Variation in eluate volume during DNA extraction does not affect eluate concentration of plasma-derived DNA

INTRODUCTION

DNA testing is used across many diverse fields, including forensics, environmental monitoring, food quality assessment, genetic research, and healthcare diagnostics^1–6. Reliable results depend on stringent requirements, such as accurate DNA quantification and robust handling of low-level DNA. Different technologies impose different demands on DNA testing, for example in relation to input DNA concentration, volume, and purity. In addition, pre-analytical factors can affect test reliability, particularly in sensitive applications like cell-free DNA (cfDNA) analysis^7–10. While some variation is inevitable, high analytical precision is undermined if sample collection and DNA extraction introduce substantial variability. Pre-analytical aspects such as sample collection, processing, storage, and DNA extraction have been studied intensively across different DNA testing fields^7–14. Here, we investigate the variation in the eluate volume after elution at the end of the DNA extraction process, using the automated QIAsymphony extraction system. We investigate how potential variation in eluate volume may affect DNA quantification using the high-precision technique of Droplet Digital PCR (ddPCR).

MATERIALS AND METHODS

In this study, we used blood samples from three healthy volunteers. Informed and written consent was obtained from all participating volunteers when blood samples were collected. The project was conducted in accordance with section 42d(2)(2) of the Danish Health Act, and as a quality assurance study no further Ethics Committee approval was needed.

The three volunteers each donated 10 blood samples (total was 30 individual samples). Blood was drawn (locally on site with no sample transportation) in 10 mL Cell-Free DNA BCT^® tubes (Streck, La Vista, Nebraska, USA), stored at RT, and processed within 24 hours. Samples were centrifuged at 2,000 × g for 10 min, and plasma was carefully collected and pooled into a single pool. The pool was homogenized using magnetic stirring for 10 min at RT. The pool was then aliquoted into 24 portions of 4.5 mL, and DNA extractions were made from each of these aliquots. DNA extraction was done from 4 mL plasma using the QIAsymphony SP instrument and the QIAsymphony DSP Virus/Pathogen Kit (Qiagen, Hilden, Germany). Elution volume was set to 60 μL. A precise measurement of elution volume was done by weighing a designated, sterile 2-mL screw cap micro tube (Sarstedt, Nümbrecht, Germany) before and after elution and then subtracting the individual tube weight to get an exact elution volume. An A&D GR-200 analytical balance was used as precision weight (A&D Instruments, Abingdon, England). Assuming an eluate density of 1.00 g/mL allowed for the following volume calculation:

$$Volume\hspace{0.17em}(mL)=(post-extraction\hspace{0.17em}mass-pre-extraction\hspace{0.17em}mass)\hspace{0.17em}[g]*(\mathit{1}\hspace{0.17em}mL/\mathit{1}\hspace{0.17em}g).$$

Genomic DNA (gDNA) from the buffy coat of the three participating individuals was tested using a panel of biallelic SNPs on quantitative PCR (qPCR), according to the previously described method¹⁵. The S63 SNP-assay for ddPCR-based DNA quantification¹⁵ was selected for which one of the participating individuals was homozygous for one allele, another was homozygous for the other allele, and the third was heterozygous. Signals for both probes were summarized to calculate the amount of total cfDNA. The QX200 ddPCR system (Bio-Rad, Hercules, CA, USA) was used to determine the concentration of cfDNA. Each eluate was tested with 6–8 replicate reactions in the ddPCR system, depending on the available eluate volume. Each ddPCR reaction with a final volume of 20 μL consisted of 10 μL of 2× ddPCR Supermix for probes (No dUTP) (Bio-Rad), 0.8 μL primers and probes at a final concentration of 900 nM and 250 nM respectively, 8 μL cfDNA (eluate), and 1.2 μL sterile H₂O. Droplets were generated using the QX200 Droplet Generator (Bio-Rad) and subjected to DNA amplification on a C1000 Touch Thermal Cycler (Bio-Rad) with the following PCR conditions: 95°C for 10 min initial denaturation, 40× (94°C denaturation for 30 s, 60°C annealing and extension for 1 min), then 98°C for 10 min for enzyme deactivation. The temperature ramp rate was 2°C/s. The ddPCR plate was held at 12°C until further analysis. Droplets were read by the QX200 Droplet Reader (Bio-Rad) and analyzed using QX Manager Software (v1.2). Direct Quantification (DQ) was used, and a threshold value was set manually before interpreting the results.

Associations between eluate measurements and DNA copy number variables were evaluated using Spearman rank correlation analysis. The plasma pool aliquots were processed and analyzed as technical replicates. As such, the analyses reflect technical variability within a pooled sample rather than biological variability between individuals.

Potential outliers were identified and excluded using the robust regression and outlier removal (ROUT) method with a false discovery rate (Q) of 1%. Spearman correlations were calculated separately for DNA copies per milliliter of eluate and total DNA copy number following outlier exclusion. All statistical tests were two-sided.

RESULTS

We investigated how potential variation in DNA extraction eluate volume might affect downstream DNA quantification. The collection tubes were weighted before and after elution to estimate the eluate weight. The tubes weighed an average of 1.513 g with a CV of 0.311% (No.=24). The calculated mean eluate volume from DNA extraction of identical pool aliquots was 63.3 μL (range: 53.2–70.8 μL) (No.=24); the CV was 6.6%. Estimated mean DNA concentration was 66.1 (range: 57.5–85.6) DNA copies per μL eluate; the CV was 7.6%.

The mean total DNA per eluate was 4,179 DNA copies (range: 3,614–4,794 DNA copies); the CV was 7.9%.

There was a positive correlation between eluate volume and total DNA per eluate (Figure 1A); however, there was no correlation between eluate volume and DNA concentration (Figure 1B). For eluate volume, we compared within-day variation (CV of 5.6%) with day-to-day variation (CV of 9.6%), revealing a higher day-to-day variation.

Figure 1. (A) Correlation between eluate volume and total DNA detected and (B) correlation between eluate volume and DNA concentration.

Data were not normally distributed (p=0.0002, Shapiro-Wilk test). (A) Significant correlation was observed; Spearman correlation test: r=0.67 (95 % CI: 0.35–0.85), p=0.0004; no outliers identified (ROUT). (B) No significant correlation observed; Spearman correlation test: r=0.35 (95 % CI: −0.08–0.67), p=0.0991; excluding one outlier identified (ROUT) marked by open circle. (If the outlier was included, Spearman r=0.19; p=0.3737). The ROUT method identified a single observation as an outlier for DNA concentration per milliliter of eluate but not for total DNA copy number, reflecting differences in variance structure between normalized and absolute DNA measures. For visualization, data were plotted with locally fitted regression lines and 95 % confidence intervals to illustrate the direction and strength of associations.

DISCUSSION

In this study, we showed that ddPCR-based quantitative measurements of DNA concentrations were unaffected by variation in eluate volume during DNA extraction. As expected, we found a correlation between eluate volume and total DNA in the eluate, but rather unexpectedly, there was no statistically significant correlation between eluate volume and DNA concentration, as measured using aliquots of pooled plasma material from which DNA was extracted using the QIAsymphony instrument. Thus, an increased elution volume did not dilute the DNA concentration, nor did a decrease in volume increase the DNA concentration, at the investigated elution volume of 60 μL.

The consequence of these findings is that we can rely on quantitative DNA data as measurement of DNA concentration. This is obviously essential for diagnostic tools that are based on DNA quantification, and it is highly relevant for methods that measure minor quantitative changes in DNA.

The study findings are limited to the specific bead-based DNA extracting instrument and protocol used. The study is also limited by low sample size. However, the data are clear, and they substantiate the precision of our quantitative ddPCR method, which we use for testing donor-derived cell-free DNA in transplanted patients¹⁵. Furthermore, we may expect similar unaffected precision in quantitative measurements from Ct-values using real-time PCR, based on the same DNA extraction instrument and protocol, as used for example in pregnant women¹⁰.

It would be interesting to assess the potential variation of other elution volumes and assess their potential impact for downstream quantitative DNA measurements. Others have compared different DNA extraction systems¹⁴, but not looked at eluate variation.

CONCLUSIONS

Variation in eluate volume during DNA extraction on a QIAsymphony did not affect downstream ddPCR-based DNA quantification. Assessment of such basic pre-analytical factors is always warranted and of paramount importance for ensuring test reliability.