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Biology of PRF: Fibrin Matrix, Growth Factor Release, and Cellular Activity

Contributors

Masako Fujioka-Kobayashi

Yufeng Zhang

Reinhard Gruber

Richard J. Miron

Chapter Highlights

 What is PRF?

 How does PRF differ from PRP at the biologic and cellular level?

 What is the role of each cell type found in PRF?

 What is the role of each GF found in PRF?

 How does centrifugation speed and time affect PRF?

 What advantages exist for horizontal centrifugation versus fixed-angle centrifugation?

Video 2-1

Much can be discussed with respect to the biology of PRF and its ability to impact tissue regeneration. During the natural wound healing process, vascularization of tissues plays a pivotal role, facilitating the invasion of incoming cells, growth factors (GFs), cytokines, and other regenerative factors. The main aim of platelet concentrates, discovered over two decades ago, is to favor new blood flow (angiogenesis) to damaged tissues, thereby improving their healing potential by delivering a supraphysiologic concentration of blood-derived cells (namely platelets) and regenerative GFs. This chapter takes a deep look into the actual separation of blood layers during the centrifugation process to provide the clinician a general overview of the cell types and GFs found in PRF, including their roles, and also discusses the effects of centrifugation speed and time on cell layer separation. Furthermore, the advantages of producing an autologous fibrin scaffold are presented as it being a key regulator of wound healing because of its autologous source and its ability to promote the slow and gradual release of GFs over time. The advantages of horizontal centrifugation versus fixed-angle centrifugation are also discussed based on recent data from various laboratories from around the world.

The wound healing process is divided into three stages: the inflammatory phase, the proliferative phase, and the remodeling phase (Fig 2-1). The inflammatory phase starts at the time of injury and generally involves a wide array of cytokines and growth factors (GFs) that are released within the first 24 to 48 hours. Accordingly, a dynamic interaction occurs between endothelial cells, angiogenic cytokines, and the extracellular matrix (ECM) in an attempt to accelerate wound healing via an orchestrated delivery of multiple GFs in a well-controlled fashion.¹

Fig 2-1 The three stages of wound healing: (a) Inflammatory phase. (b) Proliferative phase. (c) Remodeling phase.

In general, blood provides essential components to the healing process that comprise both cellular and protein products that essentially are the base components of wound healing. During the healing process, blood will undergo clotting within a few minutes to prevent further blood loss. This is an important step that will be later discussed in the PRF tube section, because in order for clotting to occur and even be improved in both speed and quality (in particular in patients taking anticoagulants), a proper understanding of the clotting cascade is required. In its simplest of forms, oxygen helps improve blood clotting, and for this reason, the simple removal of centrifugation tube lids following the spin process will lead to faster clotting of PRF and a superior fibrin mesh. One of the major roles of platelets is to assist during hemostasis through a fibrin clot formation.^1,2 Not surprisingly, the additional use of PRF for wound healing (for example, following tooth removal and extraction socket healing) in patients taking anticoagulants can drastically improve the healing outcomes simply by improving clotting. Because PRF contains many platelets and a fibrin nucleus is already formed, bleeding has been shown to be significantly reduced postoperative when PRF is utilized in patients on anticoagulant therapy.³

Tips

 Oxygen helps improve blood clotting, so the simple removal of centrifugation tube lids following the spin process will lead to faster clotting of PRF.

 In patients undergoing anticoagulant therapy, the simple addition of PRF during surgery can help favor faster clotting, thereby reducing bleeding times postoperative.

Platelets also release various GFs and cytokines that further lead to tissue regeneration but also attract macrophages and neutrophils to the defect site. These cells are responsible for clearing debris, replacing necrotic tissue, and removing bacteria from the wound site.

The proliferative phase begins by day 3, where the blood clot within the wound is further supplied with a provisional matrix typically composed in part with fibrin, which facilitates cell migration, while the clot within the vessel lumen contributes to hemostasis.² Fibroblast cells are recruited to the wound site and begin producing new collagen in a random and somewhat disorganized order. Simulta-neously, new blood vessel formation leads to new angiogenesis, and the wound gradually begins to gain initial stability.

During the third and final stage (the remodeling phase), disorganized collagen is replaced by newly organized collagen fibrils that provide enhanced stability and strength to the injured site, where tissue regeneration takes place⁴ (see Fig 2-1).

Whole blood is comprised of four main components: blood plasma, red blood cells (RBCs), white blood cells (WBC), and platelets. Initially, platelets were reported as the major responsible component for the activation and release of crucial GFs for wound healing, including PDGF, coagulation factors, adhesion molecules, cytokines, and angiogenic factors. Their role has been extremely well described in the literature, so typically the entire field has been referred to as platelet concentrates or platelet-rich plasma/fibrin. Interestingly, however, over the years more attention has been placed on leukocytes, which are not only responsible for host defense but also highly implicated in the wound healing and regenerative phases.

Table 2-1 highlights the various cell types found in blood, including their density, frequency, and surface area. Note that while platelets are the lightest of the group, WBCs and RBCs are very similar in density. For these reasons they are also harder to separate in a centrifuge based on density. Noteworthy is the fact that per µL, there are 5,000,000 RBCs when compared to only 5,000 WBCs. Therefore, RBCs outnumber WBCs in a 1,000:1 ratio, which make them difficult to separate, especially on a fixed-angle centrifugation device as discussed later in this chapter (Video 2-2).

Table 2-1 Properties of cells found in whole blood

Video 2-2

Leukocytes and RBCs are similar in density. This makes these two cell types extremely difficult to separate, especially because RBCs outnumber them 1,000 to 1.

Cells in PRF

As shown in Table 2-1, the three main cell types found in PRP and PRF are platelets, leukocytes (WBCs), and RBCs. The entire initial goal of platelet concentrates was of course to concentrate platelets. Because they are the lightest of all cells found in blood, it was possible to utilize a centrifugation device to separate these layers based on their density. Lighter cells (platelets) could therefore be accumulated to the top, followed by leukocytes. Because RBCs are the densest of the group, they tend to migrate downward during the centrifugation process. In an ideal situation, the final PRF matrix should be composed of a high concentration of platelets, leukocytes, and fibrin. It has been shown that the initially developed PRF (also termed L-PRF for leukocyte PRF) concentrates contained greater than 90% platelets and more than 50% leukocytes within a high-density fibrin network when compared to whole blood.⁵ By utilizing more advanced quantification devices and recently developed methods, our research team has been better able to harvest leukocytes specifically. The lower yield of leukocytes is typically a result of their more similar density to RBCs, making them harder to separate from and accumulate in the upper layers where PRF is collected. This is particularly difficult on fixed-angle centrifuges. Several other methods have been proposed to favor accumulation of cells, including shorter centrifugation times as well as lower centrifugation forces, as discussed later in this chapter (see section on the low-speed centri-fugation concept).⁶

Leukocytes have been shown to be an integral component of PRF therapy and play a prominent role in wound healing. Studies from basic sciences and animal research have revealed how impactful a role leukocytes play during tissue regeneration by comparing PRP/PRF therapy with and without WBCs.^7–9 In these split design studies, the contralateral side receiving leukocytes performed significantly better, promoting researchers and clinicians to develop protocols to better incorporate or harvest leukocytes. Naturally, PRF contains a higher number of leukocytes when compared to the first-generation platelet concentrates PRP and PRGF.

While the role of leukocytes has been well described as host defense against incoming pathogens, they also play a central role in immune modulation of biomaterials and participate in the wound healing process due to their ability to secrete key immune cytokines such as IL-1β, IL-6, IL-4, and TNF-α.^2,10,11 They have been highly investigated in PRF therapy, with the impact of centrifugation speed and time affecting both their concentration and location, mainly owing to the fact that they are very similar in density and size to RBCs. Previously, it was demonstrated how faster protocols initially utilized to produce L-PRF were far too high in both g-force and time (2700 rpm for 12 minutes; ~700g).⁶ This led to the histologic observation that the majority of cells were concentrated either at the buffy coat region or at the bottom of centrifugation tubes within the RBC layer component.⁶ Based on these observations, it became clear that centrifugation speeds (g-forces) were evidently too high, pushing leukocytes especially down to the bottom of centrifugation tubes and away from the PRF clot. In order to redistribute leukocyte cell numbers across the entire PRF matrix, both a change in centrifugation speed and/or time (lower) as well as a change in the centrifugation device (horizontal centrifugation as opposed to fixed-angle) were deemed necessary to further improve platelet formulations, as reviewed later in this chapter.

Advantages of a 3D Fibrin Network

Fibrin is the activated form of a plasmatic molecule called fibrinogen that converts into fibrin with thrombin. Fibrin formation is one of the first key components to tissue wound healing. When an individual cuts himself, the first event taking place prior to any regeneration is fibrin clot formation. This is why patients on anticoagulant therapy typically do not heal quite as effectively because delayed clotting leads to delayed healing. The obvious advantage of PRF therapy is its ability to accumulate various cell types including platelets without anticoagulants, thereby improving clotting properties. Once a fibrin clot is formed during the centrifugation cycle, cells and GFs are able to be trapped within the 3D fibrin matrix, favoring the slower and gradual release of GFs from PRF over time.¹² Fibrin is a soluble fibrillary molecule that is present in high quantity both in plasma itself as well as in the α-granules of platelets. Fibrin therefore plays a determining role in platelet aggregation during hemostasis and is critical to healing. Not surprisingly, the use of fibrin alone (without GFs or living cells as a fibrin glue) has been shown to lead to matrix stabilization favoring tissue stability, cellular invasion, and ultimately tissue regeneration.^13–15 However, PRF has numerous advantages in that during the fibrin clot formation, a supraphysiologic concentration of platelets, leukocytes, and GFs are also present, forming a sort of “superclot” consisting of an intimate assembly of cytokines, glycanic chains, and structural glycoproteins enmeshed within a slowly polymerized fibrin network^16,17 (Fig 2-2).

Fig 2-2 SEM examination of the fibrin clot revealing a dense and mature fibrin matrix with various cell types entrapped within its matrix.

PRF forms a “superclot” consisting of an intimate assembly of cytokines, glycanic chains, and structural glycoproteins enmeshed within a slowly polymerized fibrin network.

The fibrin scaffold produced following centrifugation has further been identified as a biologic 3D network with the ability for the fibrillar micropores to support cell migration, proliferation, differentiation, and delivery of GFs. Platelets have theoretically been described as being massively trapped within the fibrin network, and the release of GFs is largely dictated by the actual timespan in which the 3D PRF scaffold is broken down (typically within 2–3 weeks).¹⁸ One of the major advantages of PRF when compared to PRP is the fact that by simply removing anticoagulants, a fibrin matrix is formed with a natural autologous delivery system capable of slowly and gradually releasing GFs over time.^7–9 This leads to GF delivery over a period of 2 to 3 weeks as opposed to only a few hours observed in PRP.¹²

Finally, stem cells exist naturally in whole blood, albeit at extremely low levels.^19,20 Stem cells have the ability and potential to differentiate into many cell types, including adipocytes, osteoblasts, and chondrocytes. Many commercial enterprises report that mesenchymal stem cells (MSCs) exist in extremely high numbers in PRF or that only certain protocols or machinery favor their accumulation, but these reports have not been validated in any high-quality peer-reviewed journal. While future research investigating the impact of MSCs in blood is necessary, it may represent a potential future strategy to isolate MSCs relatively easily at low cost.

The commercial claims that MSCs exist in extremely high numbers in PRF or that only certain protocols or machinery favors their accumulation have not been validated.

Growth Factors in Blood

Naturally, GFs are critical to wound healing, and a variety of GFs have been commercialized as recombinant human sources once their roles were established. GFs are largely responsible for the migration of cells and also play a critical role in their adhesion, proliferation, and differentiation. While GFs exist in all tissues, it is important to note that blood serves as a main reservoir of numerous GFs and cytokines promoting angiogenesis and tissue regeneration for wound healing. It is also important to note that certain GFs may exist as inactive or partially active precursors that require proteolytic activation, or may further require binding to matrix molecules for activity or stabilization. For this reason, interfering with the natural clotting cascade such as when utilizing PRP may affect the bioactivity of certain GFs.¹² Typically, GFs also have extremely short biologic half-lives in the order of a few minutes.²¹ The body has been trained to secrete various GFs in programmed orders to activate very complex cellular processes.²² Unlike recombinant human GFs that typically only comprise a single GF, platelet concentrates create the opportunity to deliver many autologous GFs simultaneously. Furthermore, leukocytes are known immune cells capable of “sensing” their microenvironment during the regenerative phase. Together with platelets, leukocytes serve as a major cell type during the natural wound healing process. The GFs most accumulated and delivered in PRF include VEGF, PDGF, TGF-β1, EGF, and IGF.^23,24 Their individual roles are discussed below.

Unlike recombinant human GFs that typically only comprise a single GF, platelet concentrates create the opportunity to deliver many autologous GFs simultaneously.

VEGF

VEGF is secreted by activated thrombocytes and macrophages to damaged sites to promote angiogenesis. The VEGF family is related to PDGF and includes VEGF-A, -B, -C, -D, and -E. VEGF has previously been isolated and utilized as a recombinant GF described as the most potent GF for angiogenesis of tissues, stimulating new blood vessel formation, and facilitating nutrients and increased blood flow to sites of injury.^25,26 It has potent effects on tissue remodeling, and the incorporation of recombinant human VEGF into various bone biomaterials alone has been demonstrated to increase new tissue regeneration.²⁷

PDGF

PDGFs are essential regulators that mainly promote the migration, proliferation, and survival of mesenchymal cell lineages.^28–33 Platelets are of course the major source of PDGF, with various groups divided into homo- (PDGF-AA, PDGF-BB, PDGF-CC, and PDGF-DD) and heterodimeric (PDGF-AB) polypeptide dimers linked by disulfide bonds. They are mainly present in large quantities in platelet α-granules. Therefore, the ability to concentrate platelets in PRF leads to the massive accumulation and subsequent release of PDGF following centrifugation. It is important to note that because PDGF has an extremely short half-life (characterized by as little as 2 minutes), the ability for the PRF scaffold to act as a reservoir of GFs by protecting PDGF from matrix metalloproteinases actually drastically improves its release profile in living tissues when compared to PRP. Because it plays such a critical role in the mechanisms of physiologic healing, a recombinant source (rhPDGF-BB) was made commercially available with FDA clearance for the regeneration of various defects in medicine and dentistry.

Because PDGF has an extremely short half-life (characterized by as little as 2 minutes), the ability for the PRF scaffold to act as a reservoir of GFs by protecting PDGF from matrix metalloproteinases actually drastically improves its release profile in living tissues when compared to PRP.

TGF-β1

TGF-β is a large superfamily of more than 30 members that mediate tissue repair, immune modulation, and ECM synthesis.^34,35 TGF-β1 is the predominant isoform that supports cell proliferation of practically all cell types.²¹ It further plays a role in angiogenesis, re-epithelialization, and connective tissue regeneration.²¹ Platelets are known to be a major source of TGF-β production. Specific to bone healing and remodeling, TGF-β also exerts effects by being heavily released from autogenous bone.²¹

EGF

The EGF family is responsible for the chemotaxis and angiogenesis of endothelial cells and mitosis of mesenchymal cells. It further enhances epithelialization and markedly shortens the overall healing process time when administered as a recombinant source. EGF is typically secreted naturally by the body after acute injury and acts to significantly increase the tensile strength of wounds. EGF receptors exist on most human cell types, including those that play a critical role during wound repair such as fibroblasts, endothelial cells, and keratinocytes.³⁶

IGF

IGFs are positive regulators of proliferation and differentiation of most cell types.³⁷ IGF is found in high levels in PRF because it is highly released from platelets during their activation and degranulation, leading to the differentiation of mesenchymal cells. IGF is also an extremely attractive GF because it constitutes the major axis of programmed cell apoptosis regulation by inducing survival signals protecting cells from many apoptotic stimuli.³⁷

Comparative Analysis of PRP Versus PRF

Years ago, much research investigated the comparative analysis in bioactivity and tissue regenerative properties of PRP and PRF.¹² The aim was to determine if PRF was significantly better than previously utilized PRP with respect to the initial cell content, GF release, and clinical benefit. Naturally, the release profile of GFs has been an important and highly investigated research topic over the years and differs quite significantly between PRP and PRF. The typical advantage of PRF over PRP is that it enables the controlled release of GFs over an extended period of time because of its fibrin matrix. Therefore, when PRF is directly compared to PRP, a much longer GF delivery response has been observed³⁸ (Fig 2-3). In a first study on this topic performed by our research team, GF release from three different platelet concentrates including PRP, L-PRF, and A-PRF demonstrated a much higher total amount of GFs released from PRF when compared to PRP over a 10-day period. In order to precisely quantify each GF in a comparative fashion, PDGF, VEGF, TGF-β1, EGF, and IGF were compared at numerous time points including 15 minutes, 60 minutes, 8 hours, 24 hours, 3 days, and 10 days (see Fig 2-3). Interestingly, at an early time point (15 minutes), significantly higher numbers of GFs were released from PRP when compared to L-PRF or A-PRF, whereas both PRF groups had higher release at later time points and also total release of GFs. From 3 days and onward, the GF release of PRF far exceeded that of PRP.¹²

Fig 2-3 GF release from PRP and PRF at each time point for PDGF-AA, PDGF-AB, and TGF-β1 over a 10-day period. Notice that while PRP has significantly higher GF release at early time points, over a 10-day period, significantly higher levels are most commonly found with A-PRF due to the slow and gradual release of GFs following use of slower centrifugation speeds. (Adapted from Kobayashi et al.¹²)

Overall, PRP can be recommended for fast delivery of GFs, whereas PRF is better suited for long-term delivery and favors better overall wound healing potential.

The Low-Speed Centrifugation Concept

The evolution of PRF in the years 2014 to 2017 was highly focused on ways to improve centrifugation parameters in order to favor more GF release. Within the original L-PRF matrix, cells were surprisingly found gathered at the bottom of the PRF matrix or bottom of centrifugation tubes.⁶ In simple terms, the faster and longer centrifugation is carried out, the more matter (ie, cells) is moved to the bottom of centrifugation tubes. With the development of A-PRF utilizing the low-speed centrifugation concept (LSCC), cell pull-down was reduced, which increased the total number of cells left contained within the top layer of PRF.

The faster and longer centrifugation is carried out, the more cells are pushed to the bottom of centrifugation tubes.

In a cell and GF analysis study on this topic by our group, the newer protocols for the production of A-PRF+, which involves not only lower centrifugation speed but also less time (1300 rpm for 8 min), was shown to lead to even further increases in GF release of TGF-β1, PDGF-AA, PDGF-AB, PDGF-BB, VEGF, IGF, and EGF (Fig 2-4).²⁴ Therefore, an optimization of centrifugation parameters could be obtained simply by lowering the g-force and time, favoring better GF delivery.

Fig 2-4 GF release resulting from the LSCC at each time point for PDGF-AA, PDGF-AB, and TGF-β1 over a 10-day period. In general, it was found that A-PRF+ demonstrated significantly the highest GF release when compared to all other modalities after a 10-day period. (Adapted from Fujioka-Kobayashi et al.²⁴)

Not surprisingly, when cells were cultured with the various formulations of PRF, it was also observed that centrifugation utilizing these lower speeds/time further led to greater cellular bioactivity. In a study titled “Optimized platelet-rich fibrin with the low-speed concept: Growth factor release, biocompatibility, and cellular response,” our research team demonstrated that protocols at lower speeds and time not only led to significantly greater human fibroblast cell migration and proliferation compared to L-PRF but also demonstrated significantly higher mRNA levels of PDGF, TGF-β, and collagen1 at either 3 or 7 days (Fig 2-5). In addition, PRF membranes implanted subcutaneously that were fabricated using lower centrifugation speeds also generated greater and faster vascularization in vivo.²⁴

Fig 2-5 Human gingival fibroblast behavior exposed to L-PRF, A-PRF, and A-PRF+: (a) Cell migration, (b) gene expression, and (c) collagen synthesis on human gingival fibroblasts. (Adapted with permission from Fujioka-Kobayashi et al.²⁴)

Comparing Centrifugation Devices

One highly debated topic with much commercial interest has been the role of the centrifugation device on production of PRF. As such, there are general parameters that need to be respected to calculate relative centrifugal forces (RCF) effectively; this forms the basis of an entire chapter in this textbook (see chapter 4). While there are definitely quality differences between centrifugation devices, the debate between which centrifugation system to utilize is generally overhyped, leading to confusion among colleagues regarding how to accurately select, and more importantly, optimize a centrifugation system.

The debate surrounding which centrifugation system to utilize is generally overhyped, leading to confusion among colleagues regarding how to accurately select and, more importantly, optimize a centrifugation system.

As such, in 2018 our research team addressed this question in a study titled “Comparison of platelet-rich fibrin (PRF) produced using 3 commercially available centrifuges at both high (~ 700 g) and low (~ 200 g) relative centrifugation forces” with the specific aim to demonstrate that any centrifugation device could be utilized to produce PRF using the LSCC.³⁹ In that study, PRF was produced on three commonly utilized commercially available centrifuges including the IntraSpin (Intra-Lock), Duo Quattro (Process for PRF), and Salvin (Salvin Dental) devices (Fig 2-6). Two separate protocols were tested on each machine, including the original L-PRF protocol (~700g RCF-max [~400g RCF-clot] for 12 min) as well as the A-PRF+ protocol (~200g RCF-max [~130g RCF-clot] for 8 min). Each of the tested groups was compared for cell numbers, GF release, scanning electron microscopy (SEM) for morphologic differences, and clot size (both weight and length/width).

Fig 2-6 Experimental setup: Each of the centrifuges was utilized in duplicate for a total of six centrifuges. From each patient, a total of 18 tubes were drawn (2 per machine) and centrifuged accordingly at either high- or low-speed protocols. (Reprinted with permission from Miron et al.³⁹)

It was found that PRF clots produced utilizing the lower centrifugation speeds and time (1) contained a higher concentration of evenly distributed platelets, (2) secreted higher concentrations of GFs over a 10-day period, and (3) were smaller in size. This was irrespective of the centri-fugation device utilized and consistently observed on all three devices. The greatest impact was found between the protocols utilized (up to a 200% difference). Furthermore, it was revealed that centrifugation carried out at lower speeds had a slightly less dense fibrin mesh with more open spaces to allow for cellular migration (Fig 2-7). Most importantly, it was revealed for the first time that centrifugation tubes actually had a much greater impact on the final size outcome of PRF clots when compared to centrifugation devices. This was the first time our team had observed such dramatic differences in the size outcomes of PRF tubes, and since then a plethora of research has since been performed on that topic alone (see chapter 5).

Fig 2-7 SEM of PRF clots produced on three different devices at either high-speed (~700g) or low-speed (~200g) protocols. Notice that the clots produced at high g-force typically were more densely packed with fibrin. (Reprinted with permission from Miron et al.³⁹)

Research has clearly shown that PRF tubes matter much more than the centrifugation device used.

Development of i-PRF

One of the advantages of PRP is that it is liquid in nature, making it easy to utilize in combination with various bone biomaterials, most notably bone grafting materials. With PRF, as centrifugation speeds and times were being reduced further and further, a nonclotted liquid plasma layer was noticed prior to actual clot formation. This liquid-PRF layer is actually liquid fibrinogen that has not yet converted to fibrin. This liquid formulation of PRF was given the working name liquid-PRF or injectable-PRF (i-PRF) for simplicity (Fig 2-8a).⁴⁰ This layer can be quickly harvested (Fig 2-8b) and injected into a defect area. Interestingly, once liquid-PRF has converted to a solid state, it forms the standard fibrin fibrillar PRF that most are familiar with, as depicted in Fig 2-9 by SEM.⁴¹ Based on its potential for clinical applications, both preclinical and clinical studies have since been conducted to evaluate its regenerative potential.

Fig 2-8 (a) Clinical photograph of liquid-PRF. Note that this protocol separates out a small upper liquid-PRF layer about 1 mL in quantity. (b) This liquid i-PRF layer may be harvested into a syringe and utilized as an injectable platelet-rich formulation. (Reprinted with permission from Davies and Miron.⁴⁰)

Fig 2-9 The surface (a) and cross-section (b) microstructures of the i-PRF. f, fibrin; p, platelet aggregates; r, RBC. Scale bar = 10 μm. (Reprinted with permission from Zhang et al.⁴¹)

Data from our laboratories first found that GF release from PRP was typically within the first hour, whereas i-PRF had a much more widespread release of GFs over time, similar to solid-PRF^42–44 (Fig 2-10). Unlike our previous studies comparing various solid-PRF membrane formulations, however, some GFs were in fact secreted in higher levels from PRP when compared to i-PRF, whereas others were more highly released from i-PRF. In 2015, i-PRF was developed and initially studied as a very short and slow centrifugation protocol of 700 rpm (60g) for 3 to 4 minutes. Following this protocol, i-PRF remained liquid for roughly 15 to 20 minutes. This new formulation was utilized for a variety of procedures including mixing with bone grafts to form a stable fibrin graft with improved handling and graft stability.

Fig 2-10 GF release from i-PRF compared with PRP at each time point for PDGF-AA, PDGF-AB, TGF-β1, and VEGF over a 10-day period. Note the varying GF release from PRP and i-PRF. Some GFs were in fact more highly released from PRP, which posed many questions several years ago. (Adapted from Miron et al.⁴⁴)

A variety of basic research studies have since demonstrated the regenerative potential of i-PRF when compared to PRP.^44–49 While both formulations exhibited high biocompatibility of human gingival fibroblasts as well as significantly induced higher cell migration when compared to control tissue-culture plastic in vitro (Fig 2-11),⁴⁴ it was found that i-PRF induced significantly greater cell migration, mRNA levels of TGF-β, and collagen1 expression.^45–49 It wasn’t until years later that further means to improve i-PRF were developed, as highlighted later in this chapter.

Fig 2-11 Human gingival fibroblast behavior exposed to i-PRF versus PRP: (a) Cell migration, (b) gene expression, and (c) collagen synthesis. (Adapted with permission from Miron et al.⁴⁴)

Quantifying Cell Types in PRF

Pitfalls of current methods

Two pitfalls exist with quantifying cell types found within PRF research. Histologic studies have been infrequently performed and show variability and bias in results, because many times it becomes difficult to know the exact orientation of PRF relative to the histologic processing. It therefore becomes very difficult to assess where cells are actually located following cell separation and histologic preparation. Another method commonly performed has been the use of complete blood counts (CBCs). Simply, following centrifugation, the plasma layer can be harvested and sent for a CBC and compared relative to whole blood (Fig 2-12a). While this accurately reports an increased ratio with respect to platelet and leukocyte numbers, one of the main drawbacks is that the method cannot determine if the cells are evenly distributed within the according PRF layers. As illustrated in Fig 2-12b, by utilizing this method, very similar final outcomes were found with only marginal improvements of roughly 20% visible between the two previously utilized protocols of L-PRF and A-PRF, yet histologically a much better distribution of platelets was observed by Ghanaati et al.⁶ Therefore, a new method was proposed to better quantify cell types in platelet concentrates, as described next.

Fig 2-12 (a) One method to investigate cells found in PRP/PRF is to send the plasma layer following centrifugation for a full CBC. This CBC value can be compared to a control of whole blood to investigate its % increase. (b) One of the reported pitfalls of this technique is that the precise location of cells within the upper layer is not revealed. For instance, following the L-PRF protocol, the cells are gathered more precisely at the buffy coat zone, whereas the A-PRF protocol tends to more evenly distribute platelets. Therefore, this technique may not necessarily represent the most ideal scientific accuracy/data.

A novel method to quantify PRF

Over the past few years, several commercially available centrifuges have further been brought to market. These vary in many factors including protocols, RCF values, tube-rotor angulation, rotor radius size, and tube composition. Each of these plays a role in the final obtained PRF membrane, yet little data is scientifically available displaying cell numbers and content in the various layers following centrifugation.

While much commercial debate exists on this topic, there has been no accurate method to quantify/determine the precise location of cells following centrifugation, with few histologic studies performed investigating cell numbers within the fibrin clots. In a pioneering research article published in 2019, our research team proposed a novel method to quantify cell numbers and concentration within the PRF scaffolds following centrifugation by utilizing a sequential pipetting methodology.⁵⁰ Standard protocols commonly utilized for the production of PRF were investigated according to their respective manufacturer’s protocols. New to this research article, for the first time following centrifugation, 1-mL layers were sequentially pipetted from the upper layer of blood tubes toward the bottom of the tube until all 10 mL were harvested in sequential samples (Fig 2-13).⁵⁰ Each of these 10 samples from each centrifugation tube was then sent for CBC analysis to accurately quantify the cell numbers within each 1-mL blood layer and then compared according to cell numbers and concentrations. This study represented a novel experimental methodology to more accurately depict cell numbers in PRF following centrifugation using various protocols.

Fig 2-13 (a) Illustration demonstrating the proposed novel method to quantify cell types following centrifugation of PRF. Currently, one of the limitations is that whole blood is compared to the total plasma concentration following centrifugation. This, however, does not give a proper representation regarding the location of cells following centrifugation. By sequentially pipetting 1 mL of volume from the top layer downward, it is possible to send each of the 10 samples for CBC analysis and accurately determine the precise location of each cell type following centrifugation at various protocols. Notice that one layer (in this case layer 5) will contain some yellow plasma and RBCs. This is typically the location of the buffy coat, where a higher concentration of platelets is located. (b) Visual demonstration of the protocol. Following centrifugation with two 10-mL centrifugation tubes, blood layers are then separated. Thereafter, 1-mL samples are pipetted precisely from the upper layer downward. Notice that when layer 5 was drawn, it was possible to visualize the layer separation between the yellow plasma and RBC layers. This separation layer was noted for all samples. (Reprinted with permission from Miron et al.⁵⁰)

By utilizing this novel technique and investigating each 1 mL layer by layer, it was possible for the first time to investigate exactly each cell layer following centrifugation and determine the precise location of each blood cell type.

L-PRF protocol

Following centrifugation, 1-mL sequential layers were sent for CBC analysis according to Fig 2-14a. As Fig 2-14b shows, the original L-PRF protocol using the IntraSpin device (2700 rpm for 12 min; ~700g) with a 33-degree fixed-angle centrifuge revealed precisely that the number of leukocytes (control 6 × 10⁹ cells/L) and platelets was significantly concentrated in layer 5 (~17 × 10⁹ cells/L; arrows represent where the plasma and RBC layers separate). Interestingly, a threefold to fourfold increase in leukocyte number was observed specifically at this interface within the buffy coat. Notice, however, that no leukocytes were found in any of the upper 4 layers, displaying a very uneven PRF clot with respect to cell numbers. Almost all cells within the PRF clot were exclusively found within this 5th layer. Notice also that more leukocytes were found in the RBC layer below the PRF clot. A similar trend was also observed for lymphocytes, neutrophils, and monocytes (see Fig 2-14b). Naturally, all RBCs were found in layers 5 through 10 in the visually red layers. Platelets were accumulated once again precisely in layer 5 (six- to eightfold), within the buffy coat zone.

Fig 2-14 (a) Separation of the 10-mL tube into 10 1-ml layers for pipetting. (b) The concentration of cell types in each 1-mL layer utilizing the solid L-PRF protocol (2700 rpm for 12 minutes; ~700g). Notice that the majority of platelets accumulated directly within the 5th layer in the buffy coat. Furthermore, the highest concentration of leukocytes was also noted in this layer. The first 4 layers of this plasma layer were typically devoid of all cells. (Adapted with permission from Miron et al.⁵⁰)

Because the majority of cells were found in layer 5, we were interested to determine if these cells were specifically found within the yellow plasma layer (within the PRF clot) or within the RBC layer. For this, a second blood tube was utilized; 500 µL of blood volume was collected just above the RBC layer within the buffy coat, and 500 µL was taken from the RBC layer. It was revealed that the majority of platelets were found within the yellow plasma layer (> 80%), whereas the majority of leukocytes and other WBCs were found within the red blood layer (Fig 2-15). This revealed that most leukocytes were in fact not found within the PRF layers utilizing the L-PRF protocol.

Fig 2-15 Layer 5 (the zone that incorporates the buffy coat containing a plasma and RBC component) demonstrated the cell-rich zone. Analysis of this zone revealed that many of the cells were in fact located in the red zone (especially leukocytes). The cell-rich zone contains a yellow buffy coat zone, but the red portion of this buffy coat also contains many cells. Following these findings, it is generally recommended to harvest a small portion of the red zone, specifically when drawing liquid-PRF (i-PRF), because many cells are located within this region. (Adapted from Miron et al.⁵⁰)

Interestingly, the final concentration of leukocytes found using the L-PRF protocol was 4.13 × 10⁹ cells/L, whereas the control whole blood value from this patient was 6.125 × 10⁹ cells/L, representing a 33% reduction in leukocyte concentration when compared to control blood. Platelet numbers were increased 1.61-fold. The total leukocyte and platelet content represent 33% and 80% of the total blood cells found, respectively, within this 10-mL blood sample. This meant that roughly 20% of platelets and 66% of leukocytes were actually located within the RBC layer (similar to the observed histologic results by Ghanaati et al in 2014^6).

With the L-PRF protocol, the majority of leukocytes and platelets were not found within the plasma layer but rather in layer 5 within the buffy coat zone.

A-PRF protocol

Figure 2-16 depicts centrifugation following A-PRF protocols (1300 rpm for 8 minutes on a Duo Quattro centrifuge). Interestingly, the number of platelets were concentrated throughout the first four to five layers, unlike the L-PRF protocol. Here, a twofold increase in platelets was observed compared to a 1.6-fold increase utilizing the L-PRF protocol. More importantly, however, the platelets were found evenly distributed throughout the A-PRF plasma layers. When investigating leukocyte number, however, a significantly lower concentration (33% original values) as well as total numbers (9.315 vs 20.65 × 10⁹ cells/L) were found in the A-PRF group when compared to L-PRF. Therefore, it was initially suspected that either the g-force or the total time was not sufficient to adequately accumulate or separate the leukocytes utilizing the A-PRF protocol.⁵⁰ Once again, lower leukocytes in PRF were actually found when compared to whole blood.

Fig 2-16 The concentration of cell types in each 1-mL layer utilizing the solid A-PRF protocol (1300 rpm for 8 minutes; ~200g). Notice that the platelets were more evenly distributed throughout the upper 5-mL plasma layer. Noteworthy, however, is that the majority of WBCs (leukocytes, neutrophils, lymphocytes, and monocytes) were not found in the upper plasma layer. (Adapted from Miron et al.⁵⁰)

While the A-PRF protocol with LSCC led to a higher concentration of platelets, it was not effectively capable of concentrating leukocytes.

i-PRF protocol

Liquid-PRF protocols were then investigated and compared (Fig 2-17). The IntraSpin protocol (2700 rpm for 3 min; ~700g) is depicted in Fig 2-17a. Interestingly, this protocol accumulated platelets evenly throughout the PRF layer better than when utilizing the 12-minute protocol. Nevertheless, leukocytes were significantly lower once again when compared to whole blood, representing only 54% of the original control blood concentrations. This demonstrates that following centrifugation, lower numbers of leukocytes are found in L-PRF samples when compared to control blood in either L-PRF protocol. Platelet concentrates were increased 2.12-fold.

Fig 2-17 (a) The concentration of cell types in each 1-mL layer utilizing the i-PRF IntraSpin protocol (2700 rpm for 3 minutes; ~700g). Notice that most platelets are more evenly distributed utilizing this protocol when compared to the 12-minute solid-PRF IntraSpin protocol. (b) The concentration of cell types in each 1-mL layer utilizing the i-PRF Duo Quattro protocol (800 rpm for 3 minutes; ~60g). Notice that very little change in platelet or leukocyte accumulation is observed utilizing this centrifugation cycle. A slight increase in platelets and leukocytes is, however, observed when compared to the control. (Adapted from Miron et al.⁵⁰)

The i-PRF protocol recommended by Process for PRF (Duo Quattro centrifuge) produced a 1.23-fold increase in leukocyte concentration and a 2.07-fold increase in platelet concentration when compared to whole blood (see Fig 2-17b). The overall accumulation demonstrated an 18% total leukocyte content and a 31% total platelet count when compared to whole blood. This represented an extremely low platelet yield, as all other protocols produced at least 80% total yield. (Keep in mind here that this means 70% of platelets are found within the red layer following the use of this LSCC and not in the upper, liquid-PRF layer.) Most notably, the change in cell density layer by layer, as depicted in Fig 2-17b, was almost unnoticeable. These findings revealed that the i-PRF protocol displayed an inability to concentrate cells effectively, and it was clear that improvements were needed.

The use of the original i-PRF protocol only accumulated on average 18% of leukocytes and 31% of platelets. It was clear improvements were needed.

Discussion of findings

One of the most surprising findings was the observation that almost all platelets were accumulated in layer 5 using the conventional L-PRF protocols. Almost no platelets were observed in the first four layers following centrifugation, and the majority of leukocytes were found in the RBC layer, not in the PRF clot. This was a bit ironic, granted the working name leukocyte platelet-rich fibrin (as in, leukocyte-rich and platelet-rich fibrin). Previous studies have also shown that L-PRF protocols result in lower platelet and leukocyte numbers when compared to various other protocols produced at lower RCF values.^6,51

It was certainly ironic that low levels of leukocytes were found in the PRF clot of L-PRF when compared to normal blood, considering the working name leukocyte PRF, which implies that the PRF clot would be leukocyte rich.

While platelet concentrations were also lower in L-PRF vs A-PRF, it was revealed that the actual differences were not as drastic as previously reported.⁵¹ Our study demonstrated more specifically that the platelets and leukocytes are in fact found precisely at the junction between the yellow plasma and RBC interface. Previous studies likely failed to collect all the liquid within the yellow plasma layer, and as a result, extremely low platelet and leukocyte values may have been reported. As clearly shown in the present study, failure to do so, especially at a g-force of ~700g (RCF-max) or greater, results in extremely low concentration values because the upper 4 mL of plasma is practically devoid of cells. Therefore, we demonstrated in this study that L-PRF protocols are in fact quite rich in platelets (~80–90% total), but these cells are found precisely within a 1-mL layer directly above the RBCs within the buffy coat. It also demonstrates the effectiveness of the present methodologic protocol for evaluating PRF protocols.

In this study, the A-PRF protocol resulted in a more evenly distributed upper platelet layer throughout the PRF plasma layer, further validating the LSCC (Fig 2-18). Tubes at lower g-forces centrifuged for less time consistently resulted in a better distribution of platelets throughout the PRF matrix in the upper 4 to 5 mL, whereas an uneven distribution of cells was found using the 700g by 12-minute protocol. It is therefore clinically recommended to avoid utilizing original L-PRF protocols for membrane fabrication because all the cells are entirely found in a thin layer at the base of the PRF clot. However, low leukocyte yields were observed utilizing these protocols.

Fig 2-18 Summary of the findings comparing L-PRF and A-PRF protocols. While neither was typically able to collect leukocytes (reviewed later in the chapter), the lower centrifugation speeds using the A-PRF protocol allowed for the more even distribution of platelets in the upper layers. (Reprinted with permission from Miron et al.⁵²)

This study also led to the observation that the manufacturer’s recommended protocol for i-PRF (~60g for 3 minutes) was not adequately effective at separating cell types or producing high yields of platelets/leukocytes. Figure 2-17 demonstrates minimal change in cell layer changes following this short centrifugation cycle at low RCF values. Based on the data obtained within this study, a paradigm shift in our understanding of platelet concentrates with respect to the LSCC was noted. We now know that too-low RCF values/times will produce ineffective separation of blood layers, as demonstrated in these i-PRF protocols.

Based on these findings, we now know that too-low RCF values/times will produce ineffective separation of blood layers.

Pitfalls in i-PRF protocols

Several interesting findings were observed more recently with respect to the original i-PRF protocols. In 2016, when the first PRF textbook was written, Miron and Choukroun wrote the following: “Interestingly however, total growth factor release of PDGF-BB, VEGF, and TGF-β1 were significantly higher in PRP when compared to i-PRF. Methods to further understand these variations are continuously being investigated in our laboratory as well as others. It may be hypothesized that the differences in spin protocols are suggested to have collected slightly different cell populations and/or total growth factors responsible for the variations in release over time.”

Back then, it remained puzzling to our research team why higher GF content was found in PRP despite the protocols utilized. Based on the results from the newly developed quantification method, it was revealed very clearly that the reason for these lower levels of GFs released from i-PRF was in fact owing to its inability to fully shift cells to their correct blood layers because centrifugation was carried out at too low a speed/time. Figure 2-18 demonstrates that only roughly 30% of total platelets are in fact accumulated in i-PRF, with 70% remaining in the lower RBC layer.⁵² Furthermore, a separate study conducted by an independent group found only minimal improvements in platelet numbers following i-PRF protocols (less than 50% increase), with an actual reduction in leukocyte concentration as well as VEGF release.⁵³ Therefore, it must clearly be noted that with respect to the LSCC, data has now demonstrated that it is definitely possible to centrifuge too slowly and too little a time for effectiveness. It became clear that improvements could be made to these i-PRF protocols.

Optimization of i-PRF into C-PRF

Based on our findings that following L-PRF protocols, the majority of cells were massively accumulated at the buffy coat directly above the RBC layer (within 1 mL) with very few cells found throughout the upper four 1-mL layers,⁵⁰ we hypothesized that if we could specifically collect this 1-mL layer, we would create a much more liquid-PRF formulation rich in cells and GFs (Fig 2-19). In a study titled “A novel method for harvesting concentrated platelet-rich fibrin (C-PRF) with a 10-fold increase in platelet and leukocyte yield,”⁵² we addressed two specific questions: (1) In what total volume were the majority of these cells located above the RBC layer within the buffy coat? and (2) What final concentration could be harvested by collecting only the cells found within this precise buffy coat region when compared to conventional i-PRF protocols?

Fig 2-19 Proposed method to harvest C-PRF. Based on the finding that following L-PRF protocols all cells are accumulated within 300–500 µL above the RBC, this proposed method would collect this 0.3–0.5 mL of liquid C-PRF directly above the RBC junction for a highly concentrated liquid of platelets, leukocytes, and monocytes. (Reprinted with permission from Miron et al.⁵²)

Unlike the previous study, we decided to quantify PRF using 100-µL sequential layers (ie, 0.1 mL) to precisely investigate the exact location of cells (Fig 2-20). Because we had previously observed a massive cell accumulation within the buffy layer in a 1-mL sample range following the L-PRF protocol, we aimed to investigate precisely the volume in which these cells were located within this 1-mL layer. As such, we developed a novel methodologic approach whereby 100-µL sequential layers were pipetted starting from about 1.2–1.5 mL above the buffy coat down to the RBC layer (see Fig 2-20, depicted as +1 to +12 layers). Additionally, three layers were harvested within the RBC layer to determine the number of cells incorporated within this layer as well (see Fig 2-20, depicted as –1 to –3 layers). Each of these 100-µL layers was sent for CBC analysis.

Fig 2-20 A second methodologic illustration depicting the sequential harvesting technique. Because the majority of cells accumulated within the 1-mL within the buffy coat following L-PRF protocols, we sought to investigate precisely the total volume of liquid (mL) above the buffy coat in which cells are concentrated. For the L-PRF protocols, 3.5 mL were removed followed by sequential 100-µL layers pipetted followed by CBC analysis. Three layers in the RBC layer were also harvested. In comparison, all plasma layers of the i-PRF protocol were also harvested in 100-µL sequential layers. Three RBC layers (100 µL each) were also collected. (Reprinted with permission from Miron et al.⁵²)

The second tube from each group was utilized to determine the final concentration from the liquid version of the i-PRF yellow plasma layer. For L-PRF protocols, one tube was utilized to harvest 0.5 mL of a C-PRF (defined as the 0.5-mL buffy coat directly above the RBC layer). This layer was termed concentrated-PRF in reference to the harvesting of this concentrated buffy coat layer (see Fig 2-19). Similarly, 0.3 mL of C-PRF liquid was harvested from this layer as well.

To address our first question regarding the precise volume in the buffy coat in which the increased cell numbers were observed, the first 3.5 mL of the upper plasma layer was removed (acellular layer) from the centrifugation tube (leaving 1.0–1.5 mL of remaining sample above the RBC layer). The sequential pipetting methodology was then utilized with 100-µL layers to accurately determine up to what layer above the RBC the cells were precisely located (see Fig 2-20). Furthermore, 300 µL within the RBC layer was also harvested and quantified in 100-µL sequential layers. In contrast, the entire i-PRF layer was collected starting from the upper 100-µL layer and sequentially pipetted until all plasma layers were collected. Once again, 300 µL was sequentially pipetted in 100-µL layers from the RBC layer.

Figure 2-21 demonstrates the results following sequential pipetting of 100-µL layers of the i-PRF protocol. Notice how specifically in layer +1, a threefold increase (from 5 to 15 × 10⁹ cells/L) is found in leukocytes directly at the buffy coat layer (represented by arrows). Notice also the five- to sixfold increase in monocytes. The RBCs begin to accumulate at layer +1, and by layer –1 the sample is within the RBC layer. The remaining WBC and platelet levels drop within layer –1 after the yellow-red transition (see Fig 2-21). Following the i-PRF protocol, we observed a 2.5-fold increase (from baseline ~220 to ~550 × 10⁹ platelets/L) in platelets in the top 13 layers (1.3 mL) following the i-PRF protocol and only a slight increase in leukocytes.

Fig 2-21 The concentration of cell types in each 100-µL layer utilizing the i-PRF protocol (800 rpm for 3 minutes; ~60g). Notice the significant increase in leukocytes and monocytes in the buffy coat layer (+1; arrows). (Adapted from Miron et al.⁵²)

Figure 2-22 demonstrates the results following sequential pipetting of 100-µL layers of the L-PRF protocol. Interestingly, almost all the cells accumulate within the three layers (ie, 300 µL) above the RBC layer. Most surprisingly, within this layer a massive increase in platelets, monocytes, leukocytes, and lymphocytes was observed. For instance, a roughly 225 to 6,000 × 10⁹ platelets/L increase was observed, representing a > 25-fold increase in platelet concentration, specifically 100 µL above the RBC layer.

Fig 2-22 The concentration of cell types in each of the layers (100 µL each) above the RBC layer following the L-PRF protocol. Notice the massive increase in platelets (roughly a 20-fold increase) specifically at the buffy coat layer between the yellow plasma and RBC layers. Interestingly, all cells seemed to accumulate within the three to five layers (300–500 µL) above the RBC layer. (Adapted from Miron et al.⁵²)

Based on these results, we assumed that a 0.3- to 0.5-mL layer of C-PRF could be preferentially collected within this buffy coat directly above the RBC layer. Figure 2-23a demonstrates that while the i-PRF protocol increases leukocyte numbers 1.23-fold, a marked and significant increase representing a 4.62- and 7.34-fold increase was observed with both 0.5 mL and 0.3 mL of C-PRF, respectively. Even more pronounced is that while i-PRF protocols have typically been shown to increase platelet yields between 200% and 300%, the C-PRF protocols massively increased platelet yields 1138% and 1687%, respectively (Fig 2-23b). A similar trend was also observed for monocytes (Fig 2-23c). The total values following averages from six patients are summarized in Table 2-2.

Fig 2-23 Concentration of leukocytes (a), platelets (b), and monocytes (c) following centrifugation using i-PRF protocols versus collecting 0.3 to 0.5 mL of C-PRF. Notice that while i-PRF was typically responsible for a 1.2- to 2.5-fold increase in the various cell types following centrifugation, up to a 15-fold increase in platelet concentration could be achieved with C-PRF. An asterisk (*) represents a significant difference when compared to i-PRF; a double asterisk (**) represents a value significantly higher than all groups; P < .05. (Adapted from Miron et al.⁵²)

Table 2-2 Leukocyte, platelet, and monocyte concentrations in whole blood compared to i-PRF and C-PRF

Chapter 3 discusses this new method of concentrating liquid-PRF directly from the buffy coat layer in more detail and reports on the marked improvement in cellular activity when compared to original i-PRF protocols.

Horizontal Centrifugation of PRF

Until 2019, the majority of centrifugation carried out for the production of PRF was performed on fixed-angle centrifuges. Horizontal centrifugation, on the other hand, is utilized frequently in research laboratories and in medical hospitals due to its superior ability to separate layers based on their density (Fig 2-24). In fact, the original PRP systems that date back 20 years were brought to market utilizing horizontal centrifugation for this very reason⁵⁴ (see chapters 3 and 4). In an attempt to investigate cell layer separation utilizing horizontal centrifugation, the same layer-by-layer sequential pipetting (1 mL each) was utilized as previously done with the L-PRF and A-PRF protocols.

Fig 2-24 Illustrations comparing fixed-angle and horizontal centrifugation. (a) With fixed-angle centrifuges, a greater separation of blood layers based on density is achieved owing to the greater difference in RCF-min and RCF-max. (b) Following centrifugation on fixed-angle centrifuges, blood layers do not separate evenly, and as a result, an angled blood separation is observed. In contrast, horizontal centrifugation produces an even separation. (c) Because of the large RCF values (~200g–700g), cells are pushed toward the outside and downward. On a fixed-angle centrifuge, cells are pushed toward the back of centrifugation tubes and then downward/upward based on cell density. These g-forces produce additional shear stress on cells as they separate based on density along the back walls of centrifugation tubes. In contrast, horizontal centrifugation allows for the free mobility of cells to separate into their appropriate layers based on density, allowing for more optimal cell separation as well as less trauma/shear stress on cells. (Reprinted with permission from Miron et al.⁵⁰)

Figure 2-25 depicts a 700g force performed on a horizontal centrifuge for 8 minutes (the selection of this protocol is presented in chapter 3). Interestingly, it was observed that more leukocytes, platelets, lymphocytes, and monocytes were much more evenly distributed throughout the PRF layers when compared to fixed-angle centrifugation. Unlike either the L-PRF or A-PRF protocols, a general increase in leukocyte numbers was observed (127% original values), and a 2.4-fold increase in platelet concentration was observed. This represents over a fourfold increase in leukocytes when compared to A-PRF protocols and a twofold increase when compared to L-PRF. Furthermore, this method concentrated 99.7% of all platelets and 53% of all leukocytes within the plasma layer.⁵⁰

Fig 2-25 The concentration of cell types in each 1-mL layer utilizing the solid-PRF horizontal centrifugation protocol (700g for 8 minutes). Notice that most of the platelets as well as WBCs are now more evenly distributed throughout the upper plasma layer. (Adapted from Miron et al.⁵⁰)

Within that study, horizontal centrifugation was proposed as a means to better separate cell layers based on density (Video 2-3). Two advantages were noted utilizing horizontal centrifugation. First, a completely horizontal tube produced from a swing-out bucket allows for the greatest differential between the minimum and maximum radius found within a centrifugation tube (see Fig 2-24a). This allows for a greater ability to separate cell layers based on disparities between the RCF-min and RCF-max produced within a tube. Second, a fixed-angle centrifuge results in more trauma to cells. Because centrifugation typically pushes cells outward and downward, cell layer separation is always observed in an angulated fashion toward the back distal surface of PRF tubes using fixed-angle centrifugtion (see Fig 2-24b). Furthermore, during the centrifugation process, cells are pushed toward the outer wall and then typically migrate either up or down the centrifugation tube based on density. Larger cells (such as RBCs) entrap smaller cells such as platelets during the centrifugation process and drag them downward along the back centrifugation wall into the RBC layer as a result of this cell accumulation against the back wall (see Fig 2-24c). In contrast, PRF produced via horizontal centrifugation and separation allows the direct flow-through of cells (see Fig 2-24c).

Video 2-3

Therefore, horizontal centrifugation allows cells to migrate freely throughout the blood layers. This allows for better separation of cell types (along with the greater differentiation in RCF values between RCF-min and RCF-max), resulting in higher final concentrations of desired cells (platelets and leukocytes) within the appropriate final blood layers. Furthermore, cells are less likely to suffer potential damage along the back wall of centrifugation tubes produced using high g-forces (~200–700g) following fixed-angle centrifugation. We therefore introduced this concept as “gentler centrifugation,” whereby cells are more freely able to separate between layers without the necessary friction produced on the back wall of fixed-angle centrifuges such as those produced on the fixed-angle IntraSpin and Process for PRF devices.⁵⁰ This concept has been expanded with a series of research investigation launched to better optimize PRF. This is presented in greater detail in chapter 3 of this textbook.

During the centrifugation process on fixed-angle centrifuges, cells are pushed toward the outer distal wall and then typically migrate either up or down the centrifugation tube based on density. Larger cells (such as RBCs) entrap smaller cells such as platelets during the centrifugation process and drag them downward along the back centrifugation wall. These platelets and leukocytes don’t make it to the upper PRF membrane.

Biologic Activity of PRF on Immune Cells

In an extensive systematic review by Strauss et al investigating the biologic properties of PRF, 1,746 studies were identified, of which 53 were included.⁵⁵ Because PRF is capable of improving angiogenesis in vivo, it was reported that PRF enhanced the proliferation, migration, adhesion, and osteogenic differentiation of a variety of different cell types along with cell signaling activation. Furthermore, it was concluded that PRF reduced inflammation, suppressed osteoclastogenesis, and increased the expression of various GFs in mesenchymal cells.⁵⁵

Several interesting and very recent studies have investigated the effects of PRF and its involvement with macrophage polarization from proinflammatory M1 toward proresolving M2 phenotypes.⁵⁶ Macrophages are extremely important cells during the healing process and can be involved with either secretion of proinflammatory markers (M1) or proresolution markers (M2). These studies are extremely relevant to PRF.

In a study by Nasirzade et al, murine primary macrophages and a human macrophage cell line were exposed to saliva and lipopolysaccharides (LPS) with and without PRF lysates.⁵⁶ The expression of the proinflammatory M1 marker genes IL-1β and IL-6 were greatly decreased, and PRF-conditioned medium enhanced the expression of tissue-resolution markers. It was therefore concluded that PRF holds an anti-inflammatory activity and shifts the macrophage polarization from an M1 toward an M2 phenotype.⁵⁶ In a second study on this topic, it was also reported that i-PRF reduced proinflammatory M1 phenotype of macrophages along with activated dendritic cells around muscle defects injected with bacterial suspension (Figs 2-26 and 2-27).⁴¹

Fig 2-26 The expressions of CD11b in tissues from control (a and c) and i-PRF groups (b and d). Scale bar in a and b = 20 μm; scale bar in c and d = 10 μm. (e) Quantification of the immunohistochemical staining was calculated by the percentage of CD11b-positive cells in all cells in the area of the same size. ****P < .0001. Error bars indicate SD. (Reprinted with permission from Zhang et al.⁴¹)

Fig 2-27 Effects of i-PRF on the maturation of dendritic cells. (a and b) Sections of immunofluorescence with antibodies against CD11c and CD86. Scale bar = 20 μm. (c) Relative expressions of maturation-related and inflammatory-related genes in DC2.4. (d and e) Immunoblotting and relative intensity of NF-κB signaling pathway and maturation-related markers of DC2.4 stimulated by the whole blood (WB) or i-PRF in the presence of LPS. *P < .05; **P < .005; ***P < .0005; ****P < .0001; ns, no significant difference. Error bars indicate SD. (Reprinted with permission from Zhang et al.⁴¹)

While the study of PRF on various cell types such as immune cells has only begun, it appears that it acts to reduce the proinflammatory response and further decrease common oral cavity inflammatory responses to LPS. Thus, this work may in part explain the observed clinical decreases in postoperative pain reported in the clinical chapters discussed later in this textbook.

PRF appears to reduce the proinflammatory response and further decrease common oral cavity responses to LPS.

Anti-inflammatory and Antibacterial Properties of PRF

Both the anti-inflammatory and antibacterial properties of PRF have been a topic of much interest in recent years following the clinical observation that PRF seems to reduce postoperative swelling and pain. In a study titled “Effects of liquid platelet-rich fibrin on the regenerative potential of hPDLCs cultured under inflammatory conditions,” our group investigated the effects of PRF on human periodontal ligament cells (hPDLCs) under inflammatory conditions. As a model, hPDLCs were investigated using a migration and proliferation assay. To investigate hPDLC differentiation, alkaline phosphatase (ALP) assay, Alizarin Red Staining, and gene expression levels of Runx2, Col1a1, and OCN were conducted. Furthermore, cells were cultured with LPS to induce an inflammatory condition to investigate the ability of PRF to impact inflammatory resolution. All assays were compared to PRP (lower in WBCs).

Osteogenic differentiation demonstrated that liquid-PRF significantly induced greater ALP activity and more mineralized nodules when compared to PRP and controls (Fig 2-28). According to the experimental timeline, cells were pretreated with or without LPS to induce an inflammatory condition for 7 days, and then liquid-PRF was added to the culture media for an additional incubation period of 7 days (Fig 2-29a). Immunofluorescence images demonstrated that LPS induced more p65 expression (a marker for inflammation; Fig 2-29b), while the addition of liquid-PRF decreased its expression level. Furthermore, other inflammation markers including IL-1β and TNF-α were also significantly downregulated, as confirmed by real-time polymerase chain reaction (RT-PCR; Fig 2-29c). In summary, it was concluded that liquid-PRF displayed an anti-inflammatory response when hPDLCs were cultured with LPS.

Fig 2-28 Differentiation of hPDLCs. (a and b) Effects of PRP and liquid-PRF on ALP activity were detected by ALP staining and ALP activity test, respectively. (c) Alizarin Red S staining showed the mineralized nodules in each group after induction for 14 days. (d) The semiquantification of mineralization level. (e) Relative gene expression levels of Col1a1, OCN, and Runx2 after being treated with PRP or liquid-PRF for 14 days. Error bars correspond to the mean ± SD; significant differences are indicated: *P < .05; **P < .01.

Fig 2-29 Liquid-PRF can decrease the inflammation induced by LPS. (a) The timeline of experimental inflammatory condition stimulation. (b) Immunofluorescence staining of p65 in hPDLCs after being cultured with or without LPS and/or liquid-PRF. (c) The relative gene expression levels of inflammatory markers including IL-1β, TNF-α, and p65. Error bars correspond to the mean ± SD; significant differences are indicated: *P < .05; **P < .01; ns, not statistically significant vs control group.

In a final experiment, it was observed that liquid-PRF promoted the osteogenic differentiation of hPDLCs even when cultured in an inflammatory environment. Briefly, cells pretreated and cultured with LPS resulted in an intense reduction in mineralization nodule formation (Figs 2-30a and 2-30b). As the previous experiment demonstrated the ability for liquid-PRF to decrease the inflammatory response, it was further found that liquid-PRF could actually reverse a decrease in mineralization observed by LPS and resulted in a significant upregulation of expression markers Runx2, Col1a1, and OCN (Fig 2-30c). These findings indicate that the anti-inflammatory effect and regenerative potential of liquid-PRF can counterbalance the negative inflammatory effect induced by LPS. Later chapters address these findings more specific to the periodontal field, because PRF has been shown to improve the regeneration of intrabony and furcation defects not only by improving GF release but also by counterbalancing the inflammatory response induced by LPS (see chapter 10).

Fig 2-30 Liquid-PRF can promote the osteogenic potential of hPDLCs in an inflammatory environment induced by LPS. (a) Alizarin Red S staining indicated the odontoblastic differentiation of hPDLCs in the presence of LPS and/or liquid-PRF. (b) Mineralization level. (c) Gene expression levels of Runx2, Col1a1, and OCN in inflammatory condition. Error bars correspond to the mean ± SD; significant differences are indicated: *P < .05; **P < 0.01; ns, not statistically significant vs control group.

Furthermore, it has also been shown that PRF exerts potent antibacterial properties. In another study by our group, PRF was separated into solid and leachate components, and the PRF clots were divided into five equal layers to explore the specific antibacterial aspects of PRF. Both antimicrobial tests and flow cytometric analysis revealed that PRF produced using horizontal centrifugation demonstrated a significantly better antibacterial effect than L-PRF and was strongly correlative with immune cell numbers and types (Fig 2-31). In addition, our results demonstrated that the antimicrobial ability of PRF clots were less efficient than the wet PRF containing leachate, which suggest a promising application guidance to retain the liquid components of PRF for better anti-infection properties during clinical use. This study is presented in greater detail in chapter 3.

Fig 2-31 Antibacterial properties of L-PRF and H-PRF. (a) Photographs of PRF obtained after centrifugation by both protocols. Note the horizontal layer centrifugation in H-PRF versus the angled layering in L-PRF. (b) Weight and size measurements of PRF matrices. (c and d) Photographs and quantitative analysis of bacterial colony of Staphylococcus aureus and Escherichia coli incubated with L-PRF or H-PRF clots for 4 hours. (e and f) Photographs and quantification of the inhibition zone of L-PRF and H-PRF membranes incubated with S aureus or E coli after 24 hours. *P < .05; **P < .01; ***P < .001; ns, not statistically significant.

The Effect of Age, Sex, and Time on the Size Outcomes of PRF Membranes

Two topics that were heavily questioned for many years were (1) How long does the clinician have from the start of blood draw? and (2) Why do colleagues observe so much variability in clot size even from blood draws coming from the same patient? In 2019, we addressed this topic in a publication where the final PRF size outcomes were compared following centrifugation that took place after 0, 30, 60, 90, and 120 seconds in both male and female patients of different age categories. Each participant donated six vials of blood, and centrifugation was begun precisely after 0, 30, 60, 90, and 120 seconds.⁵⁷

As depicted in Fig 2-32, by 90 seconds already a drop in membrane size of 13% was observed, and by 120 seconds this dropped even further. The entire goal of centrifugation is to separate layers based on density, so when blood remains sitting in a centrifugation tube for 120 seconds, it is certain that some of the fibrinogen and thrombin are beginning to convert into fibrin. Thereafter, when centri-fugation begins, it becomes harder and harder to separate layers (and most importantly cell types as a result). Therefore, centrifugation should be carried out between 60 and 90 seconds after blood draw. It generally takes 15 seconds to fill the tube with blood to begin with, sometimes even longer (Fig 2-33).

Fig 2-32 Average size of PRF membranes from 60 patients following blood draw after an initial wait period of 0, 30, 60, 90, and 120 seconds prior to centrifugation. Notice that after 90 seconds, the PRF membranes were significantly reduced in size (by 13%). Following a 120-second wait period, these membranes were further significantly reduced in size (by 23%) compared to the control (0-second wait period). *P < .05 indicates a significant difference between 0 seconds from centrifugation and the investigated time period of 90 and 120 seconds. (Adapted from Miron et al.⁵⁷)

Fig 2-33 Required time interval to fill each manufacturer’s PRF tube. Note that in general it takes roughly 15 seconds, but some manufacturers have slower-filling tubes. (Adapted from Miron et al.³⁹)

Always remember that the entire goal of centri-fugation is to separate layers based on density. When blood sits in a centrifugation tube for 120 seconds, it is certain that some of the fibrinogen and thrombin are beginning to convert into fibrin.

One noticeable trend was that the size of the membranes produced between male and female patients was different. On average, the size of PRF membranes produced by females was 17% larger than those produced in males (Fig 2-34). As the role of centrifugation is to separate blood layers transitionally over time, these differences were due to females generally reporting lower hematocrit levels within their peripheral blood compared to males.^58,59 The same trend was also observed in older patients (Fig 2-35). This is also due to the fact that as one ages, a lower hemat-ocrit count is usually noted, and therefore the “density” of blood is lowered (fewer RBCs). Seemingly, it is easier for blood layers to separate accordingly.

Fig 2-34 (a) Comparison of the average size of PRF membranes between males and females in 60 patients. Notice that at earlier time points, the female PRF membranes were significantly larger compared to male PRF membranes. *P < .05 indicates a significant difference between male and female PRF membrane sizes (%). (b) Bar graph representing the average size of PRF membranes from males and females in 60 patients. On average, the female PRF membranes were 17% larger. *P < .05 indicates a significant difference between male and female PRF membrane sizes (%). (Adapted from Miron et al.⁵⁷)

Fig 2-35 Comparison of the average size of PRF membranes between various age groups: 21–40 years, 41–60 years, and 61–80 years. While no significant differences were noticed between the groups, in general older patients produced larger membranes. (Adapted from Miron et al.⁵⁷)

Because centrifugation separates blood based on density, it is important to note that variability will exist. Females and older patients have less hematocrit when compared to men and younger individuals. As a result, PRF clots produced in older females will be significantly larger than young males (especially young athletes or those living at high altitude). Generally, in the younger male population or patients routinely living at high altitude, a 20% increase in the RCF values of each protocol is recommended.

Conclusion

The use of GFs in dentistry has gained tremendous momentum and popularity in recent years, especially because of the easily obtainable and low-cost group of platelet concentrates. Autologous PRF is a 100% natural blood-derived tissue engineering scaffold that is totally physiologic and safe and may be utilized for the purpose of wound healing. This chapter outlined the main GFs and cell types found in PRF and further demonstrated the massive effect of centrifugation parameters on the final cell layer separation in PRF. Major advancements with respect to first utilizing the LSCC and thereafter horizontal centrifugation have more recently optimized the final production of PRF. Future research remains ongoing to further highlight all the biologic properties and advantages of PRF, such as its ability to regulate immune cells as well as participate in antimicrobial defense. In summary, PRF serves as an excellent tissue engineering scaffold by fulfilling its three main criteria: scaffold (fibrin), cells (platelets and leukocytes), and GFs (PDGF, VEGF, TGF-β).

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