Platelet-Rich Plasma in Orthopedics

Platelet-Rich Plasma in Orthopedics

Jennifer E. Woodell-May1 and William S. Pietrzak2

Abstract: Autologous platelet-rich plasma (PRP) has become a popular

clinical treatment in a variety of soft tissue and hard tissue applications.

Clinicians use PRP to harvest the platelets’ natural ability to promote hemostasis

and to release cytokines into the wound bed with hopes of stimulating

the rate of healing and improving tissue quality. Platelet-derived growth factor,

transforming growth factor β1, vascular endothelial growth factor, basic

fibroblast growth factor, epidermal growth factor, insulin-like growth factor

and connective tissue growth factor are among the more notable cytokines

released from platelets. To understand the utility of PRP in orthopedics, a

review of the preclinical studies with PRP reveals a wide breadth of models

where PRP enhances the outcomes. However, it is apparent that PRP is most

successful when paired with the appropriate matrices and/or cell therapies for

optimal results. Reviews of peer-reviewed clinical studies, ranging from Level

I evidence clinical trials to Level IV case series reports, also reveal a wide

range of clinical utility for PRP. Future clinical trials should attempt to refine

the clinical application of PRP for each indication for use. These studies must

be designed with the appropriate outcome measures and follow-up time-points

to capture the benefits of PRP. Given the data published to date, PRP appears

to be a powerful autologous therapy for surgeons looking to enhance bone and

soft tissue formation. Future investigations will further define PRP’s optimal

role in medicine.

Keywords: Platelet-rich plasma, PRP, platelet concentrate, platelet gel, AGF,

growth factors, platelets, PDGF, TGF-β, orthopedics, wound healing.

26.1. Introduction

The healing response to injury has evolved over millions of years to promote

survival of a species. While humans possess sufficient healing potential to

survive a variety of injuries, healing may take a long time to complete or the

regenerated tissue may consist primarily of nonfunctional scar. A complex

healing process at a tissue injury site is initiated by platelets, which are

responsible for arresting bleeding and providing hemostasis. These same

platelets, upon activation by mediators at the injury site, release bioactive

proteins that signal wound healing cells to clean the wound and form new

tissue. Recent attempts have been made to increase the healing rate and functionality

of deposited tissue by augmenting the natural healing process and

harnessing the healing potential of platelets.

An emerging clinical technology attempts to harvest the body’s natural healing

capacity by collecting platelets from a patient, concentrating them in a small

amount of plasma and administering the platelets to the patient at the injury site.

This concentrated platelet product is known as platelet-rich plasma (PRP). Other

names include platelet concentrate and autologous growth factor (AGF). Once

activated to form a gel, it can also be called platelet gel or autologous platelet

gel.

PRP has been used in many fields of surgery. Applications in orthopedics

include spine, joint arthroplasty, dental, craniomaxillofacial, sports medicine and

foot and ankle procedures [1–6]. PRP has also been useful in other fields such

as cardiovascular, plastic surgery and ulcer wound healing [7–9]. While many of

these studies attempt to determine if an autologous platelet approach does enhance

wound healing mechanisms, a definitive conclusion has not been reached. First,

many nuances in the production of PRP include the level of platelet concentration

and the ability to not prematurely activate the platelets and lose the active factors

during the platelet processing [10]. Second, many of the commercially available

PRP systems have not been well characterized, which makes interpretation of

results difficult [11]. Finally, few prospective, randomized clinical studies have

been performed from which to draw unambiguous conclusions.

Despite these limitations, a growing body of evidence – taken collectively –

seems to support a clinical role for PRP in wound healing. The purpose of this

chapter is to provide a basic background of platelets, their role in wound healing,

methods of concentration and administration and an objective summary

of animal and human studies that investigate the role of PRP in wound healing

for orthopedics.

26.2. Basic Science

26.2.1. Platelet Biology

Platelets are fragments of large bone marrow cells called megakaryocytes and

are formed during hematopoiesis. They are approximately 2 to 4 μm in diameter

with a volume of 10×10−9 mm3 [12, 13]. Platelets lack a nucleus, but do contain

organelles such as mitochondria, dense bodies, α-granules and lysosomal granules.

The dense bodies contain adenosine diphosphate (ADP), adenosine triphosphate

(ATP), Ca2+, serotonin, histamine, dopamine and catecholamines [14]. The

α-granules, which number about 50 to 80 per platelet, contain adhesive proteins, coagulation factors, fibrinolytic factors, antiproteases, mitogenic growth factors,

cytokines and bactericidal proteins [10, 14]. The phospholipid bilayer platelet

membrane is covered with glycoprotein receptors that mediate interactions

with surfaces, bioactive molecules and other platelets [13, 14]. Conformational

changes in platelets, which are integral to their function, are mediated by actin

and myosin fibers within the platelets [14].

Normal human platelet concentration is 200,000 to 400,000 platelets per microliter,

and they typically circulate for a 10-day lifespan [10, 12]. Platelet physiologic

function is two-fold: 1) hemostasis and 2) initiation of wound healing [10].

26.2.2. Hemostasis

Platelets maintain hemostasis by formation of a platelet plug and by modulating

fibrin formation via the coagulation cascade [13]. When a blood vessel

is injured, collagen becomes exposed. The exposed collagen interacts with

the glycoprotein receptors of the platelet membrane, resulting in a signal

transduction cascade that causes the platelets to adhere to each other and to

undergo conformational change [13]. These conformational changes, including

pseudopodia formation, are associated with release of internal stores of

Ca2+ within the platelet, which in turn stimulates the released of the contents

of the α-granules and dense granules outside of the platelet [12]. This process

is known as platelet activation (Fig. 26.1) [15]. The substances released from

the activated platelets and into the surrounding plasma, specifically ADP

and thrombin, activate adjacent platelets. These activated platelets aggregate

together, forming the platelet plug [12, 13].

Along with the platelet plug formation, another major component of a blood

clot is the fibrin mesh, which is derived through the coagulation cascade. The

coagulation cascade is the process through which a series of normally inactive

plasma proteins become activated, either by the intrinsic pathway through

interactions with a surface, or extrinsically by the presence of tissue factor

[16]. These activated factors continue a feed-forward loop that results in the

formation of a three-dimensional fibrin matrix.

Both pathways converge at the activation of Factor X, which prompts the

conversion of prothrombin to thrombin. Thrombin catalyzes the formation of

fibrin from fibrinogen, which first forms a fibrin thread and then ultimately

cross-links into a stable three-dimensional mesh [13]. The fibrin matrix,

platelet plug and trapped white and red blood cells together make up the

thrombus, a clot that stops bleeding at an injured site. Figure 26.2 details

the schematic of the coagulation cascade and illustrates the multiple levels

within the coagulation cascade during which activated platelets can directly

participate in the process.

As can be seen from Fig. 26.2 several steps in the cascade require the

presence of calcium ions. The presence of calcium forms the basis for citratebased

anticoagulants, or blood preservatives, such as citrate-phosphatedextrose

(CPD) and acid-citrate-dextrose (ACD) [10]. The added citrate ions

chelate the calcium ions, essentially disabling the calcium ion-dependent steps

of the cascade, which can be reversed by adding a calcium salt, e.g., CaCl2.

26.2.3. Wound Healing

The wound healing cascade is a temporal series of events that begins within

seconds of the injury and continues with remodeling for months following the

event. The basic nature of the cascade is similar, whether the injury occurs

in hard or soft tissue, with local signaling of precursor cells to ensure the

appropriate tissue-specific phenotype modulation. The general healing process

can be divided into three stages, including inflammatory, proliferative and

remodeling phases [10, 17, 18].

The inflammatory phase (beginning immediately after the injury) includes

initial hemostasis that involves platelet activation, aggregation and formation

of a fibrin matrix, as described above. During degranulation, or release of the

á-granule contents during platelet activation, platelets initiate the coagulation

cascade and release cytokines that are responsible for cueing the wound

healing process. The cytokines released are chemotactic for circulating white

blood cells (WBC), recruiting them to marginate out of nearby blood vessels

and migrate into the wound bed. Neutrophils, the first WBC responders, start

cleaning the site by removing bacteria and cell debris [17, 18].

During the proliferative phase (days) an influx of monocytes migrates to

the site, signaled by the growth factors released from the platelets. These

circulating monocytes differentiate into macrophages. As the platelets are

removed from the wound site, the activated macrophages take over the

signaling-modulation role from the platelets. Macrophages remove debris

by phagocytosis and also secrete factors that promote further wound healing

events. Following the macrophages, fibroblasts begin to lay down collagen

granulation tissue. New capillaries begin to feed the repair area, marking the

start of angiogenesis. During this time undifferentiated stem cells migrate to

the injury site. Depending on the signals present in the wound, the stem cells

can differentiate into tissue-specific cells such as bone, cartilage or new blood

vessels [17, 18].

The final phase of wound healing is the remodeling phase (months).

During this phase the collagen tissue contracts, bringing the edges of the

wound together. Cell density and vascularity decrease, excess repair matrix

is removed and the collagen fibers orient along lines of stress to maximize

strength [10]. The granulation tissue itself accumulates and remodels into scar

tissue or is turned over to specific tissue types like skin or bone [17, 18]. In

general, soft tissue heals by scar formation, although some components of the

original tissue may have re-formed within the scar. Bone is unique in that it

typically heals without scar; that is, healed bone cannot be distinguished from

uninjured bone [10].

26.2.4. Growth Factors Released from Platelets

The cytokines released from activated platelets signal wound healing events

to occur. Among the releasate is platelet-derived growth factor (PDGF)

(isoforms AA, AB, BB), transforming growth factor â (TGF-â1 and TGF-â2),

vascular endothelial growth factor (VEGF), basic fibroblast growth factor

(bFGF), epidermal growth factor (EGF), insulin-like growth factor (IGF),

connective tissue growth factor (CTGF) and others [10, 14; 19; 20]. Several

of these signaling proteins, in particular PDGF and TGF-â, are transformed

into active states during platelet degranulation by the addition of histones and carbohydrate side chains [10]. Table 26.1 gives an example of a growth factor

profile in whole blood and delivered in a commercial PRP preparation [19].

Since platelets modulate wound healing in every tissue in the body, it is not

expected that tissue-specific morphogens would be contained within platelet

releasate. Specifically, large amounts of bone morphogenetic proteins, capable

of differentiating mesenchymal stem cells into osteoblasts and chondrocytes,

are not found in platelets. However, small amounts of BMP-2, BMP-4 and

BMP-6 have been identified in washed platelet releasate, detected by western

blot [21]. In our laboratory we were able to detect, using the enzyme linked

immunosorbant assay method (ELISA), small amounts of BMPs from PRP

releasate, i.e., 23 pg/ml of BMP-2 and 46 pg/ml of BMP-4 (unpublished data).

No detectable amount of BMP-7 was found. These are negligible amounts

compared to the BMP content of human demineralized bone matrix (DBM),

where we found, on average per gram of DBM powder, 21 ng of BMP-2, 5 ng

of BMP-4 and 84 ng of BMP-7 [22].

The effects of many growth factors on cell behavior and wound healing

have been studied. Release of PDGF into a wound bed can have a chemotatic

effect on monocytes, neutrophils, fibroblasts, mesenchymal stem cells and

osteoblasts. PDGF is also a powerful mitogen for fibroblasts and smooth

muscle cells, and is involved in all three phases of wound healing, including

angiogenesis, formation of fibrous tissue and reepithelilization [23]. In a clinical

study pressure ulcers that were treated daily (100 ìg/g or 300 ìg/g) with a

PDGF-BB wound healing gel showed significantly decreased ulcer volume,

compared to ulcers treated with a gel lacking PDGF-BB [24].

TGF-â1 is a mitogen for fibroblasts, smooth muscle cells and osteoblasts.

Additionally, it promotes angiogenesis and extracellular matrix production

[23, 25]. In a rat tibial fracture model, injections of TGF-â (4 and 40 ng) every

other day for 40 days resulted in a dose-dependent increase in bone thickness,

with the 40 ng dose additionally increasing mechanical strength [26].

As with many growth factors, dosing is critical. Broderick, et al. injected a

much higher dose (335 ìg of TGF-â) in a canine humeral model and found a

decrease in bone mineralization [27].

Additionally, actions of other growth factors present in platelet releasate

have been described. VEGF promotes angiogenesis and can promote healing

of chronic wounds and facilitate endochondral ossification [25, 28]. However,

as seen with high doses of TGF-â, high doses of VEGF (0.5 ìg into rat

segmental defect) inhibited bone formation [29]. EGF, another platelet-derived growth factor, is a mitogen for fibroblasts, endothelial cells and keratinoctyes,

and is also useful in healing chronic wounds [25]. IGF, also found in platelets,

regulates bone maintenance, is an important modulator of cell apoptosis and,

in combination with PDGF, can promote bone regeneration [30, 31]. CTGF,

found in concentrations 20-fold higher than other growth factors, promotes

angiogenesis, cartilage regeneration, fibrosis and platelet adhesion [20].

Furthermore, many synergistic effects between these growth factors have

been found. In a rabbit limb ischemia model the combined administration of

VEGF and bFGF significantly increased angiogenesis in the ischemic limb

over either factor given alone [32]. Another synergistic mitogenic effect was

seen in vitro with the addition of bFGF, TGF-â and IGF-2 to osteoblasts [33],

while chemotactic effects were seen with the addition of TGF-â1 and PDGFBB

to osteoblasts [34]. Additionally, in a porcine wound healing model,

increased collagen content and maturity, increased angiogenesis and increased

connective tissue volume were found without an increase in inflammation

when PDGF was used in combination with IGF-1 or with TGF-á [35].

These factors are actively secreted from the á-granules within 10 minutes of

clotting, with more than 95 percent of the presynthesized growth factors secreted

within the first hour [10]. In addition the secreted growth factors remain bound

to the fibrin clot and are slowly released during clot degradation. Therefore, the

platelets local to the wound site continue to directly signal and modulate wound

healing for several days.

26.2.5. PRP Procurement Methods

Normal whole blood contains approximately 94 percent red blood cells (RBC),

0.06 percent white blood cells (WBC), and 5.9 percent platelets, by number.

A typical PRP can alter this concentration, as an example, to 52.1 percent RBC,

0.26 percent WBC and 47.6 percent platelets. These elements are collected

in the buffy coat, a layer of WBC and platelets that form between the RBC

and plasma during centrifugation. The buffy coat is suspended in a volume of

plasma and delivered back to the patient. Depending on the system used to

prepare the PRP, the concentration of platelets can typically range from four

to eight times greater than the levels in whole blood (11;36). In addition, many

systems allow the collection of a plasma layer, called the platelet poor plasma

(PPP). The platelet poor plasma (PPP) can also be delivered to the patient as

an autologous fibrin product for topical hemostasis [1].

Several commercial systems to produce PRP are commercially available.

These systems can be divided into three basic categories: the apheresis

technique, the single-spin tabletop technique and the double-spin tabletop technique.

The first and oldest technology is the creation of a PRP from the output

of an apheresis device (plateletpheresis). In these systems, whole, anticoagulated

blood is introduced into a bowl that is spinning inside of a centrifuge.

The blood separates along the walls of the bowl into the fractions described

above. Each layer is then transferred to different collection bags. In many

cases the PRP apheresis devices further concentrate the PRP with additional

centrifugation and/or filtration to remove water. These systems usually

process an entire unit of blood, so the volume of PRP produced is typically

greater than with the other methods. Additionally, the RBCs can typically be

re-infused to the patient.

The other two types of systems are considered tabletop devices requiring

only 50 to 100 cc or less of blood. With the double-spin technologies, whole

blood is introduced into a disposable unit and placed into a centrifuge. The

first centrifugation spin, called the soft spin, runs for a relatively short time at

low revolutions per minute (rpm). This spin divides the blood into the components

of RBC, buffy coat and plasma. Depending on the device, the plasma

and buffy coat are transferred, manually or automatically, into a new chamber

for the second centrifugation cycle, or hard spin. The hard spin runs for a

longer time period and at a higher rpm. This spin will create a platelet pellet

at the bottom of the chamber which, once re-suspended in plasma, becomes

the PRP.

Single-spin technologies only use one centrifugation cycle that separates the

blood into RBC, buffy coat and plasma. These devices then have methods to

capture the buffy coat from this disposable, such as a tuned density buoy that

captures the buffy coat and allows it to be extracted from the device.

Table 26.2 summarizes these three types of devices, comparing the amount

of blood drawn for processing, the volume of PRP produced, whether the

packed cell layer can be re-infused into the patient, the fold-increase in platelet

concentration in the PRP over baseline and the percent platelet recovery, which

refers to the percentage of all platelets present in the initial blood draw that are

contained in the final PRP produced. As can be seen, within a given class of

device, a wide range in values exists for most of these characteristics, especially

the fold-increase in platelet concentration and the percent platelet recovery.

The following table provides several parameters to consider when comparing

platelet concentration devices, divided into parameters associated with the

quality of the PRP product produced, the ease of use of the system and options

available with the system.

Quality of Product Ease of Use Options

• Platelet concentration • Time of preparation • Operator input

• Growth factor profile delivered • Number of steps • Volume of blood

draw

• Volume of PRP produced • Sterile barrier steps • PPP collection

availability

• Percent recovery of platelets • RBC re-infusion

capability

• Concentration of white blood

cells

• Cost

• Reproducibility of product

• Activation of platelets during

processing

One of the main challenges in characterizing a PRP is to accurately determine

the platelet concentration since levels can be much higher than most

hematology analyzers are designed to count. Woodell-May, et al. devised

a protocol for validating high PRP platelet counts made with a hematology

analyzer (Cell-Dyn 3700, Abbott Labs) using manual counts for reference,

and established a general method to ensure accuracy of the platelet count

[11]. While utilization of PRP is still being investigated, the addition of a means to expeditiously and accurately determine platelet counts can bolster

the relevance of future study conclusions [39].

The last step in platelet application is to prepare the delivery system. Typically,

activation solution must be made, which generally consists of a solution of

thrombin in 10 percent CaCl2 solution [10]. When added to PRP, calcium

ions reverse the effects of the citrate-based anticoagulant, as explained above,

while the thrombin activates the platelets as well as catalyzes the formation of

the fibrin mesh. The most common delivery system uses a dual syringe spray

apparatus [1]. The PRP is drawn into a 10 cc syringe and the activation solution

is drawn into a 1 cc syringe. Both are connected, in tandem, to the dual spray

apparatus (Fig. 26.3). Both syringe plungers are advanced in unison, with the

solutions mixed in a 1:10 volume ratio and a combined spray exiting a single

orifice. In this way the PRP becomes activated as it is applied to the wound. The

PPP can be delivered in a similar fashion. In bone grafting applications, PRP

can be mixed with bone grafting material such as demineralized bone matrix,

allograft bone chips or synthetic bone void fillers.

26.3. Review of Preclinical Studies

PRP has been studied in a variety of preclinical models including orthopedics,

spine, dental, craniomaxillofacial and sports medicine. A summary of selected

published preclinical studies using PRP can be found in Table 26.3. A (+) was

assigned to the study if any positive result was seen with the PRP, a (o) was

given if no difference was found between the test and control groups, and a (−)

was given when the PRP caused an inhibitory effect.

The varied studies described in Table 26–3 illustrate the nonspecific response

of the body to PRP application, with the local tissue environment modulating

the effect to ensure that the appropriate site-specific tissue is regenerated.

Although open to interpretation, 15 of the 18 studies (83%) appear to show

a positive influence of PRP on wound healing and tissue regeneration, two

(11%) show a neutral effect, and one (6%) shows an inhibitory effect Upon

closer inspection, however, it can be seen that some of the studies that reported

positive PRP results used PRP in conjunction with osteoconductive matrices,

so the effects of PRP in isolation may not have been tested. This is not necessarily

a limitation, however, since a multifaceted approach is often required to

attempt to match the combined osteoconductivity, osteoinductivity and osteogenicity

of the “gold standard” autograft in bone healing applications.

It appears that the mixture of PRP with an osteoconductive carrier, such as

allograft bone or ceramic bone void fillers, can enhance bone formation over

the carrier alone [45, 49, 52–53, 55]. However, in more challenging models

for bone formation, such as in a gap model around an implant or in a posterolateral

spine fusion, PRP mixed with an osteoconductive matrix could not

significantly improve the outcome [47, 55]. To overcome these challenges,

adding bone marrow aspirate or cultured mesenchymal stem cells to the graft

in addition to the PRP performed as well as autograft [41–43, 51, 54–55].

In vitro support of the combination of PRP with bone marrow was shown with

a dose-dependent increase in stem cell proliferation with the addition of PRP

[43]. This composite approach avoids the donor site morbidity associated with

autograft harvest.

Applications that take advantage of the angiogenic effect of PRP are illustrated

by the increased tissue repair seen after injection of PRP into an ACL

or Achilles tendon [56, 57]. Additionally, adding PRP to a bone repair model

with compromised vascularity, such as in a diabetic animal, improved fracture

healing [48]. In a critically sized segmental defect model, increased vascular

invasion was seen in groups implanted with PRP, cultured stem cells and allograft

bone, over groups implanted with the allograft alone [43].

The proliferative and angiogenic growth factors delivered in PRP,

when combined with DBM, might be expected to accelerate bone healing

compared to DBM alone. Ranly, et al. added the single growth factor,

PDGF, to DBM powder in an ectopic bone formation model in a nude

mouse [44]. An insignificant increase in DBM osteoinductivity was seen

with the lowest dose, 0.1 μg PDGF/10 mg DBM implant, but there was an

inhibition of osteoinductivity with the two higher doses of 1 μg PDGF/10 mg

DBM implant and 10 μg PDGF/10 mg DBM implant. The low dose compares

to that which may be present in a typical 6 cc PRP preparation [19]

( approximately 100 ng), while the inhibitory doses are one and two orders of

magnitude greater than that found in a typical PRP. In the same study, PRP

was added to DBM. After a 56-day implantation no difference in the bone

induction histology score appeared with or without PRP, but an increase in

DBM resorption with the addition of PRP was detected. Ossicles formed in

this model will remodel over time, so it may be expected that the addition of

PRP would accelerate the natural remodeling process, which would include

the resorption of the DBM particles. In a similar study a more physiologic

dose of 50 ng of PDGF was added to 25 mg of DBM in an aged rat ectopic

model and was implanted for 14 days [58]. The PDGF addition to the DBM

increased mRNA production of collagen Type II, alkaline phosphatase

activity and calcium content, which are all indicative of bone formation.

These studies illustrate the complex nature of the effect of PRP on osseous

healing, and that important study design considerations include the animal

model, dosage, outcome measures and time intervals.

While the in vivo ectopic osteoinduction model has become a standard

method and enables comparison among studies, an orthotopic model may be

more predictive of expected clinical outcomes. When DBM was mixed with

PRP in a porcine cranial defect and healing was compared to autograft, PRP

enhanced bone formation compared to autograft at the earliest time-point of two

weeks. However, at later time-points PRP had no effect on mineralization [50].

Animal studies utilize species-specific PRP. Since blood cell size varies

from species to species, it is possible that the ability for PRP systems to

concentrate platelets might also be species-dependent. In the majority of animal

studies, the actual platelet dose was not quantified, which can confound

attempts to compare studies. In our laboratory we compared the platelet concentration

and growth factor profile in various species, and compared it to

humans using a single-spin, table-top platelet concentration system that had

been previously characterized for processing human blood [19]. Table 26.4

(unpublished data) summarizes the multi-species results.

Whole blood platelet counts for all species were essentially within the

normal range of human platelet counts of 200,000 platelets/μl to 400,000

platelets/μl [12]. However, the ability of the platelet concentration system

to collect platelets varied five to eight times over baseline, depending on

species. This is probably due to variances in cell size and density. Similarly,

the amount of the growth factors PDGF and TGF-β1 also varied with the

different species preparations. It is interesting to note that the fold increase in

growth factors did not necessarily match the corresponding fold increase in

platelet concentration, a result also noted by others [19]. We were unable to

count goat platelets accurately because there is less size difference between

red blood cells and platelets than for other species, which limits usefulness of

the goat model.

From the vast range of preclinical studies using PRP in orthopedic applications,

the results appear to predict that PRP used in clinical orthopedic applications will

be successful. The next section will address the clinical use of PRP to date.

26.4. Clinical Review

The best evidence to support clinical use of PRP is prospective, randomized

and controlled studies (Level of Evidence I (blinded) or II (not-blinded))

[59]. A search has revealed only one Level I study and five Level II studies

published in orthopedic applications, with the balance being retrospective and

case studies (Level III (retrospective, controlled studies) and Level IV (case

series with no controls) ) as summarized in Table 26.5. As with the preclinical

studies a (+) was assigned to the study if any positive result was seen with the

PRP, a (o) was given if no difference was found between the test and control

groups and a (−) indicates when the PRP caused an inhibitory effect.

As seen in the preclinical studies, PRP as an adjuvant to other treatments

has resulted in excellent clinical outcomes. For example, a case series of

patients treated with PRP and cultured mesenchymal stem cells in distraction

osteogenesis had successful outcomes without taking an autograft harvest

[61]. PRP has also been used with autograft and has increased bone maturation over autograft alone [2, 70, 74, 77]. However, more surprisingly, PRP in

combination with allograft has demonstrated clinical outcomes equivalent

to autograft in a spine fusion study [4]. PRP, in combination with osteoconductive

matrices, has also improved the clinical results compared to the

osteoconductive material alone [3, 6, 60, 73, 78]. In the only Level I evidence

trial using PRP, PRP mixed with xenograft in periodontal intrabony defects

significantly increased defect fill over the xenograft alone [3].

While many favorable outcomes with PRP have been demonstrated

clinically, not all published results support the use of PRP. Specifically, in

clinically challenging spine fusion cases, three studies have published no

difference between PRP and the control groups [63–65], and one study found

increased bone resorption when PRP was used [67]. However, two of these

studies did show a decrease in pseudoarthrodesis and a faster fusion rate in the

PRP groups, though the results were not significant [64–65]. The comparisons

between PRP and control groups were compared at a 24-month follow-up time

in the Hee, et al. [65] study, and at 34 months in the Castro, et al. [64] study.

Given that the effects of PRP are suggested to speed up healing processes,

it should be expected that groups compared at this late follow-up time-point

would be equal. Similarly, in the Carreon, et al. study, no difference was found

with the addition of PRP mixed with autograft with instrumented lumbar spine

fusion at 32- to 37-month follow-up [63]. Any effect the PRP had on healing

would more likely be seen early on in the healing process, but not in the overall

amount of bone formation once the fusion is complete. In the Weiner and

Walker study the fusion rate was much lower in the PRP with autograft group

than the autograft alone [67]. As this procedure was uninstrumented, it was a

very challenging spine fusion model. Without instrumentation, micromotion

can occur more freely at the fusion site. Since PRP contains angiogenic growth

factors, one might expect that micromotion would increase fibrosis tissue formation

preferentially over bone, increasing the pseudoarthrodesis. In contrast,

a case series published with PRP mixed with autograft in an instrumented

spine fusion demonstrated 58 out of 60 patients fusing [68], and yet another

case series with PRP and autograft in an instrumented spine fusion showed no

pseuodoarthrosis [66].

In a prospective, randomized trial (Level II) comparing PRP and allograft

to autograft in spine fusions, the groups were compared at six months postoperation

and found to be equivalent [4]. These results are promising in that

PRP might find clinical utility as an adjuvant with other matrices to reduce

the need for a painful autograft harvest. However, both preclinical and clinical

results suggest that PRP can enhance biologic repair and, in conjunction with

adequate bone matrices and/or cells, can be effective in spine fusion at a much

lower cost, both in terms of material costs when compared to recombinant

BMPs, and in morbidity of the donor site when compared to an autograft

harvest.

In the field of dental and craniomaxillofacial surgery, many procedures are

bilateral. This permits PRP to be added unilaterally, allowing each patient to

be his or her own control. As mentioned earlier, in the only published Level I

evidence trial (with double-blinding), PRP mixed with xenograft in periodontal

intrabony defects significantly increased defect fill over the xenograft alone

[3]. In several other studies PRP showed enhanced results over control groups

in bilateral intrabony defects [3, 73–74]. Only in one series of three patients

in a bilateral defect did PRP show no difference when mixed with xenograft

in a sinus lift [75]. However, in a prospective, randomized study comparing

PRP with a synthetic bone void filler (â-TCP) in a sinus lift, PRP was shown

to increase bone formation [6].

In the only soft tissue application reviewed, PRP injections improved chronic

elbow tendinosis (or tennis elbow) [72]. Mishra and Pavelko found statistically

significant improvement in visual analog pain scores in patients that received

injections of buffered PRP into the tendon area of maximum tenderness. The

treatment groups received PRP that was 5.4-fold higher concentration than

their whole blood. The control groups received the same injection technique

with bupivacaine, but did not see the improvement that the PRP group did. It

can, therefore, be concluded that the injection into the tendon alone was not

enough to stimulate healing. The authors hypothesize that the concentrated

growth factors in PRP work together to initiate the healing response. Tendon

improvement could be due to increased collagen Type I production from the

tendon fibroblasts [79], or from increased angiogenesis into the site [80].

As with the preclinical studies, the actual dose of platelets delivered or the

concentration of growth factors is not always reported in the clinical studies.

This makes it difficult to determine the correct platelet dose for each surgical

application. In vitro experiments have shown a dose-dependent increase in

stem cell proliferation with a dose of up to a 10-fold increase in platelets over

baseline [81–82]. Combined with DBM in an ectopic implantation model,

a dose-dependent inhibition of bone formation was seen with recombinant

PDGF [44]. However, as mentioned earlier, at doses normally found in a PRP,

a slight increase in bone formation was seen. Many growth factors exhibit this

biphasic response, with stimulatory effects on bone formation at lower doses

and inhibitory effects at high doses [25–29].

One study evaluated the dose-dependent response of bone formation with

PRP around an implant in a rabbit model [46]. In this study, three levels of

platelet concentration were compared. The lowest concentration was 164,000

to 373,000 platelets/ìl (0.5–1.5X fold increase over whole blood), an intermediate

dose was 503,000 to 1,729,000 platelets/ìl (2-6X fold increase over

whole blood) and a high dose was 1,845,000 to 3,200,000 platelets/ìl (9–11X

fold increase over whole blood). At four weeks only the intermediate group had

significant increase of bone. From this study the authors draw the conclusion

that the optimum dose of PRP concentration is 1,000,000 platelets/ìl [46].

26.5. Future Trends and Needs

Clinical utilization of PRP will continue to evolve. Future devices will provide

PRP faster and more efficiently, with more customization of end level products

such as fibrinogen concentration or output volume. Development of better

delivery devices and methods to combine PRP with various matrices will also

continue to emerge. Concurrently, more Level I and Level II evidence studies

will need to be completed to further refine the uses of PRP, matching clinical

indications with the right matrix and cell technologies. Additionally, designing

these studies with the appropriate follow-up time-points will be critical to

capture the clinical benefits of PRP. It is clear from the review of the studies

to date that a one-size-fits-all solution for all PRP applications does not exist.

The choice of the appropriate matrix and, when required, fixation technique,

will need to be coupled to each application that PRP finds utility. In addition

to defining the clinical utilizations, better characterization of the PRP products

should be included in the studies to help elucidate the correct platelet

concentration for each application.

26.6. Conclusions

PRP has demonstrated numerous clinical benefits to patients [1, 3–4, 6]. Better

matching of clinical indication to the right matrices and/or requirement of cell

delivery will need to be determined in future evaluations. Better characterization

of the product delivered and further refinement in clinical utility will

better elucidate the dose of platelets that is optimal for each application. The

working hypothesis is that the optimal platelet dose is 1,000,000 platelets/ìl

[46]. This number was determined from one clinical indication. It is reasonable

to consider that this type of dosing study would be needed for each new

indication for use, and that each indication would have a discrete optimal

platelet dose.

PRP remains a potentially powerful autologous therapy for surgeons that

want to enhance bone formation. The addition of growth factors such as

PDGF, TGF-â1 and VEGF will promote cellular bioactivity by increasing

cell proliferation, chemotaxis and angiogenesis. The benefits seen to date with

PRP’s use will continue to encourage researchers to actively investigate the

use of PRP in orthopedics and in other fields of medicine [72].