The Effects of The Menstrual Cycle On Growth Factors In Platelet-Rich Fibrin: A Cross-sectional Observational Study

Summary

This study evaluates the effects of menstrual cycle phases on the release of growth factors from platelet-rich fibrin (PRF).

Abstract

Platelet-rich fibrin (PRF) is widely used in dental surgery for regenerative procedures; however, variability in its biological content remains a key limitation for standardization. This study aimed to implement a tightly controlled experimental workflow to evaluate how menstrual cycle phases influence the release of growth factors from PRF. Venous blood samples were obtained from 21 healthy female volunteers at three predefined menstrual phases (menstrual: days 1–5; follicular: days 6–14; luteal: days 15–28), with all collections performed within a fixed time window (11:00–11:30 a.m.) to minimize circadian variation. For each phase, separate tubes were allocated for complete blood count, hormone analysis (estrogen, progesterone, follicle-stimulating hormone [FSH], and luteinizing hormone [LH]), and PRF preparation. PRF was produced by immediate centrifugation under standardized conditions (708 × g for 12 min; fixed-angle rotor, 40 °; radius 88 mm; no anticoagulant). Clots were macroscopically verified (homogeneous, elastic structure), carefully separated from the red blood cell layer, and individually weighed. To standardize growth factor release, each clot was incubated in RPMI-1640 medium using a weight-based ratio (1 g PRF: 1 mL medium), followed by controlled incubation at 37 °C with orbital agitation (100 rpm). Supernatants were collected and stored at -80 °C prior to analysis. Growth factors (IGF-1, PDGF, FGF-2, VEGF, and TGF-β1) were quantified using enzyme-linked immunosorbent assay (ELISA) under uniform assay conditions. Within this standardized protocol, FGF-2 levels were significantly higher during the follicular phase compared to the menstrual and luteal phases. A moderate positive correlation was observed between FGF-2 and estrogen levels in this phase, whereas no statistically significant differences were detected for other growth factors. These findings indicate that menstrual cycle phase may contribute to variability in PRF-derived growth factor profiles. The outlined protocol provides a reproducible framework for minimizing pre-analytical and analytical variability in PRF studies. The clinical implications of phase-related differences warrant further investigation in studies incorporating larger cohorts and treatment outcomes.

Introduction

Historically, the use of platelet concentrates began with the development of fibrin adhesives and evolved into various formulations and preparation protocols. Platelet-rich fibrin (PRF), first introduced by Choukroun et al. in 2001, is a second-generation platelet concentrate widely used in dentistry¹. This autologous biomaterial is obtained through centrifugation without anticoagulants and forms a fibrin matrix enriched with platelets, leukocytes, and circulating immune components². The fibrin network functions as a scaffold that enables the gradual release of growth factors essential for tissue repair and regeneration³.

Platelets play a central role in wound healing and hemostasis by releasing cytokines and growth factors upon activation⁴. In PRF, this process results in the sustained release of key mediators, including vascular endothelial growth factor (VEGF), fibroblast growth factor-2 (FGF-2), platelet-derived growth factor (PDGF), insulin-like growth factor (IGF), and transforming growth factor-beta1 (TGF-β1)⁵. Despite its widespread clinical use, findings regarding the regenerative effectiveness of PRF remain inconsistent across studies^3,6. One major source of variability is the heterogeneity of PRF preparations, which is influenced by both technical and biological factors.

From a methodological perspective, differences in centrifugation protocols (e.g., relative centrifugal force, duration, rotor type), tube materials, and post-collection handling can significantly alter PRF architecture and cellular distribution⁷. In addition, analytical variability—such as lack of standardization in clot processing, inconsistent normalization strategies, and differences in incubation conditions prior to ELISA—can lead to substantial discrepancies in reported growth factor levels⁸. These inconsistencies limit comparability between studies and reduce the translational reliability of PRF research.

Biological variability further complicates interpretation. Several patient-related factors, including age, sex, platelet count, platelet distribution width (PDW), and mean platelet volume (MPV), are known to influence PRF composition⁹. Moreover, hormonal, genetic, and environmental influences may modulate platelet behavior and cytokine release¹⁰. There is both direct and indirect evidence that sex hormones affect platelet biology¹¹. Estrogen, in particular, has been shown to enhance platelet responsiveness to physiological agonists and increase thrombin-induced aggregation through receptor-mediated pathways¹². As estrogen levels fluctuate across the menstrual cycle, platelet function and associated biological outputs may also vary accordingly¹³. However, the literature remains inconsistent, with some studies reporting enhanced platelet activation and others demonstrating neutral or inhibitory effects¹⁴.

Given this combined technical and biological variability, there is a clear need for methodologically controlled workflows that minimize pre-analytical and analytical bias when evaluating PRF-derived growth factors. In contrast to many previous studies that rely on single time-point sampling, unstandardized clot handling, or non-normalized growth factor quantification, the present study employs a phase-specific, time-controlled blood collection protocol, fixed centrifugation parameters, and weight-based normalization of PRF clots prior to incubation and ELISA analysis⁸. This approach is designed to improve reproducibility and allow more reliable comparison of growth factor release across physiological states.

Alternative approaches to PRF research include the use of advanced formulations such as advanced-PRF⁷ and injectible-PRF¹⁵, horizontal centrifugation systems⁷, or direct membrane implantation without ex vivo analysis. While these methods may enhance cellular yield or clinical applicability, they often introduce additional variability in growth factor quantification or limit the ability to isolate specific biological effects. In contrast, the current workflow prioritizes analytical standardization and controlled comparison, making it particularly suitable for investigating physiological modifiers such as hormonal fluctuations.

The proposed protocol is most appropriate for controlled experimental or translational studies aiming to quantify growth factor release under standardized conditions. It may be less applicable in settings focused solely on clinical outcomes, simplified chairside preparation, or studies where immediate PRF application without processing is required. Researchers should consider adopting this workflow when the primary objective is to reduce variability and improve the reproducibility of PRF-related measurements.

Therefore, the aim of the present study was to evaluate the correlation between hormone levels across different menstrual cycle phases and the release of growth factors from PRF using a standardized and reproducible methodological framework.

Protocol

This study was conducted at the Department of Periodontology, Faculty of Dentistry, Çukurova University, Adana, Turkey, with the participation of 21 female volunteers. All participants were informed in detail about the purpose, design, and procedures of the study, and written informed consent was obtained prior to inclusion. The study protocol was reviewed and approved by the Ethics Committee of Cukurova University Faculty of Medicine (Approval No: 125/56, dated 16.09.2022). All procedures were conducted in accordance with the principles of the Declaration of Helsinki for research involving human subjects.

All procedures involving human blood, including biosafety precautions and the handling and disposal of biohazardous and chemical waste, were conducted in accordance with established biosafety guidelines as outlined by the Centers for Disease Control and Prevention and the National Institutes of Health¹⁶. Personnel used appropriate personal protective equipment (PPE) i.e gloves, lab coat, protective eyewear. All sample handling was performed in designated laboratory areas to prevent contamination and exposure. Biological waste (blood residues, PRF remnants, consumables) was disposed of in accordance with Cukurova University Medical Waste Management Protocol and national biomedical waste regulations. The sharp objects were placed in puncture-resistant containers; the liquid wastes were chemically disinfected prior to disposal and the solid biohazard wastes were autoclaved before licensed disposal.

Participant selection

Individuals meeting the following criteria were included in the study: Absence of any systemic illness, regular menstrual cycle of 27–30 day and age between 18–25 years

Individuals meeting the following criteria were excluded from the study: Presence of systemic illness, irregular menstrual cycle, use of any medication or hormonal preparation, presence of endocrine or bleeding disorders and history of excessive bleeding during menstruation

Blood collection and group allocation

The initial days of each participant's menstrual cycle were used to construct a scheduling protocol. Based on widely accepted physiological classifications, the cycle was divided into three phases: Days 1–5 as the menstrual phase (associated with endometrial shedding), Days 6–14 as the follicular phase (characterized by rising estrogen levels and follicular development), and Days 15–28 as the luteal phase (marked by increased progesterone secretion following ovulation)^12,13. Blood samples were collected from each participant during the corresponding phases. Over a single cycle, a total of nine blood tubes were collected per subject: three for complete blood count (one per phase), three for sex hormone analysis (estrogen, progesterone, luteinizing hormone (LH), follicle-stimulating hormone (FSH), and three for PRF preparation and ELISA-based growth factor analysis. All blood samples were collected between 11:00 and 11:30 a.m. at the same time each day to minimize biological variability. Venous blood samples were obtained from the antecubital vein using a sterile vacuum-based blood collection system and 21G butterfly needles to ensure atraumatic sampling. Blood was directly collected into sterile 10 mL glass tubes without anticoagulant (PRF tubes), in accordance with standard PRF preparation protocols. To minimize pre-analytical variability, all samples were collected by the same trained nurse under standardized conditions.

Immediately after collection, tubes were handled gently without agitation and transferred to the centrifuge within 60–90 s to prevent premature coagulation and ensure consistent fibrin polymerization. Tubes were positioned symmetrically within the rotor to maintain balance and avoid vibration-related variability during centrifugation. No additives or anticoagulants were used at any stage of the procedure.

As a result, three study groups were established: Group 1: Samples of the menstrual phase, Group 2: Samples of the follicular phase, and Group 3: Samples of the luteal phase

PRF preparation steps:

Blood samples were immediately centrifuged without anticoagulants using a bench-top centrifuge equipped with a fixed-angle rotor (40 °; radius: 88 mm). Centrifugation was performed under standardized conditions at 708 × g for 12 min at room temperature (22°C–24 °C). Relative centrifugal force (RCF) values were calculated using the standard formula (RCF = 1.118 × 10⁻⁵ × r × g²), taking the rotor radius (88 mm) into account, in accordance with established PRF protocols described by Choukroun et al.¹⁷ and subsequent standardization studies^1,3,18. Following centrifugation, the three distinct layers were identified: a lower red blood cell layer, an upper platelet-poor plasma layer, and an intermediate PRF clot. The PRF clot was gently removed with sterile tweezers, avoiding disruption of the erythrocyte interface. Residual red blood cell components were carefully trimmed using sterile scissors.

PRF clots were macroscopically evaluated for a smooth and homogeneous surface, elastic cylindrical structure, and dense, firm consistency according to predefined criteria^19,20. Clots that did not meet these criteria were discarded, and new blood samples were obtained when necessary. All samples were processed under standardized conditions and frozen at -80 °C within 30 min of collection to minimize proteolytic degradation. Samples were stored for a controlled, uniform duration (up to 4 weeks) before batch analysis to reduce inter-assay variability and ensure comparability of growth factor measurements.

Growth factor ELISA analysis steps:

PRF clots were carefully separated from the red blood cell fraction under aseptic conditions using sterile surgical scissors and were individually weighed using a calibrated analytical balance. RPMI-1640 culture medium was added to each sample at a fixed ratio based on clot weight (1 g PRF: 1 mL medium) to standardize growth factor extraction and minimize variability associated with differences in clot mass. Samples were incubated at 37 °C on an orbital shaker at 100 × g to facilitate homogeneous diffusion of soluble growth factors from the PRF matrix into the culture medium. Following incubation, the supernatants were collected and stored at -80 °C until analysis.

Growth factor concentrations, including insulin-like growth factor-1 (IGF-1), platelet-derived growth factor (PDGF), fibroblast growth factor-2 (FGF-2), vascular endothelial growth factor (VEGF), and transforming growth factor-β1 (TGF-β1), were quantified using commercially available enzyme-linked immunosorbent assay (ELISA) kits according to the manufacturer’s instructions. The following kits were used: Human IGF-1 ELISA Kit, Human PDGF ELISA Kit, Human FGF-2 ELISA Kit, Human VEGF ELISA Kit, and Human TGF-β1 ELISA Kit. All samples and standards were analyzed in duplicate, and absorbance was measured at 450 nm using a microplate reader. Calibration curves were generated using four-parameter logistic (4-PL) regression to determine analyte concentrations⁵.

ELISA steps

Sample dilution was determined according to the expected concentration range and the manufacturer’s instructions for each ELISA kit. Samples expected to fall within the standard curve range were analyzed without dilution. When preliminary readings or expected analyte concentrations exceeded the upper limit of the standard curve, samples were diluted using the sample diluent provided in the kit and reanalyzed. Final concentrations were calculated by multiplying the measured values by the corresponding dilution factor.

Standards (100 µL), prepared by serial dilution according to the FineTest kit instructions, and undiluted or appropriately diluted samples were added to each well of a pre-coated 96-well microplate. The plate was incubated at 37 °C for 90 min to allow antigen binding. All samples and standards were analyzed in duplicate. Following incubation, the liquid was removed without washing, and 100 µL of biotin-labeled detection antibody working solution was added to each well. The plate was incubated at 37 °C for 60 min. After incubation, the wells were aspirated and washed three times using 350 µL of 1× wash buffer per well. The wash buffer was left in the wells for approximately 1 min during each wash cycle to ensure adequate removal of unbound components. Next, 100 µL of horseradish peroxidase (HRP)–streptavidin conjugate working solution was added to each well, and the plate was incubated at 37 °C for 30 min. The wells were washed five times using 350 µL of wash buffer per well to minimize background signal. Then, 90 µL of tetramethylbenzidine (TMB) substrate solution was added to each well, and the plate was incubated at 37 °C for 10–20 min in the dark to allow color development, while the reaction was visually monitored. The enzymatic reaction was terminated by adding 50 µL of the stop solution provided with the kit to each well, resulting in a color change from blue to yellow.

Absorbance was measured at 450 nm using a microplate reader within 5 min after the stop solution was added. A standard curve was generated using the known concentrations provided in the kit, and the curve was fitted using a four-parameter logistic (4-PL) regression model. Sample concentrations were calculated accordingly and corrected for any dilution factors.

Statistical analysis

Numerical variables were summarized as mean ± standard deviation for normally distributed data, and as median with minimum–maximum values when normality assumptions were not met. Categorical variables were expressed as frequencies and percentages. The normality of numerical data was assessed using the Shapiro–Wilk test. For comparisons of repeated numerical measurements obtained from the same individuals at different time points, repeated measures analysis of variance (ANOVA) was applied when parametric assumptions were satisfied; otherwise, the Friedman test was used. Correlations between non-normally distributed numerical variables were evaluated using Spearman’s rank correlation coefficient. Statistical analyses were performed using statistical software (IBM SPSS Statistics version 26.0), and a two-sided p value < 0.05 was considered statistically significant.

Results

No missing data were observed, as all participants completed sampling at all three menstrual phases. Complete blood count parameters were within physiological reference ranges for all samples, supporting the consistency of sample quality and the reliability of PRF preparation.

Evaluation of blood parameters across menstrual phases

A comparison of hematological parameters across menstrual cycle phases is presented in Table 1. White b...

Discussion

The menstrual cycle is a complex physiological process orchestrated by hormonal fluctuations that drive changes in various systems within the female body. In this study, we investigated the content of PRF obtained from blood samples collected during different phases of the menstrual cycle and analyzed the levels of growth factors within this PRF.

The menstrual cycle was divided into three phases based on widely accepted physiological classifications: the menstrual phase (associated with endome...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Analytical balance	Precision balance (0.001 g readability)	Sartorius	Entris
Blood collection tubes	Glass tubes, no anticoagulant (10 mL)	BD	Vacutainer system
Centrifuge	Bench-top centrifuge	Hettich	Universal 320R
ELISA kit – FGF	Human FGF ELISA kit	FineTest	EH0541
ELISA kit – IGF-1	Human IGF-1 ELISA kit	FineTest	ER0030
ELISA kit – PDGF	Human PDGF ELISA kit	FineTest	ER1240
ELISA kit – TGF-β1	Human TGF-β1 ELISA kit	FineTest	EH0287
ELISA kit – VEGF	Human VEGF ELISA kit	FineTest	EH0327
HRP–streptavidin conjugate	ELISA reagent (included in kit)	FineTest	Included in kit
Incubator	Temperature-controlled incubator (37 °C)	Memmert	—
Microcentrifuge tubes	Sterile tubes (1.5–2 mL)	Eppendorf	—
Microplate reader	450 nm absorbance reader	BioTek	ELx800
Needle	21G sterile disposable needle	BD	—
Orbital shaker	Adjustable shaker (100 rpm)	IKA	KS 3000 i control
Personal protective equipment	Gloves, lab coat, protective eyewear	Various	—
RPMI-1640 medium	Cell culture medium	Gibco (Thermo Fisher Scientific)	11875-093
Statistical analysis software	Data analysis software (Version 26.0)	IBM	SPSS Statistics
Stop solution	ELISA stop reagent	FineTest	Included in kit
TMB substrate	Chromogenic substrate for ELISA	FineTest	Included in kit
Wash buffer	ELISA wash buffer (1×)	FineTest	Included in kit