMPLE 16 
Comparison of Additional 5′ Nucleases in the Invader-Directed Cleavage Assay 
A number of eubacterial Type A DNA polymerases (i.e., Pol I type DNA polymerases) have been shown to function as structure specific endonucleases (Example 1 and Lyamichev et al., supra). In this example, it was demonstrated that the enzymes of this class can also be made to catalyze the invader-directed cleavage of the present invention, albeit not as efficiently as the Cleavase® enzymes.
Cleavase® BN nuclease and Cleavase® A/G nuclease were tested along side three different thermostable DNA polymerases: Thermus aquaticus DNA polymerase (Promega), Thennus thermophilus and Thermus flavus DNA polymerases (Epicentre). The enzyme mixtures used in the reactions shown in lanes 1-11 of FIG. 34 contained the following, each in a volume of 5 μl: Lane 1: 20 mM MOPS (pH 7.5) with 0.1% each of Tween 20 and NP-40, 4 mM MnCl2, 100 mM KCl; Lane 2: 25 ng of Cleavase® BN nuclease in the same solution described for lane 1; Lane 3: 2.25 μl of Cleavase® A/G nuclease extract (prepared as described in Example 2), in the same solution described for lane 1; Lane 4: 2.25 μl of Cleavase® A/G nuclease extract in 20 mM Tris-Cl, (pH 8.5), 4 mM MgCl2 and 100 mM KCl; Lane 5: 11.25 polymerase units of Taq DNA polymerase in the same buffer described for lane 4; Lane 6: 11.25 polymerase units of Tth DNA polymerase in the same buffer described for lane 1; Lane 7: 11.25 polymerase units of Tth DNA polymerase in a 2× concentration of the buffer supplied by the manufacturer, supplemented with 4 mM MnCl2; Lane 8: 11.25 polymerase units of Tth DNA polymerase in a 2× concentration of the buffer supplied by the manufacturer, supplemented with 4 mM MgCl2; Lane 9: 2.25 polymerase units of Tfl DNA polymerase in the same buffer described for lane 1; Lane 10: 2.25 polymerase units of Tfl polymerase in a 2× concentration of the buffer supplied by the manufacturer, supplemented with 4 mM MnCl2; Lane 11: 2.25 polymerase units of Tfl DNA polymerase in a 2× concentration of the buffer supplied by the manufacturer, supplemented with 4 mM MgCl2.
Sufficient target DNA, probe and invader for all 11 reactions was combined into a master mix. This mix contained 550 fmoles of single-stranded M13mp19 target DNA, 550 pmoles of the invader oligonucleotide (SEQ ID NO:35) and 55 pmoles of the probe oligonucleotide (SEQ ID NO:32), each as depicted in FIG. 28c, in 55 pmoles of distilled water. Five μl of the DNA mixture was dispensed into each of 11 labeled tubes and overlaid with a drop of ChillOut® evaporation barrier. The reactions were brought to 63° C. and cleavage was started by the addition of 5 μl of the appropriate enzyme mixture. The reaction mixtures were then incubated at 63° C. temperature for 15 minutes. The reactions were stopped by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 90° C. for 1 minute immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA. Following electrophoresis, the products of the reactions were visualized by the use of an Hitachi FMBIO fluorescence imager, and the results are displayed in FIG. 34. Examination of the results shown in FIG. 34 demonstrates that all of the 5′ nucleases tested have the ability to catalyze invader-directed cleavage in at least one of the buffer systems tested. Although not optimized here, these cleavage agents are suitable for use in the methods of the present invention.
EXAMPLE 17
The Invader-Directed Cleavage Assay can Detect Single Base Differences in Target Nucleic Acid Sequences
The ability of the invader-directed cleavage assay to detect single base mismatch mutations was examined. Two target nucleic acid sequences containing Cleavase® enzyme-resistant phosphorothioate backbones were chemically synthesized and purified by polyacrylamide gel electrophoresis. Targets comprising phosphorothioate backbones were used to prevent exonucleolytic nibbling of the target when duplexed with an oligonucleotide. A target oligonucleotide, which provides a target sequence that is completely complementary to the invader oligonucleotide (SEQ ID NO:35) and the probe oligonucleotide (SEQ ID NO:32), contained the following sequence: 5′-CCTTTCGCTTTCTTCCCTTCCTTTCTCGCCACGTTCGCCGGC-3′ (SEQ ID NO:36). A second target sequence containing a single base change relative to SEQ ID NO:36 was synthesized: 5′-CCTTTCGCTCTCTTCCCTTCCTTTCTCGCCACGTTCGCCGGC-3 (SEQ ID NO:37; the single base change relative to SEQ ID NO:36 is shown using bold and underlined type). The consequent mismatch occurs within the “Z” region of the target as represented in FIG. 25.
To discriminate between two target sequences which differ by the presence of a single mismatch), invader-directed cleavage reactions were conducted using two different reaction temperatures (55° C. and 60° C.). Mixtures containing 200 fmoles of either SEQ ID NO:36 or SEQ ID NO:37, 3 pmoles of fluorescein-labelled probe oligonucleotide (SEQ ID NO:32), 7.7 pmoles of invader oligonucleotide (SEQ ID NO:35) and 2 μl of Cleavase® A/G nuclease extract (prepared as described in Example 2) in 9 μl of 10 mM MOPS (pH 7.4) with 50 mM KCl were assembled, covered with a drop of ChillOut® evaporation barrier and brought to the appropriate reaction temperature. The cleavage reactions were initiated by the addition of 1 μl of 20 mM MgCl2. After 30 minutes at either 55° C. or 60° C., 10 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes was added to stop the reactions. The reaction mixtures where then heated to 90° C. for one minute prior to loading 4 μl onto 20% denaturing polyacrylamide gels. The resolved reaction products were visualized using a Hitachi FMBIO fluorescence imager. The resulting image is shown in FIG. 35.
In FIG. 35, lanes 1 and 2 show the products from reactions conducted at 55° C.; lanes 3 and 4 show the products from reactions conducted at 60° C. Lanes 1 and 3 contained products from reactions containing SEQ ID NO:36 (perfect match to probe) as the target. Lanes 2 and 4 contained products from reactions containing SEQ ID NO:37 (single base mis-match with probe) as the target. The target that does not have a perfect hybridization match (i.e., complete complementarity) with the probe will not bind as strongly, i.e., the Tm of that duplex will be lower than the Tm of the same region if perfectly matched. The results presented here show that reaction conditions can be varied to either accommodate the mis-match (e.g., by lowering the temperature of the reaction) or to exclude the binding of the mismatched sequence (e.g., by raising the reaction temperature).
The results shown in FIG. 35 demonstrate that the specific cleavage event which occurs in invader-directed cleavage reactions can be eliminated by the presence of a single base mis-match between the probe oligonucleotide and the target sequence. Thus, reaction conditions can be chosen so as to exclude the hybridization of mis-matched invader-directed cleavage probes thereby diminishing or even eliminating the cleavage of the probe. In an extension of this assay system, multiple cleavage probes, each possessing a separate reporter molecule (i.e., a unique label), could also be used in a single cleavage reaction, to simultaneously probe for two or more variants in the same target region. The products of such a reaction would allow not only the detection of mutations which exist within a target molecule, but would also allow a determination of the relative concentrations of each sequence (i.e., mutant and wild type or multiple different mutants) present within samples containing a mixture of target sequences. When provided in equal amounts, but in a vast excess (e.g., at least a 100-fold molar excess; typically at least 1 pmole of each probe oligonucleotide would be used when the target sequence was present at about 10 fmoles or less) over the target and used in optimized conditions. As discussed above, any differences in the relative amounts of the target variants will not affect the kinetics of hybridization, so the amounts of cleavage of each probe will reflect the relative amounts of each variant present in the reaction.
The results shown in the example clearly demonstrate that the invader-directed cleavage reaction can be used to detect single base difference between target nucleic acids.
EXAMPLE 18 
The Invader-Directed Cleavage Reaction is Insensitive to Large Changes in Reaction Conditions
The results shown above demonstrated that the invader-directed cleavage reaction can be used for the detection of target nucleic acid sequences and that this assay can be used to detect single base difference between target nucleic acids. These results demonstrated that 5′ nucleases (e.g., Cleavase®BN, Cleavase® A/G, DNAPTaq, DNAPTth, DNAPTfl) could be used in conjunction with a pair of overlapping oligonucleotides as an efficient way to recognize nucleic acid targets. In the experiments below it is demonstrated that invasive cleavage reaction is relatively insensitive to large changes in conditions thereby making the method suitable for practice in clinical laboratories.
The effects of varying the conditions of the cleavage reaction were examined for their effect(s) on the specificity of the invasive cleavage and the on the amount of signal accumulated in the course of the reaction. To compare variations in the cleavage reaction a “standard” invader cleavage reaction was first defined. In each instance, unless specifically stated to be otherwise, the indicated parameter of the reaction was varied, while the invariant aspects of a particular test were those of this standard reaction. The results of these tests are either shown in FIGS. 38-40, or the results described below.
a) The Standard Invader-Directed Cleavage Reaction
The standard reaction was defined as comprising 1 fmole of M13mp18 single-stranded target DNA (New England Biolabs), 5 pmoles of the labeled probe oligonucleotide (SEQ ID NO:38), 10 pmole of the upstream invader oligonucleotide (SEQ ID NO:39) and 2 units of Cleavase® A/G in 10 μl of 10 mM MOPS, pH 7.5 with 100 mM KCl, 4 mM MnCl2, and 0.05% each Tween-20 and Nonidet-P40. For each reaction, the buffers, salts and enzyme were combined in a volume of 5 μl; the DNAs (target and two oligonucleotides) were combined in 5 μl of dH2O and overlaid with a drop of ChillOut® evaporation barrier. When multiple reactions were performed with the same reaction constituents, these formulations were expanded proportionally.
Unless otherwise stated, the sample tubes with the DNA mixtures were warmed to 61° C., and the reactions were started by the addition of 5 μl of the enzyme mixture. After 20 minutes at this temperature, the reactions were stopped by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes. Samples were heated to 75° C. for 2 minutes immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. The products of the reactions were visualized by the use of an Hitachi FMBIO fluorescence imager. In each case, the uncut probe material was visible as an intense black band or blob, usually in the top half of the panel, while the desired products of invader specific cleavage were visible as one or two narrower black bands, usually in the bottom half of the panel. Under some reaction conditions, particularly those with elevated salt concentrations, a secondary cleavage product is also visible (thus generating a doublet). Ladders of lighter grey bands generally indicate either exonuclease nibbling of the probe oligonucleotide or heat-induced, non-specific breakage of the probe.
FIG. 37 depicts the annealing of the probe and invader oligonucleotides to regions along the M13mp18 target molecule (the bottom strand). In FIG. 37 only a 52 nucleotide portion of the M13mp18 molecule is shown; this 52 nucleotide sequence is listed in SEQ ID NO:31 (this sequence is identical in both M13mp18 and M13mp19). The probe oligonucleotide (top strand) contains a Cy3 amidite label at the 5′ end; the sequence of the probe is 5′-AGAAAGGAAGGGAAGAAAGCGAAAGGT-3′ (SEQ ID NO:38. The bold type indicates the presence of a modified base (2′—O—CH3). Cy3 amidite (Pharmacia) is a indodicarbocyanine dye amidite which can be incorporated at any position during the synthesis of oligonucleotides; Cy3 fluoresces in the yellow region (excitation and emission maximum of 554 and 568 nm, respectively). The invader oligonucleotide (middle strand) has the following sequence:

    5′-GCCGGCGAACGTGGCGAGAAAGGA-3′ (SEQ ID NO:39).


b) KCl Titration
FIG. 38 shows the results of varying the KCl concentration in combination with the use of 2 mM MnCl2, in an otherwise standard reaction. The reactions were performed in duplicate for confirmation of observations; the reactions shown in lanes 1 and 2 contained no added KCl, lanes 3 and 4 contained KCl at 5 mM, lanes 5 and 6 contained 25 mM KCl, lanes 7 and 8 contained 50 mM KCl, lanes 9 and 10 contained 100 mM KCl and lanes 11 and 12 contained 200 mM KCl. These results show that the inclusion of KCl allows the generation of a specific cleavage product. While the strongest signal is observed at the 100 mM KCl concentration, the specificity of signal in the other reactions with KCl at or above 25 mM indicates that concentrations in the full range (i.e., 25-200 mM) may be chosen if it is so desirable for any particular reaction conditions.
As shown in FIG. 38, the invader-directed cleavage reaction requires the presence of salt (e.g., KCl) for effective cleavage to occur. In other reactions, it has been found that KCl can inhibit the activity of certain Cleavase® enzymes when present at concentrations above about 25 mM (For example, in cleavage reactions using the S-60 oligonucleotide shown in FIG. 26, in the absence of primer, the Cleavase® BN enzyme loses approximately 50% of its activity in 50 mM KCl). Therefore, the use of alternative salts in the invader-directed cleavage reaction was examined. In these experiments, the potassium ion was replaced with either Na+ or Li+ or the chloride ion was replaced with glutamic acid. The replacement of KCl with alternative salts is described below in sections c-e.
c) NaCl Titration
NaCl was used in place of KCl at 75, 100, 150 or 200 mM, in combination with the use 2 mM MnCl2, in an otherwise standard reaction. These results showed that NaCl can be used as a replacement for KCl in the invader-directed cleavage reaction, with like concentration giving like results, (i.e., the presence of NaCl, like KCl, enhances product accumulation).
d) LiCl Titration
LiCl was used in place of KCl in otherwise standard reactions. Concentrations tested were 25, 50, 75, 100, 150 and 200 mM LiCl. The results demonstrated that LiCl can be used as a suitable replacement for KCl in the invader-directed cleavage reaction (i.e., the presence of LiCl, like KCl, enhances product accumulation), in concentrations of about 100 mM or higher.
e) KGlu Titration
The results of using a glutamate salt of potassium (KGlu) in place of the more commonly used chloride salt (KCl) in reactions performed over a range of temperatures were examined. KGlu has been shown to be a highly effective salt source for some enzymatic reactions, showing a broader range of concentrations which permit maximum enzymatic activity [Leirmo et al. (1987) Biochem. 26:2095]. The ability of KGlu to facilitate the annealing of the probe and invader oligonucleotides to the target nucleic acid was compared to that of LiCI. In these experiments, the reactions were run for 15 minutes, rather than the standard 20 minutes, in standard reactions that replaced KCl 200 mM, 300 mM or 400 mM KGlu. The reactions were run at 65° C., 67° C., 69° C. or 71° C. The results showed demonstrated that KGlu was very effective as a salt in the invasive cleavage reactions, with full activity apparent even at 400 mM KGlu, though at the lowest temperature cleavage was reduced by about 30% at 300 mM KGlu, and by about 90% to 400 mM KGlu.
f) MnCl2 And MgCl2 Titration And Ability To Replace MnCl2 With MgCl2 
In some instances it may be desirable to perform the invasive cleavage reaction in the presence of Mg2+, either in addition to, or in place of Mn2+ as the necessary divalent cation required for activity of the enzyme employed. For example, some common methods of preparing DNA from bacterial cultures or tissues use MgCl2 in solutions which are used to facilitate the collection of DNA by precipitation. In addition, elevated concentrations (i.e., greater than 5 mM) of divalent cation can be used to facilitate hybridization of nucleic acids, in the same way that the monovalent salts were used above, thereby enhancing the invasive cleavage reaction. In this experiment, the tolerance of the invasive cleavage reaction was examined for 1) the substitution of MgCl2 for MnCl2 and for the ability to produce specific product in the presence of increasing concentrations of MgCl2 and MnCl2.
FIG. 39 shows the results of either varying the concentration of MnCl2 from 2 mM to 8 mM, replacing the MnCl2 with MgCl2 at 2 to 4 mM, or of using these components in combination in an otherwise standard reaction. The reactions analyzed in lanes 1 and 2 contained 2 mM each MnCl2 and MgCl2, lanes 3 and 4 contained 2 mM MnCl2 only, lanes 5 and 6 contained 3 mM MnCl2, lanes 7 and 8 contained 4 mM MnCl2, lanes 9 and 10 contained 8 mM MnCl2. The reactions analyzed in lanes 11 and 12 contained 2 mM MgCl2 and lanes 13 and 14 contained 4 mM MgCl2. These results show that both MnCl2 and MgCl2 can be used as the necessary divalent cation to enable the cleavage activity of the Cleavase® A/G enzyme in these reactions and that the invasive cleavage reaction can tolerate a broad range of concentrations of these components.
In addition to examining the effects of the salt environment on the rate of product accumulation in the invasive cleavage reaction, the use of reaction constituents shown to be effective in enhancing nucleic acid hybridization in either standard hybridization assays (e.g., blot hybridization) or in ligation reactions was examined. These components may act as volume excluders, increasing the effective concentration of the nucleic acids of interest and thereby enhancing hybridization, or they may act as charge-shielding agents to minimize repulsion between the highly charged backbones of the nucleic acids strands. The results of these experiments are described in sections g and h below.
g) Effect Of CTAB Addition
The polycationic detergent cetyltrietheylammonium bromide (CTAB) has been shown to dramatically enhance hybridization of nucleic acids [Pontius and Berg (1991) Proc. Natl. Acad. Sci. USA 88:8237]. We examined the effect of adding the detergent CTAB in concentrations from 100 mM to 1 mM to invasive cleavage reactions in which 150 mM LiCl was used in place of the KCl in otherwise standard reactions. These results showed that 200 mM CTAB may have a very moderate enhancing effect under these reaction conditions, and the presence of CTAB in excess of about 500 μM was inhibitory to the accumulation of specific cleavage product.
h) Effect Of PEG Addition
We examined the effect of adding polyethylene glycol (PEG) at 4.8 or 12% (w/v) concentrations to otherwise standard reactions. The effects of increasing the reaction temperature of the PEG-containing reactions was examined by performing duplicate sets of PEG titration reactions at 61° C. and 65° C. The results showed that at all percentages tested, and at both temperatures tested, the inclusion of PEG substantially eliminated the production of specific cleavage product.
In addition to, the presence of 1×Denhardts in the reaction mixture was found to have no adverse effect upon the cleavage reaction [50× Denhardts contains per 500 ml: 5 g Ficoll, 5 g polyvinylpyrrolidone, 5 g BSA]. Further, the presence of each component of Denhardt's was examined individually (i.e., Ficoll alone, polyvinylpyrrolidone alone, BSA alone) for the effect upon the invader-directed cleavage reaction; no adverse effect was observed.
i) Effect Of The Addition Of Stabilizing Agents
Another approach to enhancing the output of the invasive cleavage reaction is to enhance the activity of the enzyme employed, either by increasing its stability in the reaction environment or by increasing its turnover rate. Without regard to the precise mechanism by which various agents operate in the invasive cleavage reaction, a number of agents commonly used to stabilize enzymes during prolonged storage were tested for the ability to enhance the accumulation of specific cleavage product in the invasive cleavage reaction.
We examined the effects of adding glycerol at 15% and of adding the detergents Tween-20 and Nonidet-P40 at 1.5%, alone or in combination, in otherwise standard reactions. The results demonstrated that under these conditions these adducts had little or no effect on the accumulation of specific cleavage product.
The effects of adding gelatin to reactions in which the salt identity and concentration were varied from the standard reaction. The results demonstrated that in the absence of salt the gelatin had a moderately enhancing effect on the accumulation of specific cleavage product, but when either salt (KCl or LiCl) was added to reactions performed under these conditions, increasing amounts of gelatin reduced the product accumulation.
j) Effect Of Adding Large Amounts Of Non-Target Nucleic Acid
In detecting specific nucleic acid sequences within samples, it is important to determine if the presence of additional genetic material (i.e., non-target nucleic acids) will have a negative effect on the specificity of the assay. In this experiment, the effect of including large amounts of non-target nucleic acid, either DNA or RNA, on the specificity of the invasive cleavage reaction was examined. The data was examined for either an alteration in the expected site of cleavage, or for an increase in the nonspecific degradation of the probe oligonucleotide.
FIG. 40 shows the effects of adding non-target nucleic acid (e.g., genomic DNA or tRNA) to an invasive cleavage reaction performed at 65° C., with 150 mM LiCl in place of the KCl in the standard reaction. The reactions assayed in lanes 1 and 2 contained 235 and 470 ng of genomic DNA, respectively. The reactions analyzed in lanes 3, 4, 5 and 6 contained 100 ng, 200 ng, 500 ng and 1 μg of tRNA, respectively. Lane 7 represents a control reaction which contained no added nucleic acid beyond the amounts used in the standard reaction. The results shown in FIG. 40 demonstrate that the inclusion of non-target nucleic acid in large amounts could visibly slow the accumulation of specific cleavage product (while not limiting the invention to any particular mechanism, it is thought that the additional nucleic acid competes for binding of the enzyme with the specific reaction components). In additional experiments it was found that the effect of adding large amounts of non-target nucleic acid can be compensated for by increasing the enzyme in the reaction. The data shown in FIG. 40 also demonstrate that a key feature of the invasive cleavage reaction, the specificity of the detection, was not compromised by the presence of large amounts of non-target nucleic acid.
In addition to the data presented above, invasive cleavage reactions were run with succinate buffer at pH 5.9 in place of the MOPS buffer used in the “standard” reaction; no adverse effects were observed.
The data shown in FIGS. 38-40 and described above demonstrate that the invasive cleavage reaction can be performed using a wide variety of reaction conditions and is therefore suitable for practice in clinical laboratories.
EXAMPLE 19
Detection of RNA Targets by Invader-Directed Cleavage
In addition to the clinical need to detect specific DNA sequences for infectious and genetic diseases, there is a need for technologies that can quantitatively detect target nucleic acids that are composed of RNA. For example, a number of viral agents, such as hepatitis C virus (HCV) and human immunodeficiency virus (HIV) have RNA genomic material, the quantitative detection of which can be used as a measure of viral load in a patient sample. Such information can be of critical diagnostic or prognostic value.
Hepatitis C virus (HCV) infection is the predominant cause of post-transfusion non-A, non-B (NANB) hepatitis around the world. In addition, HCV is the major etiologic agent of hepatocellular carcinoma (HCC) and chronic liver disease world wide. The genome of HCV is a small (9.4 kb) RNA molecule. In studies of transmission of HCV by blood transfusion it has been found the presence of HCV antibody, as measured in standard immunological tests, does not always correlate with the infectivity of the sample, while the presence of HCV RNA in a blood sample strongly correlates with infectivity. Conversely, serological tests may remain negative in immunosuppressed infected individuals, while HCV RNA may be easily detected [J. A. Cuthbert (1994) Clin. Microbiol. Rev. 7:505].
The need for and the value of developing a probe-based assay for the detection the HCV RNA is clear. The polymerase chain reaction has been used to detect HCV in clinical samples, but the problems associated with carry-over contamination of samples has been a concern. Direct detection of the viral RNA without the need to perform either reverse transcription or amplification would allow the elimination of several of the points at which existing assays may fail.
The genome of the positive-stranded RNA hepatitis C virus comprises several regions including 5′ and 3′ noncoding regions (i.e., 5′ and 3′ untranslated regions) and a polyprotein coding region which encodes the core protein (C), two envelope glycoproteins (E1 and E2/NS1) and six nonstructural glycoproteins (NS2-NS5b). Molecular biological analysis of the HCV genome has showed that some regions of the genome are very highly conserved between isolates, while other regions are fairly rapidly changeable. The 5′ noncoding region (NCR) is the most highly conserved region in the HCV. These analyses have allowed these viruses to be divided into six basic genotype groups, and then further classified into over a dozen sub-types [the nomenclature and division of HCV genotypes is evolving; see Altamirano et al., J. Infect. Dis. 171:1034 (1995) for a recent classification scheme].
In order to develop a rapid and accurate method of detecting HCV present in infected individuals, the ability of the invader-directed cleavage reaction to detect HCV RNA was examined. Plasmids containing DNA derived from the conserved 5′-untranslated region of six different HCV RNA isolates were used to generate templates for in vitro transcription. The HCV sequences contained within these six plasmids represent genotypes 1 (four sub-types represented; 1a, 1b, 1c, and Δ1c), 2, and 3. The nomenclature of the HCV genotypes used herein is that of Simmonds et al. [as described in Altamirano et at., supra]. The Δ1c subtype was used in the model detection reaction described below.
a) Generation Of Plasmids Containing HCV Sequences
Six DNA fragments derived from HCV were generated by RT-PCR using RNA extracted from serum samples of blood donors; these PCR fragments were a gift of Dr. M. Altamirano (University of British Columbia, Vancouver). These PCR fragments represent HCV sequences derived from HCV genotypes 1a, 1b, 1c, Δ1c, 2c and 3a.
The RNA extraction, reverse transcription and PCR were performed using standard techniques (Altamirano et al., supra). Briefly, RNA was extracted from 100 μl of serum using guanidine isothiocyanate, sodium lauryl sarkosate and phenol-chloroform [Inchauspe et al., Hepatology 14:595 (1991)]. Reverse transcription was performed according to the manufacturer's instructions using a GeneAmp rTh reverse transcriptase RNA PCR kit (Perkin-Elmer) in the presence of an external antisense primer, HCV342. The sequence of the HCV342 primer is 5′-GGTTTTTCTTTGAGGTTTAG-3′ (SEQ ID NO:40). Following termination of the RT reaction, the sense primer HCV7 [5′-GCGACACTCCACCATAGAT-3′ (SEQ ID NO:41)] and magnesium were added and a first PCR was performed. Aliquots of the first PCR products were used in a second (nested) PCR in the presence of primers HCV46 [5′-CTGTCTTCACGCAGAAAGC-3′ (SEQ ID NO:42)] and HCV308 [5′-GCACGGTCTACGAGACCTC-3′ (SEQ ID NO:43)]. The PCRs produced a 281 bp product which corresponds to a conserved 5′noncoding region (NCR) region of HCV between positions −284 and −4 of the HCV genome (Altramirano et al., supra).
The six 281 bp PCR fragments were used directly for cloning or they were subjected to an additional amplification step using a 50 μl PCR comprising approximately 100 fmoles of DNA, the HCV46 and HCV308 primers at 0.1 μM, 100 μM of all four dNTPs and 2.5 units of Taq DNA polymerase in a buffer containing 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 1.5 mM MgCl2 and 0.1% Tween 20. The PCRs were cycled 25 times at 96° C. for 45 sec., 55° C. for 45 sec. and 72° C. for 1 min. Two microliters of either the original DNA samples or the reamplified PCR products were used for cloning in the linear pT7Blue T-vector (Novagen) according to manufacturer's protocol. After the PCR products were ligated to the pT7Blue T-vector, the ligation reaction mixture was used to transform competent JM109 cells (Promega). Clones containing the pT7Blue T-vector with an insert were selected by the presence of colonies having a white color on LB plates containing 40 μg/ml X-Gal, 40 μg/ml IPTG and 50 μg/ml ampicillin. Four colonies for each PCR sample were picked and grown overnight in 2 ml LB media containing 50 μg/ml carbenicillin. Plasmid DNA was isolated using the following alkaline miniprep protocol. Cells from 1.5 ml of the overnight culture were collected by centrifugation for 2 min. in a microcentrifuge (14K rpm), the supernatant was discarded and the cell pellet was resuspended in 50 μl TE buffer with 10 μg/ml RNAse A (Pharmacia). One hundred microliters of a solution containing 0.2 N NaOH, 1% SDS was added and the cells were lysed for 2 min. The lysate was gently mixed with 100 μl of 1.32 M potassium acetate, pH 4.8, and the mixture was centrifuged for 4 min. in a microcentrifuge (14K rpm); the pellet comprising cell debris was discarded. Plasmid DNA was precipitated from the supernatant with 200 μl ethanol and pelleted by centrifugation a microcentrifuge (14K rpm). The DNA pellet was air dried for 15 min. and was then redissolved in 50 μl TE buffer (10 mM Tris-HCl, pH 7.8, 1 mM EDTA).
b) Reamplification Of HCV Clones To Add The Phage T7 Promoter For Subsequent In Vitro Transcription
To ensure that the RNA product of transcription had a discrete 3′ end it was necessary to create linear transcription templates which stopped at the end of the HCV sequence. These fragments were conveniently produced using the PCR to reamplify the segment of the plasmid containing the phage promoter sequence and the HCV insert. For these studies, the clone of HCV type Δ1c was reamplified using a primer that hybridizes to the T7 promoter sequence: 5′-TAATACGACTCACTATAGGG-3′ (SEQ ID NO:44; “the T7 promoter primer”) (Novagen) in combination with the 3′ terminal HCV-specific primer HCV308 (SEQ ID NO:43). For these reactions, 1 μl of plasmid DNA (approximately 10 to 100 ng) was reamplified in a 200 μl PCR using the T7 and HCV308 primers as described above with the exception that 30 cycles of amplification were employed. The resulting amplicon was 354 bp in length. After amplification the PCR mixture was transferred to a fresh 1.5 ml microcentrifuge tube, the mixture was brought to a final concentration of 2 M NH4OAc, and the products were precipitated by the addition of one volume of 100% isopropanol. Following a 10 min. incubation at room temperature, the precipitates were collected by centrifugation, washed once with 80% ethanol and dried under vacuum. The collected material was dissolved in 100 μl nuclease-free distilled water (Promega).
Segments of RNA were produced from this amplicon by in vitro transcription using the RiboMAXTM Large Scale RNA Production System (Promega) in accordance with the manufacturer's instructions, using 5.3 μg of the amplicon described above in a 100 μl reaction. The transcription reaction was incubated for 3.75 hours, after which the DNA template was destroyed by the addition of 5-6 μl of RQ1 RNAse-free DNAse (1 unit/μl) according to the RiboMAXTM kit instructions. The reaction was extracted twice with phenol/chloroform/isoamyl alcohol (50:48:2) and the aqueous phase was transferred to a fresh microcentrifuge tube. The RNA was then collected by the addition of 10 μl of 3M NH4QAc, pH 5.2 and 110 μl of 100% isopropanol. Following a 5 min. incubation at 4° C., the precipitate was collected by centrifugation, washed once with 80% ethanol and dried under vacuum. The sequence of the resulting RNA transcript (HCV1.1 transcript) is listed in SEQ ID NO:45.
c) Detection Of The HCV1.1 Transcript In The Invader-Directed Cleavage Assay
Detection of the HCV1.1 transcript was tested in the invader-directed cleavage assay using an HCV-specific probe oligonucleotide [5′-CCGGTCGTCCTGGCAATXCC-3′ (SEQ ID NO:46); X indicates the presence of a fluorescein dye on an abasic linker) and an HCV-specific invader oligonucleotide [5′-GTTTATCCAAGAAAGGACCCGGTC-3′ (SEQ ID NO:47)] that causes a 6-nucleotide invasive cleavage of the probe.
Each 10 μl of reaction mixture comprised 5 pmole of the probe oligonucleotide (SEQ ID NO:46) and 10 pmole of the invader oligonucleotide (SEQ ID NO:47) in a buffer of 10 mM MOPS, pH 7.5 with 50 mM KCl, 4 mM MnCl2, 0.05% each Tween-20 and Nonidet-P40 and 7.8 units RNasin® ribonuclease inhibitor (Promega). The cleavage agents employed were Cleavase® A/G (used at 5.3 ng/10 μl reaction) or DNAPTth (used at 5 polymerase units/10 μl reaction). The amount of RNA target was varied as indicated below. When RNAse treatment is indicated, the target RNAs were pre-treated with 10 μg of RNase A (Sigma) at 37° C. for 30 min. to demonstrate that the detection was specific for the RNA in the reaction and not due to the presence of any residual DNA template from the transcription reaction. RNase-treated aliquots of the HCV RNA were used directly without intervening purification.
For each reaction, the target RNAs were suspended in the reaction solutions as described above, but lacking the cleavage agent and the MnCl2 for a final volume of 10 μl, with the invader and probe at the concentrations listed above. The reactions were warmed to 46° C. and the reactions were started by the addition of a mixture of the appropriate enzyme with MnCl2. After incubation for 30 min. at 46° C., the reactions were stopped by the addition of 8 μl of 95% formamide, 10 mM EDTA and 0.02% methyl violet (methyl violet loading buffer). Samples were then resolved by electrophoresis through a 15% denaturing polyacrylamide gel (19:1 cross-linked), containing 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. Following electrophoresis, the labeled reaction products were visualized using the FMBIO-100 Image Analyzer (Hitachi), with the resulting imager scan shown in FIG. 41.
In FIG. 41, the samples analyzed in lanes 1-4 contained 1 pmole of the RNA target, the reactions shown in lanes 5-8 contained 100 fmoles of the RNA target and the reactions shown in lanes 9-12 contained 10 fmoles of the RNA target. All odd-numbered lanes depict reactions performed using Cleavase® A/G enzyme and all even-numbered lanes depict reactions performed using DNAPTth. The reactions analyzed in lanes 1, 2, 5, 6, 9 and 10 contained RNA that had been pre-digested with RNase A. These data demonstrate that the invasive cleavage reaction efficiently detects RNA targets and further, the absence of any specific cleavage signal in the RNase-treated samples confirms that the specific cleavage product seen in the other lanes is dependent upon the presence of input RNA.
EXAMPLE 20 
The Fate of the Target RNA in the Invader-Directed Cleavage Reaction
In this example, the fate of the RNA target in the invader-directed cleavage reaction was examined. As shown above in Example 1D, when RNAs are hybridized to DNA oligonucleotides, the 5′ nucleases associated with DNA polymerases can be used to cleave the RNAs; such cleavage can be suppressed when the 5′ arm is long or when it is highly structured [Lyamichev et al. (1993) Science 260:778 and U.S. Pat. No. 5,422,253, the disclosure of which is herein incorporated by reference]. In this experiment, the extent to which the RNA target would be cleaved by the cleavage agents when hybridized to the detection oligonucleotides (i.e., the probe and invader oligonucleotides) was examined using reactions similar to those described in Example 20, performed using fluorescein-labeled RNA as a target.
Transcription reactions were performed as described in Example 19 with the exception that 2% of the UTP in the reaction was replaced with fluorescein-12-UTP (Boehringer Mannheim) and 5.3 μg of the amplicon was used in a 100 μl reaction. The transcription reaction was incubated for 2.5 hours, after which the DNA template was destroyed by the addition of 5-6 μl of RQ1 RNAse-free DNAse (1 unit/μl) according to the RiboMAXTM kit instructions. The organic extraction was omitted and the RNA was collected by the addition of 10 μl of 3M NaOAc, pH 5.2 and 110 μl of 100% isopropanol. Following a 5 min. incubation at 4° C., the precipitate was collected by centrifugation, washed once with 80% ethanol and dried under vacuum. The resulting RNA was dissolved in 100 μl of nuclease-free water. 50% of the sample was purified by electrophoresis through a 8% denaturing polyacrylamide gel (19:1 cross-linked), containing 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. The gel slice containing the full-length material was excised and the RNA was eluted by soaking the slice overnight at 4° C. in 200 μl of 10 mM Tris-Cl, pH 8.0, 0.1 mM EDTA and 0.3 M NaOAc. The RNA was then precipitated by the addition of 2.5 volumes of 100% ethanol. After incubation at −20° C. for 30 min., the precipitates were recovered by centrifugation, washed once with 80% ethanol and dried under vacuum. The RNA was dissolved in 25 μl of nuclease-free water and then quantitated by UV absorbance at 260 nm.
Samples of the purified RNA target were incubated for 5 or 30 min. in reactions that duplicated the Cleavase® A/G and DNAPTth invader reactions described in Example 20 with the exception that the reactions lacked probe and invader oligonucleotides. Subsequent analysis of the products showed that the RNA was very stable, with a very slight background of non-specific degradation, appearing as a gray background in the gel lane. The background was not dependent on the presence of enzyme in the reaction.
Invader detection reactions using the purified RNA target were performed using the probe/invader pair described in Example 19 (SEQ ID NOS:46 and 47). Each reaction included 500 fmole of the target RNA, 5 pmoles of the fluorescein-labeled probe and 10 pmoles of the invader oligonucleotide in a buffer of 10 mM MOPS, pH 7.5 with 150 mM LiCl, 4 mM MnCl2, 0.05% each Tween-20 and Nonidet-P40 and 39 units RNAsine (Promega). These components were combined and warmed to 50° C. and the reactions were started by the addition of either 53 ng of Cleavase® A/G or 5 polymerase units of DNAPTth. The final reaction volume was 10 μ. After 5 min at 50° C., 5 μl aliquots of each reaction were removed to tubes containing 4 μl of 95% formamide, 10 mM EDTA and 0.02% methyl violet. The remaining aliquot received a drop of ChillOut® evaporation barrier and was incubated for an additional 25 min. These reactions were then stopped by the addition of 4 μl of the above formamide solution. The products of these reactions were resolved by electrophoresis through separate 20% denaturing polyacrylamide gels (19:1 cross-linked), containing 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. Following electrophoresis, the labeled reaction products were visualized using the FMBIO-100 Image Analyzer (Hitachi), with the resulting imager scans shown in FIGS. 42A (5 min reactions) and 42B (30 min. reactions).
In FIG. 53 the target RNA is seen very near the top of each lane, while the labeled probe and its cleavage products are seen just below the middle of each panel. The FMBIO-100 Image Analyzer was used to quantitate the fluorescence signal in the probe bands. In each panel, lane 1 contains products from reactions performed in the absence of a cleavage agent, lane 2 contains products from reactions performed using Cleavase® A/G and lane 3 contains products from reactions performed using DNAPTth.
Quantitation of the fluorescence signal in the probe bands revealed that after a 5 min. incubation, 12% or 300 fmole of the probe was cleaved by the Cleavase® A/G and 29% or 700 fmole was cleaved by the DNAPTth. After a 30 min. incubation, Cleavase® A/G had cleaved 32% of the probe molecules and DNAPTth had cleaved 70% of the probe molecules. (The images shown in FIGS. 42A and 42B were printed with the intensity adjusted to show the small amount of background from the RNA degradation, so the bands containing strong signals are saturated and therefore these images do not accurately reflect the differences in measured fluorescence)
The data shown in FIG. 42 clearly shows that, under invasive cleavage conditions, RNA molecules are sufficiently stable to be detected as a target and that each RNA molecule can support many rounds of probe cleavage.
EXAMPLE 21 
Titration of Target RNA in the Invader-Directed Cleavage Assay
One of the primary benefits of the invader-directed cleavage assay as a means for detection of the presence of specific target nucleic acids is the correlation between the amount of cleavage product generated in a set amount of time and the quantity of the nucleic acid of interest present in the reaction. The benefits of quantitative detection of RNA sequences was discussed in Example 19. In this example, we demonstrate the quantitative nature of the detection assay through the use of various amounts of target starting material. In addition to demonstrating the correlation between the amounts of input target and output cleavage product, these data graphically show the degree to which the RNA target can be recycled in this assay
The RNA target used in these reactions was the fluorescein-labeled material described in Example 20 (i.e., SEQ ID NO:45). Because the efficiency of incorporation of the fluorescein-12-UTP by the T7 RNA polymerase was not known, the concentration of the RNA was determined by measurement of absorbance at 260 nm, not by fluorescence intensity. Each reaction comprised 5 pmoles of the fluorescein-labeled probe (SEQ ID NO:46) and 10 pmoles of the invader oligonucleotide (SEQ ID NO:47) in a buffer of 10 mM MOPS, pH 7.5 with 150 mM LiCl, 4 mM MnCl2, 0.05% each Tween-20 and Nonidet-P40 and 39 units of RNAsin® (Promega). The amount of target RNA was varied from 1 to 100 fmoles, as indicated below. These components were combined, overlaid with ChillOut® evaporation barrier and warmed to 50° C.; the reactions were started by the addition of either 53 ng of Cleavase® A/G or 5 polymerase units of DNAPTth, to a final reaction volume of 10 μl. After 30 minutes at 50° C., reactions were stopped by the addition of 8 μl of 95% formamide, 10 mM EDTA and 0.02% methyl violet. The unreacted markers in lanes 1 and 2 were diluted in the same total volume (18 μl). The samples were heated to 90° C. for 1 minute and 2.5 μl of each of these reactions were resolved by electrophoresis through a 20% denaturing polyacrylamide gel (19:1 cross link) with 7M urea in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA, and the labeled reaction products were visualized using the FMBIO-100 Image Analyzer (Hitachi), with the resulting imager scans shown in FIG. 43.
In FIG. 43, lanes 1 and 2 show 5 pmoles of uncut probe and 500 fmoles of untreated RNA, respectively. The probe is the very dark signal near the middle of the panel, while the RNA is the thin line near the top of the panel. These RNAs were transcribed with a 2% substitution of fluorescein-12-UTP for natural UTP in the transcription reaction. The resulting transcript contains 74 U residues, which would give an average of 1.5 fluorescein labels per molecule. With one tenth the molar amount of RNA loaded in lane 2, the signal in lane 2 should be approximately one seventh (0.15×) the fluorescence intensity of the probe in lane 1. Measurements indicated that the intensity was closer to one fortieth, indicating an efficiency of label incorporation of approximately 17%. Because the RNA concentration was verified by A260 measurement this does not alter the experimental observations below, but it should be noted that the signal from the RNA and the probes does not accurately reflect the relative amounts in the reactions.
The reactions analyzed in lanes 3 through 7 contained 1, 5, 10, 50 and 100 fmoles of target, respectively, with cleavage of the probe accomplished by Cleavase® A/G. The reactions analyzed in lanes 8 through 12 repeated the same array of target amounts, with cleavage of the probe accomplished by DNAPTth. The boxes seen surrounding the product bands show the area of the scan in which the fluorescence was measured for each reaction. The number of fluorescence units detected within each box is indicated below each box; background florescence was also measured.
It can be seen by comparing the detected fluorescence in each lane that the amount of product formed in these 30 minute reactions can be correlated to the amount of target material. The accumulation of product under these conditions is slightly enhanced when DNAPTth is used as the cleavage agent, but the correlation with the amount of target present remains. This demonstrates that the invader assay can be used as a means of measuring the amount of target RNA within a sample.
Comparison of the fluorescence intensity of the input RNA with that of the cleaved product shows that the invader-directed cleavage assay creates signal in excess of the amount of target, so that the signal visible as cleaved probe is far more intense than that representing the target RNA. This further confirms the results described in Example 20, in which it was demonstrated that each RNA molecule could be used many times.
EXAMPLE 22
Detection of DNA by Charge Reversal
The detection of specific targets is achieved in the invader-directed cleavage assay by the cleavage of the probe oligonucleotide. In addition to the methods described in the preceding examples, the cleaved probe may be separated from the uncleaved probe using the charge reversal technique described below. This novel separation technique is related to the observation that positively charged adducts can affect the electrophoretic behavior of small oligonucleotides because the charge of the adduct is significant relative to charge of the whole complex. Observations of aberrant mobility due to charged adducts have been reported in the literature, but in all cases found, the applications pursued by other scientists have involved making oligonucleotides larger by enzymatic extension. As the negatively charged nucleotides are added on, the positive influence of the adduct is reduced to insignificance. As a result, the effects of positively charged adducts have been dismissed and have received infinitesimal notice in the existing literature.
This observed effect is of particular utility in assays based on the cleavage of DNA molecules. When an oligonucleotide is shortened through the action of a Cleavase® enzyme or other cleavage agent, the positive charge can be made to not only significantly reduce the net negative charge, but to actually override it, effectively “flipping” the net charge of the labeled entity. This reversal of charge allows the products of target-specific cleavage to be partitioned from uncleaved probe by extremely simple means. For example, the products of cleavage can be made to migrate towards a negative electrode placed at any point in a reaction vessel, for focused detection without gel-based electrophoresis. When a slab gel is used, sample wells can be positioned in the center of the gel, so that the cleaved and uncleaved probes can be observed to migrate in opposite directions. Alternatively, a traditional vertical gel can be used, but with the electrodes reversed relative to usual DNA gels (i.e., the positive electrode at the top and the negative electrode at the bottom) so that the cleaved molecules enter the gel, while the uncleaved disperse into the upper reservoir of electrophoresis buffer.
An additional benefit of this type of readout is that the absolute nature of the partition of products from substrates means that an abundance of uncleaved probe can be supplied to drive the hybridization step of the probe-based assay, yet the unconsumed probe can be subtracted from the result to reduce background.
Through the use of multiple positively charged adducts, synthetic molecules can be constructed with sufficient modification that the normally negatively charged strand is made nearly neutral. When so constructed, the presence or absence of a single phosphate group can mean the difference between a net negative or a net positive charge. This observation has particular utility when one objective is to discriminate between enzymatically generated fragments of DNA, which lack a 3 phosphate, and the products of thermal degradation, which retain a 3 phosphate (and thus two additional negative charges).
a) Characterization Of The Products Of Thermal Breakage Of DNA Oligonucleotides
Thermal degradation of DNA probes results in high background which can obscure signals generated by specific enzymatic cleavage, decreasing the signal-to-noise ratio. To better understand the nature of DNA thermal degradation products, we incubated the 5′ tetrachloro-fluorescein (TET)-labeled oligonucleotides 78 (SEQ ID NO:48) and 79 (SEQ ID NO:49) (100 pmole each) in 50 μl 10 mM NaCO3 (pH 10.6), 50 mM NaCl at 90° C. for 4 hours. To prevent evaporation of the samples, the reaction mixture was overlaid with 50 μl of ChillOut® liquid wax. The reactions were then divided in two equal aliquots (A and B). Aliquot A was mixed with 25 μl of methyl violet loading buffer and Aliquot B was dephosphorylated by addition of 2.5 μl of 100 mM MgCl2 and 1 μl of 1 unit/μl Calf Intestinal Alkaline Phosphatase (CIAP) (Promega), with incubation at 37° C. for 30 min. after which 25 μl of methyl violet loading buffer was added. One microliter of each sample was resolved by electrophoresis through a 12% polyacrylamide denaturing gel and imaged as described in Example 21; a 585 nm filter was used with the FMBIO Image Analyzer. The resulting imager scan is shown in FIG. 44.
In FIG. 44, lanes 1-3 contain the TET-labeled oligonucleotide 78 and lanes 4-6 contain the TET-labeled oligonucleotides 79. Lanes 1 and 4 contain products of reactions which were not heat treated. Lanes 2 and 5 contain products from reactions which were heat treated and lanes 3 and 6 contain products from reactions which were heat treated and subjected to phosphatase treatment.
As shown in FIG. 44, heat treatment causes significant breakdown of the 5′-TET-labeled DNA, generating a ladder of degradation products (FIG. 44, lanes 2, 3, 5 and 6). Band intensities correlate with purine and pyrimidine base positioning in the oligonucleotide sequences, indicating that backbone hydrolysis may occur through formation of abasic intermediate products that have faster rates for purines than for pyrimidines [Lindahl and Karlstrom (1973) Biochem. 12:5151].
Dephosphorylation decreases the mobility of all products generated by the thermal degradation process, with the most pronounced effect observed for the shorter products (FIG. 44, lanes 3 and 6). This demonstrates that thermally degraded products possess a 3′ end terminal phosphoryl group which can be removed by dephosphotylation with CIAP. Removal of the phosphoryl group decreases the overall negative charge by 2. Therefore, shorter products which have a small number of negative charges are influenced to a greater degree upon the removal of two charges. This leads to a larger mobility shift in the shorter products than that observed for the larger species.
The fact that the majority of thermally degraded DNA products contain 3′ end phosphate groups and Cleavase® enzyme-generated products do not allowed the development of simple isolation methods for products generated in the invader-directed cleavage assay. The extra two charges found in thermal breakdown products do not exist in the specific cleavage products. Therefore, if one designs assays that produce specific products which contain a net positive charge of one or two, then similar thermal breakdown products will either be negative or neutral. The difference can be used to isolate specific products by reverse charge methods as shown below.
b) Dephosphorylation Of Short Amino-Modified Oligonucleotides Can Reverse The Net Charge Of The Labeled Product
To demonstrate how oligonucleotides can be transformed from net negative to net positively charged compounds, the four short amino-modified oligonucleotides labeled 70, 74, 75 and 76 and shown in FIGS. 45-47 were synthesized (FIG. 45 shows both oligonucleotides 70 and 74). All four modified oligonucleotides possess Cy-3 dyes positioned at the 5′-end which individually are positively charged under reaction and isolation conditions described in this example. Compounds 70 and 74 contain two amino modified thymidines that, under reaction conditions, display positively charged R—NH3+ groups attached at the C5 position through a C10 or C6 linker, respectively. Because compounds 70 and 74 are 3′-end phosphorylated, they consist of four negative charges and three positive charges. Compound 75 differs from 74 in that the internal C6 amino modified thymidine phosphate in 74 is replaced by a thymidine methyl phosphonate. The phosphonate backbone is uncharged and so there are a total of three negative charges on compound 75. This gives compound 75 a net negative one charge. Compound 76 differs from 70 in that the internal amino modified thymidine is replaced by an internal cytosine phosphonate. The pKa of the N3 nitrogen of cytosine can be from 4 to 7. Thus, the net charges of this compound, can be from −1 to 0 depending on the pH of the solution. For the simplicity of analysis, each group is assigned a whole number of charges, although it is realized that, depending on the pKa of each chemical group and ambient pH, a real charge may differ from the whole number assigned. It is assumed that this difference is not significant over the range of pHs used in the enzymatic reactions studied here.
Dephosphorylation of these compounds, or the removal of the 3′ end terminal phosphoryl group, results in elimination of two negative charges and generates products that have a net positive charge of one. In this experiment, the method of isoelectric focusing (IEF) was used to demonstrate a change from one negative to one positive net charge for the described substrates during dephosphorylation.
Substrates 70, 74, 75 and 76 were synthesized by standard phosphoramidite chemistries and deprotected for 24 hours at 22° C. in 14 M aqueous ammonium hydroxide solution, after which the solvent was removed in vacuo. The dried powders were resuspended in 200 μl of H2O and filtered through 0.2 μm filters. The concentration of the stock solutions was estimated by UV-absorbance at 261 nm of samples diluted 200-fold in H2O using a spectrophotometer (Spectronic Genesys 2, Milton Roy, Rochester, N.Y.).
Dephosphorylation of compounds 70 and 74, 75 and 76 was accomplished by treating 10 μl of the crude stock solutions (ranging in concentration from approximately 0.5 to 2 mM) with 2 units of CIAP in 100 μl of CIAP buffer (Promega) at 37° C. for 1 hour. The reactions were then heated to 75° C. for 15 min. in order to inactivate the CIAP. For clarity, dephosphorylated compounds are designated ‘dp’. For example, after dephosphorylation, substrate 70 becomes 70dp.
To prepare samples for IEF experiments, the concentration of the stock solutions of substrate and dephosphorylated product were adjusted to a uniform absorbance of 8.5×10−3 at 532 nm by dilution with water. Two microliters of each sample were analyzed by IEF using a PhastSystem electrophoresis unit (Pharmacia) and PhastGel IEF 3-9 media (Pharmacia) according to the manufacturer's protocol. Separation was performed at 15° C. with the following program: pre-run; 2,000 V, 2.5 mA, 3.5 W, 75 Vh; load; 200 V, 2.5 mA, 3.5 W, 15 Vh; run; 2,000 V; 2.5 mA; 3.5 W, 130 Vh. After separation, samples were visualized by using the FMBIO Image Analyzer (Hitachi) fitted with a 585 nm filter. The resulting imager scan is shown in FIG. 48.
FIG. 48 shows results of IEF separation of substrates 70, 74, 75 and 76 and their dephosphorylated products. The arrow labeled “Sample Loading Position” indicates a loading line, the ‘+’ sign shows the position of the positive electrode and the ‘−’ sign indicates the position of the negative electrode.
The results shown in FIG. 48 demonstrate that substrates 70, 74, 75 and 76 migrated toward the positive electrode, while the dephosphorylated products 70dp, 74dp, 75dp and 76dp migrated toward negative electrode. The observed differences in mobility direction was in accord with predicted net charge of the substrates (minus one) and the products (plus one). Small perturbations in the mobilities of the phosphorylated compounds indicate that the overall pls vary. This was also true for the dephosphorylated compounds. The presence of the cytosine in 76dp, for instance, moved this compound further toward the negative electrode which was indicative of a higher overall pI relative to the other dephosphorylated compounds. It is important to note that additional positive charges can be obtained by using a combination of natural amino modified bases (70dp and 74dp) along with uncharged methylphosphonate bridges (products 75dp and 76dp).
The results shown above demonstrate that the removal of a single phosphate group can flip the net charge of an oligonucleotide to cause reversal in an electric field, allowing easy separation of products, and that the precise base composition of the oligonucleotides affect absolute mobility but not the charge-flipping effect.
EXAMPLE 23 
Detection of Specific Cleavage Products in the Invader-Directed Cleavage Reaction by Charge Reversal
In this example the ability to isolate products generated in the invader-directed cleavage assay from all other nucleic acids present in the reaction cocktail was demonstrated using charge reversal. This experiment utilized the following Cy3-labeled oligonucleotide: 5′-Cy3-AminoT-AminoT-CTTTTCACCAGCGAGACGGG-3′ (SEQ ID NO:50; termed “oligo 61”). Oligo 61 was designed to release upon cleavage a net positively charged labeled product. To test whether or not a net positively charged 5′-end labeled product would be recognized by the Cleavase® enzymes in the invader-directed cleavage assay format, probe oligo 61 (SEQ ID NO:50) and invading oligonucleotide 67 (SEQ ID NO:51) were chemically synthesized on a DNA synthesizer (ABI 391) using standard phosphoramidite chemistries and reagents obtained from Glen Research (Sterling, Va.).
Each assay reaction comprised 100 fmoles of M13mp18 single stranded DNA, 10 pmoles each of the probe (SEQ ID NO:50) and InvaderTM (SEQ ID NO:51) oligonucleotides, and 20 units of Cleavase® A/G in a 10 μl solution of 10 mM MOPS, pH 7.4 with 100 mM KCl. Samples were overlaid with mineral oil to prevent evaporation. The samples were brought to either 50° C., 55° C., 60° C., or 65° C. and cleavage was initiated by the addition of 1 μl of 40 mM MnCl2. Reactions were allowed to proceed for 25 minutes and then were terminated by the addition of 10 μl of 95% formamide containing 20 mM EDTA and 0.02% methyl violet. The negative control experiment lacked the target M13mp18 and was run at 60° C. Five microliters of each reaction were loaded into separate wells of a 20% denaturing polyacrylamide gel (cross-linked 29:1) with 8 M urea in a buffer containing 45 mM Tris-Borate (pH 8.3) and 1.4 mM EDTA. An electric field of 20 watts was applied for 30 minutes, with the electrodes oriented as indicated in FIG. 49B (i.e., in reverse orientation). The products of these reactions were visualized using the FMBIO fluorescence imager and the resulting imager scan is shown in FIG. 49B.
FIG. 49A provides a schematic illustration showing an alignment of the invader (SEQ ID NO:50) and probe (SEQ ID NO:51) along the target M13mp18 DNA; only 53 bases of the M13mp18 sequence is shown (SEQ ID NO:52). The sequence of the invader oligonucleotide is displayed under the M13mp18 target and an arrow is used above the M13mp18 sequence to indicate the position of the invader relative to the probe and target. As shown in FIG. 49A, the invader and probe oligonucleotides share a 2 base region of overlap.
In FIG. 49B, lanes 1-6 contain reactions performed at 50° C., 55° C., 60° C., and 65° C., respectively; lane 5 contained the control reaction (lacking target). In FIG. 49B, the products of cleavage are seen as dark bands in the upper half of the panel; the faint lower band seen appears in proportion to the amount of primary product produced and, while not limiting the invention to a particular mechanism, may represent cleavage one nucleotide into the duplex. The uncleaved probe does not enter the gel and is thus not visible. The control lane showed no detectable signal over background (lane 5). As expected in an invasive cleavage reaction, the rate of accumulation of specific cleavage product was temperature-dependent. Using these particular oligonucleotides and target, the fastest rate of accumulation of product was observed at 55° C. (lane 2) and very little product observed at 65° C. (lane 4).
When incubated for extended periods at high temperature, DNA probes can break non-specifically (i.e., suffer thermal degradation) and the resulting fragments contribute an interfering background to the analysis. The products of such thermal breakdown are distributed from single-nucleotides up to the full length probe. In this experiment, the ability of charge based separation of cleavage products (i.e., charge reversal) would allow the sensitive separation of the specific products of target-dependent cleavage from probe fragments generated by thermal degradation was examined.
To test the sensitivity limit of this detection method, the target M13mp18 DNA was serially diluted ten fold over than range of 1 fmole to 1 amole. The invader and probe oligonucleotides were those described above (i.e., SEQ ID NOS:50 and 51). The invasive cleavage reactions were run as described above with the following modifications: the reactions were performed at 55° C., 250 mM or 100 mM KGlu was used in place of the 100 mM KCl and only 1 pmole of the invader oligonucleotide was added. The reactions were initiated as described above and allowed to progress for 12.5 hours. A negative control reaction which lacked added M13m18 target DNA was also run. The reactions were terminated by the addition of 10 μl of 95% formamide containing 20 mM EDTA and 0.02% methyl violet, and 5 μl of these mixtures were electrophoresed and visualized as described above. The resulting imager scan is shown in FIG. 50.
In FIG. 50, lane 1 contains the negative control; lanes 2-5 contain reactions performed using 100 mM KGlu; lanes 6-9 contain reactions performed using 250 mM KGlu. The reactions resolved in lanes 2 and 6 contained 1 fmole of target DNA; those in lanes 3 and 7 contained 100 amole of target; those in lanes 4 and 8 contained 10 amole of target and those in lanes 5 and 9 contained I amole of target. The results shown in FIG. 50 demonstrate that the detection limit using charge reversal to detect the production of specific cleavage products in an invasive cleavage reaction is at or below 1 attomole or approximately 6.02×105 target molecules. No detectable signal was observed in the control lane, which indicates that non-specific hydrolysis or other breakdown products do not migrate in the same direction as enzyme-specific cleavage products. The excitation and emission maxima for Cy3 are 554 and 568, respectively, while the FMBIO Imager Analyzer excites at 532 and detects at 585. Therefore, the limit of detection of specific cleavage products can be improved by the use of more closely matched excitation source and detection filters.
EXAMPLE 24 
Devices and Methods for the Separation and Detection of Charged Reaction Products
This example is directed at methods and devices for isolating and concentrating specific reaction products produced by enzymatic reactions conducted in solution whereby the reactions generate charged products from either a charge neutral substrate or a substrate bearing the opposite charge borne by the specific reaction product. The methods and devices of this example allow isolation of, for example, the products generated by the invader-directed cleavage assay of the present invention.
The methods and devices of this example are based on the principle that when an electric field is applied to a solution of charged molecules, the migration of the molecules toward the electrode of the opposite charge occurs very rapidly. If a matrix or other inhibitory material is introduced between the charged molecules and the electrode of opposite charge such that this rapid migration is dramatically slowed, the first molecules to reach the matrix will be nearly stopped, thus allowing the lagging molecules to catch up. In this way a dispersed population of charged molecules in solution can be effectively concentrated into a smaller volume. By tagging the molecules with a detectable moiety (e.g., a fluorescent dye), detection is facilitated by both the concentration and the localization of the analytes. This example illustrates two embodiments of devices contemplated by the present invention; of course, variations of these devices will be apparent to those skilled in the art and are within the spirit and scope of the present invention.
FIG. 51 depicts one embodiment of a device for concentrating the positively-charged products generated using the methods of the present invention. As shown in FIG. 51, the device comprises a reaction tube (10) which contains the reaction solution (11). One end of each of two thin capillaries (or other tubes with a hollow core) (13A and 13B) are submerged in the reaction solution (11). The capillaries (13A and 13B) may be suspended in the reaction solution (11) such that they are not in contact with the reaction tube itself; one appropriate method of suspending the capillaries is to hold them in place with clamps (not shown). Alternatively, the capillaries may be suspended in the reaction solution (11) such that they are in contact with the reaction tube itself. Suitable capillaries include glass capillary tubes commonly available from scientific supply companies (e.g., Fisher Scientific or VWR Scientific) or from medical supply houses that carry materials for blood drawing and analysis. Though the present invention is not limited to capillaries of any particular inner diameter, tubes with inner diameters of up to about 1⁄8 inch (approximately 3 mm) are particularly preferred for use with the present invention; for example Kimble No. 73811-99 tubes (VWR Scientific) have an inner diameter of 1.1 mm and are a suitable type of capillary tube. Although the capillaries of the device are commonly composed of glass, any nonconductive tubular material, either rigid or flexible, that can contain either a conductive material or a trapping material is suitable for use in the present invention. One example of a suitable flexible tube is Tygon® clear plastic tubing (Part No. R3603; inner diameter= 1/16 inch; outer diameter=1⁄8 inch).
As illustrated in FIG. 51, capillary 13A is connected to the positive electrode of a power supply (20) (e.g., a controllable power supply available through the laboratory suppliers listed above or through electronics supply houses like Radio Shack) and capillary 13B is connected to the negative electrode of the power supply (20). Capillary 13B is filled with a trapping material (14) capable of trapping the positively-charged reaction products by allowing minimal migration of products that have entered the trapping material (14). Suitable trapping materials include, but are not limited to, high percentage (e.g., about 20%) acrylamide polymerized in a high salt buffer (0.5 M or higher sodium acetate or similar salt); such a high percentage polyacrylamide matrix dramatically slows the migration of the positively-charged reaction products. Alternatively, the trapping material may comprise a solid, negatively-charged matrix, such as negatively-charged latex beads, that can bind the incoming positively-charged products. It should be noted that any amount of trapping material (14) capable of inhibiting any concentrating the positively-charged reaction products may be used. Thus, while the capillary 13B in FIG. 51 only contains trapping material in the lower, submerged portion of the tube, the trapping material (14) can be present in the entire capillary (13B); similarly, less trapping material (14) could be present than that shown in FIG. 51 because the positively-charged reaction products generally accumulate within a very small portion of the bottom of the capillary (13B). The amount of trapping material need only be sufficient to make contact with the reaction solution (11) and have the capacity to collect the reaction products. When capillary 13B is not completely filled with the trapping material, the remaining space is filled with any conductive material (15); suitable conductive materials are discussed below.
By comparison, the capillary (13A) connected to the positive electrode of the power supply 20 may be filled with any conductive material (15; indicated by the hatched lines in FIG. 51). This may be the sample reaction buffer (e.g., 10 mM MOPS, pH 7.5 with 150 mM LiCl, 4 mM MnCl2), a standard electrophoresis buffer (e.g., 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA), or the reaction solution (11) itself. The conductive material (15) is frequently a liquid, but a semi-solid material (e.g., a gel) or other suitable material might be easier to use and is within the scope of the present invention. Moreover, that trapping material used in the other capillary (i.e., capillary 13B) may also be used as the conductive material. Conversely, it should be noted that the same conductive material used in the capillary (13A) attached to the positive electrode may also be used in capillary 13B to fill the space above the region containing the trapping material (14) (see FIG. 51).
The top end of each of the capillaries (13A and 13B) is connected to the appropriate electrode of the power supply (20) by electrode wire (18) or other suitable material. Fine platinum wire (e.g., 0.1 to 0.4 mm, Aesar Johnson Matthey, Ward Hill, Mass.) is commonly used as conductive wire because it does not corrode under electrophoresis conditions. The electrode wire (18) can be attached to the capillaries (13A and 13B) by a nonconductive adhesive (not shown), such as the silicone adhesives that are commonly sold in hardware stores for sealing plumbing fixtures. If the capillaries are constructed of a flexible material, the electrode wire (18) can be secured with a small hose clamp or constricting wire (not shown) to compress the opening of the capillaries around the electrode wire. If the conducting material (15) is a gel, an electrode wire (18) can be embedded directly in the gel within the capillary.
The cleavage reaction is assembled in the reaction tube (10) and allowed to proceed therein as described in proceeding examples (e.g., Examples 22-23). Though not limited to any particular volume of reaction solution (11), a preferred volume is less than 10 ml and more preferably less than 0.1 ml. The volume need only be sufficient to permit contact with both capillaries. After the cleavage reaction is completed, an electric field is applied to the capillaries by turning on the power source (20). As a result, the positively-charged products generated in the course of the invader-directed cleavage reaction which employs an oligonucleotide, which when cleaved, generates a positively charged fragment (described in Ex. 23) but when uncleaved bears a net negative charge, migrate to the negative capillary, where their migration is slowed or stopped by the trapping material (14), and the negatively-charged uncut and thermally degraded probe molecules migrate toward the positive electrode. Through the use of this or a similar device, the positively-charged products of the invasive cleavage reaction are separated from the other material (i.e., uncut and thermally degraded probe) and concentrated from a large volume. Concentration of the product in a small amount of trapping material (14) allows for simplicity of detection, with a much higher signal-to-noise ratio than possible with detection in the original reaction volume. Because the concentrated product is labelled with a detectable moiety like a fluorescent dye, a commercially-available fluorescent plate reader (not shown) can be used to ascertain the amount of product. Suitable plate readers include both top and bottom laser readers. Capillary 13B can be positioned with the reaction tube (10) at any desired position so as to accommodate use with either a top or a bottom plate reading device.
In the alternative embodiment of the present invention depicted in FIG. 52, the procedure described above is accomplished by utilizing only a single capillary (13B). The capillary (13B) contains the trapping material (14) described above and is connected to an electrode wire (18), which in turn is attached to the negative electrode of a power supply (20). The reaction tube (10) has an electrode (25) embedded into its surface such that one surface of the electrode is exposed to the interior of the reaction tube (10) and another surface is exposed to the exterior of the reaction tube. The surface of the electrode (25) on the exterior of the reaction tube is in contact with a conductive surface (26) connected to the positive electrode of the power supply (20) through an electrode wire (18). Variations of the arrangement depicted in FIG. 52 are also contemplated by the present invention. For example, the electrode (25) may be in contact with the reaction solution (11) through the use of a small hole in the reaction tube (10); furthermore, the electrode wire (18) can be directly attached to the electrode wire (18), thereby eliminating the conductive surface (26).
As indicated in FIG. 52, the electrode (25) is embedded in the bottom of a reaction tube (10) such that one or more reaction tubes may be set on the conductive surface (26). This conductive surface could serve as a negative electrode for multiple reaction tubes; such a surface with appropriate contacts could be applied through the use of metal foils (e.g., copper or platinum, Aesar Johnson Matthey, Ward Hill, Mass.) in much the same way contacts are applied to circuit boards. Because such a surface contact would not be exposed to the reaction sample directly, less expensive metals, such as the copper could be used to make the electrical connections.
The above devices and methods are not limited to separation and concentration of positively charged oligonucleotides. As will be apparent to those skilled in the art, negatively charged reaction products may be separated from neutral or positively charged reactants using the above device and methods with the exception that capillary 13B is attached to the positive electrode of the power supply (20) and capillary 13A or alternatively, electrode 25, is attached to the negative electrode of the power supply (20).
EXAMPLE 25
Primer-Directed and Primer Independent Cleavage Occur at the Same Site When the Primer Extends to the 3′ Side of a Mismatched “Bubble” in the Downstream Duplex
As discussed above in Example 1, the presence of a primer upstream of a bifurcated duplex can influence the site of cleavage, and the existence of a gap between the 3′ end of the primer and the base of the duplex can cause a shift of the cleavage site up the unpaired 5′ arm of the structure (see also Lyamichev et al., supra and U.S. Pat. No. 5,422,253). The resulting non-invasive shift of the cleavage site in response to a primer is demonstrated in FIGS. 8, 9 and 10, in which the primer used left a 4-nucleotide gap (relative to the base of the duplex). In FIGS. 8-10, all of the “primer-directed” cleavage reactions yielded a 21 nucleotide product, while the primer-independent cleavage reactions yielded a 25 nucleotide product. The site of cleavage obtained when the primer was extended to the base of the duplex, leaving no gap was examined. The results are shown in FIG. 53 (FIG. 53 is a reproduction of FIG. 2C in Lyamichev et al. These data were derived from the cleavage of the structure shown in FIG. 5, as described in Example 1. Unless otherwise specified, the cleavage reactions comprised 0.01 pmoles of heat-denatured, end-labeled hairpin DNA (with the unlabeled complementary strand also present), 1 pmole primer [complementary to the 3′ arm shown in FIG. 5 and having the sequence: 5′-GAATTCGATTTAGGTGACACTATAGAATACA (SEQ ID NO:53)] and 0.5 units of DNAPTaq (estimated to be 0.026 pmoles) in a total volume of 10 μl of 10 mM Tris-Cl, pH 8.5, and 1.5 mM MgCl2 and 50 mM KCl. The primer was omitted from the reaction shown in the first lane of FIG. 53 and included in lane 2. These reactions were incubated at 55° C. for 10 minutes. Reactions were initiated at the final reaction temperature by the addition of either the MgCl2 or enzyme. Reactions were stopped at their incubation temperatures by the addition of 8 μl of 95% formamide with 20 mM EDTA and 0.05% marker dyes.
FIG. 53 is an autoradiogram that indicates the effects on the site of cleavage of a bifurcated duplex structure in the presence of a primer that extends to the base of the hairpin duplex. The size of the released cleavage product is shown to the left (i.e., 25 nucleotides). A dideoxynucleotide sequencing ladder of the cleavage substrate is shown on the right as a marker (lanes 3-6).
These data show that the presence of a primer that is adjacent to a downstream duplex (lane 2) produces cleavage at the same site as seen in reactions performed in the absence of the primer (lane 1). (See FIGS. 8A and B, 9B and 10A for additional comparisons). When the 3′ terminal nucleotides of the upstream oligonucleotide can base pair to the template strand but are not homologous to the displaced strand in the region immediately upstream of the cleavage site (i.e., when the upstream oligonucleotide is opening up a “bubble” in the duplex), the site to which cleavage is apparently shifted is not wholly dependent on the presence of an upstream oligonucleotide.
As discussed above in the Background section and in Table 1, the requirement that two independent sequences be recognized in an assay provides a highly desirable level of specificity. In the invasive cleavage reactions of the present invention, the invader and probe oligonucleotides must hybridize to the target nucleic acid with the correct orientation and spacing to enable the production of the correct cleavage product. When the distinctive pattern of cleavage is not dependent on the successful alignment of both oligonucleotides in the detection system these advantages of independent recognition are lost.
EXAMPLE 26 
Invasive Cleavage and Primer-Directed Cleavage When There is Only Partial Homology in the “X” Overlap Region
While not limiting the present invention to any particular mechanism, invasive cleavage occurs when the site of cleavage is shifted to a site within the duplex formed between the probe and the target nucleic acid in a manner that is dependent on the presence of an upstream oligonucleotide which shares a region of overlap with the downstream probe oligonucleotide. In some instances, the 5′ region of the downstream oligonucleotide may not be completely complementary to the target nucleic acid. In these instances, cleavage of the probe may occur at an internal site within the probe even in the absence of an upstream oligonucleotide (in contrast to the base-by-base nibbling seen when a fully paired probe is used without an invader). Invasive cleavage is characterized by an apparent shifting of cleavage to a site within a downstream duplex that is dependent on the presence of the invader oligonucleotide.
A comparison between invasive cleavage and primer-directed cleavage may be illustrated by comparing the expected cleavage sites of a set of probe oligonucleotides having decreasing degrees of complementarity to the target strand in the 5′ region of the probe (i.e., the region that overlaps with the invader). A simple test, similar to that performed on the hairpin substrate above (Ex. 25), can be performed to compare invasive cleavage with the non- invasive primer-directed cleavage described above. Such a set of test oligonucleotides is diagrammed in FIG. 54. The structures shown in FIG. 54 are grouped in pairs, labeled “a”, “b”, “c”, and “d”. Each pair has the same probe sequence annealed to the target strand (SEQ ID NO:54), but the top structure of each pair is drawn without an upstream oligonucleotide, while the bottom structure includes this oligonucleotide (SEQ ID NO:55). The sequences of the probes shown in FIGS. 54a-54d are listed in SEQ ID NOS:32, 56, 57 and 58, respectively. Probable sites of cleavage are indicated by the black arrowheads. (It is noted that the precise site of cleavage on each of these structures may vary depending on the choice of cleavage agent and other experimental variables. These particular sites are provided for illustrative purposes only.)
To conduct this test, the site of cleavage of each probe is determined both in the presence and the absence of the upstream oligonucleotide, in reaction conditions such as those described in Example 18. The products of each pair of reactions are then be compared to determine whether the fragment released from the 5′ end of the probe increases in size when the upstream oligonucleotide is included in the reaction.
The arrangement shown in FIG. 54a, in which the probe molecule is completely complementary to the target strand, is similar to that shown in FIG. 28. Treatment of the top structure with the 5′ nuclease of a DNA polymerase would cause exonucleolytic nibbling of the probe (i.e., in the absence of the upstream oligonucleotide). In contrast, inclusion of an invader oligonucleotide would cause a distinctive cleavage shift similar, to those observed in FIG. 29.
The arrangements shown in FIGS. 54b and 54c have some amount of unpaired sequence at the 5′ terminus of the probe (3 and 5 bases, respectively). These small 5′ arms are suitable cleavage substrate for the 5′ nucleases and would be cleaved within 2 nucleotide's of the junction between the single stranded region and the duplex. In these arrangements, the 3′ end of the upstream oligonucleotide shares identity with a portion of the 5′ region of the probe which is complementary to the target sequence (that is the 3′ end of the invader has to compete for binding to the target with a portion of the 5′ end of the probe). Therefore, when the upstream oligonucleotide is included it is thought to mediate a shift in the site of cleavage into the downstream duplex (although the present invention is not limited to any particular mechanism of action), and this would, therefore, constitute invasive cleavage. If the extreme 5′ nucleotides of the unpaired region of the probe were able to hybridize to the target strand, the cleavage site in the absence of the invader might change but the addition of the invader oligonucleotide would still shift the cleavage site to the proper position.
Finally, in the arrangement shown in FIG. 54d, the probe and upstream oligonucleotides share no significant regions of homology, and the presence of the upstream oligonucleotide would not compete for binding to the target with the probe. Cleavage of the structures shown in FIG. 54d would occur at the same site with or without the upstream oligonucleotide, and is thus would not constitute invasive cleavage.
By examining any upstream oligonucleotide/probe pair in this way, it can easily be determined whether the resulting cleavage is invasive or merely primer-directed. Such analysis is particularly useful when the probe is not fully complementary to the target nucleic acid, so that the expected result may not be obvious by simple inspection of the sequences.
EXAMPLE 27 
Modified Cleavase® Enzymes
In order to develop nucleases having useful activities for the cleavage of nucleic acids the following modified nucleases were produced.
a) Cleavase® BN/thrombin Nuclease
i) Cloning and Expression of Cleavase® BN/thrombin Nuclease
Site directed mutagenesis was used to introduce a protein sequence recognized by the protease thrombin into the region of the Cleavase® BN nuclease which is thought to form the helical arch of the protein through which the single-stranded DNA that is cleaved must presumably pass. Mutagenesis was carried out using the Transformer® mutagenesis kit (Clonetech, Palo Alto, Calif.) according to manufacturer's protocol using the mutagenic oligonucleotide 5′-GGGAAAGTCCTCGCAGCCGCGCGGGACGAGCGTGGGGGCCCG (SEQ ID NO:59). After mutagenesis, the DNA was sequenced to verify the insertion of the thrombin cleavage site. The DNA sequence encoding the Cleavase® BN/thrombin nuclease is provided in SEQ ID NO:60; the amino acid sequence of Cleavase® BN/thrombin nuclease is provided in SEQ ID NO:61.
A large scale preparation of the thrombin mutant (i.e., Cleavase® BN/thrombin) was done using E. coli cells overexpressing the Cleavase® BN/thrombin nuclease as described in Example 28.
ii) Thrombin Cleavage of Cleavase® BN/thrombin
Six point four (6.4) mg of the purified Cleavase® BN/thrombin nuclease was digested with 0.4 U of thrombin (Novagen) for 4 hours at 23° C. or 37° C. Complete digestion was verified by electrophoresis on a 15% SDS polyacrylamide gel followed by staining with Coomassie Brilliant Blue R. Wild-type Cleavase® BN nuclease was also digested with thrombin as a control. The resulting gel is shown in FIG. 61.
In FIG. 61, lane 1 contains molecular weight markers (Low-Range Protein Molecular Weight Markers; Promega), lane 2 contains undigested Cleavase® BN/throbin nuclease, lanes 3 and 4 contain Cleavase® BN/thrombin nuclease digested with thrombin at 23° C. for 2 and 4 hours, respectively, and lanes 5 and 6 contain Cleavase® BN/thrombin nuclease digested with thrombin at 37° C. for 2 and 4 hours, respectively. These results show that the Cleavase® BN/thrombin nuclease has an apparent molecular weight of 36.5 kilodaltons and demonstrate that Cleavase® BN/thrombin nuclease is efficiently cleaved by thrombin. In addition, the thrombin cleavage products have approximate molecular weights of 27 kilodaltons and 9 kilodaltons, the size expected based upon the position of the inserted thrombin site in the Cleavase® BN/thrombin nuclease.
To determine the level of hairpin cleavage activity in digested and undigested Cleavase® BN/thrombin nuclease, dilutions were made and used to cleave a test hairpin containing a 5′ fluoroscein label. Varying amounts of digested and undigested Cleavase® BN/thrombin nuclease were incubated with 5 μM oligonucleotide S-60 hairpin (SEQ ID NO:29; see FIG. 26) in 10 mM MOPS (pH 7.5), 0.05% Tween-20, 0.05% NP-40, and 1 mM MnCl2 for 5 minutes at 60° C. The digested mixture was electrophoresed on a 20% acrylamide gel and visualized on a Hitachi FMBIO 100 fluoroimager. The resulting image is shown in FIG. 62.
In FIG. 62, lane 1 contains the no enzyme control, lane 2 contains reaction products produced using 0.01 ng of Cleavase® BN nuclease, lanes 3, 4, and 5 contain reaction products produced using 0.01 ng, 0.04 ng, and 4 ng of undigested Cleavase® BN/thrombin nuclease, respectively, and lanes 6, 7, and 8 contain reaction products produced using 0.01 ng, 0.04 ng, and 4 ng of thrombin-digested Cleavase® BN/thrombin nuclease, respectively. The results shown in FIG. 62 demonstrated that the insertion of the thrombin cleavage site reduced cleavage activity about 200-fold (relative to the activity of Cleavase® BN nuclease), but that digestion with thrombin did not reduce the activity significantly.
M13 single-stranded DNA was used as a substrate for cleavage by Cleavase® BN nuclease and digested and undigested Cleavase® BN/thrombin nuclease. Seventy nanograms of single-stranded M13 DNA (New England Biolabs, Beverly, Mass.) was incubated in 10 mM MOPS, pH 7.5, 0.05% Tween-20, 0.05% NP-40, 1 mM MgCl2 or 1 mM MnCl2, with 8 ng of Cleavase® BN nuclease, undigested Cleavase® BN/thrombin nuclease, or digested Cleavase® BN/thrombin nuclease for 10 minutes at 50° C. Reaction mixtures were electrophoresed on a 0.8% agarose gel and then stained with a solution containing 0.5 μg/ml ethidium bromide (EtBr) to visualize DNA bands. A negative image of the EtBr-stained gel is shown in FIG. 63.
In FIG. 63, lane 1 contains the no enzyme control, lane 2 contains reaction products produced using Cleavase® BN nuclease and 1 mM MgCl2, lane 3 contains reaction products produced using Cleavase ® BN nuclease and 1 mM MnCl2, lane 4 contains reaction products produced using undigested Cleavase® BN/thrombin nuclease and 1 mM MgCl2, lane 5 contains reaction products produced using undigested Cleavase® BN/thrombin nuclease and 1 mM MnCl2, lane 6 contains reaction products produced using thrombin-digested Cleavase® BN/thrombin nuclease and 1 mM MgCl2, and lane 7 contains reaction products produced using thrombin-digested Cleavase® BN/thrombin nuclease and 1 mM MnCl2. The results shown in FIG. 63 demonstrated that the Cleavase® BN/thrombin nuclease had an enhanced ability to cleave circular DNA (and thus a reduced requirement for the presence of a free 5′ end) as compared to the Cleavase® BN nuclease.
It can be seen from these data that the helical arch of these proteins can be opened without destroying the enzyme or its ability to specifically recognize cleavage structures. The Cleavase® BN/thrombin mutant has an increased ability to cleave without reference to a 5′ end, as discussed above. The ability to cleave such structures will allow the cleavage of long molecules, such as genomic DNA that, while often not circular, may present many desirable cleavage sites that are at a far removed from any available 5′ end. Cleavage structures may be made at such sites either by folding of the strands (i.e., CFLP® cleavage) or by the introduction of structure-forming oligonucleotides (U.S. Pat. No. 5,422,253). 5′ ends of nucleic acids can also be made unavailable because of binding of a substance too large to thread through the helical arch. Such binding moieties may include proteins such as streptavidin or antibodies, or solid supports such as beads or the walls of a reaction vessel. A cleavage enzyme with an opening in the loop of the helical arch will be able to cleave DNAs that are configured in this way, extending the number of ways in which reactions using such enzymes can be formatted.
b) Cleavase® DN Nuclease
i) Construction and Expression of Cleavase® DN Nuclease
A polymerization deficient mutant of Taq DNA polymerase, termed Cleavase® DN nuclease, was constructed. Cleavase® DN nuclease contains an asparagine residue in place of the wild-type aspartic acid residue at position 785 (D785N).
DNA encoding the Cleavase® DN nuclease was constructed from the gene encoding for Cleavase® A/G (mutTaq, Ex. 2) in two rounds of site-directed mutagenesis. First, the G at position 1397 and the G at position 2264 of the Cleavase® A/G gene (SEQ ID NO:21) were changed to A at each position to recreate a wild-type DNAPTaq gene. As a second round of mutagenesis, the wild type DNAPTaq gene was converted to the Cleavase® DN gene by changing the G at position 2356 to A. These manipulations were performed as follows.
DNA encoding the Cleavase® A/G nuclease was recloned from pTTQ18 plasmid (Ex. 2) into the pTrc99A plasmid (Pharmacia) in a two step procedure. First, the pTrc99A vector was modified by removing the G at position 270 of the pTrc99A map, creating the pTrc99G cloning vector. To this end, pTrc99A plasmid DNA was cut with NcoI and the recessive 3′ ends were filled-in using the Klenow fragment of E.coli polymerase I in the presence of all four dNTPs at 37° C. for 15 min. After inactivation of the Klenow fragment by incubation at 65° C. for 10 min, the plasmid DNA was cut with EcoRI, the ends were again filled-in using the Klenow fragment in the presence of all four dNTPs at 37° C. for 15 min. The Klenow fragment was then inactivated by incubation at 65° C. for 10 min. The plasmid DNA was ethanol precipitated, recircularized by ligation, and used to transform E.coli JM109 cells (Promega). Plasmid DNA was isolated from single colonies and deletion of the G at position 270 of the pTrc99A map was confirmed by DNA sequencing.
As a second step, DNA encoding the Cleavase® A/G nuclease was removed from the pTTQ18 plasmid using EcoRI and SalI and the DNA fragment carrying the Cleavase® A/G nuclease gene was separated on a 1% agarose gel and isolated with Geneclean II Kit (Bio 101, Vista, Calif.). The purified fragment was ligated into the pTrc99G vector which had been cut with EcoRI and SalI. The ligation mixture was used to transform competent E.coli JM109 cells (Promega). Plasmid DNA was isolated from single colonies and insertion of the Cleavase® A/G nuclease gene was confirmed by restriction analysis using EcoRI and SalI.
Plasmid DNA pTrcAG carrying the Cleavase® A/G nuclease gene cloned into the pTrc99A vector was purified from 200 ml of JM109 overnight culture using QIAGEN Plasmid Maxi kit (QIAGEN, Chatsworth, Calif.) according to manufacturer's protocol. pTrcAG plasmid DNA was mutagenized using two mutagenic primers, E465 (SEQ ID NO:62) (Integrated DNA Technologies, Iowa) and R754Q (SEQ ID NO:63) (Integrated DNA Technologies), and the selection primer Trans Oligo AlwNI/SpeI (Clontech, Palo Alto, Calif., catalog #6488-1) according to Transformer Site-Directed Mutagenesis Kit protocol (Clontech) to produce a restored wild-type DNAPTaq gene (pTrcWT).
pTrcWT plasmid DNA carrying the wild-type DNAPTaq gene cloned into the pTrc99A vector was purified from 200 ml of JM109 overnight culture using QIAGEN Plasmid Maxi kit (QIAGEN, Chatsworth, Calif.) according to manufacturer's protocol. pTrcWT was then mutagenized using the mutagenic primer D785N (SEQ ID NO:64) (Integrated DNA Technologies) and the selection primer Switch Oligo SpeI/AlwNI (Clontech, catalog #6373-1) according to TransformerTM Site-Directed Mutagenesis Kit protocol (Clontech) to create a plasmid containing DNA encoding the Cleavase® DN nuclease. The DNA sequence encoding the Cleavase® DN nuclease is provided in SEQ ID NO:65; the amino acid sequence of Cleavase® DN nuclease is provided in SEQ ID NO:66.
A large scale preparation of the Cleavase® DN nuclease was done using E. coli cells overexpressing the Cleavase® DN nuclease as described in Example 28.
c) Cleavase® DA Nuclease and Cleavase® DV Nuclease
Two polymerization deficient mutants of Taq DNA polymerase, termed Cleavase® DA nuclease and Cleavase® DV nuclease, were constructed. The Cleavase® DA nuclease contains a alanine residue in place of the wild-type aspartic acid residue at position 610 (D785A). The Cleavase® DV nuclease contains a valine residue in place of the wild-type aspartic acid residue at position 610 (D610V).
i) Construction and Expression of the Cleavase® DA and Cleavase® DV Nucleases
To construct vectors encoding the Cleavase® DA and DV nucleases, the Cleavase® A/G nuclease gene contained within pTrcAG was mutagenized with two mutagenic primers, R754Q (SEQ ID NO:63) and D610AV (SEQ ID NO:67) and the selection primer Trans Oligo AlwNI/SpeI (Clontech, catalog #6488-1) according to the TransformerTM Site-Directed Mutagenesis Kit protocol (Clontech,) to create a plasmid containing DNA encoding the Cleavase® DA nuclease or Cleavase® DV nuclease. The D610AV oligonucleotide was synthesized to have a purine, A or G, at position 10 from the 5′ end of the oligonucleotide. Following mutagenesis, plasmid DNA was isolated from single colonies and the type of mutation present, DA or DV, was determined by DNA sequencing. The DNA sequence encoding the Cleavase® DA nuclease is provided in SEQ ID NO:68; the amino acid sequence of Cleavase® DA nuclease is provided in SEQ ID NO:69. The DNA sequence encoding the Cleavase® DV nuclease is provided in SEQ ID NO:70; the amino acid sequence of Cleavase® DV nuclease is provided in SEQ ID NO:71.
Large scale preparations of the Cleavase® DA and Cleavase® DV nucleases was done using E. coli cells overexpressing the Cleavase® DA nuclease or the Cleavase® DV nuclease as described in Example 28.
EXAMPLE 28
Cloning and Expression of Thermostable FEN-1 Endonucleases
Sequences encoding thermostable FEN-1 proteins derived from three Archaebacterial species were cloned and overexpressed in E. coli. This example involved a) cloning and expression of a FEN-1 endonuclease from Methanococcus jannaschii; b) cloning and expression of a FEN-1 endonuclease from Pyrococcus furiosus; c) cloning and expression of a FEN-1 endonuclease from Pyrococcus woesei; d) large scale preparation of recombinant thermostable FEN-1 proteins; and e) activity assays using FEN-1 endonucleases.
a) Cloning and Expression Of A FEN-1 Endonuclease From Methanococcus jannaschii 
DNA encoding the FEN-1 endonuclease from Methanococcus jannaschii (M. jannaschii) was isolated from M. jannaschii cells and inserted into a plasmid under the transcriptional control of an inducible promoter as follows. Genomic DNA was prepared from 1 vial of live M. jannaschii bacteria (DSMZ, Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany # 2661) with the DNA XTRAX kit (Gull Laboratories, Salt Lake City, Utah) according to the manufacturer's protocol. The final DNA pellet was resuspended in 100 μl of TE (10 mM Tris HCl, pH 8.0, 1 mM EDTA). One microliter of the DNA solution was employed in a PCR using the AdvantageTM cDNA PCR kit (Clonetech); the PCR was conducted according to manufacturer's recommendations. The 5′-end primer (SEQ ID NO:72) is complementary to the 5′ end of the Mja FEN-1 open reading frame with a one base substitution to create an NcoI restriction site [a fragment of the M. jannaschii genome which contains the gene encoding M. jannaschii (Mja) FEN-1 is available from GenBank as accession # U67585]. The 3′-end primer (SEQ ID NO:73) is complementary to a sequence about 15 base pairs downstream from the 3′ end of the Mja FEN-1 open reading frame with 2 base substitutions to create a SalI restriction enzyme site. The sequences of the 5′-end and 3′-end primers are: 5′-GGGATACCATGGGAGTGCAGTTTGG-3′ (SEQ ID NO:72) and 5′-GGTAAATTTTTCTCGTCGACATCCCAC-3′ (SEQ ID NO:73), respectively. The PCR reaction resulted in the amplification (i.e., production) of a single major band about 1 kilobase in length. The open reading frame (ORF) encoding the Mja FEN-1 endonuclease is provided in SEQ ID NO:74; the amino acid sequence encoded by this ORF is provided in SEQ ID NO:75.
Following the PCR amplification, the entire reaction was electrophoresed on a 1.0% agarose gel and the major band was excised from the gel and purified using the Geneclean II kit (Bio110, Vista, Calif.) according to manufacturer's instructions. Approximately 1 μg of the gel-purified Mja FEN-1 PCR product was digested with NcoI and SalI. After digestion, the DNA was purified using the Geneclean II kit according to manufacturer's instructions. One microgram of the pTrc99a vector (Pharmacia, Piscataway, N.J.) was digested with NcoI and SalI in preparation for ligation with the digested PCR product. One hundred nanograms of digested pTrc99a vector and 250 ng of digested Mja FEN-1 PCR product were combined and ligated to create pTrc99-MJFEN1. pTrc99-MJFEN1 was used to transform competent E. coli JM109 cells (Promega) using standard techniques.
b) Cloning and Expression Of A FEN-1 Endonuclease From Pyrococcus furiosus 
DNA encoding the Pyrococcus furiosus (P. furiosus) FEN-1 endonuclease was obtained by PCR amplification using a plasmid containing DNA encoding the P. furiosus (Pfu) FEN-1 endonuclease (obtained from Dr. Frank Robb, Center of Marine Biotechnology, Baltimore, Md.). DNA sequences encoding a portion of the Pfu FEN-1 endonuclease can be obtained from GenBank as accession Nos. AA113505 and W36094. The amplified Pfu FEN-1 gene was inserted into the pTrc99a expression vector (Pharmacia) to place the Pfu FEN-1 gene under the transcriptional control of the inducible trc promoter. The PCR amplification was conducted as follows. One hundred microliter reactions contained 50 mM Tris HCl, pH 9.0, 20 mM (NH4)2SO4, 2 mM MgCl2, 50 μM dNTPs, 50 pmole each primer, 1 U Tfl polymerase (Epicentre Technologies, Madison, Wis.) and 1 ng of FEN-1 gene-containing plasmid DNA. The 5′-end primer (SEQ ID NO:76) is complementary to the 5′ end of the Pfu FEN-1 open reading frame but with two substitutions to create an NcoI site and the 3′-end primer (SEQ ID NO:77) is complementary to a region located about 30 base pairs downstream of the FEN-1 open reading frame with two substitutions to create a PstI site. The sequences of the 5′-end and 3′-end primers are: 5′-GAGGTGATACCATGGGTGTCC-3′ (SEQ ID NO:76) and 5′-GAAACTCTGCAGCGCGTCAG-3′ (SEQ ID NO:77), respectively. The PCR reaction resulted in the amplification of a single major band about 1 kilobase in length. The open reading frame (ORF) encoding the Pfu FEN-1 endonuclease is provided in SEQ ID NO:78; the amino acid sequence encoded by this ORF is provided in SEQ ID NO:79.
Following the PCR amplification, the entire reaction was electrophoresed on a 1.0% agarose gel and the major band was excised from the gel and purified using the Geneclean II kit (Bio101) according to manufacturer's instructions. Approximately 1 μg of gel purified Pfu FEN-1 PCR product was digested with NcoI and PstI. After digestion, the DNA was purified using the Geneclean II kit according to manufacturer's instructions. One microgram of the pTrc99a vector was digested with NcoI and PstI prior to ligation with the digested PCR product. One hundred nanograms of digested pTrc99a and 250 ng of digested Pfu FEN-1 PCR product were combined and ligated to create pTrc99-PFFEN1. pTrc99-PFFEN1 was used to transform competent E. coli JM109 cells (Promega) using standard techniques.
c) Cloning and Expression Of A FEN-1 Endonuclease From Pyrococcus woesei 
For the cloning of DNA encoding the Pyrococcus woesei (Pwo) FEN-1 endonuclease, DNA was prepared from lyophilized P. woesei bacteria (DSMZ #3773) as described [Zwickl et al. (1990) J. Bact. 172:4329] with several changes. Briefly, one vial of P. woesei bacteria was rehydrated and resuspended in 0.5 ml of LB (Luria broth). The cells were centrifuged at 14,000×g for 1 min and the cell pellet was resuspended in 0.45 ml of TE. Fifty microliters of 10% SDS was added and the mixture was incubated at RT for 5 min. The cell lysate was then extracted three time with 1:1 phenol:chloroform and three times with chloroform. Five hundred microliters of isopropanol was added to the extracted lysate and the DNA was pelleted by centrifugation at 14,000×g for 10 min. The DNA pellet was washed in 0.5 ml of 70% ethanol and the DNA was pelleted again by centrifugation at 14,000×g for 5 min. The DNA pellet was dried and resuspended in 100 μl of TE and used for PCR reactions without further purification.
To generate a P. woesei FEN-1 gene fragment for cloning into an expression vector, low stringency PCR was attempted with primers complementary to the ends of the P. furiosus FEN-1 gene open reading frame. The sequences of the 5′-end and 3′-end primers are 5′-GATACCATGGGTGTCCCAATTGGTG-3′ (SEQ ID NO:80) and 5′-TCGACGTCGACTTATCTCTTGAACCAACTTTCAAGGG-3′ (SEQ ID NO:81), respectively. The high level of sequence similarity of protein homologs (i.e., proteins other than FEN-1 proteins) from P. furiosus and P. woesei suggested that there was a high probability that the P. woesei FEN-1 gene could be amplified using primers containing sequences complementary to the P. furiosus FEN-1 gene. However, this approach was unsuccessful under several different PCR conditions.
The DNA sequence of FEN-1 genes from P. furiosus and M. jannaschii were aligned and blocks of sequence identity between the two genes were identified. These blocks were used to design internal primers (i.e., complementary to sequences located internal to the 5′ and 3′ ends of the ORF) for the FEN-1 gene that are complementary to the P. furiosus FEN-1 gene in those conserved regions. The sequences of the 5′- and 3′-internal primers are 5′-AGCGAGGGAGAGGCCCAAGC-3′ (SEQ ID NO:82) and 5′-GCCTATGCCCTTTATTCCTCC-3′ (SEQ ID NO:83), respectively. A PCR employing these internal primers was conducted using the AdvantageTM PCR kit and resulted in production of a major band of  ̃300 bp.
Since the PCR with the internal primers was successful, reactions were attempted which contained mixtures of the internal (SEQ ID NOS:82 and 83) and external (SEQ ID NOS:80 and 81) primers. A reaction containing the 5′-end external primer (SEQ ID NO:80) and 3′-end internal primer (SEQ ID NO:83) resulted in the production of a 600 bp band and a reaction containing the 5′-end internal primer (SEQ ID NO:82) and 3′-end external primer (SEQ ID NO:81) resulted in the production of a 750 bp band. These overlapping DNA fragments were gel-purified and combined with the external primers (SEQ ID NOS:80 and 81) in a PCR reaction. This reaction generated a 1 kb DNA fragment containing the entire Pwo FEN-1 gene open reading frame. The resulting PCR product was gel-purified, digested, and ligated exactly as described above for the Mja FEN-1 gene PCR product. The resulting plasmid was termed pTrc99-PWFEN1. pTrc99-PWFEN1 was used to transform competent E. coli JM109 cells (Promega) using standard techniques.
d) Large Scale Preparation of Recombinant Thermostable FEN-1 Proteins
The Mja, Pwo and Pfu FEN-1 proteins were purified by the following technique which is derived from a Taq DNA polymerase preparation protocol [Engelke et al. (1990) Anal. Biochem. 191:396] as follows. E. coli cells (strain JM109) containing either pTrc99-PFFEN1, pTrc99-PWFEN1, or pTrc99-MJFEN1 were inoculated into 3 ml of LB (Luria Broth) containing 100 μg/ml ampicillin and grown for 16 hrs at 37° C. The entire overnight culture was inoculated into 200 ml or 350 ml of LB containing 100 μg/ml ampicillin and grown at 37° C. with vigorous shaking to an A600 of 0.8. IPTG (1 M stock solution) was added to a final concentration of 1 mM and growth was continued for 16 hrs at 37° C.
The induced cells were pelleted and the cell pellet was weighed. An equal volume of 2× DG buffer (100 mM Tris-HCl, pH 7.6, 0.1 mM EDTA) was added and the pellet was resuspended by agitation. Fifty mg/ml lysozyme (Sigma, St. Louis, Mo.) was added to 1 mg/ml final concentration and the cells were incubated at room temperature for 15 min. Deoxycholic acid (10% solution) was added dropwise to a final concentration of 0.2% while vortexing. One volume of H2O and 1 volume of 2× DG buffer was added and the resulting mixture was sonicated for 2 minutes on ice to reduce the viscosity of the mixture. After sonication, 3 M (NH4)2SO4 was added to a final concentration of 0.2 M and the lysate was centrifuged at 14000×g for 20 min at 4° C. The supernatant was removed and incubated at 70° C. for 60 min at which time 10% polyethylimine (PEI) was added to 0.25%. After incubation on ice for 30 min., the mixture was centrifuged at 14,000×g for 20 min at 4° C. At this point, the supernatant was removed and the FEN-1 proteins was precipitated by the addition of (NH4)2SO4 as follows.
For the Pwo and the Pfu FEN-1 preparations, the FEN-1 protein was precipitated by the addition of 2 volumes of 3 M (NH4)2SO4. The mixture was incubated overnight at room temperature for 16 hrs and the protein was centrifuged at 14,000×g for 20 min at 4° C. The protein pellet was resuspended in 0.5 ml of Q buffer (50 mM Tris-HCl, pH 8.0, 0.1 mM EDTA, 0.1% Tween 20). For the Mja FEN-1 preparation, solid (NH4)2SO4 was added to a final concentration of 3 M ( ̃75% saturated), the mixture was incubated on ice for 30 min, and the protein was spun down and resuspended as described above.
The resuspended protein preparations were quantitated by determination of the A279 and aliquots containing 2-4 μg of total protein were electrophoresed on a 10% SDS polyacrylamide gel (29:1 acrylamide:bis-acrylamide) in standard Laemmli buffer [Laemmli (1970) Nature 277:680] and stained with Coomassie Brilliant Blue R; the results are shown in FIG. 64.
In FIG. 64, lane 1 contains molecular weight markers (Mid-Range Protein Molecular Weight Markers; Promega); the size of the marker proteins is indicated to the left of the gel. Lane 2 contains purified Cleavase® BN nuclease; lanes 3-5 contain extracts prepared from E. coli expressing the Pfu, Pwo and Mja FEN-1 nucleases, respectively. The calculated (i.e., using a translation of the DNA sequence encoding the nuclease) molecular weight of the Pfu FEN-1 nuclease is 38,714 daltons and the calculated molecular weight for the Mja FEN-1 nuclease is 37,503 Daltons. The Pwo and Pfu FEN-1 proteins co-migrated on the SDS-PAGE gel and therefore, the molecular weight of the Pwo FEN-1 nuclease was estimated to be 38.7 kDa.
e) Activity Assays Using FEN-1 Endonucleases
i) Mixed Hairpin Assay
The Cleavase® BN nuclease has an approximately 60-fold greater affinity for a 12 base pair stem-loop structure than an 8 base pair stem-loop DNA structure. As a test for activity differences between the Cleavase® BN nuclease and the FEN-1 nucleases, a mixture of oligonucleotides having either a 8 or a 12 bp stem-loop (see FIG. 60 which depicts the S-33 and 11-8-0 oligonucleotides) was incubated with an extract prepared from E. coli cells overexpressing the Mja FEN-1 nuclease (prepared as described above). Reactions contained 0.05 μM of oligonucleotides S-33 (SEQ ID NO:84) and 11-8-0 (SEQ ID NO:85) (both oligonucleotides contained 5′-fluorescein labels), 10 mM MOPS, pH 7.5, 0.05% Tween-20, 0.05% NP-40, 1 mM MnCl2. Reactions were heated to 90° C. for 10 seconds, cooled to 55° C., then 1 μl of crude extract (Mja FEN-1) or purified enzyme (Cleavase® BN nuclease) was added and the mixtures were incubated at 55° C. for 10 minutes; a no enzyme control was also run. The reactions were stopped by the addition of formamide/EDTA, the samples were electrophoresed on a denaturing 20% acrylamide gel and visualized on a Hitachi FMBIO 100 fluoroimager. The resulting image is shown in FIG. 65.
In FIG. 65, lane 1 contains the reaction products generated by the Cleavase® BN nuclease, lane 2 contains the reaction products from the no enzyme control reaction and lane 3 contains the reaction products generated by the Mja FEN-1 nuclease. The data shown in FIG. 76 demonstrates that the Cleavase® BN nuclease strongly prefers the S33 structure (12 bp stem-loop) while the Mja FEN-1 nuclease cleaves structures having either an 8 or a 12 bp stem-loop with approximately the same efficiency. This shows that the Mja FEN-1 nuclease has a different substrate specificity than the Cleavase® BN nuclease, a useful feature for InvaderTM assays or CFLP® analysis as discussed in the Description of the Invention.
EXAMPLE 29
Terminal Deoxynucleotidyl Transferase Selectively Extends the Products of InvaderTM-Directed Cleavage
The majority of thermal degradation products of DNA probes will have a phosphate at the 3′-end. To investigate if the template-independent DNA polymerase, terminal deoxynucleotide transferase (TdT) can tail or polymerize the aforementioned 3′-end phosphates (i.e., add nucleotide triphosphates to the 3′ end) the following experiment was performed.
To create a sample containing a large percentage of thermal degradation products, the 5′ fluorescein-labelled oligonucleotide 34-078-01 (SEQ ID NO:86) (200 pmole) was incubated in 100 μl 10 mM NaCO3 (pH 10.6), 50 mM NaCl at 95° C. for 13 hours. To prevent evaporation, the reaction mixture was overlaid with 60 μl ChillOutTM14 liquid wax. The reaction mixture was then divided into two equal aliquots (A and B). Aliquot A was mixed with one-tenth volume 3M NaOAc followed by three volumes ethanol and stored at −20° C. Aliquot B was dephosphorylated by the addition of 0.5 μl of 1M MgCl2 and 1 μl of 1 unit/μl Calf Intestine Alkaline Phosphatase (CIAP) (Promega), with incubation at 37° C. for 30 minutes. An equal volume of phenol:chloroform:isomayl alcohol (24:24:1) was added to the sample followed by vortexing for one minute and then centrifugation 5 minutes at maximum speed in a microcentrifuge to separate the phases. The aqueous phase was removed to a new tube to which one-tenth volume 3M NaOAc, and three volumes ethanol was added followed by storage at −20° C. for 30 minutes. Both aliquots (A and B) were then centrifuged for 10 minutes at maximum speed in a microcentrifuge to pellet the DNA. The pellets were then washed two times each with 80% ethanol and then desiccated to dryness. The dried pellets were then dissolved in 70 μl ddH2O each.
The TdT reactions were conducted as follows. Six mixes were assembled, all mixes contained 10 mM TrisOAc (pH 7.5), 10 mM MgOAc, 50 mM KCl, and 2 mM dATP. Mixes 1 and 2 contained one pmole of untreated 34-078-01 (SEQ ID NO:86), mixes 3 and 4 contained 2 μl of aliquot A (above), mixes 5 and 6 contained 2 μl of aliquot B (above). To each 9 μl of mixes 1, 3 and 5, 1 μl ddH2O was added, to each 9 μl of mixes 2, 4, and 6, 1 μl of 20 units/μl TdT (Promega) was added. The mixes were incubated at 37° C. for 1 hour and then the reaction was terminated by the addition of 5 μl 95% formamide with 10 mM EDTA and 0.05% marker dyes. Five microliters of each mixture was resolved by electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked) with 7 M urea, in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA, and imaged using with the FMBIO Image Analyzer with a 505 nm filter. The resulting imager scan is shown in FIG. 66.
In FIG. 66, lanes 1, 3 and 5 contain untreated 34-078-01 (SEQ ID NO:86), heat-degraded 34-078-01, and heat-degraded, dephosphorylated, 34-078-01, respectively incubated in the absence of TdT. Lanes 2, 4 and 6 contain, untreated 34-078-01, heat-degraded 34-078-01, and heat-degraded, dephosphorylated, 34-078-01, respectively incubated in the presence of TdT.
As shown in FIG. 66, lane 4, TdT was unable to extend thermal degradation products which contain a 3′-end phosphate group, and selectively extends molecules which have a 3′-end hydroxyl group.
EXAMPLE 30
Specific TdT Tailing of the Products of InvaderTM-Directed Cleavage with Subsequent Capture and Detection on Nitrocellulose Supports
When TdT is used to extend the specific products of cleavage, one means of detecting the tailed products is to selectively capture the extension products on a solid support before visualization. This example demonstrates that the cleavage products can be selectively tailed by the use of TdT and deoxynucleotide triphosphates, and that the tailed products can be visualized by capture using a complementary oligonucleotide bound to a nitrocellulose support.
To extend the cleavage product produced in an InvaderTM-directed cleavage reaction, the following experiment was performed. Three reaction mixtures were assembled, each in a buffer of 10 mM MES (pH 6.5), 0.5% Tween-20, 0.5% NP-40. The first mixture contained 5 fmols of target DNA-M13mp18, 10 pmols of probe oligo 32-161-2 (SEQ ID NO:87; this probe oligonucleotide contains 3′ ddC and a Cy3 amidite group near the 3′ end), and 5 pmols of InvaderTM oligonucleotide 32 161-1 (SEQ ID NO:88; this oligo contains a 3′ ddC). The second mixture contained the probe and InvaderTM oligonucleotides without target DNA. The third mixture was the same as the first mixture, and contained the same probe sequence, but with a 5′ fluorescein label [oligo 32-161-4 (SEQ ID NO:89; this oligo contains a 3′ ddC, 5′ fluorescein label, and a Cy3 dye group near the 3′ end)], so that the InvaderTM-directed cleavage products could be detected before and after cleavage by fluorescence imaging. The probe only control sample contained 10 pmols of oligo 32-161-2 (SEQ ID NO:87). Each 3 μl of enzyme mix contained 5 ng of Cleavase® DN nuclease in 7.5 mM MgCl2. The TdT mixture (per each 4 μl) contained: 10 U of TdT (Promega), 1 mM CoCl2, 50 mM KCl, and 100 μM of dTTP. The InvaderTM cleavage reaction mixtures described above were assembled in thin wall tubes, and the reactions were initiated by the addition of 3 μl of Cleavase® DN enzyme mix. The reactions were incubated at 65° C. for 20 min. After cooling to 37° C., 4 μl of the TdT mix was added and the samples were incubated for 4 min at 37° C., Biotin-16-dUTP was then added to 100 μM and the samples were incubated for 50 min at 37° C. The reactions were terminated by the addition of 1 μl of 0.5 M EDTA.
To test the efficiency of tailing the products were run on an acrylamide gel. Four microliters of each reaction mixture was mixed with 2.6 μl of 95% formamide, 10 mM EDTA and 0.05% methyl violet and heated to 90° C. for 1 min, and 3 μl were loaded on a 20% denaturing acrylamide gel (19:1 cross-linked ) with 7 M urea, in buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA. A marker [ΦX174-HinfI (fluorescein labeled)] also was loaded. After electrophoresis, the gel was analyzed using a FMBIO-100 Image Analyzer (Hitachi) equipped with a 505 nm filter. The resulting scan is shown in FIG. 67.
In FIG. 67, lane 1 contained the probe 32-161-2 only, without any treatment. Lanes 2 and 3 contained the products of reactions run without target DNA, without or with subsequent TdT tailing, respectively. Lanes 4 and 5 contained the products of reactions run with target DNA, probe oligo 32-161-2 (SEQ ID NO:87) and InvaderTM oligo 32-161-1 (SEQ ID NO:88), without or with subsequent TdT tailing, respectively. Lanes 6 and 7 show the products of reactions containing target DNA, probe oligo 32-161-4 (SEQ ID NO:89) and InvaderTM oligo 32-161-1 (SEQ ID NO:88), without or with subsequent TdT tailing, respectively. Lane M contains the marker ΦX174-HinfI.
The reaction products in lanes 4 and 5 are the same as those seen in lanes 6 and 7, except that the absence of a 5′ fluorescein on the probe prevents detection of the released 5′ product (indicated as “A” near the bottom of the gel) or the TdT extended 5′ product (indicated as “B”, near the top of the gel). The Cy3-labeled 3′ portion of the cleaved probe is visible in all of these reactions (indicated as “C”, just below the center of the gel).
To demonstrate detection of target-dependent Invader-directed cleavage products on a solid support, the reactions from lanes 3 and 5 were tested on the Universal Genecomb® (Bio-Rad) which is a standard nitrocellulose matrix on a rigid nylon backing styled in a comb format, as depicted in FIG. 68. Following the manufacturer's protocol, with one modification: 10 μl of the Invader-directed cleavage reactions were used instead the recommended 10% of a PCR. To capture the cleavage products, 2.5 pmols of the capture oligo 59-28-1 (SEQ ID NO:90) were spotted on each tooth. The capture and visualization steps were conducted according to the manufacturer's directions. The results are shown in FIG. 68.
In FIG. 68, teeth numbered 6 and 7 show the capture results of reactions performed without and with target DNA present. Tooth 8 shows the kit positive control.
The darkness of the spot seen on tooth 7, when compared to tooth 6, clearly indicates that products of InvaderTM-directed cleavage assays may be specifically detected on solid supports. While the Universal Genecomb® was used to demonstrate solid support capture in this instance, other support capture methods known to those skilled in the art would be equally suitable. For example, beads or the surfaces of reaction vessels may easily be coated with capture oligonucleotides so that they can then be used in this step. Alternatively, similar solid supports may easily be coated with streptavidin or antibodies for the capture of biotin- or hapten-tagged products of the cleavage/tailing reaction. In any of these embodiments, the products may be appropriately visualized by detecting the resulting fluorescence, chemiluminescence, colorimetric changes, radioactive emissions, optical density change or any other distinguishable feature of the product.
EXAMPLE 31
Comparison of the Effects of Invasion Length and 5′ Label of the Probe on InvaderTM-Directed Cleavage by the Cleavase® A/G and Pfu FEN-1 Nucleases
To investigate the effect of the length of invasion as well as the effect of the type of dye on ability of Pfu FEN-1 and the Cleavase® A/G nuclease to cleave 5′ arms, the following experiment was performed. Three probes of similar sequences labeled with either fluorescein, TET, or Cy3, were assembled in reactions with three InvaderTM oligonucleotides which created overlapping target hybridization regions of eight, five, and three bases along the target nucleic acid, M13mp18.
The reactions were conducted as follows. All conditions were performed in duplicate. Enzyme mixes for Pfu FEN-1 and the Cleavase® A/G nuclease were assembled. Each 2 μl of the Pfu FEN-1 mix contained 100 ng of Pfu FEN-1 (prepared as described in Ex. 28) and 7.5 mM MgCl2. Each 2 μl of the Cleavase® A/G mix contained 5.3 ng of the Cleavase® A/G nuclease and 4.0 mM MnCl2. Six master mixes containing buffer, M13mp18, and InvaderTM oligonucleotides were assembled. Each 7 μl of mixes 1-3 contained 1 fmol M13mp18, 10 pmoles InvaderTM oligonucleotide [34-078-4 (SEQ ID NO:39), 24-181-2 (SEQ ID NO:91), or 24-181-1 (SEQ ID NO:92) in 10 mM MOPS (pH 7.5), 150 mM LiCl. Each 7 μl of mixes 4-6 contained 1 fmol of M13mp18, 10 pmoles of InvaderTM oligonucleotide [34-078-4 (SEQ ID NO:39), 24-181-2 (SEQ ID NO:91), or 24-181-1 (SEQ ID NO:92)] in 10 mM Tris (pH 8.0). Mixtures 1-6 were then divided into three mixtures each, to which was added either the fluorescein-labeled probe (oligo 34-078-01; SEQ ID NO:86), the Cy3-labeled probe (oligo 43-20; SEQ ID NO:93) or the TET-labeled probe (oligo 90; SEQ ID NO:32 containing a 5′ TET label). Each 7 μl of all mixtures contained 10 pmoles of corresponding probe. The DNA solutions described above were covered with 10 μl of ChillOut® evaporation barrier and brought to 68° C.
The reactions made from mixes 1-3 were started with 2 μl of the Cleavase® A/G nuclease mix, and the reactions made from mixes 4-6 were started with 2 μl of the Pfu FEN-1 mix. After 30 minutes at 68° C., the reactions were terminated by the addition of 8 μl of 95% formamide with 10 mM EDTA and 0.05% marker dyes. Samples were heated to 90° C. for 1 minute immediately before electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked) with 7 M urea, in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA. The products of the cleavage reactions were visualized following electrophoresis by the use of a Hitachi FMBIO fluorescence imager. Results from the fluorescein-labeled probe are shown in FIG. 69, results from the Cy3-labeled probe in FIG. 70, and results from the TET-labeled probe in FIG. 71. In each of these figures the products of cleavage by Cleavase® A/G are shown in lanes 1-6 and the products of cleavage by PfuFEN-1 are shown in lanes 7-12. In each in case the uncut material appears as a very dark band near the top of the gel, indicated by a “U” on the left. The products of cleavage directed by Invader oligonucleotides with 8, 5 or 3 bases of overlap (i.e., the “X” region was 8, 5, or 3 nt long) are shown in the first, second and third pair of lanes in each set, respectively and the released labeled 5′ ends from these reactions are indicated by the numbers 8, 5, and 3 on the left. Note that in the cleavage reactions shown in FIG. 70 the presence of the positively charged Cy3 dye causes the shorter products to migrate more slowly than the larger products. These products do not contain any additional positive charges, e.g., amino modifications as used in Example 23, and thus still carry a net negative charge, and migrate towards the positive electrode in a standard electrophoresis run.
It can be seen from these data that the Cleavase® A/G and Pfu FEN-1 structure-specific nucleases respond differently to both dye identity and to the size of the piece to be cleaved from the probe. The Pfu FEN-1 nuclease showed much less variability in response to dye identity than did the Cleavase® A/G nuclease, showing that any dye wold be suitable for use with this enzyme. In contrast, the amount of cleavage catalyzed by the Cleavase® A/G nuclease varied substantially with dye identity. Use of the fluorescein dye gave results very close to those seen with the Pfu FEN-1 nuclease, while the use of either Cy3 or TET gave dramatically reduced signal when compared to the Pfu FEN-1 reactions. The one exception to this was in the cleavage of the 3 nt product carrying a TET dye (lanes 5 and 6, FIG. 71), in which the Cleavase® A/G nuclease gave cleavage at the same rate as the Pfu FEN-1 nuclease. These data indicate that, while Cleavase® A/G may be used to cleave probes labeled with these other dyes, the Pfu FEN-1 nuclease is a preferred nuclease for cleavage of Cy3- and TET-labeled probes.
EXAMPLE 32
Examination of the Effects of a 5′ Positive Charge on the Rate of Invasive Cleavage Using the Cleavase® A/G or Pfu FEN-1 Nucleases
To investigate whether the positive charges on 5′ end of probe oligonucleotides containing a positively charged adduct(s) (i.e., charge reversal technology or CRT probes as described in Ex. 23 and 24 have an effect on the ability of the Cleavase® A/G or Pfu FEN-1 nucleases to cleave the 5′ arm of the probe, the following experiment was performed.
Two probe oligonucleotides having the following sequences were utilized in InvaderTM reactions: Probe 34-180-1: (N-Cy3)TNH2TNH2CCAGAGCCTAATTTGCCAGT(N-fluorescein)A, where N represents a spacer containing either the Cy3 or fluorescein group (SEQ ID NO:94) and Probe 34-180-2: 5′-(N-TET)TTCCAGAGCCTAATTTGCCAGT-(N-fluorescein)A, where N represents a spacer containing either the TET or fluorescein group (SEQ ID NO:95). Probe 34-180-1 has amino-modifiers on the two 5′ end T residues and a Cy3 label on the 5′ end, creating extra positive charges on the 5′ end. Probe 34-180-2 has a TET label on the 5′ end, with no extra positive charges. The fluorescein label on the 3′ end of probe 34-180-1 enables the visualization of the 3′ cleaved products and uncleaved probes together on an acrylamide gel run in the standard direction (i.e., with the DNA migrating toward the positive electrode). The 5′ cleaved product of probe 34-180-1 has a net positive charge and will not migrate in the same direction as the uncleaved probe, and is thus visualized by resolution on a gel run in the opposite direction (i.e.; with this DNA migrating toward the negative electrode).
The cleavage reactions were conducted as follows. All conditions were performed in duplicate. Enzyme mixes for the Pfu FEN-1 and Cleavase® A/G nucleases were assembled. Each 2 μl of the Pfu FEN-1 mix contained 100 ng of Pfu FEN-1 (prepared as described in Ex. 28) and 7.5 mM MgCl2. Each 2 μl of the Cleavase® A/G nuclease mix contained 26.5 ng of Cleavase® A/G nuclease and 4.0 mM MnCl2. Four master mixes containing buffer, M13mp18, and InvaderTM oligonucleotides were assembled. Each 7 μl of mix 1 contained 5 fmol M13mp18, 10 pmoles InvaderTM oligonucleotide 123 (SEQ ID NO:96) in 10 mM HEPES (pH 7.2). Each 7 μl of mix 2 contained 1 fmol M13mp18, 10 pmoles InvaderTM oligonucleotide 123 in 10 mM HEPES (pH 7.2). Each 7 μl of mix 3 contained 5 fmol M13mp18, 10 pmoles InvaderTM oligonucleotide 123 in 10 mM HEPES (pH 7.2), 250 mM KGlu. Each 7 μl of mix 4 contained 1 fmol M13mp18, 10 pmoles InvaderTM oligonucleotide 123 in 10 mM HEPES (pH 7.2), 250 mM KGlu. For every 7 μl of each mix, 10 pmoles of either probe 34-180-1 (SEQ ID NO:94) or probe 34-180-2 (SEQ IDNO:95) was added. The DNA solutions described above were covered with 10 μl of ChillOut® evaporation barrier and brought to 65° C. The reactions made from mixes 1-2 were started by the addition of 2 μl of the Pfu FEN-1 mix, and the reactions made from mixes 3-4 were started by the addition of 2 μl of the Cleavase® A/G nuclease mix. After 30 minutes at 65° C., the reactions were terminated by the addition of 8 μl of 95% formamide containing 10 mM EDTA. Samples were heated to 90° C. for 1 minute immediately before electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked) with 7 M urea, in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA and a 20% native acrylamide gel (29:1 cross-linked) in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA.
The products of the cleavage reactions were visualized following electrophoresis by the use of a Hitachi FMBIO fluorescence imager. The resulting images are shown in FIG. 72. FIG. 72A shows the denaturing gel which was run in the standard electrophoresis direction, and FIG. 72B shows the native gel which was run in the reverse direction. The reaction products produced by Pfu FEN-1 and Cleavase® A/G nucleases are shown in lanes 1-8 and 9-16, respectively. The products from the 5 fmol M13mp18 and 1 fmol M13mp18 reactions are shown in lanes 1-4, 9-12 (5 fmol) and 5-8, 13-16 (1 fmol). Probe 34-180-1 is in lanes 1-2, 5-6, 9-10, 13-14 and probe 34-180-2 is in lanes 3-4, 7-8, 11-12, 15-16.
The fluorescein-labeled 3′ end fragments from all cleavage reactions are shown in FIG. 72A, indicated by a “3”′ mark at the left. The 3 nt 5′ TET-labeled products are not visible in this figure, while the 5° Cy3-labeled products are shown in FIG. 72B.
The 3′ end bands in FIG. 72A can be used to compare the rates of cleavage by the different enzymes in the presence of the different 5′ end labels. It can be seen from this band that regardless of the amount of target nucleic acid present, both the Pfu FEN-1 and the Cleavase® A/G nucleases show more product from the 5′ TET-labeled probe. With the Pfu FEN-1 nuclease this preference is modest, with only an approximately 25 to 40% increase in signal. In the case of the Cleavase® A/G nuclease, however, there is a strong preference for the 5′ TET label. Therefore, although when the charge reversal method is used to resolve the products, a substantial amount of product is observed from the Cleavase® A/G nuclease-catalyzed reactions, the Pfu FEN-1 nuclease is a preferred enzyme for cleavage of Cy3-labeled probes.
EXAMPLE 33
The Use of Universal Bases in the Detection of Mismatches by InvaderTM Directed Cleavage
The term “degenerate base” refers to a base on a nucleotide that does not hydrogen bond in a standard “Watson-Crick” fashion to a specific base complement, i.e., A to T and G to C. For example, the inosine base can be made to pair via one or two hydrogen bonds to all of the natural bases (the “wobble” effect) and thus is called degenerate. Alternatively, a degenerate base may not pair at all; this type of base has been referred to as a “universal” base because it can be placed opposite any nucleotide in a duplex and, while it cannot contribute stability by base-pairing, it does not actively destabilize by crowding the opposite base. Duplexes using these universal bases are stabilized by stacking interactions only. Two examples of universal bases, 3-nitropyrrole and 5-nitroindole, are shown in FIG. 73. In hybridization, placement of a 3-nitropyrrole three bases from a mismatch position enhances the differential recognition of one base mismatches. The enhanced discrimination seems to come from the destabilizing effect of the unnatural base (i.e., an altered Tm in close proximity to the mismatch). To test this same principle as a way of sensitively detecting mismatches using the InvaderTM-directed cleavage assay, InvaderTM oligonucleotides were designed using the universal bases shown in FIG. 73, in the presence or absence of a natural mismatch. In these experiments, the use of single nitropyrrole bases or pairs of nitroindole bases that flank the site of the mismatch were examined.
The target, probe and InvaderTM oligonucleotides used in these assays are shown in FIG. 74. A 43 nucleotide oligonucleotide (oligo 109; SEQ ID NO:97) was used as the target. The probe oligonucleotide (oligo 61; SEQ ID NO:50) releases a net positively charged labeled product upon cleavage. In FIG. 74, the InvaderTM oligonucleotide is shown schematically above the target oligonucleotide as an arrow; the large arrowhead indicates the location of the mismatch between the InvaderTM oligos and the target. Under the target oligonucleotide, the completely complementary, all natural (i.e., no universal bases) InvaderTM oligo (oligo 67; SEQ ID NO:51) and a composite of InvaderTM oligos containing universal bases (“X”) on either side of the mismatch (“M”) are shown. The following InvaderTM oligos were employed: oligo 114 (SEQ ID NO:98) which contains a single nt mismatch; oligo 115 (SEQ ID NO:99) which contains two 5-nitroindole bases and no mismatch; oligo 116 (SEQ ID NO:100) which contains two 5-nitroindole bases and a single nt mismatch; oligo 112 (SEQ ID NO:101) which contains one 3-nitropyrrole base and no mismatch; oligo 113 (SEQ ID NO:102) which contains one 5-nitropyrrole base and a single nt mismatch; and oligo 67 (SEQ ID NO:51) which is completely complementary to the target.
The InvaderTM-directed cleavage reactions were carried out in 10 μl of 10 mM MOPS (pH 7.2), 100 mM KCl, containing 1 μM of the appropriate invading oligonucleotide (oligos 67, 112-116), 10 nM synthetic target 109, 1 μM Cy-3 labeled probe 61 and 2 units of Cleavase® DV (prepared as described in Ex. 27). The reactions were overlayed with Chill-Out® liquid wax, brought to the appropriate reaction temperature, 52° C., 55° C., or 58° C. and initiated with the addition of 1 μl of 40 mM MnCl2. Reactions were allowed to proceed for 1 hour and were stopped by the addition of 10 μl formamide. One fourth of the total volume of each reaction was loaded onto 20% non-denaturing polyacrylamide gels which were electrophoresed in the reverse direction. The products were visualized using an Hitachi FMBIO-100 fluorescent scanner using a 585 nm filter. The resulting images are shown in FIGS. 75A-C. In each panel, lanes 1-6 contain reactions products from reactions using InvaderTM oligo 67, 114, 115, 116, 112 and 113, respectively. Reactions run at 52° C., 55° C. and 58° C. are shown in Panels A, B and C, respectively.
These data show that two flanking 5-nitroindoles display a significantly greater differentiation then does the one 3-nitropyrrole system, or the all natural base hybridization, and this increased sensitivity is not temperature dependent. This demonstrates that the use of universal bases is a useful means of sensitively detecting single base mismatches between the target nucleic acid and the complex of detection oligonucleotides of the present invention.
EXAMPLE 34
Detection of Point Mutations in the Human Ras Oncogene Using a Miniprobe
It is demonstrated herein that very short probes can be used for sensitive detection of target nucleic acid sequences (Ex. 37). In this example it is demonstrated that the short probes work very poorly when mismatched to the target, and thus can be used to distinguish a given nucleic acid sequence from a close relative with only a single base difference. To test this system synthetic human ras oncogene target sequences were created that varied from each other at one position. Oligonucleotide 166 (SEQ ID NO:103) provided the wild-type ras target sequence. Oligonucleotide 165 (SEQ ID NO:104) provided the mutant ras target sequence. The sequence of these oligonucleotides are shown in FIG. 76, and the site of the sequence variation in the site corresponding to codon 13 of the ras gene is indicated. The InvaderTM oligonucleotide (oligo 162) has the sequence: 5′-GSCSTSCSASASGSGSCSACTCTTGCCTACGA-3′ (SEQ ID NO:105), where the “S” indicates thiol linkages [i.e., these are 2′-deoxynucleotide-5′-O-(1-thiomonophates)]. The miniprobe (oligo 161) has the sequence: 5′-(N-Cy3) TNH2TNH2CACCAG-3′ (SEQ ID NO:106) and is designed to detect the mutant ras target sequence (i.e., it is completely complementary to oligo 165). The stacker oligonucleotide (oligo 164) has the sequence: 5′-CSTSCSCSASASCSTSASCCACAAGTTTATATTCAG-3′ (SEQ ID NO:107). A schematic showing the assembly of these oligonucleotides into a cleavage structure is depicted in FIG. 76.
Each cleavage reaction contained 100 nM of both the invading (oligo 162) and stacking (oligo 164) oligonucleotides, 10 μM Cy3-labeled probe (oligo 161) and 100 pM of either oligo 165 or oligo 166 (target DNA) in 10 μl of 10 mM HEPES (pH 7.2), 250 mM KGlu, 4 mM MnCl2. The DNA mixtures were overlaid with mineral oil, heated to 90° C. for 15 sec then brought to a reaction temperature of 47°, 50°, 53° or 56° C. Reactions were initiated by the addition of 1 μl of 100 ng/μl Pfu FEN-1. Reactions were allowed to proceed for 3 hours and stopped by the addition of 10 μl formamide. One fourth of the total volume of each reaction was loaded onto a 20% non-denaturing polyacrylamide gel which was electrophoresed in the reverse direction. The gel was scanned using an Hitachi FMBIO-100 fluorescent scanner fitted with a 585 nm filter, and the resulting image is shown in FIG. 77.
In FIG. 77, for each reaction temperature tested, the products from reactions containing either the mutant ras target sequence (oligo 165) or the wild-type (oligo 166) are shown.
These data demonstrate that the miniprobe can be used to sensitively discriminate between sequences that differ by a single nucleotide. The miniprobe was cleaved to produce a strong signal in the presence of the mutant target sequence, but little or no miniprobe was cleaved in the presence of the wild-type target sequence. Furthermore, the discrimination between closely related targets is effective over a temperature range of at least 10° C., which is a much broader range of temperature than can usually be tolerated when the selection is based on hybridization alone (e.g., hybridization with ASOs). This suggests that the enzyme may be a factor in the discrimination, with the perfectly matched miniprobe being the preferred substrate when compared to the mismatched miniprobe. Thus, this system provides sensitive and specific detection of target nucleic acid sequences.
EXAMPLE 35
Effects of 3′ End Identity on Site of Cleavage of a Model Oligonucleotide Structure
As described in the examples above, structure-specific nucleases cleave near the junction between single-stranded and base-paired regions in a bifurcated duplex, usually about one base pair into the base-paired region. It was shown in Example 10 that thermostable 5′ nucleases, including those of the present invention (e.g., Cleavase® BN nuclease, Cleavase® A/G nuclease), have the ability to cleave a greater distance into the base paired region when provided with an upstream oligonucleotide bearing a 3′ region that is homologous to a 5′ region of the subject duplex, as shown in FIG. 26. It has also been determined that the 3′ terminal nucleotide of the invader oligonucleotide may be unpaired to the target nucleic acid, and still shift cleavage the same distance into the down stream duplex as when paired. It is shown in this example that it is the base component of the nucleotide, not the sugar or phosphate, that is necessary to shift cleavage.
FIGS. 78A and B shows a synthetic oligonucleotide which was designed to fold upon itself which consists of the following sequence: 5′-GTTCTCTGCTCTCTGGTCGCTGTCTCGCTTGTGAAACAAGCGAGACAGCGTGGTCTCTCG-3′ (SEQ ID NO:29). This oligonucleotide is referred to as the “S-60 Hairpin.” The 15 basepair hairpin formed by this oligonucleotide is further stabilized by a “tri-loop” sequence in the loop end (i.e., three nucleotides form the loop portion of the hairpin) [Hiraro, I. et al. (1994) Nucleic Acids Res. 22(4): 576]. FIG. 78B shows the sequence of the P-15 oligonucleotide (SEQ ID NO:30) and the location of the region of complementarity shared by the P-15 and S-60 hairpin oligonucleotides. In addition to the P-15 oligonucleotide shown, cleavage was also tested in the presence of the P-14 oligonucleotide (SEQ ID NO:108) (P-14 is one base shorter on the 3′ end as compared to P-15), the P-14 with an a basic sugar (P-14d; SEQ ID NO:109) and the P14 with an a basic sugar with a 3′ phosphate (P-14dp; SEQ ID NO:110). A P-15 oligo with a 3′ phosphate, P-15p (SEQ ID NO:111) was also examined. The black arrows shown in FIG. 78 indicate the sites of cleavage of the S-60 hairpin in the absence (top structure; A) or presence (bottom structure; B) of the P-15 oligonucleotide.
The S-60 hairpin molecule was labeled on its 5′ end with fluorescein for subsequent detection. The S-60 hairpin was incubated in the presence of a thermostable 5′ nuclease in the presence or the absence of the P-15 oligonucleotide. The presence of the full duplex which can be formed by the S-60 hairpin is demonstrated by cleavage with the Cleavase® BN 5′ nuclease, in a primer-independent fashion (i.e., in the absence of the P-15 oligonucleotide). The release of 18 and 19-nucleotide fragments from the 5′ end of the S-60 hairpin molecule showed that the cleavage occurred near the junction between the single and double stranded regions when nothing is hybridized to the 3′ arm of the S-60 hairpin (FIG. 27, lane 2).
The reactions shown in FIG. 78C were conducted in 10 μl 1× CFLP buffer with 1 mM MnCl2 and 50 mM K-Glutamate, in the presence of 0.02 μM S-60, 0.5 μM InvaderTM oligonucleotide and 0.01 ng per μl Cleavase® BN nuclease. Reactions were incubated at 40° C. for 5 minutes and stopped by the addition of 8 μl of stop buffer (95% formamide, 20 mM EDTA, 0.02% methyl violet). Samples were heated to 75° C. for 2 min immediately before electrophoresis through a 15% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. Gels were then analyzed with a FMBIO-100 Image Analyzer (Hitachi) equipped with 505 nm filter. The resulting image is shown in FIG. 78C.
In FIG. 78C lane 1 contains products from the no enzyme control; lane 2 contains products from a reaction run in the absence of an InvaderTM oligo; lanes 3-6 contain products from reactions run the presence of the P-14d, P-14dp, P-15 and P-15p InvaderTM oligos, respectively.
From the data shown in FIG. 78C, it can be seen that the use of the P-15 InvaderTM oligonucleotide produces a shift in the cleavage site, while the P14 InvaderTM oligonucleotide with either a ribose (P14d) or a phosphorylated ribose (P14dp) did not This indicates that the 15th residue of the InvaderTM oligonucleotide must have the base group attached to promote the shift in cleavage. Interestingly, the addition of phosphate to the 3′ end of the P15 oligonucleotide apparently reversed the shifting of cleavage site. The cleavage in this lane may in fact be cleavage in the absence of an InvaderTM oligonucleotide as is seen in lane 2. In experiments with 5′ dye-labeled InvaderTM oligonucleotides with 3′ phosphate groups these oligonucleotides have been severely retarded in gel migration, suggesting that either the enzyme or another constituent of the reaction (e.g., BSA) is able to bind the 3′ phosphate irrespective of the rest of the cleavage structure. If the InvaderTM oligonucleotides are indeed being sequestered away from the cleavage structure, the resulting cleavage of the S-60 hairpin would occur in a “primer-independent′ fashion, and would thus not be shifted.
In addition to the study cited above, the effects of other substituents on the 3′ ends of the InvaderTM oligonucleotides were investigated in the presence of several different enzymes, and in the presence of either Mn++ or Mg++. The effects of these 3′ end modifications on the generation of cleaved product are summarized in the following table. All of modifications were made during standard oligonucleotide synthesis by the use of controlled pore glass (CPG) synthesis columns with the listed chemical moiety provided on the support as the synthesis starting residue. All of these CPG materials were obtained from Glen Research Corp. (Sterling, Va.).
FIG. 79 provides the structures for the 3′ end substituents used in these experiments.







TABLE 4







Modification Studies At 3′ End Of Invader Oligo










Extension By
Effect on Invader Rxn. (As Invader)


3′-End Modification
Terminal Transferase
Enzyme:Condition - Effect





3′ phosphate
no
A:5 - inhibits reaction, no detectable activity


Glen part # 20-2900-42


3′ acridine
yes, poorly
A:5 - decrease in activity, <10%


Glen part # 20-2973-42

B:5 - decrease in activity, <10%




B:4 - decrease in activity, <10%




C:1 - decrease in activity, <10%




C:2 - decrease in activity, ~20%




C:4 - decrease in activity, ~50%




C:3 - decrease in activity, <5%


3′ carboxylate
no
A:1 - decrease in activity, ~50%


Glen part # 20-4090-42

activity shift in cleavage site




C:3 - reduces rate, <10% activity


3′ nitropyrole
yes
A:5 - increase in activity, ~2×


Glen part # 20-2143-42


3′ nitroindole
yes
A:5 - decrease in activity, ~33% activity


Glen part # 20-2144-42


3′ arabinose
yes
A:5 - decrease in activity, ~50% activity


Glen part # 10-4010-90


3′ dideoxyUTP-
no
A:5 - decrease in activity, ~40% activity


fluorescein


3′-3′ linkage
no
A:1 - equivalent cleavage


Glen part # 20-0002-01

activity shift in cleavage site




C:3 - decrease in activity, ~25% activity


3′ glyceryl
yes, very poorly
C:3 - decrease in activity, ~30% activity


Glen part # 20-2902-42

loss of specificity of cleavage (2 sites)


3′ amino modifier C7
yes
C:3 - decrease in activity, ~30%


Glen part # 20-2957-42

activity loss of specificity, multiple sites


3′ deoxy, 2′ OH
yes, very poorly
A:5 - decrease in activity, <20% activity


Glen part # 20-2104-42

B:5 - decrease in activity, <20% activity




B:3 - decrease in activity, <20% activity




C:1 - equivalent activity




C:2 - equivalent activity




C:4 - ? increase in activity




C:3 - decrease in activity, ~40% activity





Enzymes:


A) Cleavase ® DV nuclease


B) Cleavase ® BN nuclease


C) Pfu FEN-1


Condition:


1) 4 mM MnCl2, 150 mM LiC1


2) 4 mM MnCl2, 50 mM KC1


3) 7.5 mM MgCl2, no monovalent


4) 4 mM MgCl2, 50 mM KC1


5) 10 mM MgOAc, 50 mM KC1






It can be seen from these data that many different modifications can be used on the 3′ end of the InvaderTM oligonucleotide without detriment. In various embodiments of the present invention, such 3′ end modifications may be used to block, facilitate, or otherwise alter the hybridization characteristics of the InvaderTM oligonucleotide, (e.g., to increase discrimination against mismatches, or to increase tolerance of mismatches, or to tighten the association between the InvaderTM oligonucleotide and the target nucleic acid). Some substituents may be used to alter the behavior of the enzyme in recognizing and cleaving within the assembled complex.
Altered 3′ ends may also be used to prevent extension of the InvaderTM oligonucleotide by either template-dependent or template-independent nucleic acid polymerases. The use of otherwise unmodified dideoxynucleotides (i.e., without attached dyes or other moieties) are a particularly preferred means of blocking extension of InvaderTM oligonucleotides, because they do not decrease cleavage activity, and they are absolutely unextendable.
EXAMPLE 36
Effect of Probe Concentration, Temperature and a Stacker Oligonucleotide on the Cleavage of Miniprobes by InvaderTM-Directed Cleavage
The stacker oligonucleotides employed to form cleavage structures may serve two purposes in the detection of a nucleic acid target using a miniprobe. The stacker oligonucleotide may help stabilize the interaction of the miniprobe with the target nucleic acid, leading to greater accumulation of cleaved probe. In addition, the presence of this oligo in the complex elongates the duplex downstream of the cleavage site, which may enhance the cleavage activity of some of the enzymes of the present invention. An example of different preferences for the length of this duplex by different structure-specific nucleases is seen in the comparison of the Cleavase® BN nuclease and the Mja FEN-1 nuclease cleavage of 8 bp and 12 bp duplex regions in FIG. 65. Increased affinity of the enzyme for the cleavage structure also results in increased accumulation of cleaved probe during reactions done for a set amount of time.
The amount of miniprobe binding to the target is also affected by the concentration of the miniprobe in the reaction mixture. Even when a miniprobe is only marginally likely to hybridize (e.g., when the reaction is performed at temperatures in excess of the expected melting temperature of the probe/target duplex), the amount of probe on the target at any given time can be increased by using high concentrations of the miniprobe.
The need for a stacker oligonucleotide to enhance cleavage of the miniprobe was examined at both low and high probe concentrations. The reactions were carried out in 10 μl of 10 mM HEPES (pH 7.2), 250 mM KGIu, 4 mM MnCl2, containing 100 nM of both the invading (oligo 135; SEQ ID NO:112) and stacking oligonucleotides (oligo 147; SEQ ID NO:113) and 100 pM ssM13 DNA. The reactions were overlayed with mineral oil, heated to 90° C. for 15 sec then brought to the reaction temperature. Reactions were performed at 35°, 40°, 45°, 50°, 55°, 60°, and 65° C. The cleavage reactions were initiated by the addition of 1 μl of 100 ng/μl Pfu FEN-1 and 1 μl of varying concentrations of Cy-3 labeled 142 miniprobe oligonucleotide (SEQ ID NO:114). Reactions were allowed to proceed for 1 hour and stopped by the addition of 10 μl formaldehyde. One fourth of the total volume of each reaction was loaded onto 20% non-denaturing polyacrylamide gels which were electrophoresed in the reverse direction. Gels were visualized using an Hitachi FMBIO-100 fluorescent scanner using a 585 nm filter. The fluorescence in each product band was measured and the graph shown in FIG. 80 was created using a Microsoft Excel spreadsheet.
The data summarized in FIG. 80 showed that the concentration of the miniprobe had a significant effect on the final measure of product, showing dramatic increases as the concentration was raised. Increases in the concentration of the miniprobe also shifted the optimum reaction temperature upward. It is known in the art that the concentration of the complementary strands in a hybridization will affect the apparent Tm of the duplex formed between them. More significantly to the methods and compositions of the present invention is the fact that the presence of the stacker oligonucleotide has a profound influence on the cleavage rate of the miniprobe at all probe concentrations. At each of the probe concentrations the presence of the stacker as much as doubled the signal from the cleavage product. This demonstrated the utility of using the stacker oligonucleotide in combination with the miniprobes described herein.
EXAMPLE 37
The Presence of a Mismatch in the InvaderTM Oligonucleotide Decreases the Cleavage Activity of the Cleavase® A/G Nuclease
In any nucleic acid detection assay it is of additional benefit if the assay can be made to sensitively detect minor differences between related nucleic acids. In the following experiment, model cleavage substrates were used that were identical except for the presence or absence of a mismatch near the 3′ end of the InvaderTM oligonucleotide when hybridized to the model target nucleic acid. The effect of a mismatch in this region on the accumulation of cleaved probe was then assessed.
To demonstrate the effect of the presence of a mismatch in the InvaderTM oligonucleotide on the ability of the Cleavase® A/G nuclease to cleave the probe oligonucleotide in an InvaderTM assay the following experiment was conducted. Cleavage of the test oligonucleotide IT-2 (SEQ ID NO:115) in the presence of InvaderTM oligonucleotides IT-1 (SEQ ID NO:116) and IT-1A4 (SEQ ID NO:117). Oligonucleotide IT-1 is fully complementary to the 3′ arm of IT-2, whereas oligonucleotide IT-1A4 has a T→A substitution at position 4 from the 3′ end that results in an A/A mismatch in the InvaderTM-target duplex. Both the matched and mismatched InvaderTM oligonucleotides would be expected to hybridize at the temperature at which the following reaction was performed. FIG. 81 provides a schematic showing IT-1 annealed to the folded IT-2 structure and showing IT-1A4 annealed to the folded IT-2 structure.
The reactions were conducted as follows. Test oligonucleotide IT-2 (0.1 μM), labeled at the 5′ end with fluorescein (Integrated DNA Technologies), was incubated with 0.26 ng/μl Cleavase® AG in 10 μl of CFLP® buffer with 4 mM MgCl2, in the presence of 1 μM IT-1 or IT-1A4 at 40° C. for 10 min; a no enzyme control was also run. Samples were overlaid with 15 μl Chill-Out® liquid wax to prevent evaporation. Reactions were stopped by addition of 4 μl stop buffer (95% formamide, 20 mM EDTA, 0.02% methyl violet). The cleavage products were separated on a 20% denaturing polyacrylamide gel and analyzed with the FMBIO-100 Image Analyzer (Hitachi) equipped with 505 nm filter. The resulting image is shown in FIG. 82.
In FIG. 82, lane 1 contains reaction products from the no enzyme control and shows the migration of the uncut IT-2 oligo; lanes 2-4 contain products from reactions containing no InvaderTM oligo, the IT-1 InvaderTM oligo and the IT-1A4 InvaderTM oligo, respectively.
These data show that cleavage is markedly reduced by the presence of the mismatch, even under conditions in which the mismatch would not be expected to disrupt hybridization. This demonstrates that the invader oligonucleotide binding region is one of the regions within the complex in which can be used for mismatch detection, as revealed by a drop in the cleavage rate.
EXAMPLE 38
Comparison of the Activity of the Pfu FEN-1 and Mja FEN-1 Nucleases in the InvaderTM Reaction
To compare the activity of the Pfu FEN-1 and the Mja FEN-1 nucleases in InvaderTM reaction the following experiment was performed. A test oligonucleotide IT3 (SEQ ID NO: 118) that forms an Invader-Target hairpin structure and probe oligonucleotide PR1 (SEQ ID NO:119) labeled at the 5′ end with fluorescein (Integrated DNA Technologies) were employed in InvaderTM assays using either the Pfu FEN-1 or the Mja FEN-1 nucleases.
The assays were conducted as follows. Pfu FEN-1 (13 ng/μl) and Mja FEN-1 (10 ng/μl) (prepared as described in Ex. 28) were incubated with the IT3 (0.1 nM) and PR1 (2 and 5 μM) oligonucleotides in 10 μL CFLP® buffer, 4 mM MgCl2, 20 mg/ml tRNA at 55° C. for 41 min. Samples were overlaid with 15 μl Chill-Out® evaporation barrier to prevent evaporation. Reactions were stopped by addition of 70 μl stop buffer (95% formamide, 20 mM EDTA, 0.02% methyl violet). Reaction products (1 μl) were separated on a 20% denaturing polyacrylamide gel, visualized using a fluroimager and the bands corresponding to the probe and the product were quantitiated. The resulting image is shown in FIG. 83. In FIG. 83, the turnover rate per target per minute is shown below the image for each nuclease at each concentration of probe and target tested.
It was demonstrated in Example 32 that the use of the Pfu FEN-1 structure-specific nuclease in the InvaderTM-directed cleavage reaction resulted in a faster rate of product accumulation than did the use of the Cleavase® A/G. The data presented here demonstrates that the use of Mja FEN-1 nuclease with the fluorescein labeled probe further increases the amount of product generated by an average of about 50%, demonstrating that, in addition to the Pfu FEN-1 nuclease, the Mja FEN-1 nuclease is a preferred structure-specific nuclease for the detection of nucleic acid targets by the method of the present invention.
EXAMPLE 39
Detection of RNA Target Nucleic Acids Using Miniprobe and Stacker Oligonucleotides
In addition to the detection of the M13 DNA target material described above, a miniprobe/stacker system was designed to detect the HCV-derived RNA sequences described in Example 19. A probe of intermediate length, either a long mid-range or a short standard probe, was also tested. The miniprobe used (oligo 42-168-1) has the sequence: 5′-TET-CCGGTCGTCCTGG-3′ (SEQ ID NO: 120), the stacker oligonucleotide used (oligo 32-085) with this miniprobe has the sequence: 5′-CAATTCCGGTGTACTACCGGTTCC-3′ (SEQ ID NO:121). The slightly longer probe, used without a stacker (oligo 42-088), has the sequence: 5′-TET-CCGGTCGTCCTGGCAA-3′ (SEQ ID NO:122). The InvaderTM oligonucleotide used with both probes has the sequence: 5′-GTTTATCCAAGAAAGGACCCGGTC-3′ (SEQ ID NO:47). The reactions included 50 fmole of target RNA, 10 pmole of the InvaderTM oligonucleotide and 5 pmole of the miniprobe oligonucleotide in 10 μl of buffer containing 10 mM MES, pH 6.5 with 150 mM LiCl, 4 mM MnCl2, 0.05% each Tween-20 and NP-40, and 39 units of RNAsin (Promega). When used, 10 pmoles of the stacker oligonucleotide was added. These components were combined, overlaid with Chillout® evaporation barrier, and warmed to 50° C.; the reactions were started by the addition of 5 polymerase units of DNAPTth, to a final reaction volume of 10 μl. After 30 minutes at 50° C., reactions were stopped by the addition of 8 μl of 95% formamide, 10 mM EDTA and 0.02% methyl violet. The samples were heated to 90° C. for 1 minute and 2.5 μl of each of these reactions were resolved by electrophoresis through a 20% denaturing polyacrylarnide (19:1 cross link) with 7M urea in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA, and the labeled reaction products were visualized using the FMBIO-100 Image Analyzer (Hitachi). The resulting image is shown in FIG. 84.
In FIG. 84, lanes 1 and 2 show the products of reactions containing the HCV InvaderTM oligonucleotide and the longer probe (oligo 42-088), without and with the target RNA present, respectively. Lanes 3, 4, and 5 show the products of reactions containing the InvaderTM oligonucleotide and the shorter probe (oligo 42-168-1). Lane 3 is a control reaction without target RNA present, while lanes 4 and 5 have the target, but are without or with the stacker oligonucleotide, respectively.
Under these conditions the slightly longer (16 nt) probe oligonucleotide was cleaved quite easily without the help of a stacker oligonucleotide. In contrast, the shorter probe (13 nt) required the presence of the stacker oligonucleotide to produce detectable levels of cleavage. These data show that the miniprobe system of target detection by InvaderTM-directed cleavage is equally applicable to the detection of RNA and DNA targets. In addition, the comparison of the cleavage performance of longer and shorter probes in the absence of a stacker oligonucleotide give one example of the distinction between the performance of the miniprobe/stacker system and the performance of the mid-range and long probes in the detection of nucleic acid targets.
EXAMPLE 40
Effect of an Unpaired 3′ Tail on Transcription from a Complete (Un-Nicked) Promoter
In designing the method of transcription-based visualization of the products of InvaderTM-directed cleavage, it was first necessary to assess the effect of a 3′ tail on the efficiency of transcription from a full length promoter. The duplexes tested in this example are shown at the bottom of FIG. 93, and are shown schematically in FIGS. 85A-C.
Transcription reactions were performed using the MEGAshortscriptTM system from Ambion, Inc. (Austin, Tex.), in accordance with the manufacturer's instructions with the exception that a fluorescein labeled ribonucleotide was added. Each DNA sample was assembled in 4 μl of RNAse-free dH2O. Reactions 1-3 each contained 10 pmole of the copy template oligo 150 (SEQ ID NO:123); reaction 2 contained 10 pmole of the promoter oligo 151 (SEQ ID NO:124); sample 3 contained 10 pmole of the 3′ tailed promoter oligo 073-065 (SEQ ID NO:125); sample 4 had no added DNA. To each sample, 6 μl of a solution containing 1 μl of 10× Transcription Buffer, 7.5 mM each rNTP, 0.125 mM fluorescein-12-UTP (Boehringer) and 1 μl T7 MEGAshortscriptTM Enzyme Mix was added. The samples were then incubated at 37° C. for 1 hour. One microliter of RNase-free DNase 1 (2 U/μl) was added to each sample and the samples were incubated an additional 15 minutes at 37° C. The reactions were then stopped by the addition of 10 μl of a solution of 95% formamide, 5 mM Na2EDTA, with loading dyes. All samples were heated to 95° C. for 2 minutes and 4 μl of each sample was resolved by electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked) with 7M urea, in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA. The gel was analyzed with a FMBIO II fluorescence image analyzer, and the resulting image is shown in FIG. 93. The RNA produced by successful transcription appears near the middle of the panel, as indicated (“RNA”).
Examination of the products of transcription shown in lanes 2 and 3 show that the presence of the 3′ tail on the full-length promoter has an adverse affect on the efficiency of transcription, but does not shut it off completely. Because the objective of the transcription-based visualization assays of the present invention is to discriminate between uncleaved probe and the shorter products of the invasive cleavage assay (cut probe), these data indicate that production of a full-length promoter in the cleavage reaction would be difficult to resolve from the background created by transcription from promoters containing the uncleaved probe if no other oligonucleotides were included in the assay. Means of suppressing transcription from such a branched promoter are discussed in the Description of the Invention and discussed below in Ex. 43.
EXAMPLE 41
Examination of the Influence of the Position of the Nick on the Efficiency of Transcription from Partial and Complete Composite Bacteriophage T7 Promoters
In the Description of the Invention, the procedure for testing prospective promoter pieces for suitability in an invasive cleavage-linked assay is described. One aspect of the test is to examine the effect a chosen nick site has on the efficiency of transcription from the final composite promoter. In addition, the individual pieces of nicked promoter are tested for transcription activity in the presence of the full-length un-nicked strand. In this experiment, a comparison on these points is made between a composite promoter having a nick in the non-template strand between nucleotides −11 and −10 relative to the initiation site (+1), and a promoter having a nick on the same strand, but positioned between nucleotides −8 and −7. The figure numbers for the schematic representations of the contents of each reaction are indicated below each lane (e.g., 85A=FIG. 85A). The site where the nick would be in a fully assembled composite promoter using the reaction oligonucleotides is also indicated below each lane (“−11/−10” and “−8/−7”).
Transcription reactions were performed using the MEGAshortscriptTM system, in accordance with the manufacturer's instructions, but with the exception that a fluorescein labeled ribonucleotide was added. Each DNA sample was assembled in 4 μl of RNAse-free dH2O. Reaction 1 had no added DNA. Reactions 2-9 each contained 10 pmole of the copy template oligo 150 (SEQ ID NO:123). Reactions 3 and 4 contained 10 pmole of the −11 “cut” probe (oligo 073-061-01; SEQ ID NO:127) or 20 pmole of the −10 partial promoter oligo 073-061-02 (SEQ ID NO:130), respectively, and reaction 5 contained both. Reactions 6 and 7 contained either the 10 pmole of the −8 “cut” probe (oligo 073-062-01; SEQ ID NO:126) or 20 pmoles of the −7 partial promoter oligo 073-062-02 (SEQ ID NO:129), respectively, and reaction 8 contained them both. Reaction 9 contained 10 pmole of the intact promoter oligo 151 (SEQ ID NO:124).
The transcription reactions were initiated, incubated, terminated and the reaction products were resolved and imaged as described in Ex. 40. The resulting image is shown in FIG. 92. The reaction numbers correspond to the lane numbers above the image. The RNA created by successful transcription appears in the upper third of the image. Comparison to the positive control reaction (rxn. 9) shows that the full-length RNA produced by each of the composite promoters is the same size as that produced in the control reaction, indicated that transcription initiated at the same site in each reaction.
In FIG. 92, lanes 3, 4, and 5 compare transcription from the two species of partially assembled promoters (see schematics in FIGS. 86A and B) and the fully assembled composite promoter (FIG. 88B) having a nick between nucleotides −11 and −10 relative to the start of transcription. It can be seen from these data that neither partial promoter (lanes 3 and 4) is able to support transcription of the copy template, but that the composite promoter (lane 5) with this nick site is strongly transcribed. Surprisingly, comparison to the control reaction (lane 9) shows that the presence of a nick at this site (−11/−10) actually enhances transcription. While not limiting the present invention to any particular mechanism, it is believed that the enhancement of transcription is a result of both suppressing the formation of the shorter abortive transcripts and by allowing greater accumulation of the full length product. This result is highly reproducible.
In FIG. 92, lanes 6, 7, and 8 compare transcription a similar set of partial and complete promoters in which the nick is shifted 3 residues closer to the transcription start site. Examination of lane 6 shows that the presence of 3 extra bases on the −8“cut” probe (compared to the −11 “cut” probe in lane 3) allow this partial promoter to initiate transcription. This indicates that the −8/−7 site would be a poor choice for use in this embodiment of the present invention.
This experiment demonstrates the process for determining the suitable placement of a nick within a promoter assembly to achieve the desired result. Similar tests can easily be designed for testing other nicks within the bacteriophage T7 promoter tested in this example, or for testing suitable nick placement in any desired phage, prokaryotic or eukaryotic promoter.
EXAMPLE 42
Detection of the Products of InvaderTM-Directed Cleavage through Transcription from a Composite Promoter
The examples described above indicate that a small oligonucleotide can be used to complete assembly of a composite T7 promoter, thereby enabling transcription from that promoter. Earlier examples demonstrate that the invasive cleavage reaction can be used release specific small oligonucleotide products from longer probe oligonucleotides. In this example it is demonstrated that these two observations can be combined, and that the products of the invasive cleavage reaction can be used to complete a promoter and enable subsequent transcription. The schematic representations of the composite promoters tested in this example are shown in FIG. 88.
Two invasive cleavage reactions were set up, one without (rxn. 1) and one with (rxn. 2) input target DNA. The reactions (1 and 2) comprised 10 mM MOPS (pH 7.5), 0.05% Tween-20, 0.05% NP-40 and 20 pmoles probe oligo 073-067-01 (SEQ ID NO:132) and 10 pmoles InvaderTM oligo 073-073-02 (SEQ ID NO:134) in a volume of 14 μl. Reaction 2 also included 100 fmoles M13mp18 ssDNA. The samples were placed at 60° C. and 6 μl of a solution containing 20 ng of Mja FEN-1 and 40 mM Mg2Cl were added to each sample to start the reactions. The samples were incubated at 60° C. for 30 minutes and stopped by the addition of 3 μl of 2.5M NaOAc, 83 mM Na2EDTA (pH 8.0). Each sample was transferred to a 1.5 ml microcentrifuge tube and then the DNAs were precipitated by the addition of 60 μl of chilled 100% ethanol, and were stored at −20° C. for 20 minutes. The pellets were collected by microcentrifugation, washed once with 80% ethanol to remove excess salt, then dried under vacuum. The product of this invasive cleavage reaction is a 12 nt oligonucleotide having the sequence: 5′-CGAAATTAATAC-3′ (SEQ ID NO:128), termed the −12 cut probe (same sequence as oligo 073-073-03).
For transcription, the dried samples were each dissolved in 4 μl of a solution containing 1 pmole copy template oligo 150 and 2 pmoles −11 partial promoter oligo 073-073-012 (SEQ ID NO:131). Control samples 3 and 4 each contained 1 pmole of the copy template oligo 150; sample 3 also contained 1 pmole probe oligo 073-067-01 (SEQ ID NO:132) and 2 pmoles −11 partial promoter oligo 073-073-012 (see structure 88A); sample 4 contained 1 pmole −12 “cut” probe oligo 073-073-03 (SEQ ID NO:128) and 2 pmoles −11 partial promoter oligo 073-073-012 (see structure 88B). These are the structures that would be expected to exist in the transcription reactions from the two invasive cleavage reactions described above.
The transcription reactions were initiated, incubated, terminated and the products were resolved and imaged as described in Ex. 40. The resulting image is shown in the right half of FIG. 89 (lanes 6-9). Samples 3 and 4 appear in lanes 6 and 7, respectively, and the reactions 1 and 2 from the invasive cleavage reaction products (indicated by the use of the lower case “i”), appear in lanes 8 and 9, respectively. The number of the Fig. showing the schematic representation of the expected promoter structure in each reaction is indicated above each lane, and the placement of the nick is also indicated. The uppercase letters indicate which structure in the particular figure to examine for each reaction. The lowercase “i” above lanes 8 and 9 indicate that these transcriptions were derived from actual invasive cleavage reactions. These products are compared to the RNA produced in the control reaction in lane 5, the procedure for which is described in Ex. 44. The RNA created by successful transcription appears in the upper third of the panel (indicated by “RNA”).
The reaction shown in lane 6 shows no transcription. This demonstrates that a nick between nucleotides −12 and −11 in the on-template strand of the T7 promoter eliminates transcription if the promoter is assembled from uncut probe such as the 3′ end of the probe forms a branch within the promoter sequence. This is in contrast tot he results seen with the −11/−10 nick examined below. Further, the transcript apparent in lane 7 shows that an unbranched promoter with a nick at the same site (−12/−11) produces the correct RNA, with few abortive initiation products (see lanes 2 and 5 of FIG. 89, described in Ex. 44). The reactions in lanes 8 and 9 demonstrate that the same effect is observed when the invasive cleavage reaction is the sole source of the upstream piece (−12 cut probe) of the T7 promoter. It is worthy of note that the promoter that is transcribed in lane 8 is made complete by the presence of 1 pmole of a synthetic “cut” probe oligo, without any uncut probe in the mixture, while the promoter that is transcribed in lane 9 is completed by the product of an invasive cleavage reaction that had only 100 fmole of target DNA in it. This reaction also included the residual uncut probe (up to approx. 10 pmoles), which may compete for binding at the same site. Nonetheless, the transcriptions from the invasive cleavage reaction products are only slightly reduced in efficiency, and are just as free of background as is the “no target” sample (lane 8). This example clearly demonstrates that the cleavage products from the invasive cleavage reaction can be used in combination with a partial promoter oligo to promote the production of RNA, without background transcription generated by the presence of the uncut probe. This RNA product is clearly dependent on the presence of the target material in the invasive cleavage reaction.
EXAMPLE 43
Shutting Down Transcription from a “Leaky” Branched T7 Composite Promoter through the Use of a Downstream Partial Promoter Oligonucleotide Having a 5′ Tail
The previous example demonstrated that placement of a nick in the non-template strand of a bacteriophage T7 promoter between the −12 and −11 nucleotides, relative to the transcription start site, prevents transcription of the branched promoter while allowing transcription when the composite promoter is assembled using the cut probe. When the nick is placed in other locations in the T7 promoter, transcription may be initiated from either promoter, although it is usually less efficient from the branched promoter. This example demonstrates that the addition of a 5′ tail that can base pair to the uncut probe (FIG. 90A) to the downstream partial promoter piece effectively blocks transcription from that promoter, but does not prevent transcription when a cut probe completes the promoter (FIG. 90B).
Two invasive cleavage reactions were set up, one without (rxn. 7) and one with (rxn. 8) input target DNA. The reactions (7 and 8) comprised 10 mM MOPS (pH 7.5), 0.05% Tween-20, 0.05% NP-40 and 20 pmoles probe oligo 073-067-01 (SEQ ID NO:132) and 10 pmoles InvaderTM oligo 073-067-02 (SEQ ID NO:133) in a volume of 14 μl. Reaction 8 also included 100 fmoles M13mp18 ssDNA. The samples were placed at 60° C. and 6 μl of a solution containing 20 ng of Mja FEN-1 and 40 mM Mg2Cl were added to each sample to start the reactions. The samples were incubated at 60° C. for 30 minutes and then stopped by the addition of 3 μl of 2.5M NaOAc, 83 mM Na2EDTA (pH.8.0). Each sample was transferred to a 1.5 ml microcentrifuge tube and the DNAs were precipitated, washed and dried as described in Ex. 42. The product of this invasive cleavage reaction is 13 nt oligonucleotide sequence, 5′-CGAAATTAATACG-3′ (SEQ ID NO:127), termed the −11 cut probe (same sequence as oligo 073-061-01 which is referred to as the −11 “cut” probe to indicate it was not generated in an invasive cleavage reaction).
In the transcription reactions, all of the DNAs were dissolved in 4 μl of RNase-free dH2O. Sample 1 had no added DNA, samples 2-8 contained 1 pmole of the copy template oligo 150 (SEQ ID NO:123). In addition, sample 3 contained 1 pmole of −11 “cut” probe oligo 073-061-01 (SEQ ID NO:127) and 2 pmoles of −10 partial promoter oligo 073-061-02 (SEQ ID NO:130), sample 4 contained 1 pmole of probe oligo 073-067-01 and 2 pmoles of −10 partial promoter oligo 073-061-02. Control sample 5 contained 1 pmole of probe oligo 073-067-01 and 2 pmoles of partial promoter w/5′ tail oligo 073-074 (5′-TACTGACTCACTATAGGGTCTTCTATGGAGGTC-3′ (SEQ ID NO:146) (see structure in FIG. 90A) and sample 6 contained 1 pmole of −11 “cut” probe oligo 073-061-01 and 2 pmoles of partial promoter w/5′ tail oligo 073-074 (see structure in FIG. 90B). These are the structures (i.e., 90A and 90B) that would be expected to exist in the transcription reactions from the two invasive cleavage reactions described above.
The dried samples 7 and 8 from the invasive cleavage (above) were each dissolved in 4 μl of dH2O containing 1 pmole copy template oligo 150 and 2 pmoles partial promoter w/5′ tail oligo 073-074. The transcription reactions were initiated, incubated, terminated and the reaction products were resolved and imaged as described in Ex. 40. The resulting image is shown in FIG. 91.
In FIG. 91 the lane numbers correspond to the sample numbers; the number of the figure showing the schematic representation of the expected promoter structure in each reaction is indicated above each lane (“88” and “90”), and the placement of the nick is also indicated (“−11/−10”). The upper-case letters indicate which structure in the particular figure to examine for each reaction. The lower case “i” above lanes 7 and 8 indicates that these transcriptions were derived from actual invasive cleavage reactions. The RNA created by successful transcription appears in the upper third of the panel, as indicated (“RNA”).
The control reactions in lanes 1 and 2, having either no DNA or having the only the copy template, produced no RNA as expected. The product in lane 4 demonstrates that the branched T7 promoter with a nick in the non-template strand between nucleotides −11 and −10 can support transcription, albeit not as efficiently as the un-branched promoter with the nick at the same site (lane 3). Examination of lane 5 shows that the use of a partial promoter oligonucleotide with a short 5′ tail that can basepair to the uncut probe as depicted in FIG. 90A, effectively suppresses this transcription but allows transcription when the probe does not have a 3′ tail (lane 6; schematic FIG. 90B). The reactions in lanes 7 and 8 demonstrate that the same effect as observed when the invasive cleavage reaction is the sole source of the upstream piece (−11 cut probe, SEQ ID NO:127) of the T7 promoter. It is worthy of note that the promoter that is transcribed in sample 6 is made complete by the presence of 1 pmole of a synthetic “cut probe”, without any uncut probe in the mixture, while the promoter that is transcribed in sample 8 is completed by the product of an invasive cleavage reaction that had only 100 fmole of target DNA in it. This reaction also included the residual uncut probe (up to approximately 19 pmoles), which may compete for binding at the same site. Nonetheless, the transcriptions from the invasive cleavage reaction products are just as strong and just as free of background in the “no target” samples.
This example clearly demonstrates that the cleavage products from the invasive cleavage reaction can be used in combination with a partial promoter oligonucleotide having a 5′ tail to promote the production of RNA, without background transcription generated by the uncut probe. This RNA product is clearly dependent on the presence of the target material in the invasive cleavage reaction.
EXAMPLE 44
Creation of a Complete Bacteriophage T7 Promoter by DNA Polymerase-Mediated Extension of a Cut Probe Comprising a Partial T7 Promoter
As demonstrated in the examples above, transcription cannot occur from the T7 promoter unless a complete promoter region is present. In the above examples, a complete promoter containing a nick in one strand was created by annealing a cut probe generated from an invasive cleavage reaction to a copy template which was annealed to a partial promoter oligo. An alternative means of creating a complete promoter in a manner dependent upon detection of a target sequence in an invasive cleavage reaction is to anneal the cut probe to a copy template devoid of a partial promoter oligo. The 3′-OH present at the end of the annealed cut probe is then extended by a DNA polymerase to create a complete and un-nicked promoter which is transcription-competent.
In this example the promoter was made complete through the use of primer extension, rather that by the co-hybridization of another oligonucleotide. The reaction steps are diagrammed schematically in FIG. 87. Two invasive cleavage reactions were set up, one without (rxn. 1) and one with (rxn. 2) input target DNA. The reactions (1 and 2) comprised 10 mM MOPS (pH 7.5), 0.05% Tween-20, 0.05% NP-40 and 20 pmoles probe oligo 073-067-01 (SEQ ID NO:132) and 10 pmoles InvaderTM oligo 073-073-02 (SEQ ID NO:134) in a volume of 14 μl. Reaction 2 also included 100 fmoles M13mp18 ssDNA. The samples were placed at 60° C. and 6 μl of a solution containing 20 ng of Mja FEN-1 and 40 mM Mg2Cl were added to each sample to start the reactions. The samples were incubated at 60° C. for 30 minutes and stopped by the addition of 3 μl of 2.5M NaOAc, 83 mM Na2EDTA (pH 8.0). Each sample was transferred to a 1.5 ml microcentrifuge tube and then the DNAs were precipitated, washed and dried as described in Ex. 42. The product of this invasive cleavage reaction is the 12 nt oligonucleotide sequence: 5′-CGAAATTAATAC-3′ (SEQ ID NO:128), termed the −12 cut probe (same sequence as oligo 073-073-03 which is referred to as the −12 “cut” probe to indicate it was not generated in an invasive cleavage reaction).
To allow extension of these products using a template-dependent DNA polymerase, a 20 μl solution containing 20 mM Tris-HCl (pH 8.5), 1.5 mM Mg2Cl, 50 mM KCl, 0.05% Tween-20, 0.05% NP-40, 25 μM each dNTP, 0.25 units Taq DNA polymerase (Boehringer) and 2 μM copy template oligo 150 (SEQ ID NO:123) was added to each of the dried cleavage samples. The samples were incubated at 30° C. for 1 hr. The primer extension reactions were stopped by the addition of 3 μl of 2.5M NaOAc with 83 mM Na2EDTA (pH 8.0)/sample. Each sample was transferred to a 1.5 ml microcentrifuge tube and the DNAs were precipitated, washed and dried as described in Ex. 42.
Samples 1 and 2 were then dissolved in 4 μl RNase-free dH2O. Samples 3, 4 and 5 are control reactions: sample 3 was 4 μl of RNase-free dH2O without added DNA, sample 4 contained 1 pmole of the copy template oligo 150 (SEQ ID NO:123) in 4 μl of RNase-free dH2O, and sample 5 contained 1 pmole of the same copy template and 1 pmole of the complete promoter oligo 151 (SEQ ID NO:124) in RNase-free dH2O.
Transcription reactions were performed using the MEGAshortscriptTM system, in accordance with the manufacturer's instructions, but with the addition of a fluorescein labeled ribonucleotide. To each sample, 6 μl of a solution containing 1 μl of 10× Transcription Buffer, 7.5 mM each rNTP, 0.125 mM fluorescein-12-UTP (Boehringer) and 1 μl T7 MEGAshortscriptTM Enzyme Mix was added. The samples were incubated at 37° C. for 1 hour. One μl of RNase-free DNase 1 (2 U/μl) was added to each sample and they were incubated an additional 15 minutes at 37° C. The reactions were stopped by the addition of 10 μl of a solution of 95% formamide, 5 mM NaEDTA, with loading dyes. All samples were heated to 95° C. for 2 minutes and four μl of each sample was resolved by electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked) with 7 M urea, in a buffer containing 45 mM Tris-Borate (pH 8.3), 1.4 mM EDTA. The results were imaged using the Molecular Dynamics Fluoroimager 595, with excitation at 488 nm and, emission detected at 530 nm.
The resulting image is shown in lanes 1 through 5 of FIG. 89; the lane numbers correspond to the sample numbers. The figure numbers corresponding to the schematic representations of the promoters transcribed in each reaction as indicated above the lanes. The RNA product from successful transcription appears in the upper third of the panel, as indicated (“RNA”). Unincorporated labeled nucleotide appears as a dense signal near the bottom (“NTPs”). Short transcription products caused by aborted initiation events [Milligan and Uhlenbeck (1989) Methods Enzymol. 180:51] appear as bands just above the free nucleotide in the lanes showing active transcription (i.e., lanes 2 and 5).
It can clearly be seen from the data in lanes 1 and 2 that the transcription is dependent on the presence of the target material in the invasive cleavage reaction. It is shown elsewhere (see lane 3, FIG. 92) that the product of the cleavage reaction is not in itself sufficient to allow transcription from the copy template. Thus, the action of the DNA polymerase in extending the hybridized cut probe across the promoter is a necessary step in enabling the transcription in this embodiment. These data clearly demonstrate that both template-dependent extension by DNA polymerase, and extension followed by transcription are suitable methods of visualizing the products of the invasive cleavage assay. As discussed in the Description of the Invention, the products of thermal breakdown that possess 3′ terminal phosphates would not be extended, and would thus be precluded from contributing to background transcription.
EXAMPLE 45
Test for the Dependence of an Enzyme on the Presence of an Upstream Oligonucleotide
When choosing a structure-specific nuclease for use in a sequential invasive cleavage reaction it is preferable that the enzyme have little ability to cleave a probe 1) in the absence of an upstream oligonucleotide, and 2) in the absence of overlap between the upstream oligonucleotide and the downstream labeled probe oligonucleotide. FIGS. 99a-e depicts the several structures that can be used to examine the activity of an enzyme that is confronted with each of these types of structures. The structure a (FIG. 99a) shows the alignment of a probe oligonucleotide with a target site on bacteriophage M13 DNA (M13 sequences shown in FIG. 99 are provided in SEQ ID NO:163) in the absence of an upstream oligonucleotide. Structure b (FIG. 99b) is provided with an upstream oligonucleotide that does not contain a region of overlap with the labeled probe (the label is indicated by the star). In structures c, d and e (FIGS. 99c-e) the upstream oligonucleotides have overlaps of 1, 3 or 5 nucleotides, respectively, with the downstream probe oligonucleotide and each of these structures represents a suitable invasive cleavage structure. The enzyme Pfu FEN-1 was tested for activity on each of these structures and all reactions were performed in duplicate.
Each reaction comprised 1 μM 5′ TET labeled probe oligonucleotide 89-15-1 (SEQ ID NO:152), 50 nM upstream oligonucleotide [either oligo 81-69-2 (SEQ ID NO:153), oligo 81-69-3 (SEQ ID NO:154), oligo 81-69-4 (SEQ ID NO:155), oligo 81-69-5 (SEQ ID NO:156), or no upstream oligonucleotide], 1 fmol M13 target DNA, 10 mg/ml tRNA and 10 ng of Pfu FEN-1 in 10 μl of 10 mM MOPS (pH 7.5), 7.5 mM MgCl2 with 0.05% each of Tween 20 and Nonidet P-40.
All of the components except the enzyme and the MgCl2 were assembled in a final volume of 8 μl and were overlaid with 10 μl of Chill-OutTM liquid wax. The samples were heated to the reaction temperature of 69° C. The reactions were started by the addition of the Pfu FEN-1 and MgCl2, in a 2 μl volume. After incubation at 69° C. for 30 minutes, the reactions were stopped with 10 μl of 95% formamide, 10 mM EDTA, 0.02% methyl violet. Samples were heated to 90° C. for 1 min immediately before electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. Gels were then analyzed with a FMBIO-100 Hitachi FMBIO fluorescence imager. The resulting image is displayed in FIG. 100.
In FIG. 100, lanes labeled “a” contain the products generated from reactions conducted without an upstream oligonucleotide (structure a), lanes labelled “b” contain an upstream oligonucleotide which does not invade the probe/target duplex (structure b). Lanes labelled “c”, “d” and “e” contain the products generated from reactions conducted using an upstream oligonucleotide that invades the probe/target duplex by 1, 3 or 5 bases, respectively. The size (in nucleotides) of the uncleaved probe and the cleavage products is indicated to the left of the image in FIG. 100.
As shown in FIG. 100, cleavage of the probe was not detectable when structures a and b were utilized. In contrast, cleavage products were generated when invasive cleavage structures were utilized (structures c-e). These data show that the Pfu FEN-1 enzyme requires an overlapping upstream oligonucleotide for specific cleavage of the probe.
Any enzyme may be examined for its suitability for use in a sequential invasive cleavage reaction by examining the ability of the test enzyme to cleave structures a-e (it is understood by those in the art that the specific oligonucleotide sequences shown in FIGS. 99a-e need not be employed in the test reactions; these structures are merely illustrative of suitable test structures). Desirable enzymes display little or no cleavage of structures a and b and display specific cleavage of structures c-e (i.e., they generate cleavage products of the size expected from the degree of overlap between the two oligonucleotides employed to form the invasive cleavage structure).
EXAMPLE 46
Use of the Products of a First Invasive Cleavage Reaction to Enable a Second Invasive Cleavage Reaction with a Net Gain in Sensitivity
As discussed in the Description of The Invention above, the detection sensitivity of the invasive cleavage reaction can be increased by the performing a second round of invasive cleavage using the products of the first reaction to complete the cleavage structure in the second reaction (shown schematically in FIG. 96). In this example the use of a probe which when cleaved in a first invasive cleavage reaction forms an integrated InvaderTM oligo and target molecule for use in a second invasive cleavage reaction is illustrated (shown schematically in FIG. 97).
A first probe was designed to contain some internal complementarity so that when cleaved in a first invasive cleavage reaction the product (“Cut Probe 1”) could form a target strand comprising an integral InvaderTM oligonucleotide, as depicted in FIG. 97. A second probe was provided in the reaction that would be cleaved at the intended site when hybridized to the newly formed target/invader (FIG. 97). To demonstrate the gain in signal due to the performance of sequential invasive cleavages, a standard invasive cleavage assay, as described above, was performed in parallel.
All reactions were performed in duplicate. Each standard (i.e., non-sequential) invasive cleavage reaction comprised 1 μM 5′ fluorescein-labeled probe oligo 073-182 (5′ F1-AGAAAGGAAGGGAAGAAAGCGAA-3′; SEQ ID NO:157), 10 nM upstream oligo 81-69-4 (5′-CTTGACGGGGAAAGCCGGCGAACGTGGCGA-3′; SEQ ID NO:155), 10 to 100 attomoles of M13 target DNA, 10 mg/ml tRNA and 10 ng of Pfu FEN-1 in 10 μl of 10 mM MOPS (pH 7.5), 8 mM MgCl2 with 0.05% each of Tween 20 and Nonidet P-40. All of the components except the enzyme and the MgCl2 were assembled in a volume of 7 μl and were overlaid with 10 μl of Chill-OutTM liquid wax. The samples were heated to the reaction temperature of 62° C. The reactions were started by the addition of the Pfu FEN-1 and MgCl2, in a 2 μl volume. After incubation at 62° C. for 30 minutes, the reactions were stopped with 10 μl of 95% formamide, 10 mM EDTA, 0.02% methyl violet.
Each sequential invasive cleavage reaction comprised 1 μM 5′ fluorescein-labeled oligonucleotide 073-191 (the first probe or “Probe 1”, 5′ F1-TGGAGGTCAAAACATCGATAAGTCGAAGAAAGGAAGGGAAGAAAT-3′; SEQ ID NO:158), 10 nM upstream oligonucleotide 81-69-4 (5′-CTTGACGGGGAAAGCCGGCGAACGTGGCGA-3′; SEQ ID NO:155), 1 μM of 5′ fluorescein labeled oligonucleotide 106-32 (the second probe or “Probe 2”, 5′ F1-TGTTTTGACCTCCA-3′; SEQ ID NO:159), 1 to 100 amol of M13 target DNA, 10 mg/ml tRNA and 10 ng of Pfu FEN-1 in 10 μl of 10 mM MOPS (pH 7.5), 8 mM MgCl2 with 0.05% each of Tween 20 and Nonidet P-40. All of the components except the enzyme and the MgCl2 were assembled in a volume of 8 μl and were overlaid with 10 μl of Chill-OutTM liquid wax. The samples were heated to the reaction temperature of 62° C. (this temperature is the optimum temperature for annealing of Probe 1 to the first target). The reactions were started by the addition of Pfu FEN-1 and MgCl2, in a 2 μl volume. After incubation at 62° C. for 15 minutes, the temperature was lowered to 58° C. (this temperature is the optimum temperature for annealing of Probe 2 to the second target) and the samples were incubated for another 15 min. Reactions were stopped by the addition of 10 μl of 95% formamide, 20 mM EDTA, 0.02% methyl violet.
Samples from both the standard and the sequential invasive cleavage reactions were heated to 90° C. for 1 min immediately before electrophoresis through a 20% denaturing acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. The gel was then analyzed with a Molecular Dynamics FluorImager 595. The resulting image is displayed in FIG. 101a. A graph showing measure of fluorescence intensity for each of the product bands is shown in FIG. 101b. 
In FIG. 101a, lanes 1-5 contain the products generated in standard invasive cleavage reactions that contained either no target (lane 1), 10 amol of target (lanes 2 and 3) or with 100 amol of target (lanes 4 and 5). The uncleaved probe is seen as a dark band in each lane about half way down the panel and the cleavage products appear as a smaller black band near the bottom of the panel, the position of the cleavage product is indicated by an arrow head to the left of FIG. 101a. The gray ladder of bands seen in lanes 1-5 is due to the thermal degradation of the probe as discussed above and is not related to the presence or absence of the target DNA. The remaining lanes display products generated in sequential invasive cleavage reactions that contained 1 amol of target (lanes 6 and 7), 10 amol of target (lanes 8 and 9) and 100 amol of target (lanes 10 and 11). The uncleaved first probe (Probe 1; labeled “1 uncut”) is seen near the top of the panel, while the cleaved first probe is indicated as “1: cut”. Similarly, the uncleaved and cleaved second probe are indicated as “2: uncut” and 2: cut,” respectively.
The graph shown in FIG. 101b compares the amount of product generated from the standard reaction (“Series 1”) to the amount of product generated from the second step of the sequential reaction (“Series 2”). The level of background fluorescence measured from a reaction that lacked target DNA was subtracted from each measurement. It can be seen from the table located below the graph that the signal from the standard invasive cleavage assay that contained 100 attomoles of target DNA was nearly identical to the signal from the sequential invasive cleavage assay in which 1 attomole of target was present, indicating that the inclusion of a second cleavage structure increases the sensitivity of the assay 100 to 200-fold. This boost in signal allows easy detection of target nucleic acids at the sub attomole level using the sequential invasive cleavage assay, while the standard assay, when performed using this enzyme for only 30 minutes, does not generate detectable product in the presence of 10 attomoles of target.
When the amount of target was decreased by 10 or 100 fold in the sequential invasive cleavage assay, the intensity of the signal was decreased by the same proportion. This indicates that the quantitative capability of the invasive cleavage assay is retained even when reactions are performed in series, thus providing a nucleic acid detection method that is both sensitive and quantitative.
While in this example the two probes used had different optimal hybridization temperatures (i.e., the temperature empiracally determined to give the greatest turnover rate in the given reaction conditions), the probes may also be selected (i.e., designed) to have the same optimal hybridization temperature so that a temperature shift during incubation is not necessary.
EXAMPLE 47
The Products of a Completed Sequential Invasive Cleavage Reaction cannot Cross Contaminate Subsequent Similar Reactions
As discussed in the Description of the Invention, the serial nature of the multiple invasive cleavage events that occur in the sequential invasive cleavage reaction, in contrast to the reciprocating nature of the polymerase chain reaction and similar doubling assays, means that the sequential invasive cleavage reaction is not subject to contamination by the products of like reactions because the products of the first cleavage reaction do not participate in the generation of new signal in the second cleavage reaction. If a large amount of a completed reaction were to be added to a newly assembled reaction, the background that would be produced would come from the amount of target that was also carried in, combined with the amount of already-cleaved probe that was carried in. In this example it is demonstrated that a very large portion of a primary reaction must be introduced into the secondary reaction to create significant signal.
A first or primary sequential invasive cleavage reaction was performed as described above using 100 amol of target DNA. A second set of 5 reactions were assembled as described in Ex. 46 with the exception that portions of the first reaction were introduced and no additional target DNA was included. These secondary reactions were initiated and incubated as described above, and included 0, 0.01, 0.1, 1, or 10% of the first reaction material. A control reaction including 100 amol of target was included in the second set also. The reactions were stopped, resolved by electrophoresis and visualized as described above, and the resulting image is displayed in FIG. 102. The primary probe, uncut second probe and the cut 2nd probe are indicated on the left as “1: cut”, 2: uncut” and 2: cut”, respectively.
In FIG. 102, lane 1 shows the results of the first reaction with the accumulated product at the bottom of the panel, and lane 2 show a 1:10 dilution of the same reaction, to demonstrate the level of signal that could be expected from that level of contamination, without further amplification. Lanes 3 through 7 show the results of the secondary cleavage reactions that contained 0, 10, 1, 0.1 or 0.01% of the first reaction material added as contaminant, respectively and lane 8 shows a control reaction that had 100 amol of target DNA added to verify the activity of the system in the secondary reaction. The signal level in lane 4 is as would be expected when 10% of the pre-cleaved material is transferred (as in lane 2) and 10% of the transferred target material from the lane 1 reaction is allowed to further amplify. At all levels of further dilution the signal is not readily distinguished from background. These data demonstrate that while a large-scale transfer from one reaction to another may be detectable, cross contamination by the minute quantities that would be expected from aerosol or from equipment contamination would not be easily mistaken for a false positive result. These data also demonstrate that when the products of one reaction are deliberately carried over into a fresh sample, these products do not participate in the new reaction, and thus do not affect the level of target-dependent signal that may be generated in that reaction.
EXAMPLE 48
Detection of Human Cytomegalovirus Viral DNA by Invasive Cleavage
The example demonstrates the ability of the invasive cleavage reaction to detect minute quantities of viral DNA in the presence of human genomic DNA. In this example, the probe and InvaderTM oligonucleotides were designed to target the 3104-3061 region of the major immediate early gene of human cytomegalovirus (HCMV) as shown in FIG. 103. In FIG. 103, the InvaderTM oligo (89-44; SEQ ID NO:160) and the fluorescein (F1)-labeled probe oligo (89-76; SEQ ID NO:161) are shown annealed along a region of the HCMV genome corresponding to nucleotides 3057-3110 of the viral DNA (SEQ ID NO:162). The probe used in this example is a poly-pyrimidine probe and as shown herein the use of a poly-pyrimidine probe reduces background signal generated by the thermal breakage of probe oligos.
The genomic viral DNA was purchased from Advanced Biotechnologies, Incorporated (Columbia, Md.). The DNA was estimated (but not certified) by personnel at Advanced Biotechnologies to be at a concentration of 170 amol (1×108 copies) per microliter. The reactions were performed in quadruplicate. Each reaction comprised 1 μM 5′ fluorescein labeled probe oligonucleotide 89-76 (SEQ ID NO:161), 100 nM InvaderTM oligonucleotide 89-44 (SEQ ID NO:160), 1 ng/ml human genomic DNA, and one of five concentrations of target HCMV DNA in the amounts indicated above each lane in FIG. 104, and 10 ng of Pfu FEN-1 in 10 μl of 10 mM MOPS (pH 7.5), 6 mM MgCl2 with 0.05% each of Tween 20 and Nonidet P-40. All of the components except the labeled probe, enzyme and MgCl2 were assembled in a final volume of 7 μl and were overlaid with 10 μl of Chill-OutTM liquid wax. The samples were heated to 95° C. for 5 min, then reduced to 62° C. The reactions were started by the addition of probe, Pfu FEN-1 and MgCl2, in a 3 μl volume. After incubation at 62° C. for 60 minutes, the reactions were stopped with 10 μl of 95% formamide, 10 mM EDTA, 0.02% methyl violet. Samples were heated to 90° C. for 1 min immediately before electrophoresis through a 20% acrylamide gel (19:1 cross-linked), with 7 M urea, in a buffer of 45 mM Tris-Borate, pH 8.3, 1.4 mM EDTA. Gels were then analyzed with a Molecular Dynamics FluorImager 595.
The resulting image is displayed in FIG. 104. The replicate reactions were run in groups of four lanes with the target HCMV DNA content of the reactions indicated above each set of lanes (0-170 amol). The uncleaved probe is seen in the upper third of the panel (“Uncut 89-76”) while the cleavage products are seen in the lower two-thirds of the panel (“Cut 89-76”). It can be seen that the intensity of the accumulated cleavage product is proportional to the amount of the target DNA in the reaction. Furthermore, it can be clearly seen in reactions that did not contain target DNA (“no target”) that the probe is not cleaved, even in a background of human genomic DNA. While 10 ng of human genomic DNA was included in each of the reactions shown in FIG. 104, inclusion of genomic DNA up to 200 ng has slight impact on the amount of product accumulated. The data did not suggest that 200 ng per 10 μl of reaction mixture represented the maximum amount of genomic DNA that could be tolerated without a significant reduction in signal accumulation. For reference, this amount of DNA exceeds what might be found in 0.2 ml of urine (a commonly tested amount for HCMV in neonates) and is equivalent to the amount that would be found in about 5 μl of whole blood.
These results demonstrate that the standard (i.e., non-sequential) invasive cleavage reaction is a sensitive, specific and reproducible means of detecting viral DNA. It can also be seen from these data that the use of a poly-pyrimidine probe reduces the background from thermal breakage of the probe, as discussed in Example 22. Detection of 1.7 amol of target is roughly equivalent to detection of 106 copies of the virus. This is equivalent to the number of viral genomes that might be found in 0.2 mls of urine from a congenitally infected neonate [102 to 106 genome equivalents per 0.2 mls; Stagno et al. (1975) J. Infectious Disease 132:568]. Use of the sequential invasive cleavage assay would permit detection of even fewer viral DNA molecules, facilitating detection in blood (101 to 105 viral particles per ml; Pector et al. (1992) J. Clin. Microbiol. 30:2359) which carries a much larger amount of heterologous DNA.
From the above it is clear that the invention provides reagents and methods to permit the detection and characterization of nucleic acid sequences and variations in nucleic acid sequences. The InvaderTM-directed cleavage reaction and the sequential InvaderTM-directed cleavage reaction of the present invention provide ideal direct detection methods that combine the advantages of the direct detection assays (e.g., easy quantification and minimal risk of carry-over contamination) with the specificity provided by a dual or tri oligonucleotide hybridization assay.
All publications and patents mentioned in the above specification are herein incorporated by reference. Various modifications and variations of the described method and system of the invention will be apparent to those skilled in the art without departing from the scope and spirit of the invention. Although the invention has been described in connection with specific preferred embodiments, it should be understood that the invention as claimed should not be unduly limited to such specific embodiments. Indeed, various modifications of the described modes for carrying out the invention which are obvious to those skilled in molecular biology or related fields are intended to be within the scope of the following claims.

## Claims

1. A method of detecting a target polynucleotide which comprises the steps of:
a) contacting a target polynucleotide having a first portion and a second portion immediately contiguous to one another with:
i) an invader oligonucleotide, at least a part of which is capable of specifically hybridizing to the first portion of the target polynucleotide;
ii) a first probe oligonucleotide comprising a first region that is capable of specifically hybridizing to the second portion of the target polynucleotide and an unpaired region located adjacent to the first region; and
iii) a reagent that is capable of cleaving to release the unpaired region of the first probe oligonucleotide to produce a cleaved unpaired region, wherein said reagent comprises a 5′ nuclease;
producing said cleaved unpaired region by cleaving said first probe oligonucleotide, forming a cleavage structure comprising said cleaved unpaired region and a second probe oligonucleotide by hybridizing said cleaved unpaired region to said second probe oligonucleotide, or forming a cleavage structure comprising said cleaved unpaired region, said second probe oligonucleotide, and a target nucleic acid by hybridizing said cleaved unpaired region and said second probe oligonucleotide to said target nucleic acid, and generating a cleaved second probe by cleaving said second cleavage structure using the reagent;
b) detecting the accumulation of the cleaved second probe oligonucleotide; and
c) determining whether the cleaved second probe oligonucleotide accumulates exponentially over time, wherein said exponential accumulation of the cleaved second probe oligonucleotide over time is indicative of the presence of said target polynucleotide.

2. The method of claim 1 wherein said detecting the accumulation of the cleaved second probe oligonucleotide comprises detection of fluorescence or phosphorescence.

3. The method of claim 1, wherein said 5′ nuclease is a thermostable 5′ nuclease.

4. The method of claim 3, wherein said thermostable 5′ nuclease is a FEN-1 nuclease.

5. The method of claim 4, wherein said FEN-1 nuclease is a FEN-1 nuclease from an archaebacterial species.

6. The method of claim 5, wherein said FEN-1 nuclease is selected from the group consisting of Methanococcus jannaschii FEN-1 and Pyrococcus furiosus FEN-1.

7. A method for detecting the presence of a first target nucleic acid in a sample, comprising:
a) incubating a sample containing a first target nucleic acid with a first nucleic acid molecule and a second nucleic acid molecule and forming a first cleavage structure, said first cleavage structure comprising:
i) said first target nucleic acid comprising a first region and a second region, said second region upstream of and contiguous to said first region;
ii) said first nucleic acid molecule comprising a first portion that is completely complementary the second region of the first target nucleic acid;
iii) said second nucleic acid molecule comprising a 3′ portion and a 5′ portion, wherein said 5′ portion is completely complementary to said first region of said first target nucleic acid;
wherein said 5′ portion of said second nucleic acid molecule is annealed to said first region of said first target nucleic acid and wherein at least a portion of said first nucleic acid molecule is annealed to said second region of said first target nucleic acid, and wherein a 5′ portion of said first nucleic acid molecule is not annealed to said first target nucleic acid,
b) cleaving said first cleavage structure with a cleavage agent comprising a 5′ nuclease, generating a non-target cleavage product, and forming a second cleavage structure comprising:
i) said non-target cleavage product; and
ii) a probe oligonucleotide;


by hybridizing said non-target cleavage product to said probe oligonucleotide, or forming a second cleavage structure comprising:

i) said non-target cleavage product;
ii) said probe oligonucleotide; and
iii) a second target nucleic acid;


by hybridizing both said non-target cleavage product and said probe oligonucleotide to said second target nucleic acid, and
c) cleaving said second cleavage structure with said cleavage agent, generating a cleaved probe and detecting said cleaved probe at a plurality of time points, wherein said cleaved probe accumulates at an exponential rate over time, and wherein the accumulation of said cleaved probe at an exponential rate over time indicates the presence of said first target nucleic acid in said sample.

8. The method of claim 7, wherein said detecting said cleaved probe comprises detection of fluorescence.

9. The method of claim 7, wherein said detecting said cleaved probe comprises detection of mass.

10. The method of claim 7, wherein said detecting said cleaved probe comprises detection of fluorescence energy transfer.

11. The method of claim 7, wherein said detecting said cleaved probe comprises detection selected from the group consisting of detection of radioactivity, luminescence, phosphorescence, fluorescence polarization, and charge.

12. The method of claim 7, wherein said 5′ nuclease is a thermostable 5′ nuclease.

13. The method of claim 12, wherein said thermostable 5′ nuclease is a 5′ nuclease of a DNA polymerase.

14. The method of claim 13, wherein said DNA polymerase is Taq DNA polymerase.

15. The method of claim 7, wherein said 3′ portion of said second nucleic acid molecule consists of a single nucleotide.

16. The method of claim 15, wherein said single nucleotide is complementary to said first target nucleic acid.

17. The method of claim 7 further comprising providing a plurality of said first nucleic acid molecule such that said first nucleic acid molecule is in concentration excess compared to said first target nucleic acid.

18. The method of claim 7 further comprising providing a plurality of said second nucleic acid molecule such that said second nucleic acid molecule is in concentration excess compared to said first target nucleic acid.

19. The method of claim 7, wherein said first target nucleic acid and said second nucleic acid molecule form a duplex, and further comprising providing a plurality of said first nucleic acid molecule such that said first nucleic acid molecule is in concentration excess compared to said duplex.

20. The method of claim 19, wherein said non-target cleavage product is generated from said first nucleic acid molecule and is in concentration excess compared to said duplex.

21. The method of claim 12, wherein said thermostable 5′ nuclease is a FEN-1 nuclease.

22. The method of claim 21, wherein said FEN-1 nuclease is a FEN-1 nuclease from an archaebacterial species.

23. The method of claim 22, wherein said FEN-1 nuclease is selected from the group consisting of Methanococcus jannaschii FEN-1 and Pyrococcus furiosus FEN-1.