Molecular biomarkers are rapidly used in clinical practice for disease detection, particularly in oncology, where they often outperform traditional diagnostic methods (for instance: Biopsy examination and visual inspection with acetic acid in cervical cancer screening) in sensitivity and specificity (1). Generating reliable molecular biomarker data requires high-quality clinical specimens, which can be challenging to obtain, particularly in low- and middle-income countries (2,3). Persistent barriers include limited numbers of trained personnel capable of performing proper sample collection, as well as logistical constraints related to transportation, cold-chain maintenance and storage (4). These challenges may compromise specimen integrity, ultimately affecting the accuracy and reliability of downstream molecular analyses.

Liquid-based cytology clinical samples, such as cervical scrapes collected with a brush and preserved in ThinPrep PreservCyt^® solution, are widely used in human papillomavirus (HPV) detection for cervical cancer triage screening, as they efficiently preserve cellular material for both cytological and molecular analyses. However, a frequent bottleneck in downstream molecular assays, such as in viral load quantification, genotyping and sequencing, is inefficient DNA extraction, leading to a low yield or suboptimal purity of DNA (5,6).

To address this critical initial step, silica spin-column based extraction kits are widely used in both clinical and research laboratories due to their convenience, scalability and relative consistency. However, these standard protocols may not be optimal for all sample types or downstream applications, a limitation that is particularly acute for low-viral-load HPV samples. Therefore, the selection of elution parameters (particularly volume, incubation duration and the number of elutions) becomes a key optimization step, as they can significantly affect both the final DNA yield and the concentration of the eluate (7).

The optimization of protocol variables, such as incubation time, elution volume and number of elutions can significantly influence DNA yield and quality (8,9). Given that ThinPrep-preserved samples contain preservative agents, variable cell concentrations and potential inhibitors, protocol optimization is particularly relevant. Optimization research reveals trade-offs: Reduced elution volume increases the concentration, but may leave residual DNA on the column, while extended incubation improves recovery (10).

To the best of our knowledge, published data on the optimization of elution variables, specifically for the Monarch gDNA Spin Column kit (New England Biolabs, Inc.) using cervical swab clinical samples is relatively limited. Therefore, the Monarch kit was consciously selected due to its technical advantages in fundamental research, providing superior DNA purity, highly concentrated output and specialized protocols, such as HMW DNA protocols for downstream applications (11,12). The present study aimed to evaluate the effects of the following parameters on DNA concentration and purity from HPV ThinPrep cervicovaginal-scrape samples using the Monarch gDNA Spin Column kit: Incubation duration, elution volume and the number of elutions.

Mixed model analyses revealed that the evaluated experimental factors accounted for a small proportion of the variance in each response, as indicated by the R² values shown in Table I. This suggests that other unobserved factors substantially contribute to the overall variability. A notable difference in the modeling results was observed between the LMM for DNA yield and the GLMM for DNA purity. Based on the standardized coefficients and model selection using AIC and likelihood ratio tests (LRT), the removal of individual predictors from the full model did not significantly worsen model fit, as all LRT P-values exceeded the conventional threshold of 0.05. However, among the yield predictors, the number of elutions (e) exhibited the largest change in AIC when removed, suggesting its relatively stronger contribution, while the interaction term between incubation duration (t) and elution volume (v) contributed minimally (Table I).

For DNA purity, the number of elutions exhibited the largest standardized coefficient, indicating a potential role in explaining purity, although its contribution was not statistically significant. The R² values further indicated that fixed effects explained only a small to moderate proportion of the variance, with marginal R² values ranging from 0.098 to 0.0267, while the inclusion of random effects substantially increased the explained variance, with conditional R² values ranging from 0.418 to 0.485 (Table I). This suggests that unmeasured factors beyond the experimental parameters evaluated in the present study may play a substantial role in determining both DNA yield and purity.

Jackknife resampling analyses were performed to evaluate the stability of both mixed-effects models, with detailed resampling results presented in Tables SII and SIII. These analyses demonstrated high agreement with the models derived from the full dataset; specifically, 17 out of 17 resampled models for DNA yield, and 10 out of 17 models for DNA purity, were consistent with the primary full models. These analyses demonstrate that despite the limited sample size and the use of second-order modeling, the results remain robust and stable.

Subsequently, the previously described mixed-effects models were mapped using RSM. It is important to emphasize that, herein, RSM was utilized strictly as an analytical tool to visualize and extract optimal points from the existing second-order mixed models, rather than to fit new independent models. Both response variables were first optimized individually based on the two continuous parameters using their corresponding mixed-effects models. For DNA yield, the RSM analysis identified a stationary optimum at 77 min of incubation duration and an elution volume of 42 µl, producing a predicted yield of 4,420 ng/µl. Although the mixed model for DNA purity lacked statistically significant predictors, its optimum parameters were still assessed via RSM visualization. The optimum point for DNA purity was achieved at 80 min of incubation and an elution volume of 75 µl, resulting in a predicted purity probability of 0.99, indicating a very high likelihood of achieving acceptable DNA purity under these conditions (Fig. 1).

Identifying experimental parameters that satisfy both responses simultaneously, the mean values of the individual optima were calculated and presented in Table II, resulting in a ‘compromised’ condition of a 59-µl elution volume and 79 min of incubation. This averaged parameter set represents a shift from the yield optimum, specifically an increase of 17 µl in elution volume and an increase of 2 min in incubation time, as well as a shift from the purity optimum, namely a decrease of 17 µl in elution volume and a decrease of 1 min in incubation time, thereby providing a balanced trade-off between DNA yield and purity (Fig. 1). Providing a quantitative basis for the optimization, the final fitted second-order polynomial equations for DNA yield and purity obtained from the mixed-effects RSM method are given below:

The present study aimed to identify the optimal DNA extraction parameters (incubation duration (t), elution volume (v) and the number of elutions (e) and DNA integrity analysis from cervicovaginal specimens in ThinPrep buffer, a matrix often characterized by low cellularity and high dilution. The experimental data, albeit derived from a limited sample size, suggest that incubation time, elution volume and the number of elutions each contributed to extraction performance, with the number of elutions exhibiting the most consistent statistical association with DNA yield. These preliminary results may have implications for supporting more consistent DNA recovery in downstream molecular diagnostics. In clinical screening workflows, particularly for specimens with low cellularity, even minor variations in extraction efficiency can influence the success of downstream molecular detection. This is particularly relevant in resource-limited settings where specimen quality is often compromised, though further validation with larger sample cohorts is warranted.

The RSM fitted using the mixed-effects framework, evaluated the effects of incubation duration (t), elution volume (v) and the number of elution cycles (e) on DNA yield and purity. As the predicted optimum conditions for DNA yield and DNA purity occurred at different combinations of incubation time and elution volume, an operational compromise condition was determined by averaging the coordinates of the two individual optima. This approach resulted in a compromise condition of approximately 79 min of incubation and 59 µl of elution volume (Table II), providing a balanced operating condition that maintained satisfactory DNA yield while preserving DNA purity. The higher efficiency of the first elution observed in the present study is consistent with the desorption kinetics of silica membranes, where DNA release is greatest during the initial contact with the elution buffer and decreases in subsequent elution cycles. Although additional elution cycles may increase total DNA recovery the incremental gain becomes progressively smaller indicating diminishing returns (8).

Furthermore, the present study observed a trade-off between total yield and concentration governed by elution volume. Previous research has reported that low-volume elution can produce more concentrated nucleic acid eluates and may improve recovery of low-abundance targets (9). The data presented herein suggest that within the 30-80 µl range, these effects act independently, allowing for flexibility based on the specific requirements of downstream applications. Lower elution volumes which yield higher DNA concentrations are more advantageous for next-generation sequencing (NGS) workflows, whereas higher elution volumes producing more dilute eluates are better suited for PCR-based applications (25,26). This optimization aligns with a broad body of literature [e.g., Akahane et al (27); Shibata et al (6) and Naegele et al (5)] showing that the delicate balance between DNA quantity, purity and concentration is the primary determinant of success in sensitive molecular assays.

Consequently, for laboratories processing ThinPrep specimens with silica-spin kits, prioritizing an extended incubation of ~77 min over increasing elution volumes or adding redundant elution steps to maximize DNA yield may be recommended. For maximizing DNA purity, an extended incubation of ~80 min is recommended instead. While a second elution remains a viable strategy when maximum DNA mass is required for replicate assays, it should be noted that this approach provides diminishing returns and does not substantially alter purity metrics. Ultimately, elution volumes in the mid-range should be selected based primarily on the concentration requirements of the specific downstream workflow, such as NGS or qPCR, as this flexibility allows for standardized, cost-effective protocols that are compatible with existing laboratory infrastructure in resource-limited settings. By maintaining measurable DNA quality and integrity profiles suitable for molecular applications, these optimized extraction conditions may support standardized extraction workflows compatible with downstream molecular analyses.

The optimization results demonstrated that achieving high DNA purity required a relatively larger elution volume compared to conditions that maximized DNA yield or those recommended by the Monarch extraction protocol. This observation is likely influenced by both intrinsic sample quality and the physicochemical behavior of silica-based extraction columns. Low-biomass samples require higher elution volumes to improve DNA release from silica membranes, while insufficient volumes result in incomplete recovery and reduced yield and purity (28,29).

Previous research has reported that increasing elution volume enhances DNA desorption efficiency under low-salt conditions, particularly in silica column-based extraction systems, albeit at expense of DNA concentration (8). Higher elution volumes enhance DNA purity by minimizing co-elution of residual contaminants (28,30). During centrifugation, a larger elution volume can promote more effective spatial separation eluted DNA and contaminants retained within the silica membrane or column matrix, thereby minimizing carryover into the final eluate (29).

In the present study, the purity cut-off values applied for DNA quality assessment were not strictly based on the minimum input specifications recommended by the NGS library preparation kit, but were instead determined through a combination of empirical considerations and evidence from recent literature. Although manufacturers recommend A260/280 (1.8-2.0) and A260/230 (2.0-2.2) ranges, recent studies show long-read sequencing performance depends more on DNA integrity and fragment length than strict spectrophotometric thresholds (12,31). This approach aligns with recent reports suggesting that functional validation through sequencing performance provides a more robust assessment of DNA suitability than reliance on purity ratios alone (5,27).

Furthermore, the findings of the present study highlight that utilizing DNA integrity as a fundamental quality control step reveals a lack of consistent correlation with total DNA concentration. As a quality control indicator, high DIN values are highly preferred; previous research indicates that DIN strongly predicts the success of downstream analyses, especially NGS (32). On the other hand, quality control assessments demonstrating a predominance of short fragments reflect DNA degradation driven by biological and environmental processes, leading to progressive fragmentation (33).

The extraction process parameters, particularly lysis duration and temperature, play a decisive role in preserving long DNA fragments. These factors are particularly critical in PreservCyt samples, where protein-DNA crosslinking requires controlled lysis conditions to prevent additional fragmentation (34,35). In addition, the findings of the present study revealed cases of high DNA concentration that do not exhibit broad fragment size distribution, suggesting that DNA quantity does not necessarily reflect its functional quality. This condition is likely due to the accumulation of degraded DNA, emphasizing the importance of evaluating fragment size distribution alongside quantification (33,36).

Moreover, the observation that increasing elution volume and the number of elution cycles does not consistently improve DNA integrity highlights the limitations of conventional approaches that focus primarily on maximizing DNA recovery. Although yield optimization is often used as a key indicator of extraction success, particularly in limited clinical samples (37), the findings of the present study suggest that such strategies may overlook critical aspects of DNA quality. As previously reported by Simbolo et al (36) short DNA fragments may dominate even when the total yield is high. Therefore, the presnet study expands the current perspective by emphasizing that extraction optimization should balance both DNA quantity and integrity.

The observed variability in DNA quality also points to the influence of pre-analytical factors, such as sample storage and handling, which can accelerate DNA degradation and affect final analytical outcomes (38). In practice, this has a direct impact on the efficiency of downstream application, including sequencing performance. High-integrity DNA is essential for achieving optimal analytical results (35). The heterogeneous DNA integrity observed in the present study may be attributed to variations in storage conditions (4, -20 and -80˚C) and storage durations. These findings are consistent with and further extend those of previous reports indicating that DNA degradation is cumulative and strongly influenced by storage conditions and sample handling (33,38); in the other words, the stability reported in the literature does not entirely eliminate the risk of fragmentation if not accompanied by optimal extraction conditions.

Overall, DNA integrity assessment provides complementary quality-control information beyond yield and purity measurements and may assist in evaluating sample suitability for downstream molecular applications. The optimization analyses suggested that elution conditions were more strongly associated with DNA recovery than incubation duration or elution volume within the tested range. Collectively, the findings support the use of a single-elution workflow and provide a framework for optimizing DNA extraction from ThinPrep specimens, while acknowledging that further validation using downstream molecular assays is required (8). Collectively, these findings confirm that elution protocol primarily determines DNA yield while storage temperature serves as a confounding factor affecting both yield and purity. Based on these results, the recommended protocol for optimal HPV DNA extraction is using the elution 1 method, with an incubation time of 77 min and an elution volume of 41 µl to maximize yield, while storage at -80˚C is advised to maintain DNA purity. While a second elution remains a viable strategy when maximum DNA mass is required for replicate assays, it should be noted that this approach provides diminishing returns and does not substantially alter purity metrics. Ultimately, elution volumes in the mid-range should be selected based primarily on the concentration requirements of the specific downstream workflow, such as NGS or qPCR, as this flexibility allows for standardized, cost-effective protocols that are compatible with existing laboratory infrastructure in resource-limited settings (39,40). By maintaining measurable DNA quality and integrity profiles suitable for molecular applications, these optimized extraction conditions may support standardized extraction workflows compatible with downstream molecular analyses.

The present study provides a framework for optimization, several limitations remain that offer clear avenues for future research. First, while the present study focused on yield and purity, it did not directly measure downstream performance metrics, such as read length distribution, mapping rates, or HPV detection sensitivity. Establishing this direct link via qPCR Ct shifts and NGS coverage remains a critical next step to validate the clinical utility of these optimized parameters. Additionally, although spectrophotometric ratios (namely: A260/A280 and A260/A230) provide essential purity data, they do not fully account for the presence of PCR inhibitors. Future studies incorporating qPCR inhibition assays, spike-in controls, or the testing of supplemental techniques, such as elution buffer warming and the addition of carrier nucleic acids would provide more in-depth biochemical insight into enhancing recovery from samples with critically low cellularity. Finally, as laboratories move toward high-throughput diagnostic settings, a formal cost-benefit analysis will be essential for economic optimization. As noted by Neal et al (41), such low-cost optimizations are increasingly vital for preserving the high-quality DNA required for advanced molecular testing in resource-constrained environments, ensuring that procedural efficiency does not come at the expense of diagnostic accuracy.

In conclusion, the present study demonstrated that the optimization of pre-analytical extraction parameters using linear mixed models identified incubation time as a contributor to DNA yield variation from cervicovaginal samples in ThinPrep buffer without compromising purity. Incubation time and elution volume independently influenced DNA recovery, suggesting opportunities to optimize extraction conditions according to downstream concentration requirements. By prioritizing procedural optimization over increased buffer volume, this approach may improve recovery efficiency, particularly in low-cellularity samples. However, as downstream functional performance was not evaluated, these findings should be considered preliminary recommendations pending validation in diagnostic workflows. Overall, the results presented herein provide a potentially cost-effective strategy to improve DNA extraction efficiency and inform future studies on clinical and molecular performance.