The effect of leukocyte-reduced platelet-rich plasma on the proliferation of autologous adipose-tissue derived mesenchymal stem cells 1

Abstract

Clinical application of platelet-rich plasma (PRP) and stem cells has become more and more important in regenerative medicine during the last decade. However, differences in PRP preparations may contribute to variable PRP compositions with unpredictable effects on a cellular level. In the present study, we modified the centrifugation settings in order to provide a leukocyte-reduced PRP and evaluated the interactions between PRP and adipose-tissue derived mesenchymal stem cells (ASCs).

PRP was obtained after modification of three different centrifugation settings and investigated by hemogram analysis, quantification of protein content and growth factor concentration. ASCs were cultured in serum-free α-MEM supplemented with autologous 10% or 20% leukocyte-reduced PRP. Cell cycle kinetics of ASCs were analyzed using flow cytometric analyses after 48 hours.

Thrombocytes in PRP were concentrated, whereas erythrocytes, and white blood cells (WBC) were reduced, independent of centrifugation settings. Disabling the brake further reduced the number of WBCs. A higher percentage of cells in the S-phase in the presence of 20% PRP in comparison to 10% PRP and 20% fetal calf serum (FCS) advocates the proliferation stimulation of ASCs.

These findings clearly demonstrate considerable differences between three PRP separation settings and assist in safeguarding the combination of leukocyte-reduced PRP and stem cells for regenerative therapies.

Keywords

Platelet-rich plasma PRP adipose-tissue derived mesenchymal stem cells growth factor cell cycle white blood cells

1 Introduction

Growth factors and stem cells have become more and more important in regenerative medicine [12 , 24] during the last decade. Platelets contain more than 5000 proteins, of which more than 300 are released upon activation [34, 40]. Among these bioactive proteins are particularly growth factors and cytokines. This has been documented for more than 25 years [11]. Since then, platelet-rich plasma (PRP) has often been defined as at least 200.000 - 1.000.000 platelet/μL suspended in plasma [19] and found an increasing use as clinical treatment for musculoskeletal tissue regeneration in orthopaedics [5 , 36] and other fields, like plastic surgery [8, 26], and chronic wound healing [18]. As the body of literature about the regenerative effects of PRP grows, the use of stem cells as a treatment option for tissue regeneration has been endorsed at the same time due to their ability to secrete various growth factors. Adipose tissue is an abundant source of mesenchymal stem cells (ASCs) which can be harvested in a simple procedure. Moreover, ASCs have already been reported for their successful application in musculoskeletal tissue regeneration [14 , 37].

Despite widespread use in randomized clinical trials for several clinical applications, there is currently insufficient evidence to support the use of PRP for treating musculoskeletal soft tissue injuries [24, 35]. Most important, in a majority of studies the quantitative composition of cells and bioactive factors for the applied PRP product has not been examined [31]. In vitro studies revealed substantial differences in the PRP product between individuals [30], or even depending on the daytime [3]. In addition, at least 16 commercial platelet separation systems are available [38], which underlines the problem of comparability of the clinical studies and might be a reason for some inconsistencies in the literature of the reported beneficial effect. It seems impossible to develop protocols for clinical applications. Each autologous PRP preparation may produce a different PRP product with unpredictable effects on a cellular level, especially due to the presence of leukocytes, which has been extensively discussed in the literature. It has been suggested that leukocyte depletion in PRP may result in a product suitable for regeneration of tissue injuries without the formation of scar tissue [23].

Aim of the present study was to investigate the biological characteristics of a commercially available PRP preparation kit (Arthrex Double Syringe, Arthrex, Inc, Naples, Florida) after modification of centrifugation settings (speed, duration, brake) in order to provide a leukocyte-reduced PRP. Moreover, the effect of leukocyte-reduced PRP on the cell cycle kinetics of autologous ASCs was evaluated, since the combined application might be a suitable tool for regenerative therapies. This human in vitro model may assist in safeguarding the combination of PRP and stem cells for regenerative therapies. The study was based on the hypothesis that leukocyte-reduced PRP can be produced by modifying the centrifugation settings, and that ASCs treated with leukocyte-depleted PRP can be expanded in an autologous serum-free cell culture setting.

2 Methods

2.1 Ethics statement

The study was approved by ethics committee of the University of Regensburg and written informed consent was obtained from each volunteer in accordance with the declaration of Helsinki prior to blood drawing.

2.2 Subjects

Healthy male volunteers (n = 30) without medication for at least two weeks and patients undergoing body-contouring surgery (n = 4) were recruited at the University Medical Center of Regensburg. Healthy individuals were equally distributed into three age groups: 21–30 years of age (group 1), 31–40 years of age (group 2), 41–50 years of age (group 3) (Fig. 1B ).

2.3 Blood samples

Venous blood (15 ml) was drawn directly into the Arthrex Double Syringe (Arthrex, Inc, Naples, Florida, USA) for the production of autologous conditioned plasma (ACP) using a winged infusion set (Sarstedt AG & Co., Nümbrecht, Germany). In addition, two EDTA tubes and two 2-ml Eppendorf tubes of blood were collected (2 ml) for a differential hemogram analysis and protein content measurement, whereas the blood sample for differential hemogram was analyzed within 2 hours after withdrawal.

2.4 Sample preparation

The venous blood withdrawal collected in the ACP double syringe was processed using a Hettich Rotofix 32a centrifuge (Andreas Hettich GmbH, Tuttlingen, Germany). In order to determine the best centrifuge setting, 3 double syringe blood draws were obtained only from participants of group 2 and samples were prepared as follows: 1500 rpm for 5 minutes with brake enabled (Setting ACP-A); 1500 rpm for 4 minutes with brake disabled (Setting ACP-B); 3000 rpm for one minute with brake disabled (Setting ACP-C) (Fig. 1 A). The venous blood was separated into two distinct layers by centrifugation whereas a plasma layer appeared on the top and the red/white blood cell layer was apparent on the bottom. The plasma, containing the platelets, was isolated by drawing the inner syringe following manuals instructions.

For the remaining 20 subjects (group 1, group 3) and the 4 male patients undergoing elective body-contouring surgeries, ACP was prepared using 1500 rpm for 4 minutes with brake disabled (Setting ACP-B) (Fig. 1 B).

2.5 Differential hemogram analysis

Thrombocyte, erythrocyte, leukocyte, lymphocyte, monocyte and neutrophil cell counts were determined in a venous blood sample (EDTA tube) and ACP (after separation) by using a Sysmex XE-5000 fluorescence flow cytometer (Sysmex Europe GmbH, Norderstedt, Germany).

2.6 Protein content

The protein concentration of serum samples and ACPs was determined in duplicates using the Biorad DC protein assay (BioRad, Hercules, California, USA) according to the manufacturer’s instructions. Briefly, venous blood was stored in vertical position for 24 h at 4°C allowing red blood cells to sediment and subsequently to recover serum by aspiration of the upper phase. Serum and ACP samples were diluted 1:1 in a RIPA-Protease-Inhibitor-solution prepared as a 10:1 mix in RNA-free water and subsequently subjected to BioRad DC protein assay. The photometric analysis was performed with the Vario Scan flash (Thermo Scientific Inc., Waltham, Massachusetts, USA) at 750 nm.

2.7 Growth factor analysis

Growth factor concentrations were determined in duplicates for serum and ACPs using enzyme-linked immunosorbent assay (ELISA) with a customized Q-Plex Kit (Quansys Bioscience Logan, Utah, USA) according to the manufacturer’s instructions. Samples for each subject were immediately frozen (−20°C) to preserve growth factor integrity, and thawed prior to performance of the ELISA assays. FGF, HGF, PDGF and VEGF were selected for growth factor analysis given their specific roles in wound healing and tissue regeneration [1]. The color intensity of each well was digitally recorded by the Versadoc Imaging System (BioRad, Hercules, California, USA) and quantified by using Q-View^TM software (Quansys Bioscience).

2.8 Adipose-tissue derived stem cell (ASC) isolation

Human ASCs were isolated from solid subcutaneous adipose tissue, which was obtained from 4 patients undergoing elective body-contouring procedures, as described previously [13]. Briefly, subcutaneous fat tissue was washed in phosphate-buffered saline, and minced into pieces of <2mm³. Serum-free α-MEM (1 ml/1 g tissue) and Liberase Blendzyme 3 (Roche Diagnostics, Rotkreuz, Switzerland) (2 U/1 g tissue) were added and incubated under continuous shaking at 37°C for 45 minutes. The digested tissue was sequentially filtered through 100-μm and 40-μm filters (Fisher Scientific Schwerte, Germany) and centrifuged at 450 g for 10 min. The supernatant was discarded and remaining cell pellet was washed twice with Hanks’ balanced salt solution (Cellgro Manassas, Virginia, USA) and finally resuspended in α-MEM growth medium containing 20% FCS, 2 mM L-glutamine 100 U/ml penicillin, 100 g/ml streptomycin. Cells were plated at a density of 3×10⁴ cells/cm² in 175 cm² cell culture flasks (Greiner Bio-One GmbH, Frickenhausen, Germany) and incubated at 37°C in a humidified atmosphere containing 5% CO₂. All non-adherent cells were removed after 18 hours incubation by washing culture dishes with Dulbecco’s phosphate buffered saline (PBS, Sigma-Aldrich, Co., St. Louis, Missouri, USA) and ASCs received fresh growth media every other day. For further experiments and analysis, 80% confluent cell layers of passage 0 were frozen in medium containing 10% dimethyl sulfoxide (DMSO, Invitrogen Life Technologies, Darmstadt, Germany) that was supplemented with 90% FCS.

2.9 Cell culture experiment with ACP

2.5×10⁵ human ASCs at passage below 3 were seeded in 100 mm cell culture dish (Becton Dickinson and Company, Franklin Lakes, New Jersey, USA) and cultured in 7 ml of α-MEM supplemented with 20% FCS, 5 mM glutamine and 100 U/ml penicillin with 100 g/ml streptomycin for 48 hours. Afterwards, the medium was changed to serum-free α-MEM and cells were incubated for 24 h in order to synchronize the cell-cycle phase. The medium was replaced by 5 ml media for all 3 experimental groups: α-MEM containing 20% FCS, α-MEM containing 10% ACP and α-MEM containing 20% ACP. Autologous cell culture setting was performed by using the ACP and human ASCs from the same patient. Cells were incubated in corresponding medium for 48 hours and harvested by trypsination for cell cycle analysis.

2.10 Cell cycle analysis

Cells were washed twice with ice-cold PBS containing 2% FCS and incubated overnight in 70% methanol on ice. Afterwards, cells were washed twice with PBS and incubated in the presence of 10μg/ml RNAse for 30 minutes at 37°C. The DNA intercalating 4^′,6-Diamidin-2-phenylindol (DAPI) fluorochrome was added at final concentration of 1μg/ml 15 min prior to analysis to ensure quantitative DNA staining.

Flow cytometric analyses were performed on each sample with 3×10⁵ DAPI stained cells using a FACSCanto-II flow cytometer (BD Biosciences, San Jose, California, USA) equipped with a blue (488 nm), a red (633 nm), and a violet (405 nm) laser and standard optical configuration. The instrument was operated with the FACSDiva software Verion 7.0 (BD Biosciences). The DNA dye DAPI was excited with the violet excitation line and fluorescence emission was detected by the optical trigon unit equipped with a 450/50 bp filter. DNA histograms were plotted on a linear scale upon cell doublet, aggregate, and debris discrimination via pulse processing. Cell cycle fractions (i.e. percentages of cells in G0/G1-, S- and G2/M-phase) were quantified using the ModFit LT 3.2 software (Verity Software House, Topsham, Maine, USA). Treatment effects are expressed by the S-phase fraction (SPF) compared to untreated cells.

2.11 Statistics

Statistical analysis was performed using SPSS software package (version 19, IBM SPSS, Chicago, Illinois, USA) whereas all graphs were prepared by using GraphPad Prism (version 5, Statcon, La Jolla, California, USA). All data were tested for normal distribution applying the Shapiro-Wilk test. Descriptive data are expressed in terms of median (range) or mean±standard deviation.

The General Linear Model Repeated Measures and the Wilcoxon Signed-Rank tests with Bonferroni correction were used to analyse statistical differences of all parameters derived by the ACP single-spin separation methods. The paired t-test and the Wilcoxon Signed-Rank test with Bonferroni correction were applied to analyse differences of all parameters describing Serum and ACP B. Differences between the age groups were investigated by the One-Way Analysis of Variance (ANOVA) and the Kruskal-Wallis test with Bonferroni correction. The Spearman’s Correlation test was used to analyse correlations between all parameters in all age groups. The paired t-test with Bonferroni correction was applied to identify significant differences with regard to outcomes from the cell cycle analysis in the different cell culture conditions. The level of significance was set at P = 0.05 for all statistical tests.

3 Results

3.1 Brake settings modifies ACP composition

The differential hemogram analysis demonstrated considerable differences between the 3 single-spin separation settings A, B and C. Thrombocytes were enriched for all settings in comparison to venous blood (p < 0.01) independent of enabled brake or time (p = 1.0) (Fig. 2A). In addition, erythrocytes, leukocytes, monocytes and neutrophils were significantly reduced by each centrifugation mode in comparison to venous blood (p < 0.01 each). In contrast, lymphocytes were significantly reduced for setting A and B in comparison to venous blood (p < 0.01), whereas the lymphocyte count for centrifugation setting ACP-C only demonstrated a lower rate with a trend to significance in comparison to peripheral blood (p = 0.06). Disabling the brake significantly reduced the cell number of leukocytes, lymphocytes and monocytes (p < 0.05 each) when centrifuged at 1500 rpm for 4 minutes (setting ACP-B) in comparison to A (Fig. 2C–F). The mean cell counts of thrombocytes, erythrocytes, leukocytes, lymphocytes, monocytes and neutrophils contained in the single-spin separation methods A, B and C are presented in Table 1.

The protein content increased significantly after centrifugation enabling the brake in setting ACP-A when compared to serum (p < 0.01). Centrifugation without brake (ACP-B and ACP-C) did not increase significantly the protein content when compared to serum (both P = 1.0). A significant higher protein content after centrifugation was measured, when brake was enabled (ACP-A) in comparison to ACP-B and ACP-C (Fig. 3).

The concentration of plasma derived growth factors (FGF, HGF, and VEGF) did not show a significant enrichment by centrifugation in comparison to baseline serum (p > 0.16 each). Moreover, no significant difference was apparent for the concentration of FGF, HGF, or VEGF by using different centrifugation settings (all P > 0.16). In contrast, the concentration of PDGF significantly increased in comparison to serum by centrifugation with enabled brake (setting ACP-A) (P < 0.05). Centrifugation settings ACP-B and ACP-C demonstrated a similar concentration for PDGF as measured in serum (both p > 0.42) (Fig. 4).

3.2 Age specific results for ACP

Numbers of thrombocytes significantly increased in ACP (p < 0.01) due to centrifugation regardless of the setting (brake enabled or disabled) and independent of subject’s age (p > 0.24). In addition, erythrocytes, leukocytes, monocytes and neutrophils showed significantly decreased cell counts (p < 0.01 each) in all age groups after centrifugation when compared to venous blood. There were no significant differences between the three age groups regarding cell counts, except for leukocytes that showed significantly higher numbers (p < 0.01) in patients aged 41 to 50 years in comparison to younger age groups (1 and 2) (Fig. 5).

The protein content of the ACP-B in comparison to serum did not increase significantly in all age groups (p > 0.24). We identified a significant difference in the protein content between the age groups 1 and 3 (p < 0.01). Furthermore, there was no correlation between the protein content and growth factor concentration due to separation by centrifugation setting ACP-B (P > 0.13).

The concentration of FGF, HGF, or VEGF in ACP was not significantly enriched by centrifugation when compared to serum in all age groups (p≥0.49 each). However, the concentration of PDGF significantly increased in age groups 1 and 3 after centrifugation and all ACP-B samples taken together (p < 0.01 each), whereas this was not the case for age group 2 (p = 0.124). In addition, a significant positive correlation was found between the thrombocyte count in ACP-B and the corresponding PDGF concentration in the age groups 1 and 2 (p < 0.05 both), which was not apparent for the age group 3 (41–50 years) (p = 0.11).

3.3 Cell lysis contributes to growth factor release

The concentration of FGF and HGF in ACP did not increase (p≥0.10 each) after lysis with RIPA Buffer when compared to untreated ACP and serum. In contrast, the concentration of PDGF and VEGF increased significantly in ACP and serum after incubation with RIPA buffer when compared to untreated ACP and serum (each p < 0.01).

3.4 Cell cycle analysis

Flow cytometric analyses of human ASCs revealed a higher percentage of cells in the S-phase in the presence of 20% ACP in comparison to 10% ACP and 20% FCS (both P≥0.14). On the contrary, a lower percentage of cells was observed in the G1-phase in the presence of 20% ACP in comparison to 10% ACP and 20% FCS (both P≥0.07). Comparable percentages of cells in the G2-phase were found in the presence of 20% ACP in comparison to 10% ACP and 20% FCS (both P≥0.22).

4 Discussion

Aim of the study was to investigate the effects of centrifugation speed and deceleration on cell contamination and growth factor concentrations in PRP. We focused on a single commercially available PRP preparation kit, and modified the centrifugation settings. The biological characteristics of the original ACP and the products obtained after centrifugation modifications are described in detail. A significant reduction in leukocytes, lymphocytes, monocytes and neutrophils was achieved by disabling the brake at 1500 rpm. Increasing centrifugation speed up to 3000 rpm at disabled brake setting with simultaneous shortening of centrifugation time resulted in a considerably higher concentration of leukocytes, lymphocytes, monocytes and neutrophils.

In order to identify potential negative effects of the centrifugation adjustments on target stem cells in regenerative medicine, cell cycle analyses of human ASCs were performed.

Previous studies defined PRP with a therapeutic effect according to platelet content ranging from 200.000 to 1.000.000 platelets/μL [19, 22]. However, uncertainty still exists regarding an optimal platelet concentration for locally applied PRP products. Platelet concentrations of 2.5 times greater than venous blood have been reported to have a positive effect on the proliferation of musculoskeletal target cells [21]. The theory of “the more, the better” regarding the platelet concentration is not supported for PRP applications [21] due to reported adverse events [15]. In the present study, the platelet content was increased by approximately 2.3 times in comparison to venous blood independent of the centrifugation settings (speed, duration, brake) which is in accordance with previous studies using this PRP preparation kit [21, 30]. We also demonstrated a significant reduction of WBC concentration in comparison to venous blood, independent of centrifugation settings. Interestingly, a disabled brake provoked another significant reduction of all white blood cell types in comparison to the standard centrifugation settings according to the manufacturer’s instructions. All significantly reduced cell types have been reported for negative effects on tissue healing by promoting or inducing an inflammatory microenvironment due to increased levels of IL-1ß and TNF-α [23]. On the other hand, leukocytes are known to produce large amounts of VEGF, which contributes to angiogenesis and is especially required in certain musculoskeletal pathologies [39]. This is in line with the present data, which showed higher leukocyte counts for ACP-A und ACP-C associated with a trend of a higher VEGF concentrations. Thus, the influence of leukocytes on the biology of each PRP product and its potential positive or negative effects should be carefully addressed in future studies since it might explain controversial data from the literature.

The growth factors FGF, HGF and VEGF were not enriched by centrifugation in comparison to serum, which was independent of the applied centrifugation settings. In general, α-granules in platelets contain cytokines and growth factors, which are released through degranulation of the alpha granules upon activation [29]. Interestingly, the concentration of PDGF significantly increased in comparison to serum by using centrifugation settings according to the manufacturer’s instructions. This finding might be partially attributable to the freezing-thawing cycle in our study: α-granules in platelets are thereby activated with subsequent release reaction. Likewise, Perut et al. demonstrated significant differences in terms of growth factor release and biological activity in thawed PRP preparations [28]. Moreover, the PDGF concentration in the PRP product according to the manufacturer’s instructions was significantly higher than in the PRP product at 1500 rpm with disabled brake. It can be hypothesized that turbulences during the process of braking represents mechanical platelet activation and might contribute to partial degranulation of the α-granules with subsequent growth factor release. In line, Mazzocca et al. determined similar FGF-2 concentrations as in the present study, whereas HGF, VEGF and especially PDGF concentrations were reported with a multifold higher increase [20]. However, all measured growth factor concentrations showed a high variability (up to 30%). In comparison, in the present study a lower variabiliy (up to 20%) was found when using the new double syringe system (Arthrex ACP^®).

RIPA induced cell lysis of serum and the PRP product revealed a significantly increased concentration of PDGF and VEGF in both serum and the PRP product when compared to the untreated serum and PRP product. The concentrations of FGF and HGF demonstrated a trend towards higher concentrations, however without statistical significance. Nevertheless, VEGF and PDGF were reported with higher concentrations in various studies as we detected after cell lysis [7, 20]. This could be partially attributable to different methodology of the applied ELISA techniques.

The cell cycle is a series of events that occurs in a cell leading to its division and replication, and is a critical process for determining cell proliferation and senescence [16]. The G1 phase is the non-proliferative resting phase, whereas the S phase is the proliferative phase of DNA replication. In the G2 phase, the cell is ready for division, which occurs during mitosis. In the present study, autologous leukocyte-reduced ACP was used as cell culture medium supplement, thereby replacing (FCS). 48 hours of cell culture in α-MEM and 20% ACP resulted in an extended S phase in comparison to cell culture in α-MEM supplemented with 20% FCS or 10% ACP. These results indicated a dose-response relationship; supplementing 20% ACP initiated a higher DNA replication, resulting in cell growth and proliferation.

Moreover, a complete autologous cell culture system was established with evidence for ACP to stimulate cell proliferation. PRP has been successfully applied as a cell culture additive to facilitate growth of stem cells in the past [2 , 33] avoiding the risk of putative pathogens and xenogeneicmaterial.

Schallmoser et al. demonstrated obvious similarities regarding senescence-associated gene expression changes of MSC, which were expanded in the presence of FCS or PRP [32]. The combination of PRP and autologous ASCs has been recently investigated in a longitudinal cohort study and reported as a safe procedure [25]. However, further evidence is required to confirm the equivalence of MSCs expanded in media supplemented with PRP.

As the market for PRP preparations is expected to increase to more than $ 120 million by 2016 [35], PRP should be well characterized prior to injection. Moreover, future clinical trials should include concentrations of cellular components (especially platelets and white blood cells), volumes, number, and timing of applications. In our opinion, the further success of PRP products in combination with cell based therapy is depending on an accurate and rational description of their components and associated biological funcion.

The study revealed that even minor changes in the centrifugation process can lead to differences in biological characteristic of prepared PRPs for the first time.

5 Conclusion

These findings emphasize considerable differences between three PRP separation settings and assist in safeguarding the combination of leukocyte-reduced PRP and stem cells for regenerative therapies. Furthermore, our results highlight the necessity of optimizing the PRP production and its reported specification since these factors affect the biological function and the therapeutic efficacy of PRP application.

Limitations of the study

Several limitations of this study need to be noted: First, all ACP samples were stored at −20°C prior to growth factor analysis and prior to cell culture experiments. Damage to platelets during processing or storing could lead to activation of the platelets with subsequent release of growth factors [1, 6]. Therefore, the data obtained by in vitro cell culture and following cell cycle analysis may not correctly mimic in vivo environment and clinical setting. However, cell culture experiments were performed with ASCs and autologous ACP to provide a reproducible and comparable environment, and not raising concern over a graft versus host reaction.

Conflict of interest

Peter Angele is an expert advisor for Arthrex Inc. (Naples, Florida, USA). All other authors declare that there is no conflict of interests regarding the publication of this paper.

Footnotes

Acknowledgements

We would like to thank Daniela Biermeier for the support with the differential hemogram analysis, Gerhard Piendl for the technical support with the cell cycle analysis and Elke Gerstl for the support with all experiments. We would like to acknowledge Arthrex Inc. (Naples, Florida, USA) for contributing to the sampling material.

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