Molecular markers developed for plant and animal species have been instrumental in answering many biological questions about the natural world, including, for example, those focusing on mating systems, effective population sizes, speciation, and taxonomy. Within the past decade, microsatellites have often been the marker of choice for many of these types of studies; however, in most cases, development of these markers is typically time-consuming and expensive (e.g., Squirrell et al., 2003). There are several techniques for developing these markers, including the traditional approach using bacterial colonies containing microsatellite inserts or newer methods involving next-generation sequencing. Regardless of the initial approach, putative primers must eventually be designed and tested (Zane et al., 2002). One of the most costly steps in this process is the fluorescent labeling of individual microsatellite primers, with costs varying depending upon the dye used (e.g., 6-FAM, VIC, NED) and the manufacturer. Costs further increase both in terms of necessary consumables and labor if PCR and fragment analysis are conducted separately for each individual primer and for every single sample.
Here we expand upon a method, first mentioned in Culley et al. (2008), to substantially increase efficiency while reducing costs during primer development by combining custom-labeling of primers by the investigator during the PCR reaction while also multiplexing multiple primer sets together for each sample. This is similar to the technique recently published by Blacket et al. (2012) but which uses different primers. Both methods greatly expand the technique suggested by Oetting et al. (1995), which only allowed up to five primer pairs to be multiplexed using a single universal primer. Our method also substantially advances the simplified protocol for single-dye labeling proposed by Schuelke (2000) as the “poor man's approach to genotyping.” We have now extended this process to analyze multiple primers in the same reaction using four different universal primers tagged with different fluorescent dyes; this process increases speed and economy by reducing preparation time and the amount of PCR reagents. The technique also allows the investigator to alter fluorescent label colors associated with any number of primers during primer testing to develop the best possible combination of primers and associated labels within a single multiplex mix. Using this method, many different pairs of primers can be run in a single PCR reaction, with the number only limited, at least theoretically, by the number and range of fragment sizes. This method differs from previous “multiplexing” techniques (e.g., Ottewell et al., 2005; Missiaggia and Grattapaglia, 2006) in which primers were labeled in separate PCR reactions for each sample, with the resulting PCR products combined together (“pooled”) by the researcher prior to fragment analysis.
Both this method and the independently derived technique of Blacket et al. (2012) involving fluorescent labeling and multiplex PCR have since been adopted by researchers (see Table 1), indicating the utility and applicability of this overall procedure across study systems. Our report adds to that of Blacket et al. (2012) in that we have expanded their conclusions by testing different thermocycler running conditions, addressing concerns that lower primer annealing temperatures in the multistep PCR recommended in earlier protocols (e.g., Oetting et al., 1995; Schuelke, 2000) could lead to spurious peaks and inconsistent intensities of amplified fragments (Shimizu and Yano, 2011; Blacket et al., 2012). We investigated the presence and effect of such spurious peaks and background noise of the two-step PCR process recommended by Schuelke (2000) compared to the single-step PCR cycling program recommended by QIAGEN, the manufacturer of the multiplexing kit used in this protocol. In addition, we tested the effect of cycle number in the PCR reaction on peak amplitude and also provide a step-by-step protocol (see Appendix 1 (APPENDIX 1.pdf)).
METHODS AND RESULTS
In this method, forward and reverse primers are first developed for each putative microsatellite fragment (e.g., using next-generation techniques or following traditional methods such as Zane et al., 2002). To test the primer pairs, each microsatellite fragment is then amplified using three primers (a “triplet”) added to the same PCR reaction: (1) a forward tailed primer that is tagged with a specific DNA sequence (the “tail”) attached to the 5′ end; (2) a labeled tail primer consisting of the same tail sequence but with a fluorescent label attached on the 5′ end; and (3) the unaltered reverse primer (Fig. 1; Appendix 1 (APPENDIX 1.pdf)). The fragment is amplified in the first set of PCR cycles with the forward tailed primer and the reverse primer, thus incorporating the tail sequence. In the next set of PCR cycles, the labeled tail anneals to the tail sequence and, together with the reverse primer, amplifies the fragment and in doing so incorporates the desired fluorescent label. Two elements of this protocol have been recommended by Schuelke (2000) as necessary to ensure that the forward tailed primer is incorporated before the labeled tail. First, the forward tailed primer is supplied at a reduced (1/4) concentration so it is used up in the first set of cycles. Second, a higher primer annealing temperature (57°C) is suggested for the first set of cycles to enhance binding of the forward tailed primer; the temperature is then reduced (53°C) for the second set of cycles to facilitate annealing of the larger labeled tail.
Species for which the dual labeling and multiplexing method was used in the primer testing step of the microsatellite development process.
The tail can be any unique DNA sequence that is not part of the plant genome being examined and will not interfere with other primers in the reaction. Previous investigations with individual reactions have used the M13(-21) sequence (Oetting et al., 1995; Schuelke, 2000; Boutin-Ganache et al., 2001; Scott et al., 2001; Ottewell et al., 2005), human microsatellites (Missiaggia and Grattapaglia, 2006), or alterations of genome-specific primers (Neilan et al., 1997). Multiplexed approaches have also involved customized primers such as bar-coded split tags (Shimizu and Yano, 2011) or a combination of specialized universal primers with DNA adapters commonly used in 454 DNA sequencing protocols (Blacket et al., 2012). In our studies of plant species, we have successfully used M13(−21) (TGTAAAACGACGGCCAGT), T7term (CTAGTTATTGCTCAGCGGT), and two modifications of M13(−21) developed by our laboratory: M13modA (TAGGAGTGCAGCAAGCAT) and M13modB (CACTGCTTAGAGCGATGC). Each of these sequences was tagged at the 5′ end with a different color of fluorescent label (6-FAM/blue, NED/yellow, VIC/green, or PET/red). The choice of label and color is up to the investigator's discretion and the requirements of the fragment analysis facility, so that overlapping PCR products are labeled with different dye colors.
When first testing this method in a new system, it is critical for investigators to verify correct operation of the primers. Separate PCRs should use (1) unlabeled forward and reverse primers to make sure they are designed well and amplify fragments; (2) the forward tailed primer and the reverse primer only to make sure they also amplify fragments; (3) the labeled tail alone to verify that it does not amplify any product. Fragments resulting from this method should consist of the original size plus the length of the tail sequence (18–19 bp).
To expand the applicability of the earlier single-primer tagging method so investigators could fluorescently label multiple primers concurrently in the same PCR reaction and analyze multiple loci at once, we used the QIAGEN PCR Multiplexing Kit (catalog no. 206143; QIAGEN, Valencia, California, USA). This kit includes HotStarTaq DNA Polymerase and a specific PCR buffer containing QIAGEN's synthetic factor MP, which is purported to stabilize bound primers and enable efficient extension of all primers in a reaction without any optimization. For our purposes, we did not use QIAGEN's Q-Solution (included in the kit), which reportedly helps amplify GC-rich templates and might be helpful in other systems. For multiplexing, we used multiple sets of the triplet primers suggested by Schuelke (2000), with each triplet specific to a different microsatellite fragment of a predicted size (obtained during primer development). If any of these triplets in the same multiplex PCR reaction were predicted to amplify similar fragments of overlapping sizes, the tail for each triplet was selected to correspond to a unique color for that particular fragment size range (to avoid making spurious allele calls resulting from the overlapping ranges of like-colored primer labels). Multiplexing was then conducted in 10-µL reaction volumes using the QIAGEN Multiplex Kit as follows: 5 µL Multiplex PCR Master Mix, 1 µL Primer Mix (consisting of 2 µM reverse primer, 2 µM labeled tail primer, and 0.5 µM forward tailed primer per triplet set), 3.8 µL H2O, and 0.2–0.3 µL DNA.
As with other techniques, different primers may sometimes anneal to each other during multiplexing, reducing amplification success during PCR. This occurred in a few species tested (e.g., Klooster et al., 2009; Brzyski, 2010) and was corrected by redesigning primers or running conflicting primer sets in different primer groups. Potential primer-dimer binding can also be examined a priori using primer software programs, such as AmplifX (Jullien, 2013), OLIGO (Molecular Biology Insights, Cascade, Colorado, USA; http://www.oligo.net/index.html), FastPCR (Kalendar et al., 2011), or Multiplex Manager (Holleley and Geerts, 2009).
We also compared the PCR thermocycler program suggested by Schuelke (2000) to the cycling program recommended for the multiplexing kit. First, to test Schuelke's (2000) protocol, our PCR cycling conditions were as follows: initial denaturation of 95°C for 15 min, followed by 30 cycles each of 94°C for 30 s, 57°C for 45 s, and 72°C for 45 s, and then with eight cycles each of 94°C at 30 s, 53°C for 45 s, and 72°C for 45 s. A final extension consisted of 72°C for 10 min. To test the PCR cycling program recommended for the QIAGEN kit, the thermocycler cycling conditions were as follows: initial denaturation of 95°C for 15 min, followed by 35 cycles each of 94°C for 30 s, 57°C for 90 s, and 72°C for 60 s, followed by a final extension of 60°C for 30 min. Although QIAGEN recommends 35–45 cycles for optimal results with their kit, our earlier work indicated that 35 cycles was adequate for full peak expression while minimizing the time necessary for a given PCR reaction. Because differences in peak amplitude were subsequently observed between the two reaction methods (see below), we further examined the effect of cycle number (N = 35, 40, 45, 50) on peak intensity using the QIAGEN thermocycler conditions. All fragment analysis was conducted at Cornell University's Life Sciences Core Laboratory Center on an 3730x1 DNA analyzer (Applied Biosystems, Carlsbad, California, USA) using the GeneScan 500 LIZ Size Standard (Applied Biosystems). Fragments were then analyzed with GeneMarker version 1.85 (SoftGenetics, State College, Pennsylvania, USA).
To illustrate the feasibility and effectiveness of this new method of primer design, we compared it with traditional methods using primers that had already been prelabeled with the fluorescent tag and those used in (1) multiplexed reactions or (2) with each primer pair run individually. Using 24 DNA samples obtained from wild and cultivated individuals of Pyrus calleryana Decne. (see Culley and Hardiman, 2009) and the QIAGEN-recommended thermocycler program described above, we found that the new method consistently identified the same amplified fragments for four different loci with complete accuracy (i.e., identical allele calls were obtained as for individual PCR or the multiplexed PCR with prelabeled primers; Fig. 2). Multiplexed samples that underwent the two-step PCR program recommended by Schuelke (2000) generally exhibited weaker peaks of lower intensity (Fig. 2A) than if the same samples were subjected to the QIAGEN-recommended PCR cycling program (Fig. 2B–E). In several cases, the two-step PCR resulted in occasional spurious peaks and more background noise, as previously suggested (e.g., Shimizu and Yano, 2011; Blacket et al., 2012). However, the strongest peaks with the highest intensity (Fig. 2F) were still obtained using prelabeled fluorescent primers that were multiplexed together in the same PCR reaction, with fragment sizes 18–19 bp larger as expected.
Subsequent tests altering the number of cycles in the QIAGEN method (Fig. 2B–E) indicated that peak amplitude generally increased with cycle number (or remained high if substantial already), with the optimal condition obtained at 45 cycles (Fig. 2D); peak amplitude typically dropped for most samples at 50 cycles (Fig. 2E). Peak amplitude at 45 cycles often approached that of samples multiplexed with prelabeled primers (Fig. 2F) or when primers were run individually (Fig. 2G). In general, samples run at 45 cycles (Fig. 2D) exhibited slightly more background noise than when samples were multiplexed using prelabeled primers (Fig. 2F), but all peaks were still readily discernable.
To date, we have successfully used this method to develop microsatellite markers in several plant species from a variety of plant families representing different habits; it has also been used in a number of other taxa in other laboratories (Table 1). In each of the species analyzed, seven to 15 microsatellite markers were developed and successfully run in one to three primer groups in multiplexed reactions (with a maximum of four to 13 primer pairs per multiplexed primer group). The microsatellite fragments were of varying size and composition, ranging from dinucleotide to hexanucleotide repeats. In some cases, the applicability of the technique was straightforward without any modification (as with Schiedea adamantis H. St. John [Culley et al., 2008] and Rhamnus cathartica L. [Culley and Stewart, 2010]), but in other cases (Monotropa hypopitys L. [Klooster et al., 2009] and Spiraea virginiana Britton [Brzyski, 2010]), certain primer sets were found not to amplify together and had to be run individually or placed in different primer groups. In addition, there were isolated cases in which a primer with especially strong amplification appeared as small false peaks for other loci in the same reaction (i.e., peak carry-over), which can also happen with traditional labeling of primers. Therefore, some troubleshooting may be required to ensure that all primers amplify properly. In addition to its use in microsatellite development, this technique has also been used to label nonmicrosatellite primers, such as those that amplify self-incompatibility alleles in Pyrus calleryana (Culley and Hardiman, unpublished data) and Spiraea virginiana (Brzyski and Culley, unpublished data).
The dual labeling and multiplexing method presented here has been successfully applied to several plant species using primarily microsatellite markers, although it should be applicable to other types of markers as well. In addition, the method offers flexibility in tagging primers with different fluorescent dyes and/or colors during the early stages of primer development. Like Blacket et al. (2012), we found that using triplet sets of primers containing unique universal tail sequences provided an effective way to amplify multiple loci in the same PCR reaction. Furthermore, we also showed that the amplitude and fluorescent intensity of the labeled fragments generated during multiplexing will also depend in part upon the thermocycler conditions during the PCR process. We discovered that the two-step thermocycler program suggested by Schuelke (2000) previously used by our laboratory (e.g., Culley et al., 2008; Klooster et al., 2009) did not yield the best results. Instead, the thermocycler conditions optimized by QIAGEN for their Multiplex PCR Kit resulted in cleaner fragment peaks with higher intensity (Fig. 2), especially when the cycle number was increased to 45, the maximum recommended by QIAGEN. Consequently, the major objection to using M13-based primers (i.e., that they require a lower primer annealing temperature; Blacket et al., 2012) was not borne out by our study as the primer(s) performed even better when a higher standard annealing temperature was used (57°C). We therefore recommend the QIAGEN-recommended thermocycler protocol with 45 cycles if the QIAGEN Multiplexing Kit will be used by the investigator.
When applied correctly, the technique reported here can reduce the expense of primer development by decreasing the amount of time and labor spent in preparing and running multiple PCRs and by reducing the direct cost of consumable supplies and services. We found that the expense of the new method is typically less than half of the cost of the traditional technique of individually labeling each primer prior to nonmultiplexed PCR reactions ( Appendix S1 (APPS-D-13-00027_Appendix S1.pdf)). For example, testing 10 primer pairs in two primer groups using 50 samples with the new method is less than half the cost of the traditional technique (estimated currently at US$639 compared to US$1750; Appendix S1 (APPS-D-13-00027_Appendix S1.pdf)). These cost savings will increase as more primers are incorporated into the multiplex PCR reaction with fewer sets of primer mixes. Consequently, this technique would be particularly advantageous for laboratories on a limited budget. However, researchers must balance the cost savings and flexibility of this new method with the observation that multiplex PCR using prelabeled fluorescent primers, which yielded strong peaks in our tests, would require less time in terms of constructing primer mixes because one less primer per set is involved. One approach employed in our laboratory is to first use the dual labeling method to fine-tune the dyes associated with different primer mixes before ordering the prelabeled forward primers direct from a manufacturer for the full analysis; this is particularly effective for large-scale studies involving hundreds of individuals. In smaller studies, we continue to use the dual labeling method exclusively to maximize dye flexibility and minimize overall costs of primer development and application. To assist researchers, a step-by-step protocol of the dual labeling technique described above is provided as Appendix 1 (APPENDIX 1.pdf), which contains all necessary details as well as answers to frequently asked questions.