DFIs are among the most common complications of DM²⁴. The emergence of resistance to commonly used antibiotics has become a threat among patients with DFIs and can lead to treatment failure with poor prognosis^25,26,27. The development of alternative treatment methods that may help prevent or control DFIs could have substantial clinical, social and economic effects. According to some research, PRP is an effective antimicrobial agent that may be beneficial in the treatment of chronic wound or bone infections¹⁵. PRP has major advantages over conventional antibiotic treatment for DFIs, as it is less effective at inducing bacterial resistance. In addition, PRP not only reduces infections but also promotes wound healing.

Conflicting reports exist concerning the application of PRP in the management of infected wounds. Research indicates that plasma may serve as an effective medium for bacterial proliferation and may facilitate the entrapment of microorganisms within the platelet-fibrin matrix following platelet aggregation, potentially shielding pathogens from the effects of antibiotics or leukocytes^28,29. Additional research indicates that PRP exhibits bacteriostatic and/or bactericidal properties against microorganisms^30,31,32,33.

A standardized protocol for PRP preparation is currently lacking, which hinders the ability to compare efficacy across studies¹⁵. Amable et al. investigated fifteen distinct conditions, such as relative centrifugal force, centrifugation time, and temperature, to identify the optimal procedure for achieving the best platelet counts. The findings revealed an increase in platelet count ranging from 0.6 to 5.2-fold, contingent upon the specific conditions observed. This finding highlights that a crucial factor affecting PRP activity is the preparation process designed to achieve optimal platelet counts³⁴. In our study, we utilized a PRP preparation protocol that resulted in an average 3.53-fold increase in platelet count compared to whole blood.

The role of leukocytes in PRP is controversial. Some evidence suggests that adding WBCs to PRP may boost its antimicrobial properties³⁵. However, Anitua found that leukocytes did not significantly improve PRP’s antimicrobial properties³⁶. Since white blood cells release proinflammatory proteases and metalloproteases, higher leukocyte concentration may worsen the inflammatory response³⁷. Thus, we used a method to generate PRP with low leukocyte concentration.

In the present study, PRP activation was conducted immediately prior to application by incorporating 10% calcium gluconate, autologous thrombin, or a combination of both. An autologous platelet-rich gel (APG) made from whole blood contains PRP, calcium, and thrombin. Activated PRP releases GFs, antimicrobial proteins, and inflammatory cytokines. Clean DFUs have been treated with APG for years, improving ulcer healing^{22,38,39,40,41}. Bovine thrombin was initially employed as an activating agent; however, the infrequent yet significant risk of coagulopathy due to antibody formation has limited its regular application. The use of calcium gluconate in the present study offers an alternative means of in vitro activation^42,43.

In the current study, we evaluated the antibacterial efficacy of PRP against three MDR microorganisms isolated from infected DFUs, namely, MRSA, K. pneumoniae, and P. aeruginosa. The three bacterial species examined were selected because they represent the most common causes of DFIs.

To test the antibacterial effect of a substance, we can use different methods such as the disk diffusion method, broth dilution method, agar dilution method, or time‒kill assay⁴⁴. We tested the in vitro antibacterial efficacy of PRP (with and without activation) via the Kirby–Bauer disk diffusion method. Unfortunately, none of our PRP preparations inhibited the growth of the studied MRSA, MDR K. pneumoniae, or P. aeruginosa isolates after 24 h of incubation. The mean value of the inhibition zones against all the tested isolates was only 6.00 mm (disc diameter) for both non-activated and activated PRP-coated empty discs. In addition, there was no increase in the diameter of the inhibition zones for the PRP-coated cefoxitin and ceftazidime discs compared with the uncoated antibiotic discs. Cetinkaya et al. found a 2 mm increase in PRP-coated empty disc diameter against K. pneumoniae and P. aeruginosa but no increase in MRSA inhibition zones⁴⁵. In contrast, Bielecki et al. reported that PRP inhibited the growth of some MRSA strains via the disk diffusion method but had no effect on K. pneumoniae or P. aeruginosa. Moreover, PRP induced in vitro growth of P. aeruginosa⁴⁶. In the previous study, PRP was prepared manually, resulting in a lower mean platelet count (228 × 109/L) compared to the present study. Cetinkaya et al. used an automated device to obtain PRP with concentrations seven times higher than the donor’s baseline platelet count. The median platelet count in PRP was 2208 × 109/L. These findings suggest a potential correlation between elevated platelet counts in PRP solutions and enhanced antimicrobial effects.

Owing to the failure of the disk diffusion method to demonstrate any antibacterial effect of PRP preparations, we tested the broth microdilution method, which in turn showed no antibacterial activity of non-activated PRP against the three tested bacterial species. Furthermore, the activation of PRP led to the formation of a gel that precipitated at the bottom of the wells of microtiter plates. Therefore, the bacteriostatic activity of activated PRPs could not be evaluated via the broth microdilution method. Hence, the bactericidal activity of the activated PRPs was evaluated by plating the tested mixtures onto TSA plates and counting the colonies. However, no bactericidal effect was detected.

Contrary to the findings of Attili et al., which indicated a decrease in the bacterial load of P. aeruginosa due to PRP, Edelblute et al. found no antibacterial efficacy of PRP against P. aeruginosa; however, notable bactericidal activity was detected against 40% of their S. aureus isolates using the broth microdilution technique^33,47. This may be due to the fact that none of the S. aureus strains examined in their investigation were MRSA. Drago et al. indicated that PRP exhibited no efficacy against P. aeruginosa; however, it did provide growth suppression against other bacteria, including Candida albicans, Streptococcus agalactiae, Streptococcus oralis, and Enterococcus faecalis¹¹.

Most P. aeruginosa strains produce one or more extracellular pigments, including pyoverdine (yellow–green), pyocyanin (blue–green), pyomelanin (brown–black), and pyorubrin (red–brown). In our study, wells containing suspensions of P. aeruginosa incubated with PRP presented yellow–green pigmentation compared with the dark–green pigmentation of those without PRP. These findings indicate that PRP led to the production of pyoverdine at the expense of pyocyanin. According to Abdelaziz et al., the synthesis of extracellular pigments is affected by the composition of incubation media, such as nitrogen and carbon sources. Hence, the production of pyoverdine in the present study might be attributed to the presence of organic nitrogen sources in PRP⁴⁸.

Another trial was performed via checkerboard synergy testing to demonstrate the effect of PRP in combination with antibiotics. This method also had indifferent effects on most of the tested isolates (58.49%). Although still non bacteriostatic, it had a synergistic effect on only 12.50% of the MRSA isolates when combined with cefoxitin and on 37.50% of the P. aeruginosa isolates when combined with ceftazidime. However, it showed antagonistic activity against 26.42% of the tested isolates. No further studies that used checkerboard synergy testing of PRP with antibiotics were found in the literature. However, Cetinkaya et al. demonstrated that PRP exhibited a synergistic effect on MRSA when used in conjunction with vancomycin in a rat model of surgical wound infection⁴⁹.

The last trial was performed via a time-kill assay. Finally, this method showed a highly statistically significant antimicrobial effect of PRG (activated PRP) against the three tested MDR strains compared with the controls after 1, 2, and 5 h of incubation. The peak point of effectiveness differed according to the studied microorganism. The efficiency of the PPR decreased after the second hour of incubation for MRSA and MDR P. aeruginosa, and after the first hour for MDR K. pneumoniae. PRP had a peak effectiveness of about 99.50%, 91.35%, and 97.23% against the growth rates of MRSA, MDR K. pneumoniae, and MDR P. aeruginosa, respectively. PRP’s antimicrobial impact lasted up to 5 h. When all of the tested strains were examined after 24 h of incubation, they all demonstrated regrowth and the elimination of the inhibitory effect of PRP, as compared to the controls. Cetinkaya et al. also discovered that PRP had antimicrobial effects on MRSA, ESBL-producing K. pneumoniae, and carbapenem-resistant P. aeruginosa; however, the effects were statistically significant only on MRSA and P. aeruginosa, and only in the first hour of their study, which is consistent with the findings of the current study⁴⁵. Moojen et al. found similar findings, stating that PRP had a strong antibacterial impact that was limited to the first hour after application and could maintain a proportional reduction in bacteria of roughly 99% compared to the control for up to 8 h⁵⁰. According to Hasan et al., the maximal antibacterial activity of PRP against S. aureus was limited to the first hour after application, and the antimicrobial effect peaked at 4 h with an 84.2% reduction in bacterial count. This rate dropped to 76.3% after 24 h¹⁷.

Compared with planktonic bacteria, bacterial biofilms are known to exhibit significantly greater resistance to antimicrobial agents. Therefore, they demand substantially higher concentrations of antimicrobial factors to exert any effect. This heightened resistance is due to the ability of biofilm-producing bacteria to shield themselves within the extracellular polymeric substance (EPS). Antimicrobial agents released by substances find it challenging to penetrate biofilm cultures because of their diffusion-limited transport through biofilm EPS. As these agents pass through, they become diluted, making it difficult for them to reach bacterial colonies^51,52.

In our study, 69.81% of the studied isolates were biofilm producers, and PRP had no effect on biofilm formation or established biofilms. Similar findings were reported by Smith et al.⁵³. In contrast, Gilbertie et al. found that PPR and amikacin had synergistic efficacy against aminoglycoside-tolerant biofilm formations, with stronger activity against gram-positive bacteria⁵⁴. This could be due to a specific “window of opportunity” when a treatment can effectively target biofilms, occurring between the initial adhesion and irreversible binding of bacteria. During this phase, bacteria are vulnerable, but once a biofilm is formed, it becomes resistant to penetration. Consequently, topical PRP application appears ineffective against well-established biofilms. In clinical settings, PRP might be more effective when combined with antibiotic therapy against these firmly established colonies⁵⁵.

In vivo experiments yielded better findings for PRP’s antibacterial action than in vitro investigations. Sun et al. successfully eliminated the infection and achieved wound closure using autologous PG and negative pressure wound therapy in a case involving a diabetic patient and an infected amputation caused by MDR Acinetobacter baumannii that remained unresolved despite extended antibiotic treatments⁵⁶. In a rabbit model of implant-associated spinal infection, Li et al. found that PRP treatment significantly reduced bacterial colonies in bone samples and improved bone healing post-operatively⁵⁷. However, our study indicated that the bacterial inhibition observed, when present, was temporary, suggesting a possible depletion of plasma antibacterial factors. This could be attributed to bacteria being less susceptible to direct topical application of PRP without an immune response providing additional antimicrobial support. These findings have practical implications for the clinical use of blood-derived biomaterials. Particularly noteworthy is the potential exploration of whether regular PRP application to wounds could mitigate the risk of bacterial regrowth and infections. As a result, more rigorous clinical evidence is required to justify the use of PRP in DFIs. Future large-scale prospective randomized controlled clinical trials should be conducted to develop uniform guidelines for the indications, contraindications, and particular methods for PRP use in clinical settings.

In conclusion, only activated PRP effectively inhibited microbial growth, indicating that coagulation activation is a crucial step. Bacterial growth inhibition by PRP was apparent immediately after activation; however, it was temporary and persisted for up to 5 h, varying by bacterial strain, indicating differential susceptibility to or consumption of the antibacterial factors present in the PRP suspension. Consequently, PRP can function synergistically with antibiotics and serve as an additional therapy for infections. Variations in study designs, including PRP preparation, platelet activation, bacterial kind, and the antibiotic resistance profile of bacteria, may account for the disparate results. Nevertheless, the majority of in vitro and in vivo investigations have demonstrated that there are no contraindications for the application of PRP in infected wounds.

Methods

An in vitro experimental study was conducted from April 2021 to May 2022 at the HIPH and the Clinical Pathology Laboratory at AMUH. The study proposal was approved by the Ethics Committee at the HIPH, Alexandria University, Egypt. Informed consent was obrained from each participant in this study. All methods were performed in accordance with the relevant guidelines and regulations for ethics approval and consent to participate.

To collect the isolates required for the study, a total of 78 swab samples were randomly collected from clinically suspected infected DF lesions of patients admitted to the Vascular Surgery and Diabetic Foot Unit at the Surgery Department in AMUH (Fig. 6). Bacterial isolates other than MRSA, MDR K. pneumoniae, and MDR P. aeruginosa were excluded from the PRP experimental study.

Sampling and processing of samples

No antimicrobial agent or antiseptic agent was introduced into diabetic foot wounds before samples were collected. Swab samples were collected via Levine’s technique⁵⁸. The swabs were then replaced into their containers and transported directly without refrigeration to the Microbiology Laboratory at the HIPH.

The collected swabs were inoculated onto blood agar and MacConkey agar plates. The plates were incubated in aerobic conditions at 37 °C for 24 to 48 h. All isolated colonies on the blood and MacConkey agar plates were identified following conventional microbiological procedures⁵⁹. All isolates of S. aureus, K. pneumoniae, and P. aeruginosa underwent antibiotic susceptibility testing using the Kirby-Bauer disk diffusion method on MHA plates⁶⁰. Isolates of S. aureus exhibiting resistance to cefoxitin (30 µg) were classified as MRSA⁶¹. K. pneumoniae and P. aeruginosa isolates exhibiting resistance to at least one agent across three or more antibiotic classes were classified as MDR⁶².

All MDR K. pneumoniae isolates were tested for ESBL production, and bacterial suspensions were prepared via the direct colony suspension method. The bacterial suspensions were subsequently inoculated onto MHA plates. Subsequently, cefotaxime and ceftazidime discs, both individually and in combination with clavulanate (cefotaxime-clavulanate 30/10 µg and ceftazidime-clavulanate 30/10 µg), were placed on each plate, which were then incubated at 35–37 °C for a duration of 16–18 h. Positive ESBL production was noted when a > 5 mm increase in the zone diameter occurred for either antimicrobial agent tested when combined with clavulanate compared to the zone diameter of the agent tested individually (Figs. 7 and 8)⁶¹.

All MDR P. aeruginosa isolates were evaluated for carbapenemase production using the modified carbapenem inactivation method (mCIM), as per the guidelines of the Clinical and Laboratory Standards Institute (CLSI) (Fig. 9)⁶¹.

Determination of the antibacterial activity of PRP

PRP preparation

Blood collection

For PRP preparation, 8–16 mL of venous blood samples were collected from each of 16 adult volunteers into sterile tubes containing the anticoagulant citrate dextrose-adenine (ACD-A). All the volunteers were predominantly in good health. The exclusion criteria included the administration of antimicrobials and/or anti-inflammatory drugs within the preceding 10 days, smoking, pregnancy, systemic diseases, infections, hemoglobin levels below 10 g/dL, and/or platelet counts of 100 × 10^3/µL or lower^12,63.

Procedure of PRP preparation

• Whole blood in the ACD-A tubes was centrifuged at 200–250 × g (1100–1200 rpm) for 10 min (Hettich, Germany). This initial centrifugation, termed ‘soft spin’, facilitates the separation of blood into three distinct layers: a bottom layer of red blood cells (RBCs) comprising 55% of the total volume, a top acellular plasma layer known as platelet-poor plasma (PPP) accounting for 40% of the total volume, and an intermediate layer referred to as the ‘buffy coat’, which contains PRP and constitutes 5% of the total volume.

• The PPP, PRP and some RBCs, specifically the upper two layers and a negligible ‘unavoidable’ quantity from the bottom layer, were placed into a separate tube devoid of anticoagulant.

• This tube was subjected to a secondary centrifugation at 300 × g (1300 rpm) for 15 min, referred to as the ‘hard spin’. This enabled the platelets (PRP) to accumulate at the tube’s base with minimal RBCs, whereas PPP constituted 80% of the volume at the top.

• The majority of the supernatant PPP was extracted using a syringe, and the platelet pellet was reconstituted in a minimal volume of PPP (1.5 ml) using vortexing (Nickel Electro LTD, England)^15,64,65. The produced plasma was identified as the PRP utilized in the study (Fig. 10).

Determination of platelet and leukocyte counts

The count of WBCs and platelets in the PRP for each participant was assessed using a complete blood cell analyzer (Abbott, USA). The outcomes were then compared with the whole blood results of the same volunteer to assess the efficacy of PRP preparation.

Preparation of autologous thrombin

For thrombin preparation, approximately 4 ml of whole blood from each donor was drawn into a serum clot activation tube. The serum clot activation tubes were kept at room temperature for 10 min until clotting. The tubes were subsequently centrifuged at 3000 ×g for 3 min. The top layer (supernatant) was considered thrombin and was transferred to a new tube. This thrombin solution was subsequently used for PRP activation^45,66.

PRP activation

Platelet activation was performed immediately before the use of PRP by adding the following:

Or

Or

• A mixture of autologous thrombin and 10% calcium gluconate at a ratio of 2:1:8 (0.8 ml of autologous thrombin and 0.4 ml of 10% calcium gluconate to each 3 ml of PRP) to produce a platelet-rich gel^17,45.

The experimental samples of PRP were divided into four types:

• Type 1: non-activated PRP.

• Type 2: calcium gluconate-activated PRP.

• Type 3: thrombin-activated PRP.

• Type 4: calcium gluconate with thrombin-activated PRP.

Kirby–Bauer disk diffusion method

The antibacterial activity of PRP was assessed using the Kirby–Bauer disk diffusion method on MHA plates⁴⁵. Using sterile micropipettes, empty discs were coated with one of the following: 10 µL of the experimental PRP or an equal volume of PBS, which was used as a control. The discs were subsequently placed on inoculated MHA plates. Standard 6 mm antibiotic discs of cefoxitin, as well as others coated with the experimental PRP, were placed on the same MHA plates using separate micropipettes for MRSA testing. For MDR K. pneumoniae and MDR P. aeruginosa, one antibiotic from each class to which the tested strain exhibited resistance was applied, both individually and in combination with the experimental PRP, on MHA plates. The test was conducted separately for each of the four PRP types.

The test was conducted multiple times utilizing varying volumes of PRP: 20, 50, and 100 µL. Each type of PRP was tested through direct inoculation onto the media surface using a micropipette, ensuring contact with the microorganisms to minimize variables that could affect their antibacterial actions⁴⁷. The plates were incubated in an inverted position at a temperature range of 35 to 37 °C for duration of 16 to 18 h. The diameters of the inhibition zones were measured using a ruler, and the antimicrobial activity of PRP was evaluated by comparing the zone diameter of the antibiotic disc tested in combination with the experimental PRP to the zone diameter of the disc tested independently (Fig. 2)^45,47.

Broth microdilution method

The antibacterial effect of PRP was assessed via broth inhibition via the microtiter method⁶⁷. The assay was performed as follows:

1. 1.

   Fifty microlitres of each of the four types of PRP (a non-activated type and 3 activated types) were added to each of 4 wells (for each tested isolate).

2. 2.

   Fifty microlitres of bacterial suspension of the tested isolate (1 × 10⁶ CFU/mL) were inoculated into each well containing different types of PRP.

3. 3.

   The following controls were used: sterility control (100 µL of Mueller–Hinton broth (MHB)), growth control (50 µL of MHB with 50 µL of bacterial suspension), and PRP control (50 MHB with 50 µL of PRP).

4. 4.

   The microtiter plate was incubated at 37 °C for duration of 16 to 20 h.

5. 5.

   After incubation, the antibacterial effect of PRP was evaluated by observing the inhibition of visible growth of bacteria.

6. 6.

   The observed turbidity or sediment (button size) indicated that there was no inhibition of visible growth of bacteria.

7. 7.

   Approximately 100 µL from each tested mixture (PRP plus bacterial suspension) and the growth control were plated onto TSA plates and incubated at 37 °C for 16 to 20 h.

8. 8.

   A comparison between the colony counts of the tested mixtures and those of the growth controls on TSA plates was used to determine the bactericidal activity of PRP.

Checkerboard synergy testing

The two-dimensional microdilution method in a 96-well microtiter plate was used to evaluate the antibacterial activity of PRP in combination with antibiotics against resistant isolates. The methodology for checkerboard synergy testing was adapted from previous studies but with some modifications^68,69.

One antibiotic agent to which the bacterial isolates were resistant was tested. Cefoxitin sodium was tested against MRSA isolates, whereas ceftazidime pentahydrate was tested against K. pneumoniae and P. aeruginosa isolates. The minimal inhibitory concentration (MIC) was defined as the lowest antibiotic concentration that inhibited visible growth of the tested isolate, as observed with the naked eye, as shown in Figs. 11 and 12⁶⁷. The experiment was conducted in duplicate for each isolate tested, and if the two MICs varied by more than two wells, the assay was repeated. The MIC of each antibiotic was assessed individually and in combination with PRP.

The combination of PRP with antibiotics was considered synergistic when the MIC of an antibiotic in combination with PRP was lower than the MIC of the antibiotic independently, indifferent when the MIC of an antibiotic in combination with PRP was the same as the MIC of the antibiotic independently, and antagonistic when the MIC of an antibiotic in combination with PRP was higher than the MIC of the antibiotic independently.

Time-kill assay

A time–kill assay was conducted to test the antibacterial effect of activated PRP against MRSA, K. pneumoniae, and P. aeruginosa isolates, where viable bacterial counts were determined for 10 µL aliquots taken from PRP and control tubes at 1, 2, 5, and 24 h⁷⁰. The bacterial growth rate in the PRP tubes was compared with the simultaneous bacterial growth rate in the control tubes. The peak point of effectiveness for PRP against each bacterium was defined as the duration of assessment in the time-kill assay that exhibited the highest rate of bacterial growth inhibition.

Time-kill test procedure

• From freshly grown blood agar cultures, 3 to 5 colonies with a single morphology were touched lightly with a sterile wire loop and suspended in a tube containing 5 mL of prewarmed MHB.

• The bacterial suspension was incubated in a shaker incubator (150 rpm) at 37 °C until turbidity was reached.

• The turbidity of the actively proliferating broth culture in the exponential growth phase was calibrated to achieve a visual comparison with 0.5 McFarland turbidity standards (about 1 × 10^8 CFU/mL).

• Within 15 min of preparation, the adjusted suspension was diluted to 1:100 by adding 100 µL of the adjusted suspension to 9.9 mL of MHB, resulting in approximately 1 × 10⁶ CFU/mL.

• The adjusted dilution (containing 1 × 10⁶ CFU/mL) was further diluted by a factor of 1:10 by adding 100 µl of bacterial suspension to 900 µL of sterile MHB, resulting in a bacterial suspension containing about 1 × 10⁵ CFU/mL.

• One hundred microliters of each bacterial isolate, with controlled colony counts of 1 × 10^5 CFU/mL, were placed into sample tubes. Subsequently, 700 µL of MHB, 160 µL of PRP, and 40 µL of platelet activator (comprising autologous thrombin and 10% calcium gluconate in a 2:1 ratio) were introduced into the tubes, achieving a bacterial concentration of 1 × 10^4 CFU/mL.

• The tubes were placed in a shaker incubator at 37 °C.

• After 1, 2, 5 and 24 h of incubation, the tubes were vortexed, and 10 µL of each mixture was added to 90 µL of sterile saline in a sterile Eppendorf tube via a sterile pipette.

• The mixtures were incubated into TSA plates and incubated at 37 °C for 24 h.

• The same processes employed for the PRP tubes were applied to the control tubes, utilizing sterile saline in place of PRP.

Colony counts

Colonies were enumerated using a colony counter following 24 h of incubation at 37 °C. Colony counts beyond 1000 were documented as 1000 CFU/mL (Fig. 13).

Determination of the antibiofilm activity of PRP

Detection of the biofilm-forming ability of the tested isolates                                                               The biofilm-production ability of the identified isolates was evaluated using the tissue culture plate method, commonly referred to as the microtiter plate (MtP) assay (Figs. 14 and 15). The procedure involved culturing the samples in 96-well microtiter plates, followed by optical density (OD) measurement using a microplate enzyme-linked immunosorbent assay (ELISA) reader after staining the wells^71,72,73. The cutoff OD (ODc) was determined by adding three standard deviations to the mean OD of the negative control. The bacterial isolates were classified according to the measured ODs as follows:

• OD ≤ ODc: non-biofilm producer (0).

• ODc ˂ OD ≤ 2×ODc: Weak biofilm producer (+).

• 2×ODc ˂ OD ≤ 4×OD: Moderate biofilm producer (++).

• 4×ODc ˂ OD: Strong biofilm producer (+++).

Effect of PRP on biofilm production (biofilm Inhibition assay)

The inhibition of biofilm formation (by the biofilm-producing isolates) was assessed via the same procedure for detection of biofilm-production ability, except that 100 µL of activated PRP was added to 200 µL of diluted bacterial suspensions in each well (Figs. 16 and 17)^71,72.

The absorbance values were measured with a microtiter plate reader at 630 nm. The extent of biofilm inhibition was determined in relation to the biofilm quantity developed without PRP (designated as 100% biofilm) and the media sterility control (designated as 0% biofilm).

It was observed that by inverting the plate to remove fluid from the wells, thick platelet gels remained at the bottom (leading to deep staining of most of the wells). Accordingly, abovementioned test was repeated but without staining, and assessment of viable biofilm cells by colony count was performed as follows⁷¹:

1. 1.

   Two hundred microliters of tryptic soy broth (TSB) were added to the wells, and biofilm cells were suspended by vigorous pipetting.

2. 2.

   The suspended biofilm was transferred to a new 96-well flat bottom microtiter plate.

3. 3.

   One hundred microliters from each well were plated onto a TSA plate and incubated for 24 h.

4. 4.

   The plates were then examined, and the colony count was recorded.

5. 5.

   The number of colonies from the PRP wells were compared with those from the corresponding control wells (defined as 100% biofilm) and the negative control wells (defined as 0% biofilm).

Effect of PRP on mature biofilms (biofilm eradication assay)

This test was conducted to assess the impact of PRP on existing biofilms, following the methodologies outlined by Cruz et al. and Haney et al.^71,72. The assay was conducted as outlined below:

1. 1.

   The biofilms were allowed to mature for 24 h as previously described.

2. 2.

   The wells were aspirated and washed 3 times with distilled water.

3. 3.

   One hundred microliters of activated PRP was added to each well, and the plate was incubated at 37 °C for 24 h.

4. 4.

   The contents were then discarded, and the plates were washed with distilled water.

5. 5.

   Approximately 200 µL of TSB was added each well, and the mixture was incubated for another 24 h.

6. 6.

   After incubation, the biofilms were scraped from the wells via sterile tips, and 100 µL from each well was inoculated into a TSA plate and incubated for 24 h⁷³.

7. 7.

   The plates were then inspected, and the number of colonies was counted (Fig. 18).

8. 8.

   Colony counts from the PRP wells were compared with those from the corresponding control wells and the negative control wells.

9. 9.

   The test was repeated except that the plate was incubated after the addition of PRP for 2 h instead of 24 h.

Statistical analysis

The data were analyzed using version 25.0 of the Statistical Package for Social Sciences (SPSS) software. Qualitative data were expressed in numerical form and percentages. The Kolmogorov-Smirnov test was employed to assess the normality of the distribution. The quantitative data were given as ranges (minimums and maximums), means, standard deviations, and medians. The significance of the acquired results was assessed at the 5% level.