When we think of molecular diagnostic tests based on nucleic acid amplification methods (PCR or otherwise), several common amplicon detection methods come to mind. These include the previously covered topics of real-time fluorescent (probe- or binding dye-based); endpoint size resolved (such as agarose gel electrophoresis, or higher resolution capillary electrophoresis); or endpoint array hybridization (liquid-phase bead arrays or conventional 2D solid-phase printed microarrays). Another method, hybridizing probe-based luminescence detection, was briefly covered in February’s installment of The Primer. While those are the most common modalities for amplicon detection in today’s molecular diagnostics laboratory, a range of other methods do exist in some test systems and include some remarkably clever and simple approaches. In this month’s column we’ll consider two representative examples of these alternative technologies.
Magnesium pyrophosphate detection
Our first example is magnesium pyrophosphate detection. Recall that in extending a nascent DNA strand such as from an oligonucleotide primer annealed to its specific complementary target, DNA polymerases incorporate dNTPs as nucleotide monophosphates in the nascent strand. The remaining two phosphates of the “TP” (triphosphate) are released as a single pyrophosphate (“PPi”) moiety.
In a living cell, this is quickly broken down to two individual phosphates by the action of pyrophosphatase enzymes. This reaction is highly thermodynamically favorable, and it acts as a major energy source for DNA strand growth through thermodynamic coupling. (If it’s been a while since you thought of that term, recall that a reaction can be driven forward by a second reaction which uses up products of the first reaction).
In an in vitro reaction however, there’s no pyrophosphatase around unless it’s been added to the reaction, and so with each dNTP added to a nascent DNA, there’s an additional PPi released into solution. Recall also that DNA polymerases require the presence of a divalent cation (usually Mg++, less commonly Mn++) in their buffer for effective enzyme activity. This cation doesn’t bind to the enzyme per se; rather, its role is to bind to the triphosphate groups of each dNTP in a chelation binding. This binding is essential in maintaining the structure and reactivity of the dNTPs (and thus the reason why metal ion chelating agents can be effective inhibitors of PCR or other polymerase-based amplification methods).
Not surprisingly, pyrophosphate is also capable of chelating Mg++ ions on its own, and it does so as PPi levels build up in the course of significant dNTP conversion to new DNA through an amplification strategy. Of importance to us today is that this Mg-PPi complex is insoluble; that is, it forms microscopic particulate precipitates in the reaction as the PPi levels increase. A second factor is that these precipitate particles are extremely small and over the time scale of an hour or so, Brownian motion effectively keeps the Mg-PPi particulates in suspension. This results in a readily visible increase in turbidity of the reaction, which can be monitored by simple optics normally tuned to look through the reaction tube walls for decreasing transparency at a 650 nM wavelength.
This approach shares many of the advantages inherent in fluorescent-based real-time PCR: reaction monitoring can be done as the reaction proceeds; time to turbidity crossing a detection threshold correlates well with starting template levels across several logs of template concentration, allowing for accurate quantitative results against a standard; and the closed tube chemistry helps avoid dreaded amplicon contamination of the laboratory. Significant differences from conventional real-time PCR include the lack of need for additional reagents (such as fluorescent dyes or hybridizing probes, making this approach intrinsically cheaper), but also a lack of specificity. Any polymerase product(s), whether correct or spurious, will lead to signal accumulation in this approach. While the same holds true for intercalating dye-based real-time detection, in that format a post-amplification melt curve analysis is possible to evaluate amplification products; such an approach is not possible with Mg-PPi turbidity determination.
Primarily because of this, the method has not generally been adopted for use in conventional PCR applications where significant risk of non-target amplification is possible. It remains more suited for use in conjunction with non-PCR amplification methods with higher intrinsic specificity such as the loop-mediated isothermal DNA amplification (LAMP) approach. While space considerations and the complexities of LAMP precluded it from detailed coverage in our previous sections on non-PCR approaches, suffice it to say it employs four target specific primers with a total of six separate regions of target homology. This results in higher specificity of amplification than conventional PCR, and LAMP-based assays are for this reason commonly paired to Mg-PPi turbidity detection. The combination of an isothermal amplification and simple optical detection makes LAMP/Mg-PPi assay systems robust and economical, with relatively low reagent costs and simple instrumentation capable of both qualitative and quantitative nucleic acid assays. There are presently a number of commercially available clinical assays in this format.
Lateral flow detection systems
While Mg-PPi-based amplification detection is instrument-based, still simpler non-instrumental methods exist for detection of nucleic acid amplification products. Lateral flow detection systems (LFAs), in which a filter or membrane surface is spotted with a solution containing analyte and read for analyte presence through the appearance of colored bands, are familiar to laboratorians primarily in immunoassay applications. They can be applied for molecular tests as well. Although sometimes referred to as “dipstick” tests, LFAs and true dipstick or immunospot tests have mechanistic differences and can most readily be distinguished by considering where sample is applied compared to where bands are read. In a true dipstick test, the sample is applied directly to the reporting band or spot, while in an LFA the sample is applied at one end of a membrane strip separated by some distance to the reporter bands, and is drawn past the reporting bands by capillary action (thus, “lateral flow”). We’ll concern ourselves here just with true LFA devices, familiar not only to laboratorians but also to much of the general public through applications such as home pregnancy test kits. This widespread application speaks to the low complexity and ease of result interpretation of tests in this format.
In the simplest application of LFA for molecular tests, consider the example of a classical PCR reaction for a single target species, but with each of the amplifying primers covalently attached to an immunological tag or hapten—with different tags for the forward and reverse primers. These tags need to be small enough to not greatly impact the hybridization efficiency or kinetics of the primer they are linked to, and should not interfere with primer binding and extension by the system DNA polymerase (this latter requirement is often met by linking the tag to the primer 5′ end, thus placing it as far as possible from the 3′ end where extension will occur).
In addition to these requirements, the tag should be one for which there are purified highly specific, high-affinity antibodies available, usually monoclonal; or tags for which some similar specific, sensitive binding interaction is possible. Examples of tags meeting these criteria include fluorescein paired to anti-fluorescein antibodies, and biotin paired to streptavidin. For our example here, let’s assume those are the tags used, with fluorescein attached to the forward primer and biotin to the reverse primer. If we employ these primers in a conventional PCR and presence of the specific target species leads to successful amplification, at PCR endpoint we have a dual-labeled PCR product—fluorescein at one end, and biotin at the other. We apply our PCR reaction to a sample pad at one end of a disposable, single-use LFA test strip, adjacent to a “conjugate pad” and followed further along the strip by a test band “T,” a control band “C,” and finally at the other end of the strip, an absorbent matrix pad (Figure 1). Application of the PCR product mixture to the sample pad starts its wicking toward the absorbent pad by capillary action, and sample absorption by the absorbent pad maintains this slow sample flow through the sequential sections of the LFA test strip.
| Figure 1. Direction of wicking in lateral flow detection |
The sample first encounters the conjugate pad containing a dried-down detection reagent, which in our example could be collodial gold nanoparticles coated with the protein streptavidin. This protein has a high specific binding affinity for biotin, and so will enter solution and bind to biotin molecules which may be present either on complete PCR product molecules (if the PCR was successful), or unreacted biotinylated reverse primer alone (if the PCR was unsuccessful). This now gold nanoparticle-labeled mixture continues to flow down the strip until it reaches the T band. Here, anti-fluorescein antibodies have been bound to the strip, which allows the strip to trap all of the fluorescein-coupled forward primers flowing past—on their own (if the PCR was unsuccessful) or as part of a PCR product (if the PCR was successful). Note that in this latter case only, this also traps gold nanoparticle label at the T strip, via its streptavidin-biotin-reverse primer linkage. Significant accumulations of gold nanoparticles are directly visible to the naked eye, making a colored band appear at the T line in the case of a positive PCR reaction.
Continuing onward, the mixture next reaches the C band, where a reagent confirmation control is bound; in our hypothetical example, this could be anti-streptavidin antibody. This band would then bind to unused (or excess) streptavidin-coated gold nanoparticle reagent, developing a color band in either a positive or negative PCR reaction case and validating the performance of the detection reagent and that sample flow has occurred. Note that this does not act as a control for the underlying PCR reaction in our example, and this holds true for the C band in many real LFA designs.
Limited multiplexing is possible in the context of LFA detection. In our example case, consider a second potential target amplified by a forward primer with an additional label such as digoxigenin paired to a biotin-labeled reverse primer. On the LFA strip, addition of a “T2” band with bound anti-digoxigenin antibody next to the T band but before the C band allows for independent detection of this second amplification product. A good application of this could be to use a PCR internal control target and primer set in the reaction, allowing for validation of the PCR reaction (T2 band positivity) as well as LFA control (C band positivity) while evaluating T band for primary assay target positivity or negativity. At present most LFA-type assays as described above are limited to at most three targets, due to the lack of suitable primer tag/antibody pairs.
More complex LFA designs are possible, in particular ones based on single-stranded amplicon capture and the use of immobilized bands of potential hybridizing oligonucleotides on the strip; however, these are in effect now essentially conventional arrays, requiring more complex handling (controlled denaturation and annealing) and losing out on the inherent simplicity of the basic LFA approach in return for greater mulitplexing capacity.
Major advantages of the LFA approach are the low unit cost and simplicity of use of the disposable detection strips, and their outstanding shelf life in ambient storage conditions. When combined with ideas such as lyophilized amplification master mix pre-aliquoted into reaction tubes, an entire test system (assay plus detection device) is well suited to use in low-complexity or even poorly industrialized or field-test environments without refrigeration. In some commercial implementations further enhancements have been included, such as designing the LFA to occur internal to a small, single-use cassette where the unopened amplification reaction tube is placed inside and cassette closure initiates reaction tube puncture and LFA running—thereby reducing the chances for amplicon contamination of the testing site. Disadvantages are a lack of high levels of multiplexing, and that the approach is capable only of qualitative results in the context described.
As with our coverage of non-PCR amplification technologies, we are limited here by space from a more exhaustive list of all extant “alternative” detection strategies. The two methods covered here are ones likely to be encountered in the MDx lab in some context, and should serve to show that methods beyond the most commonly employed not only exist but have particular strengths and weaknesses, making them suitable in particular application settings. This holds true for a number of other amplification product detection strategies which may be encountered in particular products or applications.
John Brunstein, PhD, a member of the MLO Editorial Advisory Board, is President and CSO of British Columbia-based PathoID, Inc.
