An improved rapid method for DNA recovery from cotton swabs

Preliminary experiments to improve free DNA recovery from cotton swabs using combinations of cellulases based on the experiments described by Voorhees et al. [] to digest the cotton matrix and leave the DNA in solution proved to be ineffective requiring long digestion times and poor recovery (data not shown). This result for free DNA contrasts with the findings of Voorhees et al. [] who found that cellulase digestion increased sperm head recovery suggesting that free and cellular DNA have different interactions with the cotton swab which is not surprising given the difference in form and chemical composition.

A method for extracting PCR quality DNA from crude leaf homogenates with paper disks [] provided a useful starting point for method development. In summary the authors found that DNA from crude leaf homogenates could be bound to filter paper disks, was not eluted by rapid water washing and could be eluted from the paper with PCR reaction mix. They did not however, determine which components of the reaction mix were essential or contributed to the elution of the DNA. Therefore, an experiment to test the PCR reaction mix components in all possible combinations was conducted as follows, 50 ng of purified free DNA was spotted onto 4 mm diameter Whatman No1 cellulose filter paper disks and allowed to air dry for 12 h. Duplicate filter disks were placed in 1.5 ml Eppendorf tubes and 50μl of each of the extraction mixes added as detailed in the Table 1. The tubes were vortexed for 5 s and left to stand for 3 min before another 5 s vortex and centrifugation at 16000xg for 2 min. DNA was quantified by Qubit using 10μl of the supernatant from the centrifugation step.

From the data presented in Fig. 1, shown as percentage recovery of DNA as estimated by Qubit assay, it can be seen that the presence of sodium chloride has a deleterious effect on the elution of the DNA whereas BSA, Tween 20 and dNTP mix all have a positive effect. These effects are not synergistic or fully additive but do have a greater effect when used in combination which can be more easily seen when this data is extracted from Fig. 1 as shown in Fig. 2. Based on this data we tested whether or not this method could be applied to elution of DNA from cotton swabs. An initial experiment repeating the elution with BSA (bsa), nucleotides (nuc), Magnesium (mg) and Tween 20 (tw) in combination was carried out. 50 ng of free DNA was applied onto cotton swabs and dried overnight followed by a short 5 min elution. Extracted DNA was estimated by Qubit assay which gave variable results (data not shown). This was probably due to the increased volume of elution buffer required for the larger cotton swab, 250μl as opposed to 50μl for the paper disks, and incomplete equilibration of DNA throughout the elution volume due to the high volume trapped in the swab. This problem was addressed by repeating the experiment and centrifuging the swabs in a spin basket, as described in Materials and Methods, to increase the extract recovery and make it more consistent. Control blank swabs were included, and deoxy nucleotide blanks were also Qubit assayed to assess any interference from free nucleotides (none was detected). The data presented in Fig. 3 clearly shows that DNA is eluted from cotton swabs in an almost identical manner to the filter paper disks (Fig. 2). In all subsequent experiments swabs were eluted using a spin basket step which was centrifugation at 16,000xg for 2 min. Due to the high cost of adding dNTP’s at a concentration of 1 mM each and the small effect the addition of the dNTP’s appears to have on recovery, ATP, which is much cheaper was assessed as a substitute based on the work of Tanaka et al. []who used dATP to effectively elute DNA from nano particles. These authors found dATP to be most effective in the 5–10 mM range therefore the effect of 5 mM and 10 mM ATP alone and in conjunction with 1 mg/ml BSA was tested for effectiveness in eluting DNA from cotton swabs. The results presented in Fig. 4 show that ATP alone does enhance release of DNA from the swab although there appears to be little difference between 5 and 10 mM ATP. The addition of BSA at 1 mg/ml increases the DNA recovery as would be expected from the data presented in Fig. 1 which shows BSA has a major effect in increasing recovery. Given the marginal effect of the addition dNTPs or ATP on the recovery of free DNA they were no longer considered as additives to the extraction buffer. The slightly higher recovery of DNA from the filter paper disks (comparing Fig. 2, Fig. 3) is probably due to a more efficient extraction from the paper due to the fact that the volume of extraction buffer to paper is 11:1 whereas the volume ratio for the swab extraction is approximately 1:1.

Having tested various constituents of an extraction mix and using spin baskets for elution of DNA from cellulose paper and cotton swabs it is observed that around 40% free DNA recovery was obtained in a five minute incubation by the most efficient extraction mixes. Whilst this incubation period is sufficient to test the efficacy of various extraction mixes we conducted a time course of extraction in order to determine whether an increase in incubation time would increase DNA yields. As previously 50 ng of free DNA was deposited directly onto cotton swabs in triplicate and dried overnight. Extraction was carried out as described in the legend to Fig. 5 with the incubation in extraction mix being carried out for the time points indicated. The data presented clearly shows an initial rapid release of the DNA followed by a slower linear phase significantly increasing the recovery. The final extraction mixture used in the next phase of the method development consisted of 1 mg/ml BSA, 1% tween 20 in 20 mM TrisHCl pH 8.0 referred to subsequently as extraction buffer.

In order to test the utility of the extraction buffer method it was compared to the Qiagen QIAamp kit using the rapid method described in the kit manual referred to as Protocol 1. Quadruplicate dried swabs with 50 ng DNA were subjected to the Qiagen protocol as laid out in the instruction manual which includes a 10 min digestion in proteinase K at 56 °C. To maintain parity between the two techniques one set of swabs was incubated in extraction buffer at 56 °C and a third set at room temperature for 10 min. The results which are shown in Fig. 6 as % recovery based on the input DNA amount of 50 ng (Protocol 1), indicate that direct elution using extraction buffer is much more efficient than the Qiagen kit and surprisingly incubation at elevated temperature does not seem to increase the DNA recovery by the extraction buffer. There is however a longer swab protocol described in the Qiagen manual specifically for swabs involving a longer incubation (1 h) in proteinase K at 56 °C (Protocol2) but since there are no cells present on the test swabs this should not make any difference, however the increased incubation time may enhance the Qiagen recovery. Therefore, a repeat of the Qiagen comparison experiment described above was carried out using the longer protocol 2. This includes a 1 h incubation at 56 °C and a 10 min incubation at 70 °C which, to maintain parity, was included in a set of elution buffer extractions. Standard extraction buffer incubations were for 1 h at room temperature with the results of this experiment given in Fig. 6 (protocol 2). Using this extended protocol the yield of DNA from the Qiagen kit has increased but so has the yield from the extraction buffer elution method which produced around 95% recovery. This appears to be mostly due to the increased incubation time rather than the increased temperature although there is a marginal increase at the elevated temperature. We are aware that there are other variants of the standard Qiagen protocols which are used by many laboratories, but for the purposes of this paper we are comparing to the manufacturer’s recommended protocols rather than modifications which may be used by individual labs.

There are no cells present in the free DNA applied to the swabs which is unlikely to be the case in a swab from a crime scene surface[]. However, the relative proportions of free and cellular DNA normally found on a swab are not known with any certainty since it is impossible to distinguish the origin of a PCR signal as being from free or cellular DNA as both will be amplified by any human primer set. Experiments have been conducted to differentiate between free and cellular DNA [] however these experiments relied on direct recovery of material from hands by buffer washing followed by centrifugal separation of cellular material from free DNA. This methodology is not applicable to material dried onto cotton swabs. It is possible that DNA from cellular material may be recovered more efficiently from the swabs by the Qiagen method. To investigate this, swabs were prepared with free DNA, cells and a mixture of both and allowed to dry for 24 h. DNA was then recovered from the swabs using the Qiagen QIAamp long swab method as detailed in the kit instructions or extraction buffer.

The cellular input DNA was estimated by adding cells directly to the Qiagen extraction buffer or directly to the swab elution buffer and processing as for swabs. For cellular DNA mouse embryonic fibroblasts were suspended in PBS at a concentration of 1.4 × 10⁶/ml and 10μl added to Qiagen extraction buffer containing proteinase K or 250μl of swab extraction buffer. Given the size of the mouse genome (6 pg/cell) [] 10μl containing 14,000 cells should contain 84 ng DNA. The estimate by Qubit for the direct to buffer Qiagen extraction was 93.2 ng and by extraction buffer 50 ng. In calculating the recovery of DNA, 93 ng was used as the value for input cellular DNA. This was chosen as it is unlikely that the extraction buffer causes complete lysis and release of the nuclear DNA. However, it should be noted that the extraction of cells with extraction buffer was carried out at room temperature. It is possible that heating would increase the release of nuclear DNA. Free DNA input was quantified by Qubit as 60.6 ng per sample. In calculating recovery from mixed cellular and free samples the sum of the two individual components was used as the input DNA at 153.6 ng per sample. Control blank swabs were processed by both methods.

Triplicate swabs were processed by the Qiagen QIAamp protocol or the extraction buffer protocol including the 1 h 56 °C incubation. The results are shown in Fig. 7. The data clearly shows that the swab elution buffer is more efficient. However, it is worth noting that the recovery using the swab elution buffer is higher (65 ng compared to 50 ng) from the swab than from cells added direct to the buffer without a swab. This is likely due to the fact that the swab extractions were heated to 56 °C for 1 h whereas the cells without the swab were incubated at room temperature. It is likely that the increased temperature in the presence of 1% tween 20 is sufficient to cause complete nuclear lysis. This was assessed as described below.

In order to assess the integrity of nuclei in extraction buffer cells were incubated in either PBS or swab buffer at room temperature or 56 °C for 1 h and then stained with DAPI to visualise nuclei by fluorescence microscopy. It was observed that after 1 h in PBS at room temperature the nuclei and cells are still visible and mostly intact (Fig. 8a) but the numbers are decreased at elevated temperature (Fig. 8c) whereas at room temperature the number of visible nuclei in extraction buffer is markedly reduced (Fig. 8b) and were also observed to show marked swelling relative to the PBS buffer at higher magnification (Fig. 8e and f). Elevated temperature in swab buffer resulted in nearly all nuclei being destroyed (Fig. 8d).

The next stage in assessing the development of a swab extraction protocol was to determine the optimum conditions for swabbing a surface seeded with a known amount of DNA and/or cells and estimating the recovery. It is well known that high salt can enhance the binding of DNA to cellulose as used in the technique of southern blotting[].

Given that high salt over 40 mM can interfere with subsequent PCR reactions and 150 mM results in complete inhibition [], experiments were conducted using swabs seeded with free and cellular DNA (52.5 and 60 ng respectively) to determine the effect of salt on DNA binding and a washing protocol developed to remove the salt before elution. Having a wash step in the procedure would also be beneficial to remove contaminants picked up from surfaces in the case of actual crime scene samples, provided this does not remove the DNA from the swab. These experiments were conducted by seeding swabs with known amounts of free DNA, cellular DNA or a mixture of both and washing in spin baskets then extracting the bound material before quantifying by Qubit. Washing with water or 10 mM TrisHCl buffer resulted in almost complete removal of seeded cells and DNA with less than 5% of the seeded samples remaining (data not shown). However, washing with 70% ethanol removed most of the salt, as confirmed by the third ethanol wash giving only a faint reaction with silver nitrate (little no or silver chloride precipitate), and had only a marginal effect on the recovery of material as shown in Fig. 9. Triplicate swabs were seeded with 52.5 ng of free DNA (10μl), 60 ng of cellular DNA (10μl) or a mixture of 10μl of each onto cotton swabs pre-treated with either 50μl of water or 50μl of 250 mM NaCl and allowed to dry overnight. Swabs were then either left unwashed or washed in spin baskets 3 times with 400μl of 70% ethanol followed by elution in 250μl of extraction buffer at 56 °C for 1 h. The data (Fig. 9) shows that the introduction of a washing step may have slightly beneficial effect in increasing DNA recovery even in the absence of salt in the swab wetting solution.

Having confirmed that any added salt can be removed without significantly compromising the recovery of DNA, the effect of adding salt to the swab wetting agent on DNA recovery from the surface of glass slides was investigated. Known amounts of free and cellular DNA as per the previous experiment were seeded onto glass slides and allowed to dry overnight. Swabbing was conducted with swabs wetted with either 50μl of water or 50μl of 250 mM salt. Swabs were processed using extraction buffer as in the previous experiment (250μl buffer for 1 h at 56 °C). The results show (Fig. 10) that the recovery of DNA from the glass slides was marginally enhanced by the addition of salt to the swab wetting solution.

The usefulness of this elution technique depends on the ability to reliably PCR the extracted DNA from the elution buffer. Experiments were conducted to test the suitability of the eluted extract for amplification of both free and cellular DNA. In practice touch DNA is recovered in the range of 0.06 ng/μl to 0.01 ng/μl in a volume of 50μl which is 3.0 ng to 0.5 ng in total (personal communication from practitioners) and is in agreement with the published values for glass[]. Given a recovery efficiency of about 30% as estimated from the data of Wood et al. [] DNA on the surface would have to be in the range of 10 ng to 1.6 ng to give these final values of recovered DNA. Therefore, DNA was spotted onto glass slides in the range of 30 ng to 0.1 ng and allowed to dry overnight. DNA was subsequently recovered with swabs wetted with 100μl 250 mM NaCl which were also dried overnight. Swabs were washed 3 times with 70% ethanol and then extracted with shaking at 56 °C for one hour in 250μl swab extraction buffer. Samples were then quantified for DNA by qPCR in triplicate. The qPCR standards were run in standard Qiagen elution buffer and extraction buffer for comparison. As can be seen from the data plotted in Fig. 11 there was no significant difference in the standard curves indicating that the extraction buffer components do not have a significant effect on the qPCR reaction. The data for the swabbing experiment is presented in Fig. 12 and shows that the DNA can be recovered from the glass surface and eluted from the swab with around 60% efficiency in the range over the range of input DNA that would be expected in realistic samples. Lower amounts of input DNA resulted in unreliable quantitation (data not shown) as indicated by the qPCR melt curves showing a significant primer dimer peak which results in an over estimate of the DNA present.

When the recovery experiment was repeated with cellular DNA, very little DNA was detected by qPCR but was still detected by Qubit quantitation as in previous experiments (data not shown). Even though the extraction buffer destroys nuclei effectively (Fig. 8) the Qubit dye can still bind effectively to the DNA to give a Qubit signal. However, the remaining proteins may interfere with the PCR reaction. Possibly protein remains bound to the DNA (probably histones) preventing effective primer binding for qPCR or the released cellular proteins may be digesting the nucleotides and/or primers. This second potential source of interference is unlikely since heat treatment at 56 °C will destroy most enzyme activity. However, it is possible that the proteins although denatured by the heating step may still be capable of binding the primers or nucleotides. In the Qiagen extraction method these proteins are destroyed by proteinase K digestion, however since a considerable degree of the effectiveness of the extraction buffer relies on the presence of BSA, proteinase K digestion is not an option in the extraction buffer as this would destroy the BSA. The initial rationale for the use of BSA was its inclusion in some PCR buffers and in other molecular biology applications, in particular southern blotting and other hybridisation protocols, where it acts as a blocking agent to prevent non-specific binding[]. The compound polyvinylpyrrolidone (PVP) has often been used in hybridisation buffers as a cheaper substitute for BSA and in PCR reactions to combat interference from inhibitors[]. The extraction buffer modified with the replacement of BSA with PVP at a range of concentrations was tested in a direct comparison with the normal BSA containing extraction buffer for its ability to recover DNA seeded onto cotton swabs. Triplicate swabs had 50 ng free DNA applied which were dried overnight. The swabs were extracted in 250μl of extraction buffer at 56 °C for 1 h in standard buffer (1 mg/ml BSA, 1% tween 20 in 20 mM TrisHCl) or in PVP buffer (1–3% PVP, 1% tween 20 in 20 mM TrisHCl) and the extracted DNA quantified by Qubit assay. It was found that PVP was able to substitute for BSA at 1% however higher concentrations were less efficient (Fig. 13). Given these results the final composition of the extraction buffer used in all subsequent experiments was 1% PVP,1% tween 20 in 20 mM TrisHCl containing 20 μg/ml proteinase K.

An experiment was conducted to test the extraction/recovery efficiency, from swabs and a glass surface, of the PVP proteinase K extraction buffer using free trout DNA and cellular mouse DNA. The protocol is shown diagrammatically in Fig. 14. Free DNA was diluted to 4.83 ng/μl and seeded at 10μl and 5μl giving 48.3 ng and 24.1 ng respectively as estimated by Qubit. Mouse cells were resuspended in PBS at approx. 0.72 × 10⁶/ml equivalent to 4.32 ng/μl and seeded at 10μl and 5μl approximating to 43.2 and 21.6 ng DNA respectively. Samples with a 1:1 mix of cells and free DNA were also used. A sample ID key is given in the legend to Fig. 14. All samples were extracted at 56 °C for 60 min and then spin basket collected (in the case of swabs).

The input DNA in this experiment was quantified by qPCR of the input samples added directly to the extraction buffer. The recovery of DNA is expressed as percentage recovery of the input DNA as determined by direct to buffer extraction. The data presented in Fig. 15 clearly demonstrates efficient recovery from swabs 80–90% for free DNA and 60–80% for cellular DNA. From swabbed glass slides free DNA is recovered in the range of 70–80% for free DNA and 60–70% for cellular DNA. For the mixed depositions of free and cellular DNA the recoveries are in the range of 50–70% overall.

In order to demonstrate the increased recovery of the PVP extraction method it was compared to the Qiagen QIAamp kit. The experimental protocol was as for the recovery from direct to buffer, direct to swab and from glass slide as described above and in Fig. 14 except the PVP and Qiagen methods were used in parallel in the swab extraction step. The Qiagen method for swabs as recommended by the manufacturer includes an extended extraction step at 70 °C and an increased incubation time of 1 h at 56 °C. In order to maintain parity the PVP extraction method was modified to include this elevated temperature step. The Qiagen protocol was also modified to include a spin basket step, to maximise recovery, before applying the extracts to the Qiagen columns.

The results of this comparison are presented in Fig. 16. As in the previous experiment the data is presented as % recovery with 100% defined as the direct to buffer samples. In this case however since the recovery using the Qiagen method is substantially lower than that obtained by PVP extraction the higher PVP extraction value is used as the 100% recovery. The data shows that recovery obtained using the PVP method is comparable to that obtained in the previous experiment but the recovery by QIAamp kit extraction barely reaches 20% in most cases. These findings regarding efficiency of recovery by Qiagen are in broad agreement with those reported by Kemp et al. [] although these authors used the Qiagen MinElute Purification Kit rather than the QIAamp kit used in this study. However both these kits use the same general principals and buffers to extract and purify the DNA.

The discrepancy in the recovery for the two methods could be due to losses incurred during the column purification steps of the Qiagen samples. This was investigated using the Investigator Casework Go kit which is an equivalent Qiagen kit in that the extraction buffer is added to the swab, incubated at 56 °C and the extract used directly in a PCR reaction.

Free DNA and cellular DNA were diluted serially two-fold in 10 mM TrisHCl buffer pH 8.0 to give a range of concentrations from 20 ng/μl to 6.125 ng/μl. Free DNA and cells were deposited directly to cotton swabs in a volume of 10 μl and allowed to dry overnight giving a range of 200 ng to 61.25 ng deposits. The DNA concentrations of free and cellular DNA stocks were verified by the addition of samples direct to their respective extraction buffers followed by qPCR. Again to maintain parity between the two methods, swabs were extracted using the Qiagen Investigator Go protocol for both the Qiagen and PVP samples with respect to incubation times and temperatures. This includes an incubation at 60 °C for 25 min followed by a 5 min incubation at 80 °C to kill proteinase K activity. Also samples from both methods were spin basket centrifuged to maximise recovery. The results presented in Fig. 17 show that the recovery of free DNA using the Qiagen Investigator Go kit is much improved over the QIAamp kit (Fig. 16) at higher DNA concentrations but performs poorly at low DNA concentrations compared to PVP extraction. The data also shows (Fig. 17) that recovery of cellular DNA is also improved but is still less efficient than the PVP extraction buffer.

The Qiagen Investigator Go kit was also tested in comparison to the PVP extraction method in recovery of free and cellular DNA from surfaces. Free DNA was diluted in 10 mM TrisHCl buffer pH 8.0 to give a concentration of 10 ng/μl and deposited directly in a volume of 5μl. Similarly, a cell suspension was also created at a concentration 30 ng/μl (determined by subsequent qPCR). Mixed cell and free DNA samples were also deposited using an equal (5μl) volume of both. Samples were spotted directly to cotton swabs, glass slides or drinks cans and allowed to dry for 3 h. The drinks cans and glass slides were then swabbed using cotton swabs moistened with 50μl TrisHCl buffer pH 8.0 and allowed to dry overnight.

Control direct to extraction buffer samples were treated in an identical manner to the swab samples with respect to incubation duration and temperature and subjected to qPCR giving an accurate determination of the input DNA. The swabs were processed as per the directions for the Qiagen Investigator Go kit with the exception of the spin basket step (16,000xg 2 min) using an extraction volume of 250μl. After the heat treatment step to kill proteinase K activity, 5μl samples were taken directly into 5μl of the qPCR reaction containing the appropriate mouse (cellular material) or trout (free DNA) primers.

The data is shown in Fig. 18 as % DNA recovered relative to the DNA estimated from samples added direct to the appropriate buffer. The data shows the mean and SD of the % DNA recovered from duplicate swabs assayed by qPCR in duplicate i.e.: 4 qPCR determinations from duplicate swabs. The data shows that the PVP extraction is more efficient than the Qiagen method. There are no significant differences between the surface extraction and direct to swab extractions as would be expected since the difference is between the efficiencies of extraction from the swabs and not the recovery of the DNA onto the swabs from the surfaces.

In practical terms the drawbacks of both the PVP extraction method and the Investigator Go kit are that the extraction results in the DNA being at a relatively low concentration due to the volume of extraction buffer required. Also any contaminating PCR inhibitors swabbed from the surfaces of interest will be present in the extracts. Both of these concerns can be addressed using a simple purification and concentration step using paramagnetic beads to capture the DNA by Solid Phase Reversible Immobilization (SPRI). Commercially available SPRI beads such as AMpure XP(Beckman Coulter) would be prohibitively expensive as the protocols for the pre-mixed beads requires 1.8 times the volume of beads to be added to the solution being extracted in order to give the correct PEG concentration for DNA binding to the beads.

The capacity of the beads for DNA binding is in excess of 5 ug per 1μl of beads, therefore the addition of excess beads is unnecessarily expensive. Examination of the original paper describing SPRI beads [] shows that the required beads and PEG/NaCl mix can be added separately. This allows the addition of a small volume of beads and concentrated PEG/NaCl solution which results in the minimum quantity of beads being added along with a volume of PEG/NaCl giving their required final concentrations for efficient DNA binding. The full details are given in materials and methods. Fig. 19 shows the results of testing this method for free and cellular DNA. The recovery efficiency of free DNA was assessed by adding 50 ng of free DNA (quantified by Qubit assay) to 250μl of PVP extraction buffer and purifying the DNA using 2–10μl of beads and the recovered DNA quantified by Qubit assay. Since cellular DNA cannot be accurately quantified by Qubit assay due to the contaminating cellular material, the cellular DNA was quantified by qPCR after normal extraction protocol including proteinase K digestion (250 ng input DNA). The cellular DNA recovered from the bead purification was also quantified by qPCR. The data presented in Fig. 19 for the free DNA shows that due to the high binding efficiency of the beads 2μl already gives around 70% recovery of the input DNA increasing to around 85% recovery with 8μl of beads. Since the elution volume from the beads was in this case 50μl, the DNA concentration has increased from 0.34 ng/μl to 1.3 ng/μl, a 3.8 fold increase in concentration. The data for cellular DNA shows a more gradual increase in binding. We speculate that this is due to the presence of peptides, from the proteinase K digestion, and perhaps cellular RNA competing for binding to the beads. However, the addition of 8–10μl of beads results in a recovery of 70–72%. Since the input DNA was at a concentration of 1 ng/μl this equates to 3.5 fold increase in concentration, similar to that observed for free DNA.