Growth factor and cell content in Platelet-Rich Plasma (PRP), Leukocyte- and Platelet-Rich Plasma (L-PRP), Platelet-Rich Fibrin (PRF) in patients with long bone defects from combat injuries

Abstract

Background

In contemporary regenerative medicine, platelet concentrates (PCs) are actively used as a promising method to support the regenerative process. However, the lack of standardized preparation protocols and the selection of PC types limits their broad clinical implementation, particularly in patients with non-unions or bone defects. Therefore, the aim of this study was to determine the concentration of platelets, leukocytes, and growth factors such as vascular endothelial growth factor (VEGF-A), transforming growth factor beta (TGF-β1), platelet-derived growth factor (PDGF-BB) in three types of PC, such as platelet-rich plasma (PRP), leukocyte- and platelet-rich plasma (L-PRP), platelet-rich fibrin (PRF) using a single donor model in patients with large bone defects after combat trauma compared to healthy individuals.

Methods

Blood for PRP, L-PRP and PRF was collected from 30 participants. 19 healthy volunteers and 11 patients with long bone defects after combat injuries. For the production of three types of PC, 15 ml of blood was taken from each participant. The cellular composition was determined using an automated hematological analyzer. The concentration of growth factors VEGF-A, TGF-β1, PDGF-BB was determined by ELISA.

Results

Differences in cellular composition and growth factor concentration between PC types were identified in all study participants. The concentration of platelets in PCs was distributed as follows: L-PRP > PRP > PRF; however, this did not affect the concentration of growth factors. The concentration of growth factors in PCs from patients with bone defects did not differ from that of healthy individuals. In patients with bone defects, it was not possible to achieve an enrichment of leukocyte concentration in L-PRP compared to the baseline level of whole blood; however, this parameter did not differ from that of healthy individuals.

Conclusions

Growth factor concentrations were similar in patients and healthy individuals, but patients had differences in L-PRP leukocyte enrichment and lower platelet recovery in PRF. This study highlights the need to consider platelet concentrate characteristics when selecting products for regenerative therapy.

Clinical trial number

Not applicable.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12891-025-09346-9.

Background

Bone defects can occur after tumor resection, open fractures, infections, hip replacement and osteomyelitis [1]. Gunshot-induced defects result from high-energy trauma, involving extensive bone and soft-tissue damage as well as bacterial contamination. Prior to reconstruction, patients usually require multiple surgical interventions [2]. There are some main methods of defects reconstruction: bone grafting, distraction osteogenesis, the Ilizarov method, and the Masquelet technique. Nevertheless, the rate of unsatisfactory outcomes with these approaches may reach 25%, highlighting the need for more effective and less invasive treatment strategies [3]. The use of platelet concentrates (PCs) has demonstrated efficacy in the treatment of non-unions and delayed bone healing [4] and may be a promising approach for improving the treatment of patients with bone defects following high-energy trauma [5]. However, this topic has not been sufficiently studied yet. The initial step is to evaluate the cellular composition and the concentrations of growth factors in PCs, since these are crucial regulators of osteogenesis. In such patients, both the body’s reactivity and bone regeneration are significantly altered [6].

PCs derived from the patient’s whole blood are autologous products with a higher-than-normal concentration of platelets and growth factors. PCs initiate tissue regeneration by releasing numerous biologically active growth factors, cytokines, and adhesion proteins [7]. The primary growth factors in PCs include vascular endothelial growth factor (VEGF), transforming growth factor beta (TGF-β), platelet-derived growth factor (PDGF), epidermal growth factor (EGF), and basic fibroblast growth factor (b-FGF) [8]. PCs are classified based on their leukocyte and fibrin content into the following types: (i) pure platelet-rich plasma (pure PRP); (ii) leukocyte- and platelet-rich plasma (L-PRP); (iii) pure platelet-rich fibrin without leukocytes (pure PRF); (iv) fibrin enriched with platelets and leukocytes (L-PRF) [9]. PCs are classified as platelet- and red blood cell-enriched plasma (Red-PRP) as well as platelet-, red blood cell- and white blood cell-enriched plasma (Red-L-PRP) if the red blood cell count in it exceeds 10% [10].

Both laboratory [11–14] and clinical [15] studies have demonstrated that the concentrations of platelets, leukocytes, and growth factors in PCs depend on the specific preparation method. In particular, they vary depending on the use of commercial systems or laboratory test tubes [16, 17], the relative centrifugal force (RCF), or the number of revolutions per minute (RPM) during centrifugation, where RPM is a less accurate parameter due to the failure to account for the radius of the centrifuge rotor [18], among other factors [19].

The concentration of platelets and growth factors in various PCs preparation protocols has been studied primarily in healthy individuals [12, 20–26], often using a single-donor model [27–29]. Similar studies have been conducted involving patients with osteoarthritis, tendinopathy [11, 30], and intervertebral disc herniation [31]. However, this topic remains insufficiently explored in patients with bone defects and non-union of long bone fractures [4], and there is a lack of comparative data with healthy individuals. Only Jiang et al. [32] analyzed the content of growth factors (PDGF-BB, TGF-β1, insulin-like growth factor I (IGF-1), EGF in autologous platelet lysate from a 64-year-old woman with a non-union of the tibia.

Leukocytes in PCs may be significant for the treatment of bone defects due to the specific characteristics of local dysregeneration. Leukocytes in L-PRP and PRP can activate the local immune response and inflammation, thereby promoting tissue regeneration, especially in the treatment of chronic wounds [33] and degenerative spinal diseases [31]. However, their concentration is assessed less frequently compared to platelets and growth factors [12, 34].

In this study, we determine the concentrations of growth factors (PDGF-BB, TGF-β1, VEGF-A) and cellular composition (platelets, leukocytes, erythrocytes) in PRP, L-PRP, and PRF obtained from patients with long bone defects, and to compare these with corresponding parameters in healthy individuals in order to identify differences in the bioactive profile of platelet concentrates.

Materials and methods

The laboratory study was approved by the Local Bioethics Committee (Protocol No. 244 dated 29 Apr 2024) in accordance with the Declaration of Helsinki and conducted from May to November 2024. All participants signed an informed consent form.

Study design

Using a single-donor model, PRP, L-PRP, and PRF were prepared from the blood of each individual donor to minimize the impact of potential variables in the final PCs [27–29] (Fig. 1).

Fig. 1.

Schematic illustration of blood collection and preparation of autologous platelet concentrates (PCs) in single-donor model. Illustration of the steps of whole blood collection, centrifugation, and separation of blood components (leucocytes (WBCs), platelets) from healthy individuals and patients with long bone defects. Platelet-rich plasma (PRP) and Leukocyte- and platelet-rich plasma (L-PRP) were obtained by two-stage centrifugation followed by removal of platelet-poor plasma. Platelet-rich fibrin (PRF) was obtained by one-step centrifugation with the removal of red blood cells. Cellular composition and growth factor levels (PDGF-BB, TGF-β1, VEGF-A) were analyzed for all platelet concentrates

All participants in the study were of Caucasian descent. All underwent screening, which included a complete blood count, serological tests and fibrinogen level measurement. Persons under the age of 18 were not admitted to the study; individuals with cancer, autoimmune diseases; blood dyscrasias; hepatitis B, C, and human immunodeficiency virus; those undergoing treatment with anticoagulants, non-steroidal anti-inflammatory drugs, growth factors, or hormonal medications; as well as pregnant or breastfeeding women.

The group of patients with bone defects included individuals with long bone defects following mine blast injuries: tibia (6), femur (3), humerus (1), ulna (1), all with a duration exceeding 9 months. According to the classification of K. D. Tetsworth et al. [35], these wounds corresponded to critical bone defects: type D3h (large, 4–8 cm; n = 6) and type D3i (massive, >8 cm; n = 5). All patients had infected wounds and associated soft tissue defects. The wounds were closed after several treatments with a negative pressure system. Blood samples for PCs were taken after confirmation of the absence of infection. The patients received treatment at the Department of Musculoskeletal Traumatology of the Institution.

Blood collection and preparation of PCs

For screening via complete blood count, 2 mL of peripheral venous blood was collected from 31 study participants into a Vacumed tube containing K₃EDTA (13 × 75 mm, 2 mL, F.L. Medical S.r.l., Italy) by venipuncture using a 21G needle (Vacumed 0.8 × 38 mm, F.L. Medical S.r.l., Italy). One healthy female participant was found to have thrombocytosis and was therefore excluded from the study.

The PCs study included 30 participants: 19 systemically healthy volunteers (10 men, 9 women) aged 18 to 64 years, and 11 men with long bone defects that formed after a mine blast injuries, aged 21 to 68 years.

To prepare PCs (PRP, L-PRP, PRF), 15 ml of venous blood was collected from each participant in the same manner as for the clinical blood analysis:

1 st and 2nd samples for the preparation of PRP and L-PRP: 5 ml each was collected into vacuum tubes S-Monovette Citrat Na 3.2% (Sarstedt AD&Co.KG). Sodium citrate is the most common anticoagulant for blood collection for PRP. It preserves the viability and functionality of platelets during concentrate preparation [10].

3rd sample for PRF preparation: 5 ml was collected into a Vacumed 13 × 100 mm vacuum blood collection tube with clot activator (CAT) (sterile, red cap, 5 ml, F.L. Medical S.r.l., Italy).

Blood samples were centrifuged using a Nuve NF 800R centrifuge (Nuve, Turkey) equipped with a swing-bucket rotor for horizontal centrifugation at 18 °C. Blood was centrifuged immediately after collection. PRP and L-PRP were obtained by a two-step centrifugation method, while PRF was prepared using single-step centrifugation.

The first centrifugation step was performed at a constant acceleration of RCF 600×g for 10 min to separate the blood into several layers and isolate erythrocytes from the rest of the whole blood volume. In the 3rd sample, after the first centrifugation step, the blood separated into two layers: the upper layer (PRF, ≈ 2.5 ml), a fibrin rich in platelets, was immediately separated before the formation of a fibrin clot, and the lower layer with erythrocytes was discarded (Fig. 2).

Fig. 2.

Visual comparison of three platelet concentrates in tubes: leukocyte- and platelet-rich plasma (L-PRP), platelet-rich plasma (PRP), and platelet-rich fibrin (PRF). PRP and L-PRP appear more transparent due to the absence of fibrin. PRP has a smaller leukocyte-rich layer than L-PRP (thin layer under red color at the bottom of the tube). PRF is denser and opaque because of its fibrin matrix

In the tubes with the 1 st and 2nd samples (for PRP and L-PRP), after the first centrifugation step, the blood separated into three layers: the upper layer with platelets and leukocytes, the intermediate thin layer (buffy coat) with leukocytes, and the lower layer with erythrocytes. To prepare PRP from the 1 st sample, only the upper layer (≈ 2.5 ml) was collected. For L-PRP from the 2nd sample, both the upper and intermediate layers (≈ 2.6 ml) were collected, consisting of the thin leukocyte ring with inclusion of a thin layer of erythrocytes (RBCs). The two collected samples were transferred into empty sterile plastic Falcon-type tubes of 15 ml (Plasti Lab, Lebanon) for the second centrifugation step.

The second centrifugation step was performed at a constant acceleration of RCF 1600×g for 7 min to concentrate the cells in the collected samples (PRP and L-PRP). After centrifugation, platelet-poor plasma, which made up 2/3 of the volume, was removed from both samples. The bottom 1/3 layer was PRP (≈ 0.9 ml) containing platelet granules and a low leukocyte concentration or L-PRP (≈ 0.9 ml) containing leukocytes and platelets, depending on the sample (Fig. 2).

The obtained PRP, L-PRP, and PRF were suspended and gently shaken for immediate platelet and leukocyte counting using a hematology analyzer and/or hemocytometer. For enzyme-linked immunosorbent assay, the samples were separated, then activated with 10% CaCl₂ in a 1:10 ratio to initiate the release of bioactive molecules and the formation of a matrix by fibrinogen cleavage [11], incubated for 30 min at 37 °C, and then frozen at − 20 °C according to the manufacturer’s instructions for immunoassay.

Blood cell counting

The total number of blood cells was determined in whole blood, PRP, L-PRP, and PRF. Whole blood analysis and platelet (PLT) and leukocyte (WBC) counts in PRP and L-PRP were performed using an automated hematology analyzer (Mindray BC-5000, Shenzhen Mindray Bio-Medical Electronics Co., Ltd., China) and re-counted using a hemocytometer. PRP and L-PRP (20 µl each) were diluted with Diluent M-52 D at a 1:25 ratio [36] and counted in pre-dilution mode (PD-CBC/pre-dilution count blood cells). The internal consistency of platelet concentration measurements obtained using manual and automated methods was evaluated by Cronbach’s α, which yielded a value of 0.84, indicating good reliability.

Platelet counting in PRF samples was performed in the fluid mechanically separated from the fibrin clot (clot exudate) using a hemocytometer (KS-NEU-001, Hemocytometer Neubauer Counting Chamber, Kyrios-Soter Scientific, Germany) and a light microscope of the Granum L20 series (Ningbo Yongxin Optics Co., China) equipped with a phase-contrast microscopy set at ×400 magnification. An automated hematology analyzer was not used due to the high viscosity of PRF. WBC and RBC counts were not performed in PRF, as these cells are only present in minimal amounts and are not reliably quantifiable by our method.

The platelet recovery rate (PRR, %) was calculated in PRP and L-PRP to assess the relative increase in platelet concentration compared to whole blood in corresponding volumes using the formula [37]:

VEGF-A, TGF-β1, PDGF-BB concentration

All samples were thawed at room temperature. Quantitative analysis of TGF-β1, PDGF-BB, and VEGF-A in PCs was performed using enzyme-linked immunosorbent assay (ELISA) kits (IBL International GmbH, a Tecan Group company, Hamburg, Germany) according to the manufacturer’s instructions.

Statistical analysis

Data are presented as median and interquartile range (IQR) (Q1-Q3) and were analyzed using SPSS 23 (SPSS Inc., Chicago, USA). Distribution in samples was assessed using the Shapiro–Wilk test. To assess the internal consistency of the platelet concentration measurements by manual and automated method, Cronbach’s alpha analysis was performed. Differences between PCs were determined using Friedman’s ANOVA (analysis of variance). Related-samples Wilcoxon signed rank test was performed for two-group comparison. Comparisons between the healthy men and patients with bone defects were performed using the Mann–Whitney U test. Linear correlations between platelets, leukocytes, and growth factors in PCs were determined by Spearman’s test (r). The strength of correlation was interpreted according to the following thresholds: correlations < 0.20 were considered very weak, 0.20–0.39 weak, 0.40–0.59 moderate, 0.60–0.79 strong, and > 0.80 very strong. Effect size (r) was calculated as rank-biserial correlation based on the Mann–Whitney U test (for independent samples) and the Wilcoxon signed-rank test (for related samples). Values of r were interpreted as small (0.1–0.3), medium (0.3–0.5), and large (> 0.5) according to Cohen’s guidelines. The effect size was predominantly large (r > 0.5) for all parameters when comparing different platelet concentrates. In comparisons between healthy men and patients, the effect size was medium for blood cell counts in L-PRP and VEGF in PRF, and small for all other comparisons. A detailed table of effect sizes is provided in the Additional file 1 (Table S1-S2). A significance level of p < 0.05 was considered.

Results

Subject characteristics

All patients with bone defects were male (n = 11), so the comparison group consisted of 10 healthy males (aged 18 to 45 years; 35 (IQR 27,5–42,5)) of a similar age (p = 0.07) (Table 1). Body mass index was normal in all groups and showed no significant difference between healthy men and patients (p = 0.39).

Table 1.

Demographic characteristics

Parameters	Groups			p-value
	Healthy individuals, n = 19	Healthy men, n = 10	Patients, n = 11
Sex, female, %	9 (47%)	0	0	–
Age	31 (25–40.5.5)	27.5 (24.3–30.3)	35 (27.5–42.5)	0.07
Height, cm	175 (168–187)	187 (182.8–187.8.8.8)	178 (176–184)	0.02
Weight, kg	71 (60.5–82)	82 (74–84.5.5)	75 (70.5–81)	0.56
BMI, kg/m2	23 (20.4–24)	23.9 (22.5–24)	23.9 (22.9–24.6)	0.39

Data are presented as median and interquartile range. U Mann–Whitney test: p, patients vs. healthy men. BMI-body mass index

Cell composition of the platelet concentrates

The clinical blood test parameters in the 30 study participants were within the reference range (Table 2).

Table 2.

Cellular composition of blood and platelet concentrates

Parameters	Groups			p1-value
	Healthy individuals, n = 19	Healthy men, n = 10	Patients, n = 11
Whole blood
HGB, g/l	149 (136.5–159.8.5.8)	159.5 (153.3–162)	132 (119.5–151)	0.01
MPV, Fl	9.6 (8.7–10.1)	9.5 (9.2–9.8)	8.1 (7.4–8.7)	0.04
FG g/L	3.9 (3.5–5.2)	3.6 (2.9–4.9)	4 (3.2–5.3)	0.35
Platelets,×106/ml
Whole blood	236 (220–252)	230.5 (198–247.8.8)	290 (212–369.5.5)	0.15
PRP	987 (857.5–1235.5.5.5)	899.5 (850.5–1140)	1022 (950.5–1197.5.5.5)	0.31
L-PRP	1170 (983.8–1275)	1134.5 (965.4–1199.8.4.8)	1337 (1078.5–1510)	0.17
PRF	456 (422–531.7.7)	478.5 (412.6–560.4.6.4)	424 (310–502.5.5)	0.28
p2-value	0.001	0.001	0.001
р3-value	0.001	0.001	0.001
p4-value	0.047	0.047	0.03
р5-value	0.01	0.01	0.001
р6-value	0.01	0.01	0.001
Leucocytes,×106/ml
Whole blood	5.6 (4.6–6.2)	5.5 (4.3–6.3)	6.7 (5.8–9.6)	0.02
PRP	1.5 (0.3–4.7)	3.7 (0.6–4.5)	1.5 (0.2–4.1)	0.67
L-PRP	13.5 (10.3–19.2)	14.4 (9.4–19.7)	10 (6.8–14.3)	0.11
р3-value	0.001	0.001	0.002
p4-value	0.01	0.01	0.001
Erythrocytes, ×109/ml
Whole blood	4.7 (4.6–5.2)	5.2 (4.8–5.5)	5.1 (4.3–5.4)	0.28
PRP	0.03 (0.02–0.1)	0.04 (0.02–0.1)	0.03 (0.03–0.1)	0.86
L-PRP	0.3 (0.3–0.4)	0.4 (0.3–0.4)	0.2 (0.2–0.3)	0.04
р3-value	0.001	0.001	0.001
p4-value	0.001	0.01	0.06

Data are presented as median and interquartile range. U Mann–Whitney test: p1, patients vs. healthy men; Friedman ANOVA: p2, differences between platelet concentrates; p3, differences vs. whole blood; p-values from related-samples Wilcoxon signed-rank test: p4, PRP vs. L-PRP; p5, PRP vs. PRF; p6, L-PRP vs. PRF. HGB–Hemoglobin; MPV–Mean Platelet Volume, FG–Fibrinogen, Patients–patients with long bone defects; PRP–platelet-rich plasma, L-PRP–leukocyte and platelet-rich plasma; PRF–platelet-rich fibrin

The platelet concentration differed among the three platelet concentrates L-PRP > PRP > PRF in healthy volunteers (p < 0.001) and patients (p < 0.001) (Table 2). In L-PRP and PRP of healthy individuals and patients, the platelet concentration was significantly higher (p < 0.001) compared to PRF, and it was higher in L-PRP than in PRP (p = 0.02; p = 0.03). In patients, the platelet concentration in all concentrates did not differ from that in healthy men (Fig. 3c), whereas the platelet concentration in whole blood was 1.2-fold lower (p = 0.04). The platelet concentration in all concentrates increased compared to whole blood: L-PRP > PRP > PRF (p < 0.001): in healthy individuals by 4.9-fold (p < 0.001), 4.1-fold (p < 0.001), and 1.9-fold (p < 0.001); in healthy men by 4.9-fold (p = 0.01), 3.9-fold (p = 0.01), and 2.1-fold (p = 0.01); and in patients by 4.6-fold (р=0.003), 3.5-fold (р=0.003), respectively, the platelet concentration in PRF was comparable to that in whole blood (p = 0.06) (Table 2; Fig. 3a, b).

Fig. 3.

Concentration of platelets and leukocytes in platelet-rich plasma (PRP), platelet- and leukocyte-rich plasma (L-PRP), platelet- and leukocyte-rich fibrin (PRF) in healthy individuals and patients with long bone defects after combat trauma. а in healthy individuals the platelet concentration in PRP, L-PRP, and PRF was higher than in whole blood; b), in patients platelet concentration was higher in PRP and L-PRP than in whole blood; c) no difference in platelet concentration was observed between healthy men and patients in PRP, L-PRP, and PRF; d) leukocyte concentration was higher in L-PRP and lower in PRP than in whole blood; e) leukocyte levels in patients were lower in PRP than in whole blood; f) no difference in leukocyte concentration was observed between patients and healthy men in PRP and L-PRP. Related-samples Wilcoxon signed rank test between two related groups (a, b, d, e); Mann-Whitney test comparison between healthy men and patients (c, f). **p < 0.01; **p < 0.001

The platelet recovery rate in healthy individuals was 1.14 times higher in L-PRP compared to PRP (89,4 (IQR 78,9–89,4)% vs. 78,7 (IQR 65,7–79,7)%; p = 0.013), while in PRF (78,7 (IQR 65,7–79,7)%), it did not differ from L-PRP or PRP (p = 0.18, p = 0.69). In patients, the platelet recovery rate in L-PRP was 1.3 times higher (p = 0.03) compared to PRP and 1.6 times higher (p = 0.02) compared to PRF (78,8 (IQR 47,1–156,6)% vs. 58,6 (IQR 45,6–100,2)% vs. 59 (IQR 43,4–81,8)%), while PRP and PRF showed similar values (p = 0.08). The platelet recovery rate in patients did not differ from healthy men in PRP and L-PRP (PRP: 58,6 (IQR 45,6–100,2)% vs. 82,2 (IQR 67,3–92)%, p = 0.28; L-PRP: 78,8 (IQR 47,1–156,6)% vs. 90,9 (IQR 84–92,8)%, p = 0.31), while in PRF it was 1.5 times lower (59 (IQR 43,4–81,8)% vs. 88 (IQR 70,2–101,4) %, p = 0.01).

The leukocyte concentration in L-PRP compared to whole blood was 2.4 times higher (p < 0.001) in healthy individuals, while in patients, it did not differ (p = 0.25); in PRP, it was 3.7 times lower (p = 0.001) in healthy individuals and 4.5 times lower (p = 0.01) in patients (Table 2; Fig. 3d, e). In L-PRP compared to PRP, the leukocyte concentration was 9 times higher (p = 0.01) in healthy individuals, and 6.7 times higher (p = 0.001) in patients. The leukocyte concentration in whole blood was 1.2 times higher (р=0.02) in patients, while in L-PRP and PRP (p = 0.11; p = 0.67) it did not differ from healthy men (Table 2; Fig. 3f).

In all groups, the erythrocyte concentration was lower in PRP and L-PRP compared with whole blood (healthy individuals: p = 0.001; p = 0.001; healthy men: p = 0.01; p = 0.01; patients: p = 0.003; p = 0.003) (Table 2). In patients, the erythrocyte concentration in L-PRP was 2-fold lower (p = 0.04) compared with healthy men. The hemoglobin concentration in patients’ whole blood was 1.2-fold lower (p = 0.01) than in healthy men (Table 2).

Growth factors

The VEGF-A concentration in L-PRP and PRF was higher compared to PRP in both healthy individuals (p = 0.001, p = 0.01) and patients (p = 0.02, p = 0.04), while L-PRP did not differ from PRF (p = 0.97, p = 0.93) (Fig. 4a, d). The VEGF-A concentration in all PCs from patients did not differ from healthy men (p = 0.59; p = 0.18; p = 0.31) (Table 3; Fig. 4d).

Fig. 4.

Vascular endothelial growth factor (VEGF-A), transforming growth factor beta (TGF-β1), platelet-derived growth factor (PDGF-BB) in platelet-rich plasma (PRP), leukocyte- and platelet-rich plasma (L-PRP), platelet-rich fibrin (PRF). Healthy individuals (a-c): (a) VEGF-A is significantly higher in L-PRP and PRF compared to PRP; (b) PDGF-BB is significantly higher in L-PRP compared to PRP; (c) TGF-b1 is significantly higher in PRF than in L-PRP. Healthy men vs. patients (d-f): no significant differences for all growth factors. Patients: d-e) VEGF-A and PDGF-BB are significantly higher in L-PRP and PRF compared to PRP; f) TGF-b1 did not differ between PRP, L-PRP, and PRF. Related-samples Wilcoxon signed rank test between two related groups (a-f). Mann-Whitney test comparison between healthy men and patients. *p < 0.05; **p < 0.01; ***p < 0.001

Table 3.

Concentrations of growth factors (pg/ml) in the platelet concentrates (PCs)

PCs	Groups			p1-value
	Healthy individuals, n = 19	Healthy men, n = 10	Patients, n = 11
VEG-А, pg/ml
PRP	38.4 (28.3–60.3)	47.6 (31.1–84.3)	61.2 (48.3–93.3)	0.31
L-PRP	115.7 (55.3–303.4.3.4)	173.3 (45.5–212.5.5.5)	203.4 (71.1–348.5.1.5)	0.59
PRF	99.8 (51.7–165.3.7.3)	73.8 (48.9–148.4.9.4)	195.8 (62.9–238.6.9.6)	0.18
p2-value	0.001	0.001	0.15
р3-value	0.001	0.03	0.02
p4-value	0.01	0.20	0.04
p5-value	0.97	0.29	0.93
PDGF-BB, pg/ml
PRP	342 (291–585)	585 (414.5–1184)	508 (258–768)	0.38
L-PRP	930 (586.5–1225)	1000 (784–1646.5.5)	1634 (531–2793)	0.86
PRF	962 (527–1682)	794 (405.5–1064)	1586 (801–1748)	0.27
p2-value	0.02	0.01	0.01
р3-value	0.04	0.67	0.01
p4-value	0.14	0.79	0.03
P5-value	0.97	0.45	0.66
TGF-β1, pg/ml
PRP	750 (616.5–993)	757.5 (669–983.3.3)	975 (730.5–1798.5.5.5)	0.75
L-PRP	940 (763.5–1278)	1218 (793.5–1300.5.5.5)	1659 (940.5–1905)	0.35
PRF	1752 (885–2128.5.5)	1804.5 (1205.3–2578.5.3.5)	1809 (1611–2985)	0.88
p2-value	0,10	0.27	0.69
р3-value	0.17	0.88	0.79
p4-value	0.06	0.51	0.08
p5-value	0.01	0.047	0.33

Data are presented as median and interquartile range. U Mann–Whitney test: p1, patients vs. healthy men; Friedman ANOVA: p2, differences between platelet concentrates; p-values from related-samples Wilcoxon signed-rank test: p3, PRP vs. L-PRP; p4, PRP vs. PRF; p5, L-PRP vs. PRF. VEGF-A–vascular endothelial growth factor-A, TGF-β1–transforming growth factor beta1, PDGF-BB–platelet-derived growth factor-BB, PRP–platelet-rich plasma, L-PRP–leukocyte-platelet-rich plasma; PRF – platelet-rich fibrin

The concentration of PDGF-BB in L-PRP compared to PRP was higher by 2.7 times (p = 0.040) in healthy individuals, while in patients, it was higher by 3.2 times (p = 0.01) and 3.1 times (p = 0.03) compared to PRF (Fig. 4b, e). In healthy individuals, the concentration of PDGF-BB in PRF did not differ from PRP or L-PRP (p = 0.14; p = 0.97), while in patients, it differed only from L-PRP (p = 0.66). In patients, PDGF-BB concentrations in all PCs did not differ from healthy men (p = 0.86; p = 0.28; p = 0.38) (Table 3; Fig. 4e).

The concentration of TGF-β in healthy individuals was 2.3 times higher (p = 0.01) in PRF compared to L-PRP, and in PRP, it did not differ from L-PRP or PRF (p = 0.17; p = 0.06) (Fig. 4c). Meanwhile, in patients, no differences were found between the PCs (p = 0.79; p = 0.08; p = 0.33), and no differences were observed with healthy men (p = 0.75; p = 0.35; p = 0.86) (Table 3; Fig. 4f).

Correlations

No correlation between platelet concentrations and growth factors in all three PCs was found in both healthy individuals and patients (Table 4). A significant correlation between WBC and VEGF (r = 0.48, p = 0.04) was found for PRP in healthy individuals and healthy men (r = 0.77, p = 0.01) (Fig. 5), whereas for other platelet concentrates and growth factors, the correlations were not significant (Table 4).

Table 4.

Spearman’s correlation (r) between platelets, leucocytes and growth factors in platelet concentrates (PCs)

	PCs	VEGF-A		PDGF-BB		TGF-b1
		r	p-value	r	p- value	r	p- value
Healthy individuals, n = 19
Platelets	PRP	−0,18	0,457	0,29	0,221	−0,26	0,283
	L-PRP	0,12	0,616	0,22	0,059	0,23	0,983
	PRF	−0,25	0,293	−0,21	0,383	−0,06	0,819
Leucocytes	PRP	0,48	0,037	0,16	0,729	0,45	0,163
	L-PRP	0,08	0,732	0,29	0,219	0,40	0,088
Healthy men, n = 10
Platelets	PRP	0,18	0,627	−0,47	0,174	−0,37	0,293
	L-PRP	0,06	0,881	0,43	0,214	0,79	0,829
	PRF	−0,14	0,700	−0,27	0,446	−0,22	0,533
Leucocytes	PRP	0,77	0,009	0,54	0,108	0,46	0,187
	L-PRP	0,29	0,405	0,01	0,987	0,33	0,347
Patients, n = 11
Platelets	PRP	0,12	0,729	−0,07	0,905	0,04	0,832
	L-PRP	0,12	0,719	0,48	0,137	−0,22	0,509
	PRF	−0,64	0,853	−0,25	0,467	0,61	0,47
Leucocytes	PRP	0,03	0,937	0,49	0,537	−0,21	0,120
	L-PRP	−0,02	0,958	0,29	0,385	0,26	0,264

VEGF-A Vascular endothelial growth factor, PDGF-BB Platelet-derived growth factor, TGF-β1 Transforming growth factor beta

Fig. 5.

Spearman’s correlation (r) between leukocytes and platelet vascular endothelial growth factor (VEGF-A) in platelet-rich plasma (PRP). a Healthy individuals (r = 0.48, p = 0.04); (b) Healthy men (r = 0.77, p = 0.01)

Discussion

Bone regeneration is a complex multiphase biological process involving the coordination of cellular and molecular mechanisms. TGF-β and bone morphogenetic proteins (BMPs) play an important role in differentiation, cell division, adhesion, migration, organization and apoptosis. VEGF, when administered exogenously, participates in the early stages of blood vessel formation and causes rapid vascular growth. PDGF-BB enhances the proliferation of fibroblasts, which affects collagen synthesis. IGF-I improves bone formation through proliferation and differentiation of osteoblasts. EGF stimulates the division of epithelial, endothelial and fibroblast cells. It participates in wound healing and tissue regeneration [38, 39]. PCs may be promising for bone and tissue regeneration, especially in patients with gunshot wounds, due to their high content of growth factors [5]. There are some studies demonstrating the successful use of PCs for the treatment of wounds [40, 41] However, to our knowledge, no studies have been conducted on its use for the treatment of bone defects. This is also the first study comparing platelet cell composition and growth factors in such patients.

Protocols for obtaining platelet concentrates (PCs) using commercial systems or manual methods allow for the production of products with different compositions in various medical fields [27, 42–44]. Comparing different types of PCs is crucial for developing scientifically grounded protocols with high clinical efficacy and for the standardization of this therapy in medical practice. A clear definition of the types of PCs will enable clinical trials involving patients with specific diseases, thereby enhancing the accuracy of the results obtained.

In our study, using a single donor model, we prepared three platelet concentrates: PRF obtained by single centrifugation (600 g; 10 min; without anticoagulant), PRP and L-PRP using two-step centrifugation (600 g/1600 g; 10/7 minutes; with anticoagulant). In all healthy volunteers, we achieved a platelet recovery rate of almost 80% or higher from baseline levels in PRP, L-PRP, and PRF, which meets the generally accepted quality standards for PCs [24]. However, in patients with bone defects, the recovery rate was above 80% only in L-PRP, while it was lower in PRP and PRF. The platelet recovery rate in healthy individuals and patients in L-PRP was higher compared to PRP. Meanwhile, in healthy individuals, the recovery rates in L-PRP and PRF were identical, whereas in patients, the recovery rate in L-PRP was higher than in PRF. Accordingly, in PRF, the platelet recovery rate in patients was 1.5 times lower compared to healthy men. Although MPV and haemoglobin values in patients were within normal limits, they were lower than in healthy individuals in the control group. These minor differences could have affected platelet recovery rates in PRF and led to greater platelet destruction [45].

In a study with a PC preparation protocol similar to ours, Kushida et al. [29] used two-step centrifugation (600 g/2000 g; 7 min/5 min) and achieved a threefold increase in platelet concentration in PRP using the commercial Kyocera system. We obtained the highest increase in platelet concentration in healthy individuals and patients with bone defects in L-PRP (4.9-fold and 4.6-fold), lower in PRP (4.1-fold and 3.5-fold), and the lowest in PRF (1.9-fold and 1.5-fold), respectively. In other studies involving healthy individuals [12, 22] and patients with degenerative spinal disorders [31] and osteoarthritis [30], L-PRP and PRP did not differ in platelet concentration.

Depending on the protocol and the tubes used, the increase in platelet concentration during PRP preparation can range from 1 to 6 times [13]. In similar laboratory studies involving healthy volunteers, an increase in platelet concentration from whole blood to PRP was reported to be 1.8 times [20], 2.2 times [28], 5 times [23], 6 times [22], 6.4 times [24], and 8.8 times [25], while in L-PRP, the increase was 4 times [12], 6 times [22], and in PRF, it was 1.5 times [21]. Similar results were obtained in patients with osteoarthritis (PRP and L-PRP: 6.7 times) [30], various orthopedic pathologies (osteoarthritis, tendinopathies) (PRP: 5 times) [11], and degenerative spinal diseases (PRP and L-PRP: 5.7 and 5.9 times) [31]. In our study, we observed an increase in platelet concentration in patients with bone defects in L-PRP by 4.6 times, and in PRP by 3.5 times, which was consistent with the results obtained in healthy men and aligned with the findings of other researchers.

In healthy individuals, the leucocyte concentration in L-PRP was 2.4 times higher than in whole blood and 9 times higher compared to PRP. In patients with intervertebral disc herniation, Zhang et al. [31] reported a similar increase in leucocyte concentration in L-PRP by 3.6 times. However, in patients with bone defects, we did not observe a significant increase in leukocytes compared to whole blood in L-PRP, although the leukocyte concentration in L-PRP was higher than in PRP. It is possible that patient-specific haematological or systemic factors that are not reflected in routine clinical blood tests may influence the L-PRP preparation process. Although patients had normal haematological parameters, standard blood tests may not detect minor changes in leukocyte function, activation status, or distribution between circulating and marginal pools. Such functional or microenvironmental changes may affect the efficiency of separation during centrifugation and explain the lack of leukocyte enrichment in L-PRP obtained from the patient. Recently, machine learning-based models have been developed for rapid assessment of PRP quality, which have shown significant advantages over traditional measurements of blood cells and growth factors [46]. Similar results regarding the difference in leukocyte content between L-PRP and PRP were obtained in other studies involving healthy individuals, where these two concentrates were compared [12, 22], and in patients with osteoarthritis [30].

Growth factors

The concentration of VEGF-A in both healthy individuals and patients was highest in L-PRP and PRF, and lower in PRP. The concentration of TGFβ1 in healthy individuals was highest in PRF; however, in patients, there was no difference between the three PCs. Yin et al. [22] did not find any difference in the concentration of growth factors (VEGF, PDGF-AB, TGF-β1) between PRP and L-PRP in healthy individuals, whereas Y. Kobayashi et al. [12] reported the highest levels of VEGF and PDGF-BB in L-PRP compared to PRP, which is similar to our results for patients with bone defects and healthy individuals. At the same time, the concentration of PDGF-BB in patients was highest in L-PRP and PRF compare to PRP, although this was not found in healthy men. Kobayashi et al. [26] found a higher concentration of PDGF-BB, TGFβ1, and VEGF in PRP compared to PRF in healthy volunteers, while no such difference was observed in our study. Overall, despite the lower platelet recovery in PRF in patients with bone defects compared to healthy men, this did not negatively affect the concentration of the measured growth factors. Platelet recovery is not always related to growth factor levels and depends on the type of anticoagulant. Carvalho A. et al. [47] found in PRP that the level of platelet recovery with EDTA was higher than with citrate as anticoagulant, but the VEGF concentration was lower.

We did not observe any correlation between the platelet concentration in PCs and growth factors (VEGF, TGF-β, PDGF) in all participants. Similar results were obtained in two other studies involving PRP and a larger number of healthy volunteers (>100) than ours [23, 48]. The correlation between growth factors and platelet concentration may be age-dependent in PRP, as observed in a Japanese population [20], but was not detected in healthy Polish volunteers of similar age to those included in our study [49]. This suggests possible variability in platelets across populations. In other studies involving healthy individuals, authors found a correlation in PRP between platelet count and VEGF [22, 27, 49], PDGF, and TGF-β [20, 22, 27]; in L-PRP for VEGF, PDGF, TGF-β [22]. However, Serafini et al. [21] found a correlation between platelets and TGF-β1, PDGF, but not with VEGF in PRF. Lee et al. [11] also observed a correlation between platelets and PDGF-AB, TGF-β in their study of PRP in patients with orthopedic pathology. In our study, although the platelet concentration was higher in L-PRP and PRP compared to PRF, we did not observe a low level of growth factors in PRF. It can be explained by the fact that growth factors in PRF are released not only into the exudate but also remain in the fibrin matrix, which acts as a natural reservoir for their gradual release. Therefore, even with a lower degree of platelet recovery, the structural properties of the fibrin scaffold compensate for this deficiency and allow to maintain stable concentration of growth factors to be maintained [25].

We found a correlation between leucocytes and VEGF for PRP in healthy individuals, but not for L-PRP. Meanwhile, in healthy men, the magnitude of the correlation was higher than in the group of healthy individuals (0.48 vs. 0.77). Similar to our results in three studies [12, 27, 49] showed a correlation for leucocytes and VEGF in PRP in healthy volunteers, and Kobayashi et al. [12] showed it also for PDGF, which we did not observe. It is known that leukocytes contain VEGF, with neutrophils having the highest levels [50]. The predominance of certain leukocyte subpopulations in the concentrates may explain the lack of correlation in L-PRP.

Our study has several limitations. The first limitation of our study was the small sample size. This is due to the uniqueness of the cohort of patients who had long bone defects resulting from mine-blast wounds and measuring more than 4 cm with a follow-up period of 9 months, while such injuries comprise only 12% of long bone fractures with defects [51]. Another limitation of study is the absence of female patients in the defect group. This is mainly due to the fact that most of the patients who participated in the study were male military personnel with combat injuries. The few female patients who were hospitalised during the study period did not meet the criteria for inclusion in this study. This limitation reflects the demographic structure of the treated population. However, we prepared all PCs in a one-donor model [27] to eliminate the influence of physiological or other biological factors on the final product (PRP, L-PRP, PRF). Additionally, we maintained the same conditions for all volunteers regarding the timing of blood collection for PCs, centrifugation parameters, sample processing times, and the type of anticoagulant. This allowed for a clear determination of the differences in cellular composition and growth factor concentrations in PRP, L-PRP, and PRF despite the small sample size. We also did not count the number of leukocytes and erythrocytes in PRF due to their sporadic presence. Another limitation is the lack of leukocyte subpopulation counts. Although we evaluated the leukocyte concentration in PCs, the lack of a complete analysis of the leukocyte composition limits the understanding of the potential impact of different leukocyte subpopulations, such as neutrophils, monocytes, and lymphocytes, as well as the varying composition and proportions of cells on VEGF levels. Future studies should include comprehensive profiling of leukocyte subpopulations to clarify these relationships and better understand how the immune profiles of specific patients influence the biological composition and clinical efficacy of L-PRP.

Conclusions

Growth factor concentrations did not differ significantly between healthy subjects and patients with long bone defects caused by combat injuries, but differences in cell composition were found. In patients, L-PRP did not show leukocyte enrichment, and PRF showed lower platelet recovery rates. These results suggest that although the biological potential in terms of growth factor content is largely preserved, the cellular characteristics of PCs vary and should be considered when selecting the most appropriate product for reconstructive and regenerative therapy. Furthermore, the observed differences between PRP, L-PRP and PRF highlight the importance of evaluating both the cellular composition and growth factor composition in relation to the pathological conditions of the individual patient in order to optimise clinical outcomes.

Abbreviations

BMPs

Bone morphogenetic proteins

b-FGF

Basic fibroblast growth factor

EGF

Epidermal growth factor

FG

Fibrinogen

HGB

Hemoglobin

IGF-1

Insulin-like growth factor I

L-PRP

Leukocyte and platelet-rich plasma

MPV

Mean Platelet Volume

PCs

Platelet Concentrates

PDGF

Platelet-derived growth factor

PLT

Рlatelet

PRF

Platelet-rich fibrin

PRP

Platelet-rich plasma

RBC

Red Blood cells

TGF-β

Transforming growth factor beta

VEGF

Vascular endothelial growth factor

WBC

White Blood Cells

Authors’ contributions

V.H. and V.M.: conceived the study, performed statistical data analysis, and wrote the manuscript. V.M.: data visualization. O.P.: contributed to data collection and interpretation of the results. K.R, P.V., and P.R.: interpreted the data. All authors reviewed and substantially revised the final version of manuscript.

Funding

Not applicable.

Data availability

The datasets analysed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

Approval was obtained from the Bioethics Committee of Sytenko Institute of Spine and Joint Pathology National Academy of Medical Sciences of Ukraine, Kharkiv, Ukraine (Protocol No. 244 dated 29 Apr 2024).

Consent for publication

Not applicable to this study.

Competing interests

The authors declare no competing interests.