PRP with or without Citrate

Platelet-rich plasma (PRP) is obtained from one's blood where the concentration of platelets is at least five times the amount present in a similar volume of whole blood. Think of it as concentrated Platelets. PRP has now been used in multiple conditions, such as preventing infections after cardiac surgery, cartilage degeneration, joint pains, tendon strains, hair loss, cosmetic applications, and dry eyes. 

A key question is, "What is the best way to make PRP."

The below is a good review article. Citrate is usually used to make activated PRP noting the platelets seem to work better when activated and more platelets can be extracted when using Citrate. 

However, a patient of mine noted he felt better when using PRP without Citrate. He found an interesting article below (Ref 2) noting citrate can cause long term damage to epithelial cells IN VITRO. This was curious as he was the first to notice a difference. It could be he is particularly sensitive to the citrate. The article below notes that IN VIVO studies are lacking on the long term risk of citrate, but they do note they only noticed a negative effect with very concentrated citrate over a long time period.

For now, we make all our PRP with Citrate. But if we do have a patient who feels it did not help or it burns (which is very, very rare), I might consider making the PRP without citrate. 

SLC

Ref 1

Stem Cells International
Volume 2016, Article ID 7414036, 11 pages
http://dx.doi.org/10.1155/2016/7414036

Research Article

Platelet-Rich Plasma Obtained with Different Anticoagulants and Their Effect on Platelet Numbers and Mesenchymal Stromal Cells Behavior In Vitro

Ronaldo José Farias Corrêa do Amaral,1,2 Nemias Pereira da Silva,1 Natália Ferreira Haddad,1 Luana Siqueira Lopes,1 Fábio Dias Ferreira,1 Ricardo Bastos Filho,3 Paola Alejandra Cappelletti,1 Wallace de Mello,1,4 Eric Cordeiro-Spinetti,2 and Alex Balduino1,2

1Excellion Serviços Biomédicos, Amil/UnitedHealth Group, 25651-000 Petrópolis, RJ, Brazil
2Laboratório de Biologia e Tecnologia Celular, Universidade Veiga de Almeida, 20270-150 Rio de Janeiro, RJ, Brazil
3Universidade Federal Fluminense, 24033-900 Niterói, RJ, Brazil
4Centro Universitário Celso Lisboa, 20950-091 Rio de Janeiro, RJ, Brazil

Received 8 January 2016; Revised 9 April 2016; Accepted 27 April 2016

Academic Editor: Jiabing Fan

Copyright © 2016 Ronaldo José Farias Corrêa do Amaral et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

There are promising results in the use of platelet-rich plasma (PRP) for musculoskeletal tissue repair. However, the variability in the methodology for its obtaining may cause different and opposing findings in the literature. Particularly, the choice of the anticoagulant is the first definition to be made. In this work, blood was collected with sodium citrate (SC), ethylenediaminetetraacetic acid (EDTA), or anticoagulant citrate dextrose (ACD) solution A, as anticoagulants, prior to PRP obtaining. Hematological analysis and growth factors release quantification were performed, and the effects on mesenchymal stromal cell (MSC) culture, such as cytotoxicity and cell proliferation (evaluated by MTT method) and gene expression, were evaluated. The use of EDTA resulted in higher platelet yield in whole blood; however, it induced an increase in the mean platelet volume (MPV) following the blood centrifugation steps for PRP obtaining. The use of SC and ACD resulted in higher induction of MSC proliferation. On the other hand, PRP obtained in SC presented the higher platelet recovery after the blood first centrifugation step and a minimal change in MSC gene expression. Therefore, we suggest the use of SC as the anticoagulant for PRP obtaining.

1. Introduction

Platelet-rich plasma (PRP) is a blood-derived product in which platelets are concentrated at least five times in plasma above the baseline of that in the whole blood [1]. PRP is being investigated as an autologous product to improve tissue repair in different conditions and lesions, especially for musculoskeletal tissues, such as chondral lesions [2–4], tendinopathies [5–7], muscle strains [8, 9], and bone repair [10, 11]. Besides its clinical application, PRP may be an efficient substitute to fetal bovine serum in cell culture [12–15]. Its therapeutic potential is based mainly on the growth factors present in platelet’s alpha granules [16], such as transforming growth factor beta (TGF-β) [17], vascular endothelial growth factor (VEGF) [18], and platelet-derived growth factor (PDGF) [19], which have already been demonstrated to play important roles in tissue repair. When platelets are concentrated and activated, it is expected that the concentration of the factors released reaches three to five times of that found in the plasma [16].

The general methodology to obtain PRP involves the collection of whole blood with anticoagulants, followed by one or two centrifugation steps. After a first low-speed centrifugation, erythrocyte-free platelet concentrated plasma is recovered and submitted to high-speed centrifugation. Platelet-poor plasma is then discarded and the remaining platelet pellet is homogenized into what is regarded as PRP [16]. Several aspects on this method are still under debate, such as number of centrifugations, presence of leukocytes, and use and type of platelet activator and anticoagulants [20]. When no anticoagulant is used, a blood clot will form, and serum can be obtained but without increase in platelet concentration [21]. In the case of PRP obtaining, coagulation is not intended to occur prior to platelet concentration; hence, blood must be collected in the presence of anticoagulants.

For transfusion purposes, blood is usually collected in bags containing citrate phosphate dextrose adenine (CPDA-1) solution, as anticoagulant [22, 23], from which a platelet concentrate is obtained by double centrifugation of the whole blood or apheresis. Platelets concentrates obtained by such methodologies may also be used for promoting tissue repair [24]. On the other hand, recent PRP formulations for autologous applications are usually prepared in collection tubes containing citrate solutions, in the form of sodium citrate [25–29] or ACD-A [30–32]. This last, ACD, is present in the majority of available commercial kits for PRP production [33, 34]. In other cases, heparin [35, 36] or EDTA [37] can be used. For clinical investigations, EDTA is commonly used in hematology tests, SC in coagulation tests, and ACD in plasma levels measurement of platelet-derived components [38]. Therefore, our goal was to analyze how the choice of anticoagulant for blood collection would modulate PRP characteristics as well as its effects on mesenchymal stromal cell culture.

2. Materials and Methods

2.1. Ethics Statement

All of the experimental procedures were approved by the Ethics Research Committee of the Pró-Cardíaco Hospital, Rio de Janeiro (CAAE: 14878813.4.0000.5533), and all donors signed an informed consent.

2.2. PRP Obtaining

PRP was obtained as previously described, with minor modifications [39]. Peripheral blood was collected from nine volunteer donors (6 men and 3 women) using blood collection tubes containing sodium citrate (SC) (Vacutainer®, Ref: 369714; BD Biosciences, San Jose, CA), ethylenediaminetetraacetic acid (EDTA) (Vacutainer, Ref: 367861; BD Biosciences), or anticoagulant citrate dextrose (ACD) solution A (Vacutainer, Ref: 364606; BD Biosciences) solution. The blood collected in one ACD tube was maintained in the same tube or divided into three polypropylene tubes containing no anticoagulant (Falcon™, Ref: 352063; BD Biosciences), termed as ACD-2.

Tubes were centrifuged at 300 g for 5 minutes (Megafuge® 40, Thermo Fisher Scientific, Waltham, MA). Supernatant containing plasma and platelets, termed as platelet-rich plasma 1 (PRP1), was collected from each tube and transferred to new polypropylene tubes containing no anticoagulant. In the case of ACD and ACD-2, after platelet counting, PRP1 from the same donors was mixed for the next experiments. Then, PRP1 was centrifuged at 700 g for 17 minutes. Supernatant was collected, namely, platelet-poor plasma (PPP). Part of the PPP from each tube was used to resuspend the platelet pellet, forming the platelet-rich plasma 2 (PRP2), in order to achieve the expected concentration of 106 platelets/µL. The platelets in PRP2 were activated by adding 1 M CaCl2 (final concentration of 20 mM) and incubated at 37°C for 1 hour. After clot formation, tubes were maintained at 4°C during 16 hours to allow clot contraction. Finally, tubes were centrifuged at 3000 g for 20 minutes and the supernatant was collected, termed as platelet-rich plasma releasate (PRPr). The PRPr was freezed at −80°C until thawing for experimental use.

2.3. Hematological Analysis

Counting of platelets, red blood cells, white blood cells, and analysis of mean platelet volume (MPV) were determined in whole blood, PRP1, PRP2, and PPP fractions. Those analyses were performed with a hematological analyzer (Mindray BC 2800, Perdizes, SP, Brazil). Platelet recovery after the first centrifugation step, expressed as a percentage, was calculated by dividing the total number of platelets in PRP1 by the total number of platelets in whole blood.

2.4. Quantification of Growth Factors

PRPr-derived TGF-β1 and VEGF were quantified using ELISA kits (Ref: KAC1688 and Ref: KHG0111; Invitrogen™, Thermo Fisher Scientific) according to manufacturer’s instructions. Absorbance was determined using a microplate reader (Multiskan GO, Thermo Fisher Scientific).

2.5. Bone Marrow-Derived Mesenchymal Stromal Cell (BM-MSC) Isolation and Culture

120 mL of bone marrow was obtained after donation with informed consent from two donors (a 41-year-old woman, whose cells were used for cell viability assays, and a 60-year-old man, whose cells were used for gene expression analysis). Mononuclear cells were separated using the Sepax system (Biosafe, Eysins, Switzerland), according to manufacturer’s instructions, and plated at 4 × 105 cells/cm2 in Minimum Essential Medium Eagle Alpha Modification (alpha MEM) (Cultilab, Campinas, SP, Brazil) supplemented with 10% fetal bovine serum (FBS) (Gibco) in T 150 cm2 culture flasks (Corning Incorporated, Corning, NY) and maintained in a 5% CO2 incubator at 37°C. After 5 days, medium was changed and nonadherent cells were discharged. Medium was changed every two days. This was termed as “primary culture.” After approximately 10 days, 70–80% confluence, cells were detached from culture flasks using 0.05% trypsin solution (Gibco®, Thermo Fisher Scientific) and replated onto new culture flasks at a density of 8 × 103 cells/cm2. After first trypsinization, culture was termed as at “passage #1.” Experiments were performed until “passage #5.”

2.6. Cell Viability Assay

The analysis of cell viability was performed by incorporation with thiazolyl blue tetrazolium bromide (MTT assay) (Sigma Aldrich, São Paulo, SP, Brazil). Cells (passage #3) were plated at a density of 5 × 103 cells/cm2 in duplicate in 48-well plates (Corning Incorporated) in alpha MEM (Cultilab) supplemented with 10% FBS (Gibco), 1% PRPr, 2.5% PRPr, or 5% PRPr. Four different PRPr donors were used. In another group, the four samples were pooled with equal proportions of each donor, namely, PRPr MIX. After 8 days of culture, 0.5 mg/mL MTT was added. Medium was removed after 4 hours of incubation, and 400 µL/well of DMSO was added to dissolve the reduced formazan product. The volume in each of the 48 wells was split into two wells in a 96-well plate (Corning Incorporated). Finally, the plate was read in a microplate reader (Multiskan GO, Thermo Fisher Scientific) at 570 nm. Cell culture medium was not changed during this experiment.

2.7. Gene Expression Evaluation

Cells (passage #5) were cultured in alpha MEM (Cultilab) supplemented with 10% FBS (Gibco), 1% PRPr, 2.5% PRPr, or 5% PRPr. The PRPr was used as a pool of four different donors. After five days of culture, total RNA was extracted using TRIzol® (Ambion®, Thermo Fisher Scientific). RNA concentration was determined using a Nanodrop 2000 UV-Vis spectrophotometer (Thermo Fisher Scientific) and 2 µg was reverse-transcripted into complementary DNA (cDNA) using SuperScript® First-Strand Synthesis System for RT-PCR (Invitrogen, #11904-018) in a total reaction volume of 20 µL, following manufacturer’s protocol. Oligonucleotides and probes for qPCR were purchased from Applied Biosystems (TaqMan Gene Expression Assay, #4331182): HPRT1 (Hs02800695_m1), which was analyzed as the housekeeping gene, SOX9 (Hs00165814_m1), RUNX2 (Hs00231692_m1), PPARG (Hs01115513_m1), and POU5F1 (Oct-4) (Hs0099634_9H). qPCR reactions were performed in an Applied Biosystems 7500 Standard Time PCR System in a 20 µL reaction volume using TaqMan® Universal Master Mix II, with UNG (Applied Biosystems, #4440038), according to manufacturer’s instructions. Analysis was performed using the ΔΔCt method [40].

2.8. Statistical Analysis

Data were analyzed using a two-tailed paired -test for the hematological analysis, where a group of the same donors were analyzed with different anticoagulants. In the case of growth factor quantification and cell culture experiments, where pairing of samples did not necessarily occur, a two-tailed unpaired -test was performed. Statistical significance was considered when .

3. Results

3.1. Effect of Different Anticoagulants on Initial Platelet Counting and Recovery

Blood samples were collected from five different donors in tubes containing EDTA, SC, or ACD, and platelets were counted in an automated system. Blood samples collected with EDTA yielded higher numbers of platelets, followed by SC and ACD (Figure 1(a)). In average, platelet counting in SC was 16.28% lower than that in EDTA, while that in ACD was 23.01% lower than in EDTA and 7.94% lower than in SC. However, platelet recovery, regarding the total number of platelets obtained after the first centrifugation step, was higher in the presence of SC compared to EDTA and ACD. The average of platelet recovery in EDTA and SC was 76.15% and 81.21%, respectively. Strikingly, platelet recovery in samples collected with ACD was 45.71%, almost half of those when using EDTA or SC. All three anticoagulants tested herein were purchased in commercially distributed tubes. ACD containing tube was bigger—taller and larger—compared to EDTA and SC.

Figure 1: Platelet yield and recovery in blood collected with different anticoagulants. Blood was collected in EDTA, SC, and ACD in five different donors and platelet concentration was quantified (a) as well as platelet recovery after the first centrifugation step (b). An individual analysis between ACD and ACD-2 of platelet recovery was also performed (c). Data are expressed as bar (a), box (b), and dot (c) plots. Similar symbols in (b) correspond to statistic similarity among groups ().

In order to verify if the lower platelet recovery was related to the tube format, PRP was obtained from blood samples anticoagulated in ACD using tubes of similar size compared to EDTA and SC (ACD-2). Platelet recovery improved (49.82%) but remained much lower than those recovered when using EDTA (76.15%) and SC (81.21%). Values from ACD-2 were statistically different from those obtained using SC () but not when compared to EDTA () (Figure 1(b)). If analyzed separately, it was possible to observe that platelet recovery has increased in three of the five donors when using ACD-2 instead of ACD, especially in donor 2, with an increase of 63.74%, while it has decreased in two donors, especially in donor 1, with a decrease of 25.48% after the distribution of blood into the smaller tubes (Figure 1(c)). In average, platelet concentration in PRP2 was 1,009 ± 57 × 103/µL in EDTA samples, 582 ± 108 × 103/µL in SC samples, 726 ± 200 × 103/µL in ACD samples, and 664 ± 170 × 103/µL in ACD-2 samples. All values were statistically similar between each other (), except between EDTA and SC () (data not shown).

Although the mean platelet volume (MPV), which is related to platelet size and indicates its degree of activation, was similar when whole blood (WB) samples were anticoagulated in all three anticoagulants tested, it increased progressively following the two centrifugation steps in EDTA group in all donors (in average an increase of 11.60% after the first centrifugation step and an additional increase of 2.84% after the second centrifugation step, totaling 14.44% increase compared to whole blood). This was not observed when WB was anticoagulated in SC and ACD (Figure 2).

Figure 2: Mean platelet value quantification of samples containing different anticoagulants. Mean platelet value was quantified in five different donors in whole blood (WB), PRP1, and PRP2, in tubes containing EDTA, SC, or ACD solution. The average values of the five different donors are also represented in the figure. “” corresponds to statistical difference between EDTA and SC groups as well as EDTA and ACD groups ().

3.2. TGF-β1 and VEGF Release from Platelet-Rich Plasma in Different Anticoagulants

Up to this point, it was clear that the anticoagulant has an impact on platelet recovery after blood centrifugation. However, we questioned if it would also change growth factors release from recovered platelets. For that, we quantified TGF-β1 and VEGF levels in an ELISA assay. Growth factors concentrations were similar between anticoagulant groups () for both TGF-β1 and VEGF. TGF-β1 concentration was 18,146.99 ± 2,370.33 pg/mL in EDTA; 48,559.10 ± 12,839.86 pg/mL in SC; and 30,786.15 ± 6,654.49 pg/mL in ACD (Figure 3(a)). VEGF concentration was 278.88 ± 71.78 pg/mL in EDTA, 143.65 ± 71.63 pg/mL in SC, and 362.70 ± 77.95 pg/mL in ACD (Figure 3(b)).

Figure 3: Growth factor quantification in PRPr obtained in different anticoagulant. TGF-β1 (a) and VEGF quantification (b). Data are expressed as mean, and error bars correspond to standard error.

3.3. Bone Marrow-Derived Mesenchymal Stromal Cell Culture

In order to show the effects of factors released from platelets obtained using different anticoagulants on modulating cell expansion in vitro, we used the MTT cell viability assay to analyze BM-MSC proliferation in the presence of different concentrations of PRPr. FBS-supplemented culture medium was used as reference (Figure 4). PRPrs were tested separately and mixed (MIX). All concentrations of PRPr tested from all donors were able to stimulate cell proliferation in vitro. As expected, the higher concentration tested (5%) stimulated the higher proliferative rate in vitro, regardless of the anticoagulant used. However, for this concentration, in average, cell proliferation was lower in the presence of EDTA derived PRPr compared to SC and ACD (Figure 4). In addition, cells maintained their fibroblast-like morphology regardless of the anticoagulant type (Figure 5).

Figure 4: BM-MSC viability in PRPr obtained with different anticoagulant. Absorbance at 570nm was measured after MTT viability assay of cells cultivated in different PRPr concentrations, obtained with EDTA (a), SC (b), and ACD (c), as well as 10% FBS (control). Data are expressed as mean, and error bars correspond to standard error. “” corresponds to statistical similarity with 10% FBS ().

Figure 5: Photomicrography of BM-MSC. Cells cultivated for eight days in medium supplemented with 10% FBS (a) or a pool from four donors of 5% PRPr obtained from collection tubes containing EDTA (b), SC (c), or ACD (d). Phase contrast, 200x magnification, and scale bars: 50 µm.

3.4. Modulation of Bone Marrow-Derived Mesenchymal Stromal Cell Gene Expression by Platelet-Rich Plasma Culture

We also analyzed gene expression of cells expanded in vitro (Figure 6). Only cells cultured in 5% PRPr were tested. Using 10% FBS as reference, RUNX2 was slightly upregulated in EDTA group and downregulated in SC group. PPARγ2 was slightly upregulated in EDTA group and downregulated in SC and ACD groups. SOX9 was downregulated in all groups. Oct-4 was upregulated in EDTA and ACD groups and downregulated in SC group. Although, in general, the gene expression was similar between the PRPr groups, especially when observing the maximum and minimum relative quantification of gene expression, the SC group presented the smallest variation compared to the control group, by analyzing the average relative expression of the four genes in the PRPr groups compared to the FBS group. In average, SC relative gene expression was 24.73% different from the control group, while EDTA was 46.79% and ACD was 29.74% different.

Figure 6: Relative gene expression. RUNX2, PPARγ2, SOX9, and Oct-4 gene expression in cells cultured in medium supplemented with 10% FBS (control group) or 5% PRPr obtained with EDTA, SC, or ACD. Data are expressed as relative quantification of gene expression (RQ). Upper and lower error bars correspond to RQ maximum and RQ minimum, respectively.

4. Discussion

Platelet-rich plasma (PRP) is currently one of the main strategies to promote musculoskeletal tissues repair. There are several reports in the literature evidencing its potential in clinical trials [2–11] as well as in vitro analysis [12–15]. As a cost-effective source of autologous growth factors that can affect stem cells proliferation and differentiation, it is being increasingly investigated as a supplement, adjuvant, carrier, or scaffold for stem cells-based therapeutics [41–45]. However, the lack of standardization between the methodology to obtain and use PRP among different groups may hamper the development of this technology [20]. The use of anticoagulant to collect blood is a major issue. The present work aimed to verify PRP obtaining with three types of commercially available blood collection tubes containing EDTA, SC, or ACD as anticoagulants.

Platelet counting was higher in blood collected in tubes containing EDTA, followed by SC and ACD. Indeed, it has been previously shown that platelet count in EDTA can be higher than in citrate anticoagulants [46]. Moreover, when EDTA is added to citrated samples, it can enhance platelet count in whole blood [47]. Platelet recovery after the first centrifugation step was diminished in ACD tubes compared to EDTA and SC. Additionally, a higher concentration of PDGF-BB was found in PRP obtained with EDTA compared to ACD [48]. In our case, we tried to enhance platelet recovery in ACD tubes by dividing its content into smaller tubes (12 × 75 mm × 5 mL) with no additional anticoagulant. Although no statistical difference has been detected between those two ACD forms, the splitting of blood in the smaller tubes resulted in a similar platelet recovery compared to EDTA group. Since ACD tube is bigger (16 × 100 mm × 8.5 mL) than EDTA (13 × 75 mm × 4.0 mL) and SC tubes (13 × 75 mm × 4.5 mL), it is possible that the lower platelet recovery is due not only to the type of anticoagulant itself but also to the tube format. Particularly, the tube format may have superior influence on whole blood/plasma than serum centrifugation, in view of its higher viscosity [49]. For the following experiments, we decided to mix ACD and ACD-2 PRP1 from the same donors, since it is comprised of the same type of anticoagulant and since the following centrifugation to prepare PRP2 was performed in the same type of tube for all groups analyzed.

In order to verify if the centrifugation steps influenced platelets morphology, we quantified the mean platelet volume (MPV) in whole blood, PRP1, and PRP2 obtained with different anticoagulants. The MPV in the EDTA group, but not in SC and ACD groups, has increased after the centrifugation steps, which may be an indicator of platelet activation [50, 51]. Indeed, a higher MPV is expected in whole blood collected in EDTA compared to citrated samples [46]. In addition, EDTA may change platelet morphology from a discoid to an irregular spherical shape [52]. Indeed, the use of EDTA faces ethical issues, such as being pointed to as a persistent pollutant in natural environments [53], and impediments of use in certain countries [54]. ACD and citrate-theophylline-adenosine-dipyridamole (CTAD) are also more efficient in maintaining platelet morphology than heparin and SC. In that case, it has also been shown that PRP obtained with ACD and CTAD resulted in higher TGFβ-1 concentration and induction of MSC proliferation [55]. In another previous study, EDTA, SC, and ACD were compared as maintainers of platelet responsiveness to aggregation inducers. ACD was the most capable to maintain intraplatelet signal transduction mechanisms during PRP formulation [56]. The same group, lately, showed that ACD was also capable of maintaining platelet functions for periods of time superior to SC [57]. In our case, we could not find any difference in the capability of platelet activation and clot formation in PRP2 obtained with the three different anticoagulants.

Our next step was to evaluate the effect of PRP obtaining in cell culture. For that, we normalized the platelet concentration to ideally 1000 × 103/µL in all groups, in a way that the results would not correlate to platelet concentration but to the type of the anticoagulant. This platelet concentration in PRP is being pointed to as a therapeutic concentration for in vivo purposes [1]. In bone, for example, lower concentrations are unsatisfactory and higher concentrations are inhibitory to promote tissue repair [11]. The only statistical difference observed in platelet concentration in PRP2 was between EDTA and SC group. The lower value in SC group may be attributed to a difficulty to resuspend platelet pellet. Platelet concentrate bags prepared with citrate samples, in form of ACD, contain more aggregates than bags prepared with EDTA [52]. Moreover, the effects of PRPr on cell culture were always compared to 10% FBS medium supplementation. In spite of being a xenogeneic serum with ethical and scientific issues [58], FBS is still widely used in cell culture [59] and MSC in vitro expansion [60].

In our study, we could not find differences in TGFβ-1 and VEGF concentration among the anticoagulants. It could be possibly due to the similar platelet concentration between groups. In average, TGF-β1 concentration varied from 18.15 ng/mL in EDTA to 48.56 ng/mL in SC. Other literature reports present concentrations superior to our finding: 120 ng/mL [61], 169 ng/mL [62], and 89 ng/mL in a PRP with platelet concentration 4.69 times superior to whole blood baseline but 20 ng/mL in a PRP with platelet concentration 1.99 times superior to whole blood baseline [63]. VEGF concentration ranged from 143.65 pg/mL in SC to 362.70 pg/mL in ACD. Other reports present concentrations varying from 50 pg/mL [64] to 155 ng/mL [61]. Nevertheless, our results were superior to some commercially available kits [65].

As expected, PRPr induced cell proliferation. Although there were variations comparing the influence on cell proliferation between donors, possibly due to an inherent difference on growth factors and other molecules content among each platelet granule, an evident pattern emerged: the concentration of 5% PRPr in cell culture medium was sufficient to induce cell proliferation in a similar level to 10% FBS. Additionally, SC and ACD-derived PRPr presented greater effects over cell proliferation compared to EDTA group. Cell morphology was not changed among groups. BM-MSC maintained their fibroblastic morphology regardless of the anticoagulant. Although there are still controversies in the literature regarding PRP effects on BM-MSC differentiation, there is a consensus on its effect as an inducer of proliferation [66].

As a final analysis of PRPr effects on BM-MSC, we observed slight modulations in the expression of the master genes for the osteogenic (RUNX2), adipogenic (PPARγ2), and chondrogenic (SOX9) lineages [67], as well as Oct-4, a gene related to maintenance of stemness [68, 69]. Particularly, SOX9 expression was downregulated. Indeed, there is a discussion in the literature regarding the PRP effects on chondrogenesis, with some groups claiming its inducing effects [70, 71] and others its inhibitory effects [72, 73]. Recently, our group has showed that it can be dependent on the concentration of PRPr used in cell medium, with lower concentrations inducing chondrogenesis and higher concentrations inhibiting it [74]. Oct-4 expression presented an intense variability, with upregulation in EDTA and ACD groups and downregulation in SC group. Whether this can be an indicator of the maintenance of cells stemness or not, it must be further investigated. In general, gene expression was similar between PRP groups, although SC was the group that presented the smaller variation compared to 10% FBS culture, evidencing that cells presented the lightest changes in their phenotype with this treatment.

5. Conclusion

The literature provides a variety of methodologies to obtain PRP. The first variation is the methodology to collect blood and the anticoagulant used. In this paper, we analyzed the effects of three different anticoagulants, obtained in commercially available tubes, on PRP obtaining. Although no significant change in the amount of growth factors released was observed, some features could be highlighted. The blood collection in tubes containing EDTA resulted in higher platelet yield in the whole blood. However, this was accompanied by an increase of MPV following the centrifugation steps, which is an indicator of change in platelet morphology. On the other hand, the use of tubes containing citrate solutions resulted in a greater induction of MSC proliferation. Particularly, the obtaining in SC resulted in the higher platelet recovery after the first centrifugation step. If ACD is used, the reduction of the tube size may increase platelet recovery. In addition, the PRP obtained in SC presented the smallest variation in MSC gene expression compared to cells cultured in the presence of 10% FBS. Therefore, in order to obtain a bigger amount of platelets and induce MSC proliferation without dramatically interfering with their phenotype, we suggest the use of SC as anticoagulant for PRP acquisition.

Competing Interests

The authors declare no competing interests.

Acknowledgments

The authors would like to thank all the staff at Amil Life Sciences for support.

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REF 2


In summary, both phosphate and citrate buffers revealed significant cytotoxic effects at high concentrations and longer incubation times in ocular epithelial monolayers. Transferring the results of this study to a clinical setting requires thorough in vivo preclinical data, as topical application involves factors such as blinking and tear production that need to be considered. Thus, to translate these finding to humans, further experiments with stratified cells in vitro and preclinical in vivo studies in animals are planned to discern the actual effects of the buffer concentration in eye drops on the ocular surface. This study highlights the relevance of maintaining ocular homeostasis to ensure ocular surface health.

Cornea. 2017 Jun; 36(6): 712–718.

Published online 2017 Apr 10. doi: 10.1097/ICO.0000000000001199

PMCID: PMC5421463

PMID: 28399036

Implications for Ophthalmic Formulations: Ocular Buffers Show Varied Cytotoxic Impact on Human Corneal–Limbal and Human Conjunctival Epithelial Cells

Nadine Schuerer,* Elisabeth Stein,* Aleksandra Inic-Kanada,* Marion Pucher,† Christine Hohenadl,† Nora Bintner,* Ehsan Ghasemian,* Jacqueline Montanaro,* and Talin Barisani-Asenbauer*

Author information Article notes Copyright and License information Disclaimer

Copyright © 2017 The Author(s). Published by Wolters Kluwer Health, Inc.

This is an open-access article distributed under the terms of the Creative Commons Attribution-Non Commercial-No Derivatives License 4.0 (CCBY-NC-ND), where it is permissible to download and share the work provided it is properly cited. The work cannot be changed in any way or used commercially without permission from the journal.

Associated Data

Supplementary Materials

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Abstract

Purpose:

To investigate toxicity associated with buffers commonly used in topical ocular drug formulations using a human corneal–limbal epithelial (HCLE) and a human conjunctival epithelial (HCjE) cell model.

Methods:

HCLE and HCjE cells were incubated for 10, 30, or 60 minutes with 4 different buffers based on borate, citrate, phosphate, and Tris-HCl at 10, 50, and 100 mM concentrations. To detect possible delayed effects on cell viability, after 60 minutes of buffer incubation, cells were further incubated for 24 hours with a cell medium. Cell viability was determined using a colorimetric XTT–based assay. The morphology of cells was also investigated.

Results:

HCjE cells showed more sensitivity to buffer incubation than HCLE cells. The 100 mM phosphate buffer displayed significant delayed effects on cell viability of HCLE 16.8 ± 4.8% and HCjE 39.2 ± 6.1% cells after 60 minutes of exposure (P < 0.05). HCjE cell viability was reduced after 60 minutes incubations with 50 and 100 mM citrate buffer to 42.8 ± 6.5% and 39.3 ± 7.9%, respectively, and even lower percentages at the delayed time point (both P < 0.05). HCLE cell morphology was distinctly altered by 100 mM phosphate and Tris buffers after 30 minutes, whereas HCjE cells already showed marked changes after 10 minutes of exposure to 100 mM citrate and phosphate buffers.

Conclusions:

We observed a time-dependent decrease of viability in both HCLE and HCjE cells exposed to higher buffer concentrations. Therefore, we propose further in vivo studies to translate these finding to humans to discern the real effects of the buffer concentration in eye drops on the ocular surface.

Key Words: ocular epithelium, topical drug delivery, biocompatibility, eye drops

Ophthalmic drugs are frequently administered topically, facing special challenges at the ocular surface concerning tolerability, drug permeability, and safety. Eye drop formulations often contain buffering agents. The buffer system of eye drops is often a compromise between the necessities of chemical stabilization to achieve acceptable shelf life and the physiological requirements of the product.1 Ideally, a formulated buffer system should mimic the natural system of the tear film. Tear fluid consists of 3 main elements: a bicarbonate buffer system, various proteins with dissociable groups, and phosphate compounds. The buffering capacity highly varies between individuals, yet the average physiological pH of lacrimal fluid is 7.4,2 ranging from 6.5 to 7.6.3,4

The most common buffer systems used in ophthalmic formulations are citrate, phosphate, Tris-HCl (Tris), and borate buffer (Table ​(Table1).1). No complications after the application of citrate, borate, and Tris buffering systems in humans are reported in the literature. However, there have been selected case studies in which calcific band keratopathy, a calcific degeneration of the superficial cornea characterized by calcium hydroxyapatite deposition, has appeared after instilling phosphate buffer containing ophthalmic medicinal products.5,6 Especially in eye drops applied after corneal injuries, the buffer composition has been shown to influence the healing process and the development of corneal calcification.5 As phosphate naturally occurs in the eye, it has been the buffer of choice for a long time.6 More recently, borate buffers were introduced as ocular buffers because of their antimicrobial activity and were deemed better suited than phosphate buffers for ophthalmic products.7 Tris buffers were originally tested for experimental ocular acid burns in rabbits and were found to be effective at treating ocular acid burns when the time of exposure to acid was short.8

TABLE 1.

Buffer Systems in Use for Ophthalmic Formulations

Although literature on the toxicity of preservatives as components in eye drops is available,9 there are no recent studies dealing with the comparative toxicity of buffer systems on a cellular level. Suitable in vitro systems for the evaluation of novel compounds are a critical step in the process of safety screenings for ocular formulations. We decided to use a corneal and conjunctival epithelial model because the integrity of the ocular epithelium is crucial for visual function and one of the first points of contact for eye drops.10 In addition, these specific cell lines were chosen because they have been thoroughly characterized11 and have been used in previous studies to determine cytotoxic effects.12,13 The purpose of this study was to investigate toxicity of conventional buffer systems used in ophthalmic medicinal products and devices using both human corneal–limbal epithelial (HCLE) and human conjunctival epithelial (HCjE) cell line models.

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MATERIALS AND METHODS

Cell Culture

Immortalized HCLE and HCjE cells were kindly provided by Ilene Gipson (Schepens Eye Research Institute, Harvard Medical School, Boston, MA) and their characterization has been reported.11 Both cell lines were maintained in monolayers in a serum-free keratinocyte growth medium (Life Technologies, Paisley, United Kingdom) at 37°C, 5% CO2, and 95% humidity. The medium was changed every second day, and the cells were passaged at 70% confluence. Cells were harvested by trypsinization (0.05% trypsin/0.02% EDTA in PBS; PAA Laboratories GmbH, Pasching, Austria) and seeded while within passage 2 to 6.

Preparation of Buffer Solutions

The 4 different buffers were prepared at concentrations of 10, 50, and 100 mM, all with a pH of 7.4 ± 0.1. The osmolarity of the 4 buffer solutions was set to 301 ± 18 mOsm/kg using addition of NaCl, and osmolarity was measured using the Osmomat 030-D (Gonotec GmbH, Germany), according to the manufacturer's instructions.

Cytotoxicity Assay

Cell viability was measured to assess the in vitro toxicity of 3 different concentrations of borate, citrate, Tris, and phosphate buffers through mitochondrial lactate dehydrogenase production. Cells were seeded at 1 × 104 cells/well in 96-well plates and incubated at 37°C, 5% CO2 for 24 hours for HCLE cells and 48 hours for HCjE cells. After aspiration of the cell culture medium, 100 μL of the respective buffer at a concentration of 10, 50, and 100 mM, respectively, were applied to each well and incubated at 37°C, 5% CO2 for 10, 30, or 60 minutes. Subsequently, the test buffer was aspirated and replaced with a cell culture medium. Control wells were exposed only to the cell culture medium without growth factors for the same time, as experimental wells were exposed to buffers. For immediate readout, 50 μL of XTT/PMS (N-methyl dibenzopyrazine methyl sulfate) solution (AppliChem GmbH, Germany) was added and incubated for 3 hours under standard conditions. During this period, metabolically active cells reduce the XTT tetrazolium salt to formazan.14 Then, 100 μL of the cell culture medium was transferred from each well into a new plate, and absorption was measured at 450 nm (reference wave length 630 nm) with a photometer (Tecan GENios). To detect possible delayed cell death, cells that had been cultured for 24 hours after incubation using buffers for 60 minutes were analyzed as well.

Calculations Performed

Abs_control: absorbance of corneal or conjunctival cells exposed only to the medium without growth factors.

Microscopy

Cells were seeded at 1 × 105 cells/well into 4 chamber slides and incubated for 24 hours in the case of HCLE cells and for 48 hours for HCjE cells at 37°C/5% CO2 to reach the required degree of confluency. Cells were then incubated with the various buffer solutions at a concentration of 10 or 100 mM for 10 or 30 minutes at standard conditions. The cells were fixed with 4% PFA, stained with hematoxylin and eosin (Sigma-Aldrich, St Louis, MO), mounted, and then examined by microscopy (Zeiss Axiovert 200 Cell Observer; Carl Zeiss GmbH, Vienna, Austria).

Statistics

All experiments were performed 3 times, and a type I 2-way analysis of variance with the Tukey multiple comparisons test was used to compare cell viability data considering the 2 factors buffer type and the time of exposure as well as the 2 factors concentration and time of exposure (GraphPad Prism 6).

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RESULTS

Viability of HCLE Cells Is Not Significantly Affected by the Choice of Buffer in Low Concentrations and Short Incubation

All buffers were well tolerated at concentrations of 10 and 50 mM at 10 minutes with average percentage viability between 88.5% and 100%. Incubation times over 30 minutes showed a significant decrease in cell viability at all concentrations. Generally, the longer the HCLE cells were exposed to the various buffer solutions, the significantly lower the viability of the cells (Figs. ​(Figs.1A,1A, ​A,1B,1B, ​B,1C;1C; P < 0.01).

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FIGURE 1.

Viability of HCLE cells incubated with distinct buffer solutions. HCLE cells were incubated with borate, phosphate, citrate, or Tris buffers at the indicated concentrations for 10, 30, 60 minutes, and 60 minutes followed by a recovery period of 24 hours. Cell viability (XTT assay) was evaluated in comparison with cells incubated in the serum-free keratinocyte cell culture medium, lacking growth factors for the same time. (A) 100 mM; (B) 50 mM; (C) 10 mM. At 10 minutes, all buffers were statistically different from all other time points (♦ P < 0.01). Statistical significance of determined differences was tested by a 2-way analysis of variance with the Tukey multiple comparisons test (n = 3; *P < 0.05).

Apart from a decreased viability of 16.8 ± 4.8% at 24 hours in HCLE cells exposed to 100 mM phosphate buffer for 60 minutes, other buffers showed no continued effects on late stage cell viability at 24 hours (Fig. ​(Fig.1A;1A; P < 0.05).

Statistical analysis between concentrations of the different buffers showed significant changes in the phosphate, citrate, and Tris buffer concentrations of P < 0.05 (see SDC 1, Supplemental Digital Content 1, http://links.lww.com/ICO/A516).

HCjE Cell Viability Is Significantly Affected by Citrate Buffer at High Concentrations and Longer Incubation Times

Lower viability of HCjE cells was observed with all 4 buffers at a high buffer concentration; in addition, the HCjE cells showed lower viability than HCLE cells after 10 minutes of incubation with all 4 buffers with average viability levels of 58.3% to 79.2%. Citrate buffer was observed as the most cytotoxic buffer to this cell line with 100 mM citrate buffer showing a significant reduction in the percentage of HCjE cell viability after 30 and 60 minutes to average percentages of 60.0 ± 7.1% and 39.3 ± 7.9%, respectively, compared with borate and Tris buffers at the same time points (Fig. ​(Fig.2A;2A; all P < 0.05). Furthermore, at the highest concentration, a significantly higher viability was observed after 30 minutes of incubation with Tris buffer compared with phosphate buffer (P < 0.05). HCjE cells at 24 hours that had been exposed to 100 mM citrate or phosphate buffer for 60 minutes also showed high levels of cytotoxicity of 30.4 ± 1.1% or 39.2 ± 6.1% compared with the Tris buffer levels of 84.5 ± 12.1% (both P < 0.05).

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FIGURE 2.

Viability of HCjE cells incubated with distinct buffer solutions. HCjE cells were incubated with borate, phosphate, citrate, or Tris buffers at the indicated concentrations for 10, 30, 60 minutes, and 60 minutes followed by a recovery period of 24 hours. Cell viability (XTT assay) was evaluated in comparison with cells incubated in the serum-free keratinocyte cell culture medium, lacking growth factors for the same time. (A) 100 mM; (B) 50 mM; a statistical difference was seen at the 24-hour time point compared with 10 minutes (♦ P < 0.05), (C) 10 mM; the 24-hour time point was statistically different from 30 minutes (♦ P < 0.05). Statistical significance of determined differences was tested by a 2-way analysis of variance with the Tukey multiple comparisons test (n = 3; *P < 0.05).

After 60 minutes of incubation and 24 hours after treatment with 50 mM citrate buffer, viability of HCjE cells was 42.6 ± 6.5%, significantly lower than the borate and Tris buffers (Fig. ​(Fig.2B;2B; all P < 0.05). The citrate buffer at 24 hours after the 60-minute exposure also showed greater cytotoxicity than at 10 minutes (P < 0.05).

For the 10 mM buffer concentration, differences were observed between the 24-hour time points after 60 minutes of treatment with citrate buffer compared with all other buffers (P < 0.05), showing that already at this low concentration, citrate buffer can produce a cytotoxic effect. In addition, the 24-hour time point of 10 mM citrate buffer showed significantly lower viability of 33.3 ± 0.4% than at the 30-minute time point at which viability was 66.8 ± 8.0% (Fig. ​(Fig.2C,2C, P < 0.05).

Statistical analysis of the different buffer concentrations showed significant changes in the borate, phosphate, and citrate buffer concentrations of P < 0.05 (see SDC 1, Supplemental Digital Content 1, http://links.lww.com/ICO/A516).

HCLE and HCjE Cell Morphology Is Altered at Higher Buffer Concentrations and Longer Incubation Times

When compared with control cells, all buffers tested showed mild changes in HCLE and HCjE cell morphology at the lowest concentration and time points, with rounding of cells and changes in cell-to-cell contact indicated by prominent cell processes and clearing areas around individual cells (Figs. ​(Figs.3,3, ​,4).4). These changes were exacerbated with longer exposure to the higher concentration buffers.

FIGURE 3.

Morphological evaluation of HCLE cells treated with ocular buffers. HCLE cells were incubated in 10 mM or 100 mM borate (A), citrate (B), phosphate (C), or Tris (D) buffers for 10 or 30 minutes; the serum-free cell culture medium was used as a control. After staining with hematoxylin and eosin, images were taken with a Zeiss AxioObserver. Original magnification ×20.

FIGURE 4.

Morphological evaluation of HCjE cells treated with ocular buffers. HCjE cells were incubated in 10 or 100 mM borate (A), citrate (B), phosphate (C), or Tris (D) buffers for 10 or 30 minutes; the serum-free cell culture medium was used as a control. After staining with hematoxylin and eosin, images were taken with a Zeiss AxioObserver. Original magnification ×20.

The higher concentration buffers showed marked differences in cytopathic effects with dark granular bodies and blebbing of epithelial processes. The 30-minute incubation with 100 mM borate buffer showed some condensation of nuclei, whereas the cytoplasm is even with some clearing or thinning between the edges of the cells (Figs. ​(Figs.3A,3A, ​A,4A).4A). The 100 mM citrate buffer at 30 minutes in HCLE cells showed that visible cell-to-cell contact was maintained and showed mild vacuolation of the cytoplasm (Fig. ​(Fig.3B).3B). However, in the HCjE cells, this buffer concentration already showed loss of cytoplasm after 10 minutes, and this was intensified after 30 minutes with very marked pyknosis (Fig. ​(Fig.4B).4B). Both cell lines treated with 100 mM phosphate buffer showed some condensation of nuclei and loss of cytoplasm after 10 minutes, whereas HCjE cells also displayed dark basophilic blebbing at the margins of the cytoplasm. Marked changes were seen after 30 minutes of exposure to phosphate buffer in almost half of the HCLE cells (Fig. ​(Fig.3C)3C) and in all the HCjE cells (Fig. ​(Fig.4C).4C). These changes included pyknotic nuclei, loss of cell-to-cell contact, reduction in cytoplasm, and shrinkage. Incubation with 100 mM Tris buffer showed some clearing between cell margins and condensation of nuclei after 10 minutes, whereas after 30 minutes, moderate changes with some pyknotic nuclei and many prominent dark basophilic bodies in the cytoplasm and in cell processes were observed (Fig. ​(Fig.3D).3D). Interestingly, HCjE cells showed minimal changes in response to the Tris buffer with only mild blebbing after 30 minutes at the 100 mM concentration (Fig. ​(Fig.44D).

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DISCUSSION

Topically administered ocular formulations have specific requirements for their buffering system. Buffers are needed to stabilize the pH at a level at which drugs are soluble, active, and tolerable. Because buffer capacity is regulated by its concentration, some formulations use higher dosages to enhance the drug's performance.

In this study, we investigated the toxicity of conventional buffer systems used in ophthalmic medicinal products and devices using 2 ocular cell line models. The 2 cell lines reacted differently to incubations with buffers. In lower concentrations, all tested buffers showed significant loss in HCLE cell viability after 30 minutes of incubation compared with 10 minutes, which is to be expected as the cells were not being kept in the ideal culture conditions needed for survival. Interestingly, this phenomenon was not visible in HCjE cells, in which the 10-minute time point already showed a reduction in viability. These differences could be explained by the different functions these cells fulfill in vivo, in which the avascular corneal epithelium has mostly a barrier function, whereas the conjunctival epithelium acts as a barrier and crucial mediator of the immune response triggered by inflammation.15 Therefore, we assume that conjunctival cells are quicker to respond to stimuli and are more sensitive than corneal epithelial cells.

No differences were observed in both cell lines between 10 mM buffers for up to 60 minutes of incubation, which suggests that the reduced ability of the cells to survive is due to an absence of culture medium rather than a specific buffer formulation. However, the highest concentration of phosphate buffer resulted in significantly lower HCLE and HCjE cells long-term viability and affected their morphology after 30 minutes of incubation.

The concentration of phosphate buffer has been investigated in ophthalmic products and in animal models. In a published study on rabbits, adverse effects of higher concentrations of phosphate buffers was reported, in which highly concentrated phosphate-buffered solution permanently altered physiological pH, even after discontinuation.16 Although some studies showed partly severe sequelae, an evaluation by the European Medicines Agency showed hardly any risk to patients without preexisting corneal defects.17 In Germany alone, 37% of ocular medicinal products contain phosphate buffers, which are most commonly used in antiglaucoma medications and prostaglandin-containing formulations.18 The concentration of phosphate in tested artificial tears ranged from <0.1 mM to 68.8 mM. Forty-four percent of the samples revealed phosphate concentrations above the physiological level (1.45 mM), and in 5% of the products, there were phosphate concentrations higher than 50 mM.19 Within the antiglaucoma medications analyzed, 47% of the formulations showed phosphate concentrations higher than the physiological level, and 19% had concentrations above 100 mM. The concentrations of the tested devices ranged from <0.1 to 160 mM.20 Unfortunately, most patient information leaflets do not report the concentration of the buffers used.

Our investigation also revealed cytotoxic effects of citrate buffer on HCjE cells; this was observed both in the XTT assay and morphologically and was shown to be enhanced by the time of exposure and higher concentration. It might be that the mechanism behind the cytotoxicity seen in citrate and phosphate buffers, reported to be implicated in the development of calcific band keratopathy,6,7 is due to the ocular surface having calcium-dependent channels that maintain the osmotic balance.21–23 Both citrate and phosphate buffers are not recommended for systems that are highly calcium-dependent, as citric acid and its salts act as calcium chelators, whereas phosphates react with calcium thereby producing insoluble calcium phosphate that precipitates out of the system.

Earlier reports suggested that borate-buffered contact lens multipurpose solutions showed increased cytotoxicity compared with phosphate-buffered multipurpose solution with mere cells24 or with contact lenses on cells.25 However, more recent investigations are in agreement with our findings that borate buffer is not toxic to HCLE and HCjE cells. Good biocompatibility was observed both in vitro with a 1% borate buffer (approximately 162 mM) for up to 1 hour,26 as well as in a rabbit model.27

Although we found no information in the literature on Tris buffers for ocular administration, our results indicate that this buffer is the least toxic to HCjE cells from the 4 buffers tested in this study and shows similar effects as borate buffer on HCLE cells.

In summary, both phosphate and citrate buffers revealed significant cytotoxic effects at high concentrations and longer incubation times in ocular epithelial monolayers. Transferring the results of this study to a clinical setting requires thorough in vivo preclinical data, as topical application involves factors such as blinking and tear production that need to be considered. Thus, to translate these finding to humans, further experiments with stratified cells in vitro and preclinical in vivo studies in animals are planned to discern the actual effects of the buffer concentration in eye drops on the ocular surface. This study highlights the relevance of maintaining ocular homeostasis to ensure ocular surface health.

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Supplementary Material

SUPPLEMENTARY MATERIAL:

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ACKNOWLEDGMENTS

The authors thank Jelena Marjanovic for excellent technical assistance.

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Footnotes

Supported by the “Laura Bassi Centers of Expertise” program of the Austrian Federal Ministry of Economy through the Austrian Research Promotion Agency (FFG Project Number: 822768).

M. Pucher and C. Hohenadl are employees of Croma-Pharma, Austria. The remaining authors have no conflicts of interest to disclose.

Supplemental digital content is available for this article. Direct URL citations appear in the printed text and are provided in the HTML and PDF versions of this article on the journal's Web site (www.corneajrnl.com).

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Evaluation of autologous platelet rich plasma for cardiac surgery: outcome analysis of 2000 patients

• Amit N. Patel, 
• Craig H. Selzman, 
• Ganesh S. Kumpati, 
• Stephen H. McKellar & 
• David A. Bull 

Journal of Cardiothoracic Surgeryvolume 11, Article number: 62 (2016) | Download Citation

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Abstract

Background

Deep and superficial sternal wound infections (DSWI & SWI) following cardiac surgery increase morbidity, mortality and cost. Autologous platelet rich plasma (PRP) derived from the patient’s own blood has been used in other surgical settings to promote successful wound healing. The goal of this study was to analyze the addition of PRP using a rapid point of care bedside system to standard wound care in all patients undergoing sternotomy for cardiac surgical procedures.

Methods

Over a 7 year period, 2000 patients undergoing open cardiac operations requiring sternotomy were enrolled. One thousand patients received standard of care sternal closure. The other 1000 patients received standard of care sternal closure plus PRP applied to the sternum at the time of closure. The outcomes related to wound healing, infection, readmissions, and costs were analyzed.

Results

In the 2000 patients, there were more ventricular assist device implants/heart transplants and emergency operations in the PRP group; otherwise there were no significant differences. The use of PRP reduced the incidence of DSWI from 2.0 to 0.6 %, SWI from 8.0 to 2.0 %, and the readmission rate from 4.0 to 0.8 %. The use of PRP reduced the costs associated with the development of deep and superficial wound complications from $1,256,960 to $593,791.

Conclusions

The use of PRP decreases the incidence and costs of sternal wound complications following cardiac surgery. The routine use of platelet rich plasma should be considered for all patients undergoing sternotomy for cardiac surgical procedures.