A matrix-adapted RNA extraction strategy for targeted RT-qPCR analysis of periodontal ligament fibroblasts cultured on human amniotic membrane scaffolds

Abstract

Introduction

Human amniotic membrane (HAM) is a promising scaffold for periodontal tissue engineering, supporting favorable cellular responses of human periodontal ligament fibroblasts (HPDLFs). However, molecular characterization of cells cultured on HAM is frequently hindered by extracellular matrix–derived contaminants, leading to poor RNA yield and compromised purity that limit downstream gene expression analyses. This study aims to develop and validate a matrix-adapted RNA extraction strategy to enable reliable molecular profiling of HPDLFs cultured on HAM scaffolds.

Methods

HPDLFs were seeded onto de-epithelialized HAM, and cell viability was confirmed prior to RNA isolation. A systematic, multi-phase optimization of a phenol-based extraction protocol (GENEzol™ TriRNA Pure Kit) was undertaken to mitigate matrix-associated interference. RNA yield and purity were assessed spectrophotometrically, with functional validation performed using reverse transcription quantitative polymerase chain reaction (RT-qPCR).

Results

The optimized protocol significantly enhanced RNA integrity and yield compared to the conventional method. Purity indices improved (A260/230: 0.73 ± 0.20 to 2.15 ± 0.40; A260/280: 1.41 ± 0.27 to 2.03 ± 0.18), while RNA concentration increased markedly. The proportion of samples meeting stringent quality criteria improved from 0% to 50%. Notably, mean RNA purity parameters were not significantly different from standard monolayer cultures for the selected measures assessed (p > 0.05).

Conclusions

The approach provides a practical framework for improving RNA recovery and targeted RT-qPCR validation in HAM-based culture systems which could set precedent for improving molecular investigations in periodontal tissue engineering and other scaffold-based models.

Graphical abstract

Highlights

• •Matrix-adapted protocol improves RNA extraction from HAM-based cultures.
• •RNA extraction from HPDLFs on HAM is hindered by polysaccharide-rich matrix , leading to low yield and contamination.
• •An optimized GENEzol™ TriRNA extraction protocol was developed through six iterative phases and validated using RT-qPCR.
• •Optimized method improved RNA purity and yield, with selected purity parameters comparable to standard monolayer cultures.

1. Introduction

The extraction of total ribonucleic acid (RNA) from natural biomaterial scaffolds used in regenerative periodontal applications presents significant technical challenges that limit comprehensive molecular characterization of cellular responses. Human amniotic membrane (HAM), recognized as one of the most promising biomaterials for periodontal regeneration, exemplifies these challenges due to its complex polysaccharide and protein composition [1] that interferes with conventional RNA isolation techniques.

HAM possesses unique biological properties including biocompatibility, anti-inflammatory effects, antimicrobial properties, and regenerative potential [1,2]. Its basement membrane is rich in collagen IV, laminin, and fibronectin, while the stromal matrix contains various growth factors and cytokines that promote tissue repair while minimizing scar formation [2,3]. These characteristics have established HAM as an effective scaffold for human periodontal ligament fibroblasts (HPDLFs), which are critical for periodontal regeneration [4] through their synthesis of extracellular matrix components and formation of the tooth-bone attachment apparatus.

Gene expression analysis has become essential for understanding cellular responses to scaffold materials, differentiation processes, and regenerative mechanisms in tissue engineering applications [5]. Such molecular characterization requires RNA of functional suitability as the foundation for reverse transcription quantitative polymerase chain reaction (RT-qPCR), RNA sequencing, and other downstream applications [6]. However, the use of three-dimensional biological matrices presents a significant obstacle for the acquisition of sufficient functionally competent RNA for biological evaluation.

Most commercial RNA isolation kits utilize acid guanidinium thiocyanate-phenol-chloroform extraction reagents pioneered by Chomczynski and Sacchi [7]. While these methods are effective for monolayer cell cultures, complex biological matrices like HAM present unique challenges. Large polysaccharide fragments can physically entrap nucleic acids and be co-purified during phase separation, leading to low yields [8]. Moreover, small polysaccharide fragments can partition into the aqueous phase and co-precipitate with RNA, resulting in contamination that manifests as poor A260/230 ratios and inhibition of downstream enzymatic reactions [8,9].

Previous studies have documented similar challenges with other scaffold systems. Research on cell/agarose constructs demonstrated that polysaccharide contamination resulted in unusably low RNA integrity numbers (RIN) and required specialized extraction protocols [10]. Studies with collagen matrices and other decellularized tissues have similarly reported the need for modified protocols to achieve adequate RNA quality for molecular analysis [11,12].

Despite the growing interest in HAM for scaffold-based tissue engineering applications, no reliable methods have been established for RNA extraction from HPDLFs cultured on HAM scaffolds. Standard protocols provided by kit manufacturers are optimized for two-dimensional cell cultures and do not address the unique challenges posed by complex natural matrices. Preliminary investigations in our laboratory using conventional extraction methods consistently yielded poor RNA quality, with A260/230 ratios below 1.0 and A260/280 ratios outside the acceptable range for molecular applications (1.8-2.2).

The absence of standardized RNA extraction procedures for HAM-based constructs represents a significant limitation for advancing periodontal tissue engineering research. Without reliable molecular characterization capabilities, researchers are unable to fully evaluate cellular responses to HAM scaffolds, assess expression of genes critical for periodontal regeneration, or optimize scaffold properties for enhanced therapeutic outcomes.

The aim of this study was therefore to develop and validate an optimized RNA extraction protocol specifically designed for HPDLFs cultured on de-epithelialized HAM scaffolds. We systematically evaluated extraction parameters to achieve RNA quality suitable for downstream molecular applications, thereby facilitating more reliable gene expression analysis in HAM-based periodontal regeneration research.

2. Materials and methods

2.1. HAM scaffold preparation and de-epithelialization

Glycerol-preserved human amniotic membrane (5 × 5 cm) was obtained from the Tissue Bank, Universiti Sains Malaysia (USM). Enzymatic de-epithelialization was performed using thermolysin from Geobacillus stearothermophilus (Sigma-Aldrich, USA) prepared in phosphate-buffered saline (PBS) solution from PBS tablets (Sigma-Aldrich, USA). HAM scaffolds were prepared using a modified protocol [13,14] optimized for cell seeding applications. Glycerol-preserved HAM sections were thoroughly washed with 1× PBS until complete removal of glycerol residue was achieved. The membrane was placed in sterile petri dishes and treated with 3 mL of thermolysin solution (125 μg/mL in PBS) for enzymatic de-epithelialization.

Enzymatic treatment was performed on a plate shaker for 10 min at room temperature, followed by mechanical removal of epithelial cells using a sterile cell scraper. The basement membrane side was identified by the presence of cellular debris during scraping and was thoroughly cleaned until complete de-epithelialization was confirmed. Membranes were washed extensively with PBS to remove enzymatic residues and cellular debris.

De-epithelialized HAM was sectioned into uniform 2 × 2 cm pieces using sterile scalpel blades and a standardized template to ensure consistent dimensions. Cut sections were transferred to fresh sterile petri dishes, washed again to remove cutting debris, and pre-equilibrated in complete growth medium for 30 min at 37°C prior to cell seeding.

2.2. Cell culture and scaffold seeding

Human periodontal ligament fibroblasts (HPDLFs, item code CC-7049) were purchased from Lonza Bioscience (Walkersville, MD) and maintained at passage 8-12 for this study. Cell culture was performed using α-Modified Eagle's Medium (α-MEM) with l-glutamine (Gibco™, USA), 10% fetal bovine serum (FBS, Gibco™, USA), and 1% penicillin-streptomycin (Gibco™, USA). Cell detachment was accomplished using TrypLE™ Express (Gibco™, USA). Cells were maintained in T-75 flasks at 37°C in a humidified atmosphere containing 5% CO₂, with medium changes performed every 2-3 days until 70-80% confluency was achieved.

For scaffold seeding experiments, cells were detached using TrypLE™ Express and viability was assessed using trypan blue exclusion in a hemocytometer. Cells were seeded at a density of 2.5 × 10⁵ cells/cm² onto both HAM scaffolds and standard tissue culture plastic as controls. HAM sections (2 × 2 cm) were placed in 6-well culture plates, and 250 μl of cell suspension was carefully pipetted onto the stromal surface of each scaffold to ensure uniform cell distribution. Seeded constructs were incubated for 12 h at 37°C and 5% CO₂ to allow initial cell attachment before adding 1.25 mL of complete medium to each well.

Viability of seeded cells was assessed using a PrestoBlue™ cell viability reagent (Invitrogen™, USA) at days 1, 3, 7, 14, and 21. Following culture periods, HAM scaffold samples and monolayer controls were washed with PBS. PrestoBlue™ reagent was added to HPDLFs and cell-scaffold constructs seeded in 12-well plates and incubated for 2 h at 37°C in a 5% CO₂ humidified incubator. Following incubation, triplicate samples were removed and transferred to a 96-well plate. Complete growth medium without cells was used as background control, and absorbance readings were taken spectroscopically at wavelengths of 570 nm and 600 nm using Sunrise ELISA Reader (DKSH, Germany). The viability of cells was determined by calculating the percentage reduction of PrestoBlue™ reagent, with higher percentage reduction indicating greater numbers of viable cells.

2.3. RNA extraction

The GENEzol™ TriRNA Pure Kit (Geneaid Biotech, Taiwan) was used as the base extraction system with extensive modifications. GENEzol™ reagent is a phenol, chloroform and guanidine isothiocyanate based solution for isolation of total RNA [7]. RNase-free water, molecular-grade absolute ethanol, and diethyl pyrocarbonate (DEPC) were obtained from standard suppliers. RNA quantification and purity assessment were conducted using an Eppendorf BioPhotometer Plus with μCuvette® G1.0 (Eppendorf, Germany), measuring absorbance at 230, 260, and 280 nm.

RNA extraction protocol optimization was conducted through systematic iterative refinement, as conventional protocols designed for monolayer cultures were inadequate for HAM-based constructs. Initial attempts using the standard GENEzol™ TriRNA Pure Kit protocol consistently yielded poor RNA quality characterized by low A260/230 ratios and variable yields, necessitating extensive protocol modifications. Optimization was performed using a sequential and cumulative approach. Each phase was modified based on the RNA yield and purity obtained in the preceding phase, with particular emphasis on improving A260/230 ratios because this parameter was most affected by HAM-associated matrix contaminants. The major procedural variables evaluated included scaffold washing conditions, GENEzol™ volume, cold lysis incubation time, mechanical scraping, debris removal, ethanol incubation, column loading strategy, additional ethanol washing, drying duration, and elution conditions. A summary of the protocol evolution is provided in Table 1.

Table 1.

Summary of sequential RNA extraction protocol modifications.

Protocol phase	Main modification	Primary purpose
Baseline	PBS wash; 400 μL GENEzol™; 5 min room-temperature lysis	Initial scaffold-adapted extraction
Phase 1	RNase-free water wash; 5 min cold lysis	Reduce salt/protein carryover and RNase activity
Phase 2	500 μL GENEzol™; 15 min cold/dark lysis; repeated debris removal	Improve scaffold penetration and debris clearance
Phase 3	Two RNase-free water washes; 30 min cold/dark lysis; 20 min ethanol incubation; additional wash/drying steps	Improve removal of matrix-associated contaminants
Phase 4	700 μL GENEzol™; column loading in aliquots; revised wash/drying order	Improve lysis efficiency and reduce column overload
Optimized protocol	Immediate 80% ethanol rinse following wash buffer; 5.5 min final dry spin	Improve A260/230 purity and reduce carryover contaminants

For the final phase of RNA extractions, HAM-seeded constructs were retrieved, and culture medium was carefully removed. Samples were washed twice with RNase-free water to eliminate residual culture medium components, including serum proteins, salts, and phenol red that interfere with RNA extraction. 700 μL of GENEzol™ reagent was applied directly to each 2 × 2 cm HAM scaffold for direct lysis of seeded cells. This volume, larger than typically recommended for surface area, was necessary to ensure complete penetration of the three-dimensional scaffold matrix. Samples were incubated for 1 min at room temperature followed by 30 min on ice in dark conditions to prevent oxidation and enhance RNase inhibition.

Following lysis incubation, scaffolds were thoroughly scraped using sterile cell scrapers to physically dislodge cells embedded within the HAM matrix. This mechanical disruption was critical for recovering cells that had migrated into the scaffold structure. Scraped lysates were transferred to 1.5 mL RNase-free microcentrifuge tubes and centrifuged at 11,000 rpm for 1 min to precipitate residual scaffold debris. Clear supernatants containing solubilized RNA were carefully transferred to fresh RNase-free tubes.

An equal volume of molecular-grade absolute ethanol was added to cleared lysates and mixed thoroughly. The mixture was incubated on ice for 20 min to enhance precipitation and reduce RNase activity. Ethanol-lysate mixtures were applied to RB spin columns in 700 μl aliquots to prevent membrane overloading, with each aliquot centrifuged at 13,000 rpm for 1 min.

Columns underwent an enhanced washing sequence: 400 μl Pre-Wash Buffer centrifuged at 13,000 rpm for 30 s, followed by two sequential 600 μl Wash Buffer applications, then 500 μl freshly prepared 80% ethanol centrifuged for 2 min, and finally a dry spin at 13,000 rpm for 5.5 min. The additional ethanol wash and extended drying steps were critical modifications that consistently improved RNA purity compared to standard protocols.

For RNA elution, 30 μl of RNase-free water was added to column membranes and incubated at room temperature for 1 min, followed by 3 min on ice. Final centrifugation was performed at 13,000 rpm for 1 min, and eluted RNA was immediately stored at −80°C.

2.4. RNA quality assessment

RNA concentration and purity were assessed using spectrophotometry with the Eppendorf BioPhotometer Plus. A260/280 ratios were used to evaluate protein contamination, with values between 1.8 and 2.2 considered indicative of pure RNA. A260/230 ratios assessed polysaccharide, phenol, and salt contamination, with values ≥ 2.0 considered optimal and values < 1.8 indicating significant contamination [15]. Accordingly, samples with A260/280 ≥ 1.8 and A260/230 ≥ 2.0 were classified as meeting the predefined stringent spectrophotometric purity criteria for this study. RNA integrity was assessed qualitatively by 1% agarose gel electrophoresis based on the presence of 28S and 18S ribosomal RNA bands. Electropherogram-based RNA integrity number (RIN) analysis was not performed in this study.

Functional validation and downstream suitability were evaluated using RT-qPCR amplification of the glyceraldehyde 3-phosphate dehydrogenase (GAPDH) housekeeping gene and selected periodontal or extracellular matrix-associated target genes, POSTN (periostin), PDLs17 (periodontal ligament-associated protein 17), S100A4 (S100 calcium-binding protein A4), and FMOD (fibromodulin). Primer sequences, amplicon sizes, and accession numbers are listed in Table 2. Amplification was carried out using Step-one Plus thermal cycler (Applied Biosystems, Thermo Fisher Scientific) and qPCRBIO SyGreen® 1-Step Detect Hi-ROX Kit (PCR Biosystems, UK). Cycle conditions for RT-qPCR amplification were 40 cycles 95 for 5 s, 60 for 10 s, and 75ᵒ C of 25 s. A master mix containing 5.0 μl 2× SyGreen One-Step Mix, 0.4 μl forward primer (10 μM), 0.4 μl reverse primer (10 μM), 0.5 μl 20× RTase with RNase inhibitor, and 2.7 μl RNase-free water was prepared. For each sample, 1.0 μl of template RNA (0.01 μg) was added to make a total volume of 10 μl. The final primer concentration was 400 nM for both forward and reverse primers.

Tabe. 2.

Primer sequences used for RT-qPCR validation.

Gene	Primer sequence (5′- 3′)	Amplicon size (bp)	Accession number
GAPDHa	F: TGAAGGTCGGAGTCAACGGATTTGGT R: CATGTGGGCCATGAGGTCCACCAC	148	NM_002046.7
POSTNb	F: TCACTTCACACTCTTTGCTCCCAC R: TCCCATAATAGACTCAGAACACTGGAGA	151	NM_001286666.2
PDLs17b	F: TGAGACTAAGGCCCTCACTCCAG R: AATGGAGGAGACTGTGTGTTGAACT	152	NM_001330489.2
S100A4b	F: TTCCACAAGTACTCGGGCAAA R: TGTTGCTGTCCAAGTTGCTCAT	151	NM_019554.3
FMODb	F: AAGGAGGCCAGACAGAGGGA R: GGTCATCTTCATACTGGGCC	128	NM_002023.5

^aHousekeeping gene.

^bTarget gene.

2.5. Statistical analysis

RNA extraction experiments were conducted with biological replicates (n = 6) for each condition tested. Results are presented as mean ± standard error (SE). For validation studies, HPDLFs cultured in standard T-75 flasks served as positive controls, with RNA extracted using the manufacturer's standard protocol. Independent samples t-tests were performed to compare RNA quality parameters between standard monolayer cultures (n = 6) and optimized HAM extractions (n = 6) using SPSS version 26. Levene's test assessed variance equality. Two-way repeated measures ANOVA with Bonferroni corrections was used for PrestoBlue™ cell viability assay. Post hoc tests applied to determine significance between groups (n = 3) at each time point. Statistical significance was set at p < 0.05.

3. Results

3.1. Cell viability

The PrestoBlue™ results demonstrated that viable HPDLFs were present on all HAM scaffold constructs as well as in the monolayer control cultures (Fig. 1). On day 1, similarly low levels of metabolic activity were observed between HAM-seeded constructs (2.05 ± 0.21) and control cultures (2.02 ± 0.22). Peak viability was recorded on day 3, with HAM constructs showing significantly higher (p < 0.001) PrestoBlue™ activity (58.08 ± 2.65) compared to control cultures (45.26 ± 5.44). At later time points, metabolic activity declined in both groups but remained consistently detectable through day 21, with HAM constructs (36.92 ± 3.99) and control cultures (35.13 ± 3.64) showing comparable viability levels.

Fig. 1.

Cell viability assessed by PrestoBlue™ assay demonstrating metabolic activity of HPDLFs cultured on HAM scaffolds and standard monolayer controls over 21 days. Peak viability was observed at day 3 for both groups, with significantly higher activity in HAM-scaffold constructs compared to cell-only controls (***p < 0.001, two-way ANOVA with Bonferroni correction). Data are presented as mean ± SE (n = 3 per group).

Within-group analysis revealed significant increases from day 1 to day 3 (p < 0.001) followed by stabilization through day 21, indicating that HAM scaffolds successfully supported HPDLF viability throughout the culture period. There were no significant differences between HAM constructs and control cultures on days 7, 14, and 21, demonstrating that the scaffold material maintained adequate cellular metabolic activity and viability.

3.2. Protocol optimization and RNA assessment

Through systematic optimization of the RNA extraction protocol, we achieved significant improvements in both RNA yield and purity from HPDLF-seeded HAM constructs. Our comprehensive analysis of 36 independent extraction attempts revealed substantial variation in RNA quality depending on the protocol modifications employed.

3.3. RNA concentration, quality, and purity

Our optimization attempts yielded RNA concentrations ranging from 28.0 to 251.0 μg/mL, with the highest yields achieved using the complete optimized protocol including enhanced washing steps and extended incubation times. The standard manufacturer protocol consistently yielded lower concentrations with poor purity metrics.

3.4. Quality categorization analysis

To systematically evaluate protocol performance, we categorized the 36 RNA samples into six quality groups (protocol phase 1-6) based on combined assessment of concentration, A260/280, and A260/230 ratios.

3.5. Purity assessment

A260/280 ratios, which indicate protein contamination [16], showed marked improvement with protocol optimization. While early extraction attempts yielded ratios as low as 1.25 (indicating significant protein contamination), our optimized protocol achieved ratios ranging from 1.83 to 2.38, with most samples falling within the acceptable range of 1.8-2.2 for pure RNA. Notably, some samples exhibited A260/280 ratios above 2.2 (up to 2.66), which may indicate RNA degradation or DNA contamination. However, these instances were relatively rare and occurred primarily in early optimization attempts before protocol refinement (see Fig. 2).

Fig. 2.

Systematic optimization of RNA extraction from HPDLF-seeded HAM constructs. (A) A260/230 purity ratios showing progressive contamination removal and improvement toward the optimal threshold (≥2.0). (B) A260/280 ratios show improvement toward the acceptable range for RNA purity (1.8-2.2). (C) RNA concentration yields showing increased RNA recovery across the six protocol phases. (D) Success rates at standard and stringent criteria, with cumulative improvement through six-phase protocol development. Data: mean ± SE (n = 6 per group). *p < 0.05, **p < 0.01, ***p < 0.001 vs baseline (protocol phase 1) (Tukey HSD).

The A260/230 ratio proved to be the most critical quality parameter for HAM-based extractions, as it reflects contamination from polysaccharides, phenol, and other organic compounds commonly associated with complex biological matrices [15,16]. Initial extraction attempts using standard protocols resulted in severely compromised A260/230 ratios, with many samples showing values below 1.0, indicating substantial contamination.

Our protocol optimization achieved progressive improvements in A260/230 ratios, with the best results reaching 2.97 - well above the 2.0 threshold considered optimal for pure RNA. The enhanced washing protocol, particularly the additional 80% ethanol wash and extended dry spin, was instrumental in achieving these improvements.

While all 30 samples from Groups 1-5 (baseline through phase 4 protocols) failed to meet stringent quality criteria (A260/280 ≥ 1.8 AND A260/230 ≥ 2.0), the optimized protocol achieved a 50% success rate at these criteria.

The optimization process demonstrated continuous improvement in RNA purity, with mean A260/230 ratios increasing systematically: 0.73 → 1.25 → 1.48 → 1.67 → 1.91 → 2.15, representing a 194% overall improvement.

3.6. Protocol-specific performance analysis

Our optimization process identified several interdependent steps that were critical for successful RNA extraction from HAM-seeded constructs (Table 3). Thorough pre-washing of the scaffold was essential to eliminate residual culture medium components and prevent downstream contamination. Cell lysis efficiency within the dense 3D matrix was markedly improved by increasing the volume of GENEzol™ reagent to 700 μl and extending the incubation period to 30 min on ice, which enhanced both RNA yield and purity compared to standard room temperature protocols. Mechanical cell harvesting through scaffold scraping was necessary to ensure recovery of embedded cells. A subsequent debris removal step was crucial for preventing scaffold fragments from interfering with column binding. Furthermore, an enhanced washing protocol consisting of ethanol wash proved to be one of the most impactful modifications, significantly improving A260/230 ratios. Finally, an extended dry spin of 5.5 min ensured complete solvent removal, further contributing to RNA purity. Together, these modifications contributed to an improved extraction approach that yielded RNA functionally suitable for targeted downstream RT-qPCR analysis.

Table 3.

Systematic protocol development overview.

Protocol phase	n	RNA conc (μg/mL) (mean ± SD)	A260/280 (mean ± SD)	A260/230 (mean ± SD)	Success rate (%) stringent criteria
Baseline	6	46.17 ± 49.12	1.41 ± 0.27	0.73 ± 0.20	0
Phase 1	6	54.33 ± 58.82	1.78 ± 0.11	1.25 ± 0.07	0
Phase 2	6	72.50 ± 78.30	1.86 ± 0.26	1.48 ± 0.09	0
Phase 3	6	134.50 ± 85.24	1.87 ± 0.12	1.67 ± 0.05	0
Phase 4	6	157.00 ± 42.22	2.10 ± 0.34	1.91 ± 0.02	0
Optimized	6	193.00 ± 35.05	2.03 ± 0.18	2.15 ± 0.40	50

Success defined as A260/280 ≥ 1.8 and A260/230 ≥ 2.0.

The optimized protocol improved RNA purity compared with earlier protocol phases, with 3 of 6 optimized samples meeting the combined stringent criteria of A260/280 ≥ 1.8 and A260/230 ≥ 2.0 (Table 4). Although this represents a clear improvement compared with the 0% success rate observed across the earlier protocol groups, the 50% success rate also indicates that some sample-to-sample variability remained. Therefore, the optimized protocol should be interpreted as a substantially improved extraction approach rather than a fully standardized method that consistently yields RNA meeting all stringent purity criteria.

Table 4.

Quality achievement analysis.

Quality threshold	Groups 1-5	Optimized protocol	Improvement
A260/230 ≥ 1.8	2/30 (7%)	6/6 (100%)	Threshold achieved
A260/230 ≥ 2.0	0/30 (0%)	3/6 (50%)	Partially achieved
Combined criteria	0/30 (0%)	3/6 (50%)	Partial combined attainment

3.7. Reproducibility assessment

The optimized protocol demonstrated good reproducibility when applied consistently. Replicate extractions using identical conditions showed coefficient of variation (CV%) values below 15% for both concentration and purity metrics, indicating acceptable technical reproducibility for routine use.

3.8. Quality validation

To validate the optimized protocol, RNA quality was compared with standard monolayer HPDLFs cultures. Standard monolayer extraction consistently yielded RNA of adequate purity with A260/280 ratios of 1.97 ± 0.04 and A260/230 ratios of 2.10 ± 0.06, achieving 100% success rate (6/6 samples meeting stringent criteria).

Statistical comparison revealed no significant differences between optimized HAM and standard monolayer extractions for A260/280 ratios (2.03 ± 0.18 vs 1.97 ± 0.04, p = 0.456) or A260/230 ratios (2.15 ± 0.40 vs 2.10 ± 0.06, p = 0.777) (Table 5; Fig. 4A.). While HAM extraction demonstrated higher variability, reflecting the increased complexity of scaffold-based protocols, mean quality parameters were not significantly different, demonstrating that the optimized method achieves conventional quality standards.

Table 5.

Statistical comparison of RNA quality outcomes between standard monolayer and optimized HAM extraction.

Parameter	Standard Monolayer (n = 6)	Optimized HAM (n = 6)	p-value
A260/280 Ratio	1.97 ± 0.04 (SE: 0.017)	2.03 ± 0.18 (SE: 0.075)	0.456
A260/230 Ratio	2.10 ± 0.06 (SE: 0.026)	2.15 ± 0.40 (SE: 0.164)	0.777

Independent samples t-test. Data presented as Mean ± SD (SE: standard error).

Fig. 4.

Method validation and functional demonstration of the optimized HAM extraction protocol. (A) RNA quality validation against standard monolayer controls showing no significant difference in A260/280 (p = 0.456) and A260/230 (p = 0.777) ratios. (B) RT-qPCR functional validation demonstrates successful GAPDH amplification from both extraction methods based on Ct values (Cell-only: 19.48 ± 0.22, HAM-scaffold: 20.47 ± 0.23 Ct, p = 0.011). Data are presented as mean ± SE (n = 6 per group). ns = not significant, *p < 0.05 (independent t-test).

3.9. RNA integrity and targeted RT-qPCR validation

Agarose gel electrophoresis was used as a qualitative assessment of RNA integrity. Distinct 28S and 18S ribosomal RNA bands were observed in both cell-only controls and HAM-scaffold samples, confirming successful recovery of RNA from both sample types (Fig. 3A). Control monolayer samples showed sharper and more clearly defined ribosomal RNA bands, whereas HAM-scaffold samples showed slightly reduced 28S band intensity and mild background smearing. Amplification products were observed at the expected molecular weight (∼148 bp) in both control and HAM-scaffold groups, confirming that despite minor degradation, the extracted RNA retained functional suitability for downstream RT-qPCR analysis. RT-qPCR functional validation confirmed successful GAPDH amplification from both extraction methods (Standard: 19.48 ± 0.22, HAM-Scaffold: 20.47 ± 0.23 Ct, p = 0.011), demonstrating that the optimized protocol produces RNA suitable for quantitative gene expression analysis despite the 1-cycle difference in template efficiency (Fig. 4B).

Fig. 3.

RNA integrity and functional validation analysis. (A) Agarose gel electrophoresis showing 28S and 18S ribosomal RNA bands from cell-only controls (C1–C6) and HAM-scaffold constructs (H1–H6). While control samples demonstrate superior visual integrity with sharp bands and optimal 28S:18S ratios, scaffold samples show minor degradation but retain structural RNA bands. (B) Agarose gel visualization of GAPDH RT-qPCR amplification products (∼148 bp) confirms successful amplification from both extraction methods despite visual differences. L = 1 kb DNA ladder.

Amplification products were observed at the expected molecular weight (∼148 bp) in both control and HAM-scaffold groups (Fig. 3B), confirming that despite minor degradation, the extracted RNA retained functional suitability for RT-qPCR analysis.

To further evaluate the suitability of RNA extracted using the optimized HAM protocol for downstream gene expression analysis, RT-qPCR validation was extended beyond the housekeeping gene to include periodontal and extracellular matrix-associated targets: POSTN, PDLs17, S100A4, and FMOD. Standard curve analysis showed acceptable linearity across the tested targets, with R² values ranging from 0.917 to 0.995. Amplification efficiencies ranged from 89.412% to 109.451% (Table 6.), indicating generally acceptable amplification performance for targeted RT-qPCR analysis. Melt curve analysis (Fig. 5.) demonstrated dominant target-specific melting peaks for each gene, supporting amplification specificity. These findings strengthen the functional validation of the optimized RNA extraction protocol and demonstrate its compatibility with targeted multi-gene RT-qPCR analysis. The additional target genes were only used to assess amplification performance and specificity of the scaffold-extracted RNA, not to compare biological expression levels between HAM-scaffold and monolayer culture conditions.

Table 6.

RT-qPCR amplification performance and melt curve validation using RNA extracted from HPDLFs cultured on HAM scaffolds.

Gene	R2 value	Efficiency (%)	Melting Tm (°C)
GAPDH	0.995	96.045	86.9–87.0
POSTN	0.953	109.451	80.5
PDLs17	0.917	99.186	79.7
S100A4	0.995	101.876	83.4–83.5
FMOD	0.922	89.412	83–84

Fig. 5.

Melt curve analysis of scaffold-extracted RNA following RT-qPCR validation. Melt curve profiles (n = 3) were generated for (A) GAPDH, (B) POSTN, (C) PDLs17, (D) S100A4, and (E) FMOD using RNA extracted from HPDLFs cultured on HAM scaffolds with the optimized protocol. The dominant target-specific melting peaks support amplification specificity for targeted multi-gene RT-qPCR analysis.

4. Discussion

The extraction of RNA from cells cultured on HAM scaffolds presents unique technical challenges that are not adequately addressed by standard extraction protocols designed for monolayer cell cultures. Conducting systematic optimization approach able to identify and resolve several critical issues that consistently compromised RNA quality in preliminary attempts.

This current work is built upon previous research addressing RNA extraction challenges from polysaccharide-rich scaffolds. Ogura et al. demonstrated similar challenges with agarose-based constructs, where standard TRIzol-based extraction resulted in undetectable RNA integrity numbers due to polysaccharide contamination [11]. Their successful approaches using NTC buffer and plant-specific extraction kits provided valuable insights that informed our optimization strategy.

However, our study addresses a previously unexplored application - the combination of HAM scaffolds with periodontal ligament fibroblasts. This represents a critical gap in literature, as HAM has gained increasing attention for periodontal tissue engineering applications [21,22], yet molecular characterization of cellular responses has been limited by the lack of reliable RNA extraction protocols. The optimized protocol achieved markedly improved results compared to previous studies. This current approach yielded RNA with A260/230 ratios up to 2.97, which likely reflects both our systematic optimization approach and the specific modifications developed for HAM's unique matrix composition. One-way ANOVA revealed significant differences between protocol phases for all parameters (p < 0.001). Post-hoc analysis demonstrated that the optimized protocol achieved significantly higher A260/230 ratios compared to all previous phases (p < 0.001 for Groups 1-4, p = 0.273 for Group 5). Systematic protocol optimization resulted in significant improvements across all RNA quality parameters (Table 3). One-way ANOVA revealed significant differences between protocol phases for RNA concentration [F(5,30) = 5.906, p = 0.001)], A260/280 ratios [F(5,30) = 6.703, p < 0.001)], and A260/230 ratios [F(5,30) = 41.590, p < 0.001)]. The most notable improvement was observed in A260/230 ratios, which increased from 0.73 ± 0.20 (Baseline) to 2.15 ± 0.40 (Optimized), representing a 194% improvement (p < 0.001). Post-hoc analysis revealed that significant improvements were achieved from phase 3 onwards, with the final optimized protocol showing significantly higher A260/230 ratios compared to all previous phases except Modification 4 (p = 0.273). The optimized protocol achieved a 50% success rate (3/6 samples) at stringent quality criteria (A260/280 ≥ 1.8 and A260/230 ≥ 2.0), compared to 0% success (0/30 samples) across all previous protocol variations.

While this outcome represents a substantial improvement over earlier protocol phases, it also indicates that extraction from HPDLF-seeded HAM constructs remains variable. The most significant challenge encountered was contamination from HAM matrix components, particularly polysaccharides and residual proteins from the amniotic membrane's complex extracellular matrix composition. This contamination manifested primarily as poor A260/230 ratios, often below 1.0, indicating the presence of organic compounds that absorb at 230 nm. Such contamination is particularly problematic for downstream applications, as it can inhibit reverse transcriptase and polymerase enzymes used in RT-qPCR and other molecular biology techniques [16]. This variability may reflect differences in scaffold thickness, residual extracellular matrix composition, cell distribution within the scaffold, efficiency of mechanical scraping, and the extent of co-extracted matrix-derived contaminants. These factors are difficult to control fully in natural biological scaffolds and may influence RNA yield and purity even when the same extraction procedure is applied. The systematic approach also revealed that protocol optimization required multiple coordinated modifications rather than single-parameter changes. The Levene's test results (p = 0.008 for A260/230) indicate significant variance heterogeneity between groups, reflecting the reduction in variability achieved through optimization.

Our solution involved a multi-pronged approach addressing each stage of the extraction process. The enhanced washing protocol, particularly the additional 80% ethanol wash followed by extended dry spinning, proved crucial for removing polysaccharide contaminants while preserving RNA integrity. This modification was inspired by similar approaches used for plant tissue RNA extraction, where polysaccharide contamination presents comparable challenges [[17], [18], [19]].

The phase-wise optimization suggests that no single modification alone was sufficient to overcome HAM-associated interference. Instead, improvement appeared to result from the cumulative effect of several coordinated changes targeting different sources of contamination. The scaffold washing and debris removal steps likely reduced carryover of culture medium components and insoluble matrix fragments, while increased GENEzol™ volume and prolonged cold lysis likely improved reagent penetration into the three-dimensional HAM scaffolds. Unlike monolayer cultures where cells are readily accessible for lysis, HPDLFs cultured on HAM migrate into the scaffold matrix, requiring physical disruption to ensure complete cell recovery. Our protocol's mechanical scraping step, combined with increased lysis buffer volume and extended incubation times, addressed this issue. Early modifications, including replacement of PBS with RNase-free water, increased cold lysis incubation, and debris removal, improved purity but did not achieve the predefined stringent criteria. Larger GENEzol™ volume, extended ethanol incubation, additional wash steps, and modified column drying further improved A260/230 values. The final optimized phase, in which the 80% ethanol rinse was applied immediately after the wash buffer steps followed by an extended 5.5 min dry spin, produced the highest mean A260/230 ratio and allowed 3 of 6 samples to meet the combined stringent purity criteria while all samples met acceptable standard criteria. The additional ethanol rinse and extended drying step appeared crucial for improving A260/230 ratios, consistent with reduced carryover of phenol, guanidinium salts, and other organic contaminants. This effect suggests that contaminant removal efficiency is time-dependent and requires immediate intervention before contaminants can re-associate with RNA. However, because the modifications were introduced cumulatively rather than tested independently in a factorial design, the relative contribution of each individual step cannot be quantified.

While monolayer cultures represent the gold standard for RNA extraction with predictable, high-quality yields, scaffold-based systems have historically posed significant challenges that rendered molecular analysis unreliable or impossible. The lack of significant difference in selected purity parameters, despite higher variability in HAM samples (CV: 9.1% vs 2.1% for A260/280), supports the feasibility of obtaining RNA for targeted molecular applications from complex tissue engineered constructs using an optimized extraction approach. This improvement sets precedent for further molecular studies in HAM-based periodontal tissue engineering and understanding cellular responses in scaffold-based regenerative therapies. The systematic approach that has been employed in protocol development provides a model for addressing similar challenges in other scaffold-based tissue engineering applications. The categorization of results into quality groups based on multiple parameters allows for objective assessment of protocol performance and identification of acceptable quality thresholds. However, because formal equivalence testing was not performed, these findings should not be interpreted as evidence of statistical equivalence between scaffold-derived and monolayer-derived RNA.

The apparent discrepancy between visual RNA quality and functional performance revealed in our gel analysis highlights a critical distinction in RNA extraction methodology validation. While HAM-scaffold samples showed minor degradation compared to monolayer controls, with reduced 28S:18S ratios and increased smearing, all samples successfully underwent reverse transcription and PCR amplification with comparable efficiency. This finding demonstrates that spectrophotometric quality metrics (A260/280, A260/230) and functional performance are not always perfectly correlated, particularly in complex scaffold-based systems. The successful RT-qPCR amplification from all samples from the optimized group, despite minor visual degradation, indicates that the optimization protocol preserves RNA functionality. Further quantitative integrity assessment would strengthen its broader application in tissue engineering research. The minor degradation observed may result from prolonged exposure to scaffold matrix components, enzymatic activity during the extended extraction process, or mechanical stress during cell harvesting [20,21].

A limitation of the present study is that RNA integrity was assessed using agarose gel electrophoresis rather than electropherogram-based RIN analysis. Although visualization of 28S and 18S ribosomal RNA bands provided qualitative evidence of RNA integrity, RIN values would have offered a more objective and quantitative assessment of degradation. Therefore, RNA integrity in this study should be interpreted in the context of spectrophotometric purity, agarose gel-based qualitative assessment, and functional RT-qPCR validation. Future studies could incorporate Bioanalyzer or TapeStation based integrity analysis to further confirm the quality of RNA extracted from HAM-based scaffold systems.

5. Conclusions

This study presents a matrix-adapted RNA extraction protocol that improved RNA yield and purity from HPDLFs cultured on HAM scaffolds compared with earlier extraction conditions. The optimized protocol supported qualitative RNA integrity assessment and targeted multi-gene RT-qPCR validation, indicating its functional suitability for selected downstream gene expression analyses. However, further refinement and validation using larger sample sets and quantitative RNA integrity assessment are warranted to improve reproducibility and broaden the applicability of this approach, particularly given the inherent variability associated with RNA extraction from natural scaffold-based systems.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have influenced the work reported in this manuscript.