What does “good platelet quality” in PRP actually mean?

Platelets are not passive containers for growth factors. They respond to activation stimuli, change their shape and surface receptors, can aggregate and release contents from α-granules and dense granules. For a PRP preparation, it is therefore relevant not only how many platelets are present, but also what state they are in.^[1]

In research, this state is assessed, for example, using activation markers such as P-selectin (CD62P), aggregation capacity, response to defined agonists, morphology or release of selected mediators. Such tests are not a routine requirement for every practice. They do, however, show why the equation “more platelets = better PRP” is biologically too simplistic.

The DEPA classification systematised this idea as early as 2016: Dose, Efficiency, Purity and Activation are considered separately. MIBO additionally calls for a traceable description of starting material, preparation and final product. More recent consensus work likewise stresses that platelet count, recovery, cell profile, activation and application should be documented together.^[2][3][4]

The key idea: PRP quality develops along a process chain

In PRP, the centrifuge is not the only critical point. Pre-analytical variables can alter the sample even before centrifugation; after centrifugation, the collected fraction, waiting time, activation and application determine which product is actually used. A comprehensive 2025 review describes precisely this variability in isolation, cell composition, recovery, activation and application as a central standardisation problem.^[1]

1. Starting blood: the same protocol does not produce the same PRP in every person

PRP is autologous. Variability therefore begins before any technical step. Mazzocca and colleagues showed differences between preparation methods in the same individuals as well as relevant inter-individual variation in the resulting cell and growth-factor profiles.^[5] More recent reviews likewise list patient factors and starting blood as part of product variability.^[1]

For quality assessment, this means that a manufacturer claim such as “x-fold enrichment” is not a guaranteed individual laboratory value. Anyone wishing to compare scientifically needs both a baseline value and a final-product measurement. That is why the baseline platelet count belongs in a sound PRP characterisation.

2. Blood collection: pre-analytics begins before the tube

Platelets are sensitive to their environment. In platelet function testing, blood collection, stasis, anticoagulant, time to analysis and sample handling are therefore considered relevant pre-analytical variables.^[6] An older human study, for example, found that even a delay after puncture could increase measured activation.^[7]

This does not mean that every smaller collection needle automatically destroys platelets. Lippi et al. compared 21G, 23G and 25G needles during blood collection. Small statistical differences in platelet count occurred, but the authors judged the mean differences not to be clinically significant.^[8] A comparison between a standard 21G needle and a 21G butterfly likewise found no relevant differences in several platelet-function tests.^[9]

3. Anticoagulant and PRP tube: the system is more than a container

The tube does not determine PRP quality on its own. It does, however, define technical boundary conditions: blood volume, type and amount of anticoagulant, geometry, possible separation barriers and the fraction accessible after centrifugation. A technical and biological review of 50 commercial PRP systems found substantial heterogeneity among available systems and sufficiently detailed biological characterisation for only a small proportion of them.^[10]

The anticoagulant is not neutral either. Human studies show that different anticoagulants can alter cell counts, morphology, activation markers or growth-factor profiles. ACD-A, sodium citrate, EDTA and heparin should therefore not be regarded scientifically as freely interchangeable variants.^[11][12]

The literature is not entirely consistent: EDTA, for example, can yield high recovery values but also changes platelet morphology and activation parameters. A paired human study published in 2026 found markedly higher size parameters under EDTA than under sodium citrate and interpreted this as an anticoagulant-dependent artefact.^[13] This does not produce a blanket ranking of all anticoagulants, but rather a more useful practical rule: the anticoagulant is part of the validated preparation system and should not be assessed in isolation from the rest of the protocol.

4. Centrifugation: more force is not automatically worse – and less force is not automatically better

Centrifugation is intended to separate blood components according to their physical properties. RCF, run time, rotor radius, rotor design, tube position and number of spins influence where platelets, leukocytes and erythrocytes are found after the run. An RPM value alone is therefore not transferable between different centrifuges.

It is particularly important to distinguish between concentration and unwanted pre-activation. A human study of pre-analytical activation found markedly lower P-selectin values in PRP prepared at 100–250 × g for ten minutes than in plasma after 2,000 × g.^[14] This is a strong signal that centrifugation conditions can affect activation status.

Another validation study achieved high recovery with a two-step protocol that included 1,600 × g in the second step and reported no loss of integrity or viability.^[15] This is exactly why the statement “above X × g platelets are damaged” would be scientifically unsound. Force and time must always be considered together.

The rotor may also matter. A small 2025 human study compared fixed-angle and horizontal rotors and, under the tested optimised conditions, found differences in recovery, platelet content and PDGF-BB, without a significant difference in CD62P.^[16] This does not prove that one rotor type is generally superior, but it illustrates why a protocol is more than “RPM × minutes”.

5. The collected fraction: after the spin, plasma is not the same everywhere

After centrifugation, platelets are not distributed homogeneously throughout the entire plasma column. A human study examining distribution in different fractions found marked differences in platelet concentration and mean platelet volume; together, the buffy coat and lower third of the plasma contained a large proportion of the measured platelet mass.^[17]

This makes collection technique an independent process parameter. Collecting closer to the interface may capture not only more platelets but, depending on the system, also more leukocytes or erythrocytes. The term “PRP” therefore does not automatically describe which cell population actually ends up in the syringe.

This does not lead to a universal instruction on the height from which material should be collected. The validated procedure of the specific system is decisive. For studies and quality documentation, however, it should be recorded which fraction was obtained.

6. Time and handling: freshly prepared PRP is not biologically timeless

Platelets change over time. Bausset et al. examined activation status, reactivity, aggregation, morphology and growth-factor release and, under their preparation and storage protocol, reported preserved bioactivity for several hours; thereafter, functional parameters declined.^[18] A study published in 2026 observed a progressive increase in activation and degranulation markers over 1, 6 and 24 hours, along with changes in TGF-β release.^[19]

These studies provide no universal clinical time limit, because protocols, temperature, containers, activation and measurement methods differ. They do clearly show, however, that “PRP simply stays the same until it is eventually used” is not a robust assumption. Time to application belongs in a reproducible protocol.

7. Activation: intended activation and unwanted pre-activation are two different things

PRP can be activated ex vivo before application or applied without an added activator. Calcium, thrombin, collagen contact and local tissue signals affect activation, coagulation, fibrin formation and release in different ways. A study of three PRP systems showed, for example, that release of PDGF-AB and TGF-β after thrombin activation depended on whether platelets had already been pre-activated during preparation.^[20]

For the quality discussion, the simplified question “activated or not?” is therefore not the key issue. What matters is when, by what and to what extent activation takes place. A platelet that is ready to respond before application and a platelet that has already undergone extensive degranulation are not biologically the same.

8. Needles and injection: does a very fine needle destroy platelets?

This statement is frequently repeated, but direct PRP data do not support it as a blanket rule. Bausset et al. examined PRP function after passage through different needle gauges and found no relevant effect of the tested needle sizes on platelet function. In the setting they studied, the authors recommended 30G as an option for less painful injection.^[21]

This is not a blanket clearance for arbitrarily fine needles. The study provides direct data up to 30G, not automatically for 32G, 33G or 34G. A finer needle also changes flow resistance. Internal diameter, length, viscosity, syringe geometry and desired flow rate together determine how much plunger pressure is required.

A more scientifically accurate statement is therefore:

A second finding from the same study is also interesting: the local anaesthetics tested reduced platelet aggregation capacity even though measured growth-factor release was not significantly changed.^[21] This illustrates why a single value such as “growth factor release” is not a complete description of function.

Four common PRP myths in a quality check

What is directly supported – and where should wording remain cautious?

How can PRP quality be documented meaningfully?

A practice does not need to test every PRP sample using research-laboratory methods such as flow cytometry. For scientific comparability and clean technical documentation, however, process data and product data should not be mixed. The MIBO recommendations and current consensus work point in the same direction.^[3][4]

Why this matters so much for PRP studies

A large part of the PRP literature is only partly comparable because preparation and final-product data are incompletely reported. The original MIBO work therefore developed minimum reporting requirements for PRP studies.^[3] Even years later, systematic reviews still identify substantial reporting gaps in starting blood, recovery, final-product measurement and activation.

This explains part of the apparently contradictory PRP results: two studies can both investigate “PRP” while applying biologically different preparations. Comparing only the term PRP, the RPM figure or the concentration factor overlooks a large part of the actual product.

The most important consequence: do not optimise one factor – control the chain

The current literature argues against simple quality rules. Neither the highest platelet concentration nor the lowest centrifugal force, the finest injection material or a specific tube feature is, by itself, a universal quality criterion.

A more useful approach is to know the starting material, use a validated tube/centrifuge system reproducibly, define the collected fraction, avoid unnecessary delays and characterise the PRP actually used as completely as necessary. This turns “PRP” from a catch-all term into a traceable preparation.

Professional note: This article provides scientific and technical information for qualified professional users. It is not a treatment recommendation, manufacturing instruction, centrifugation protocol or injection protocol. Study parameters must not be transferred to other tubes, centrifuges, rotors or applications without review. The intended purpose, instructions for use, regulatory requirements, validated workflows and the professional judgement of the user are decisive.