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. 2020 Sep 18;19(5):403–412. doi: 10.2450/2020.0100-20

Analysis of the mechanism of damage produced by thiazole orange photoinactivation in apheresis platelets

Portia Gough 1,✉, Todd Getz 2, Silvia De Paoli 1, Stephen Wagner 2, Chintamani Atreya 1,✉
PMCID: PMC8486609  PMID: 32955423

Abstract

Background

Pathogen Reduction Technologies (PRTs) are broad spectrum nucleic acid replication-blocking antimicrobial treatments designed to mitigate risk of infection from blood product transfusions. Thiazole Orange (TO), a photosensitizing nucleic acid dye, was previously shown to photoinactivate several types of bacterial and viral pathogens in RBC suspensions without adverse effects on function. In this report we extended TO treatment to platelet concentrates (PCs) to see whether it is compatible with in vitro platelet functions also, and thus, could serve as a candidate technology for further evaluation.

Material and methods

PCs were treated with TO, and an effective treatment dose for inactivation of Staphylococci was identified. Platelet function and physiology were then evaluated by various assays in vitro.

Results

Phototreatment of PCs yielded significant reduction (≥4-log) in Staphylococci at TO concentrations ≥20 μM. However, treatment with TO reduced aggregation response to collagen over time, and platelets became unresponsive by 24 hours post-treatment (from >80% at 1 h to 0% at 24 h). TO treatment also significantly increased CD62P expression (<1% CD62P+ for untreated and >50% for TO treated at 1 h) and induced apoptosis in platelets (<1% Annexin V+ for untreated and >50% for TO treated at 1 h) and damaged mitochondrial DNA. A mitochondria-targeted antioxidant and reactive oxygen species (ROS) scavenger Mito-Tempo mitigated these adverse effects.

Discussion

The results demonstrate that TO compromises mitochondria and perturbs internal signaling that activates platelets and triggers apoptosis. This study illustrates that protecting platelet mitochondria and its functions should be a fundamental consideration in selecting a PRT for transfusion units containing platelets, such as PCs.

Keywords: platelet, thiazole orange, mitochondria

INTRODUCTION

A major goal of pathogen reduction technologies (PRT) is to enhance the safety of the blood supply by reducing the risk of transfusion-transmitted infections (TTIs) from both known and unknown (newly emerging) infectious agents1. The ideal PRT would accomplish pathogen inactivation in a manner that is uncomplicated to execute and cost-effective, without reducing the quality or efficacy of the transfusion product1,2. The absence of an effective and balanced PRT renders blood transfusion products vulnerable to emerging and potentially lethal infectious agents3–5.

Platelet concentrates (PCs) are the blood component most frequently implicated in transfusion-associated bacterial sepsis, as they are stored at room temperature. Standard screening procedures reduced the rate of bacterial infections from PCs by 50% to 1 in ~50,000 units3. Globally, there are multiple PRTs in routine use for PCs: Intercept (Cerus Corporation, Concord, CA, USA), Mirasol (Terumo BCT, Lakewood, CO, USA), and Theraflex (Macopharma, Mouvaux, France)6,7. The clinical applications and detailed mechanisms associated with these PRTs has been reviewed extensively4,5,8. Briefly, the Intercept system uses UVA and Mirasol uses UVA+UVB for photoactivation of a sensitizing dye that binds nucleic acids and damages bound substrate, while Theraflex technology does not require sensitizing dye and utilizes UVC light. Theoretically, these methods inhibit pathogen replication while leaving anucleate cells (platelets and erythrocytes) intact6,7,9,10. The Intercept system is the only PRT currently cleared for use in PCs in the United States11. For PCs, each of these treatments results in detectable loss of in vitro platelet functions, and lower corrected count increments in patients11–15. The Cochrane review of Estcourt et al highlights the most important differences in transfusion outcomes when comparing control vs pathogen-inactivated platelets16.

Thiazole Orange (TO) is a photosensitizing dye with a flexible chemical structure with bonds capable of rotation on its axis as illustrated previously17 that targets nucleic acid with the dye in the rigid state only, which like the previously discussed PRTs, presumably inactivates pathogens without significant adverse effects on anucleate cells. The nucleic acid specific photochemicals used in Mirasol and Intercept systems have a rigid structure that can bind non-specifically to membrane lipids or proteins and cause damage to other components within bound cells, whereas the flexible structure of TO makes it less likely to activate during phototreatment when not bound to its substrate16. Treatment of RBC units with 80 μM TO and exposure to cool fluorescent light significantly inactivated select viral and bacterial pathogens without significant haemolysis or ATP loss during 42-day storage17,18.

In this report we extended TO treatment to platelet concentrates (PCs) to see whether it is compatible with platelet functions. However, we also recognised that unlike RBCs, platelets have functional mtDNA; therefore, TO effects on platelets functions could be different. To test this hypothesis, platelet functions were evaluated under treatment conditions sufficient for ≥4-log inactivation of Staphylococcus epidermidis and S. aureus, two common platelet contaminants implicated in many cases of transfusion associated bacterial sepsis. This study utilised apheresis PCs, as this is the predominant product used for platelet transfusions in the US. Based on the mechanisms outlined in this study, we would expect comparable results if experiments were conducted with WB derived platelets suspended in plasma.

MATERIALS AND METHODS

Collection and Treatment of PC

The purpose of this study was to evaluate TO (1-methyl-4-[(3-methyl-2(3H)-benzothiazolylidene] Methyl) quinolinium-p-tosylate, from Sigma Aldrich (Milwaukee, WI, USA) as a PRT for PCs, based on its inactivation of bacterial pathogens and its effects on platelet function during storage. Apheresis collection from consenting healthy donors was performed with a Fenwal Amicus instrument (Fenwal Inc., Lake Zurich, IL, USA) by the Research Blood Department at the Holland Laboratory of the American Red Cross. Apheresis platelets, which are predominantly used in the US were collected following the typical protocol in 200 mL concurrent plasma with a target yield of 3.6×1011 platelets, and donors were free from aspirin for 5 or more days. Purity of platelet apheresis products was confirmed using CBC-Differential measurements from a Sysmex haematology analyzer (Sysmex, Kobe, Japan). The human platelet work described here was approved by the ARC IRB and FDA-RISC #18-046B.

After resting for 24 hours, PCs in the bag were treated with 0–80 μM TO (dissolved in PBS on day of treatment) as follows: after addition of TO PCs were incubated for 15 minutes before phototreatment with 6.3±1.2 mW/cm2 cool fluorescent white light for 20 minutes. This process is referred to as “treatment” throughout the text. All incubations of PC bags were at room temperature with horizontal reciprocal agitation (Model LPR-3, Melco Engineering, Glendale, CA). Samples referred to as “untreated” had equivalent amount of PBS (vehicle) added. Bacterial inactivation, platelet metabolism, localisation of TO in platelets, markers of activation and apoptosis, aggregation, activation of intracellular signaling, and mtDNA damage were then examined at various timepoints following this treatment.

Bacterial Inactivation

Staphylococcus aureus (PEI-B-P-63-01) and S. epidermidis (PEI-B-P-06-01) were grown on Tryptic Soy Agar (TSA) plates from frozen stock, and multiple colonies were grown overnight in Tryptic Soy Broth to obtain overnight cultures. To establish dosing of TO that is sufficient to significantly inactivate the bacteria, PC was aliquoted into 6-well plates (4 mL/well) and inoculated with 100 μL overnight culture of S. aureus or S. epidermidis, 0–80 μM TO was added to each well and samples were subsequently phototreated as described. Samples for enumerating bacteria were taken by removing 1 mL from each well immediately following addition of bacteria (0 min), after incubation with TO (15 min), and after 20-minute phototreatment (post-inactivation). These samples were serially diluted in PBS and a 0.5 mL sample was added to top agar, which was poured over TSA. Colony forming units (CFU) were enumerated after overnight incubation. Additional experiments using 40 mL platelet aliquots contaminated with S. aureus in cryogenic storage bags were performed to confirm 6 well plate results.

Confocal microscopy

To identify localisation of TO within platelets, PC were incubated for 10 minutes at room temperature with MitoTracker Deep Red FM (ThermoFisher, Waltham, MA, USA), according to manufacturer instructions. TO was then added to PC (20 μM) and incubated for an additional 10 minutes. Platelets were fixed with 4% PFA and cytospun onto glass slides. Samples were mounted with Gold Antifade and images were acquired via laser scanning confocal microscopy on a Zeiss LSM (White Plains, NY, USA).

Platelet Apoptosis and Activation

For assessment of platelet function and activation, 40 mL of PC were aliquoted into Cryogenic storage bags (180 mL capacity, Charter Medical, Salem, NC, USA) and treated with 20 μM TO as described previously. Briefly, bags were incubated for 15 minutes after addition of TO, then treated with cool fluorescent white light for 20 minutes. Untreated samples had an equivalent amount of vehicle (PBS) added. For some experiments the mitochondrial reactive oxygen species (ROS) scavenger, MitoTempo (Sigma Aldrich), was also added to PC at 10 μM, 10 minutes prior to addition of TO.

Populations of activated and apoptotic platelets were measured using APC anti-human CD62P antibody conjugate (BD Biosciences, San Jose, CA, USA) or APC Annexin V (ThermoFisher), respectively. Following treatment with TO, samples were taken from bags at indicated timepoints and labeled with either APC-CD62P or APC-Annexin V, according to manufacturer instructions and using the manufacturer’s binding buffer. Acquisition was performed on a BD FACSCalibur (BD Biosciences) and FlowJo 10 software was used for analysis.

Platelet Function

Platelet metabolism (glucose and lactose concentrations) was measured using a Cobas b 221 blood gas analyzer (Roche, Indianapolis, IN, USA). Platelet aggregation in response to 10 μg/mL Collagen was evaluated via light transmission aggregometry (LTA) assay using a Chrono-Log 700 (Chrono-log Corp., Havertown, PA, USA).

Western Blot

The effects of TO treatment on intracellular signaling in platelets was evaluated by collection of 1 mL samples from bags before and after phototreatment. Samples were pelleted by centrifugation at 800 g for 10 minutes and lysed using RIPA buffer (Abcam, Cambridge, MA, USA) with protease inhibitor cocktail (Roche). Lysates were subjected to SDS-PAGE in Novex 12% Tris-Glycine mini gels (ThermoFisher) and transferred on to nitrocellulose membranes (iBlot, ThermoFisher). Membranes were probed using either Phospho-PKC Substrate Motif Rabbit mAb #6967 or Phospho-p38 MAPK antibody #9211 (Cell Signaling Technology, Danvers, MA, USA) with HRP-Cyclophilin B conjugate antibody (Abcam) used as loading control. Enhanced chemiluminescence (ThermoFisher) was used to develop membranes, which were then imaged on a CareStream Image Station 4000 MM Pro (Molecular Bioimaging, Bend, OR, USA).

Assay for mitochondrial DNA damage

Mitochondrial DNA (mtDNA) was isolated from 400 μL PC using the Qiagen DNA Blood Mini Kit (Qiagen, Germantown, MD, USA), according to manufacturer instructions for isolation of mtDNA from platelets. The mtDNA was quantified via qPCR, with SYBR Green PCR Master Mix (Applied Biosystems, Waltham, MA, USA) according to manufacturer instructions, of a 143 bp fragment using previously published primers (TRNK, ATP8 region, Fwd: 5′-CCA CTG TAA AGC TAA CTT AGC ATT AAC C-3′ and Rev: 5′-GTG ATG AGG AAT AGT GTA AGG AGT ATG G-3′)18. Reactions and analysis were performed by QuantStudio6 Flex Real-Time PCR System and QuantStudio Real-Time PCR Software, respectively (Applied Biosystems). Quantification of samples from TO treatment was relative to a commercial mtDNA standard from human HL-60 DNA (standard reference material 2392-1, National Institute of Standards & Technology, Gaithersburg, MD, USA), as described by Kim and colleagues19.

Damage to mtDNA was evaluated using a long-range PCR inhibition assay. Equal amounts of template were added to reactions that were amplified with KAPA HiFi HotStart ReadyMix (Roche, Wilmington, MA, USA), according to manufacturer instructions, using previously published primers for a 8.9 kb fragment of mtDNA (Accession #J01415 region, Fwd: 5′-TCT AAG CCT CCT TAT TCG AGC CGA-3′ and Rev: 5′-TTT CAT CAT GCG GAG ATG TTG GAT GG-3′)19. Reactions were performed under conditions described by Sanders and colleagues on a LabNet Multigene Thermal Cycler (LabNet International, Cary, NC, USA)20. Long-range PCR products were analysed using agarose gel electrophoresis in a 0.75% agarose gel.

Statistical Analysis

Graphing and statistical analysis were performed using GraphPad Prism software (GraphPad, San Diego, CA, USA). Paired t-test was applied and p-values of <0.05 were considered significant. Please note that for most of the assays described in this report, 3–4 individual platelet donations were analysed (n=3 or 4). However, for platelet aggregation and apoptosis assays, since donor to donor variation was observed, 7 individual donations were analysed (n=7).

RESULTS

TO inactivates staphylococci in PC

The bactericidal activity of a range of concentrations of TO was evaluated by treatment of PC spiked with S. aureus or S. epidermidis. The reason for selecting Staphylococci is these bugs are a frequent contaminant of platelets and S. aureus is often involved in septic transfusion reactions. All units were subjected to the same treatment, including 0 μM concentration, to which only vehicle (PBS) was added. Concentrations at or above 10 μM were sufficient to inactivate bacteria (Figure 1A and 1B). Comparison of CFU before and after light treatment (0 min vs post-inactivation) showed that 20 μM and above consistently reduced staphylococci by ≥4-log CFU (Figure 1C), and concentrations at or below 5 μM did not show appreciable bactericidal activity (Figure 1D). Units with 0 μM TO have no bacterial inactivation, showing that light treatment alone did not inhibit bacterial growth. Similar results were obtained with experiments of 40 mL aliquots in cryogenic storage containers (data not shown). The lowest effective concentration (20 μM) was used for evaluation of how TO affects platelet physiology and function.

Figure 1. Thiazole orange (TO) photoinactivates staphylococcal species in PCs.

Figure 1

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CFUs of (A) S. aureus or (B) S. epidermidis were enumerated via pour plating of serially diluted 0.5 mL samples taken before treatment (0 min) after addition of TO (15 min), and after phototreatment (post-inactivation). Graphs are from one representative donor (n=4), with samples plated in quadruplicate. (C) Log reduction of staphylococcal species was dose-dependent, and 20 μM was sufficient to consistently reduce bacteria by ≥4-log CFU. (D) Treatment with less than 10 μM inactivated bacteria by ≤1-log CFU. Data shown is mean +SD.

TO localises to mitochondria and increases platelet metabolism

LCSM images show colocalisation of TO with labeled mitochondria; however, this colocalisation is not selective, and the fluorescence signal from TO elsewhere in the platelets revealed that the dye is also binding other structures in the cell (Figure 2A). Taken together, this suggested that once activated by phototreatment, TO would adversely affect platelets alongside its inactivation of pathogens. Blood gas measurements of glucose and lactose concentrations in PC following treatment show that, by 24 hours post-treatment, there was a ~40% reduction in glucose and a ~170% increase in lactose, indicating that TO was rapidly accelerating platelet metabolism relative to untreated platelets (Figure 2B and 2C).

Figure 2. TO localises to mitochondria and alters platelet metabolism.

Figure 2

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Platelets were incubated with 20 μM TO followed by photoexcitation and MitoTracker Deep Red to label mitochondria. (A) Confocal microscopy shows colocalisation of signal from TO and MitoTracker, though the colocalisation is not selective (n=3). Blood gas measurements were taken at indicated timepoints to measure glucose (B) and lactose (C) concentration. Data shown is mean+SD for n=4 donors.

Platelet activation and apoptosis induced by TO treatment

Activation and apoptosis in TO treated platelets was measured via flow cytometry using APC conjugated reagents to minimize crosstalk in fluorescence signal from TO, which has excitation and emission wavelengths (λex=480 nm, λem=530 nm) similar to fluorescein. Because additional fluorescent markers could not be used in a single assay, the purity of PC units was confirmed prior to staining and platelet population was selected based on FSC vs SSC dot plots (data not shown). TO treatment caused a significant increase in surface CD62P expression by 1-hour post-treatment (Figure 3A and 3B). To elucidate the role of ROS and mitochondrial damage in the effects of TO treatment, PC were also pre-treated with the antioxidant MitoTempo, a nitroxide conjugated with a cationic moiety that targets mitochondria based on membrane potential21. TO did not significantly increase CD62P expression in platelets that were pre-treated with MitoTempo (Figure 3A and 3B).

Figure 3. Flow cytometry shows that TO treatment increases a population of activated and apoptotic platelets.

Figure 3

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At indicated times following treatment, platelets were labeled with an APC-conjugated antibody for CD62P (A and B) or Annexin V-APC (C and D). (A) Representative histogram overlays for CD62P expression. (B) Graph shows mean (with SD, n=3 donors) %CD62P+ cells. (C) Representative histogram overlays for Annexin V labeling. (D) Graph shows mean (with SD, n=3 donors) %Annexin V+ cells.

Similarly, apoptosis, measured by binding of Annexin V to platelets, showed that TO significantly increased the population of apoptotic platelets by 1-hour post-treatment, and this was also mitigated by treatment with MitoTempo (Figure 3C and 3D). The abrogation of these effects by MitoTempo indicates that TO-induced activation and apoptosis at these early timepoints are driven by ROS in the mitochondria.

LTA assay of TO-treated platelets

Aggregation of TO-treated platelets in response to collagen, an agonist chosen for its physiological relevance22, was assayed via Light Transmission Aggregometry (LTA). For all PC units tested (n=7), the percent maximum aggregation of TO-treated platelets declined over time, and by 24 hours post-treatment maximum aggregation was <5% (Figure 4A). MitoTempo partially mitigated the effects of TO treatment at early timepoints (1- and 4-hours post-treatment), but by 24 hours post-treatment these platelets also reached <5% max aggregation (Figure 4B). A similar trend for PS exposure and CD62P expression was observed in MitoTempo treated platelets by 24 hours as well (data not shown). Diminishing protective effects conferred by MitoTempo beyond 4 hours following TO-treatment suggests that ROS production and mitochondrial damage may accumulate over time and MitoTempo is eventually consumed by this process.

Figure 4. TO significantly decreases aggregation.

Figure 4

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(A) Platelets were evaluated in a light transmission aggregation (LTA) assay using 10 μg/mL collagen as an agonist. (B) Platelets were treated with 10 μM MitoTempo ± 20 μM TO and evaluated in LTA assay alongside samples in (A). Treatment with MitoTempo delayed the adverse effects of TO treatment on aggregation, indicating that these effects are primarily driven by mitochondrial ROS. For (A) and (B): representative trace at 4 h, with data in graph from duplicate measurements of one representative donor (n=7, **p<0.01, ***p<0.001 by t-test). Data shown is mean +SD.

TO treatment activates PKC and p38 MAPK

Intracellular signaling generated during TO treatment was evaluated by western blot, comparing samples before and after phototreatment. PKC and p38 MAPK were of interest because of their central roles in platelet integrin exposure, aggregation and apoptosis22–26. To evaluate activation of PKC, an antibody for phosphorylation of the motif on PKC substrates was used. This approach shows bands for all substrates that have been phosphorylated by PKC (Figure 5A). Before phototreatment, there was no difference in activation of PKC or p38 MAPK in the presence or absence of either TO or MitoTempo in platelet lysates (Figure 5A and 5B). However, immediately after phototreatment, both PKC substrates and p38 MAPK were activated (Figure 5A and 5B).

Figure 5. TO treatment activates PKC isoforms and p38 MAPK.

Figure 5

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Western blot of platelet lysates collected pre- and post-phototreatment (immediately following light exposure) probed with (A) anti-PKC substrate motif, which binds to any protein with a phosphorylated motif unique to PKC isoforms, or (B) anti-phospho-p38 MAPK antibodies. Blots shown are representative of results from three different donors.

Treatment with MitoTempo partially decreased activation; the remaining signal generated could, again, be due to the scavenging activity of MitoTempo being overwhelmed by the amount of ROS generated. This would also account for the inability of MitoTempo to rescue the increased metabolic activity of treated platelets, even shortly after treatment with TO (Figure 2B and 2C, 1 and 4 hours). Phototreatment with TO is sufficient to activate PKC and p38 MAPK signaling pathways within platelets, without external agonist, and the consequences of this signaling can be observed downstream as a high proportion of TO-treated platelets have surface CD62P expression and PS exposure.

mtDNA damage caused by TO treatment

The observed effects of TO on platelet function, its localisation to mitochondria and its affinity for binding nucleic acids indicated that mtDNA in platelets was likely damaged by TO treatment. Therefore, a PCR inhibition assay was used to assess damage to mtDNA19,20. Mitochondrial DNA was isolated immediately before and after phototreatment of PCs and quantified via qPCR using primers that targeted to a short-conserved region of the mitochondrial genome. The Threshold cycle (Ct) value of mtDNA from each sample across three different donors was relatively consistent and allowed normalisation of template quantity used for long-range PCR (Figure 6A).

Figure 6. Inhibition assay for mtDNA damage.

Figure 6

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Mitochondrial DNA was isolated from platelets before and after light exposure in the presence or absence of TO and/or MitoTempo. (A) Threshold cycle values for amplification of short fragments via qPCR of platelet mtDNA. (B) Agarose gel of PCR products from long-range PCR reaction of mtDNA shows inhibition of PCR for TO-treated samples in presence/absence of Mito-Tempo (Lanes 6 and 8). Data shown is mean+SD representative of results from three different donors.

Primers for the long-range PCR reaction amplified a large region (8.9 kb) of the mitochondrial genome so that damage to the genome would be likely to inhibit PCR, resulting in little or no PCR product. When these samples were subjected to agarose gel electrophoresis, there was no visible band for mtDNA from TO-treated platelets, including those treated with MitoTempo (Figure 6B). Inhibition of PCR in this assay confirms that TO treatment damages platelet mtDNA, and MitoTempo was unable to completely protect mtDNA from TO-induced damage. However, a PCR inhibition assay that utilizes a smaller region of the mitochondrial genome, may allow for more sensitive, nuanced detection of damage to mtDNA.

DISCUSSION

Currently, all PRTs for blood transfusion products target nucleic acids, with the goal of broadly neutralizing pathogen replication while leaving anucleate erythrocytes and platelets unaffected. These technologies were developed at a time when mtDNA was not considered to be essential to platelet function27. Subsequently, multiple studies have shown that PRTs decrease the quality and efficacy of transfused platelets1,28–30. And, a growing body of evidence has demonstrated the central role of mitochondria in key platelet functions, including aggregation and apoptosis31–36. Here, TO was evaluated as a potential PRT for PCs because it was shown to be effective in microbial reduction for RBCs, without adverse effects on function17. TO treatment effectively inactivated S. aureus and S. epidermidis, the common causes of TTI, in spiked PCs1–3. The concentrations used in the RBC study and the present PC study differ, 80 μM and 20 μM, respectively17. It is not unusual that photochemicals require different concentrations in RBCs and PCs to produce adequate pathogen inactivation, as units containing RBCs have higher opacity. Red cell membranes potentially bind a pathogen reduction agent more or less than platelets, requiring different concentrations of an agent to be available to bind and inactivate pathogens with minimum component damage. The amount of TO required to inactivate the pathogens evaluated here in PC units caused enough damage to platelets to render this PRT untenable for use in units intended for transfusion.

When activated by cool fluorescent white light, TO produces ROS capable of diffusion to other parts of the cell36. This contrasts with psoralens used in other PRTs which undergo electron transfer and adduct formation at the binding site, thereby producing far less ROS37. Based on these previous reports, in PCs, our analysis suggests that this photochemical acts via binding to and producing reactive oxygen species from nucleic acids of not only microbes, but also to the platelet mitochondrial DNA and perhaps to RNA present in platelets.

The fluorescence of TO allowed visualisation of its localisation within platelets that were also labeled with MitoTracker Deep Red. LCSM showed significant TO localisation to mitochondria. This study also demonstrated that TO activated key signaling pathways (PKC and p38 MAPK) and had adverse effects on platelet function. Specifically, TO caused significant increases in CD62P expression and phosphatidylserine exposure, detected by 1 hour after treatment, and abrogation of aggregation to relevant agonists by 24 hours post-treatment. Additionally, platelet mtDNA was found to be damaged as detected by a PCR inhibition assay, indicating ROS production within platelet mitochondria. The varying degrees of mitigation of these effects by treatment with a mitochondrial antioxidant and ROS scavenger, MitoTempo, further indicates the involvement of mitochondrial localisation and damage by TO. Although platelets are anucleate, they contain active mitochondria that serve multiple functions. Not only do mitochondria make significant contributions to the energy demands of platelets38, they are also required for activation via formation of the mitochondrial permeability transition pore, collapse of membrane polarisation, and increased ROS production33–35,39–41. Mitochondria also mediate platelet apoptosis via membrane depolarisation and generation of ROS36,41. Therefore, photodynamic agents that bind nucleic acids could adversely affect platelets by binding and damaging mtDNA, which causes mitochondria to generate ROS in response42. Given that mitochondrial ROS production plays a central role in activation of signaling that leads to platelet activation and apoptosis, mtDNA damage can have profound consequences for platelet function31.

Such consequences were observed in vitro in the present study, as TO-treated platelets no longer aggregated in response to collagen. Other PRT currently in use for PCs have similar effects on platelets, though they are less pronounced. Treatment with Mirasol, which utilizes riboflavin and UVB light, triggers ROS production and activates signaling by p38 MAPK, leading to increased activation and apoptosis23,25,36,43. Mirasol has also been demonstrated to damage platelet mtDNA43,44. Thus, while Mirasol provides protection from TTI, it has been shown to have undesired effects on platelet efficacy24. Similar effects on platelet function and efficacy have been observed for Intercept treatment, which uses Amotosalen, a psoralen, and UVA illumination45,46.

CONCLUSIONS

PRT is beneficial for improving the safety of blood transfusion products. The current strategy for inactivating pathogens relies on targeting nucleic acids for destruction; however, platelets appear to provide a unique challenge for preserving efficacy of transfused cells because of unavoidable photochemical damage. Part of this challenge stems from the central role of mitochondria in platelet metabolism, activation and apoptosis31–36. Mitochondria contain DNA that is critical to their function; and, once mtDNA is damaged, mitochondria produce ROS42. Thus, ROS produced by photosensitisation can have significant effects on platelet physiology. The effects of TO and the mitigation of these effects by MitoTempo demonstrated the role of mitochondrial ROS production in platelet activation and apoptosis in response to this PRT. In terms of platelet damage, TO effects are close to that of previously published studies with Mirasol and Intercept systems; however, the degree of damage is more severe with TO14,15,27,45,46. The currently available PRTs utilize additive solutions for treatment of platelet units47; the effects of TO on bacterial inactivation and platelet damage when suspended in additive solution are unknown but we would suspect similar kinds of damage to platelets but not perhaps similar extents of damage. On the other hand, it is also possible to speculate that due to the severity of damage induced by TO treatment that we observed, an additive solution may not be able to protect TO-treated platelets. Increasing the safety of PCs for transfusion, while preserving the function of platelets, may require additional or different approaches that minimize or avoid the consequences of ROS in platelets. Overall, this study illustrates that protecting platelet mitochondria and their functions from oxidative damage should be a fundamental consideration in selecting any new pathogen inactivation method for stored platelets or to improve the quality of platelets with existing PRTs.

ACKNOWLEDGMENTS

We appreciate the assistance of the Blood Research Group and Lenora Abel and Cheryl Anne Hapip of the American Red Cross Holland Laboratory for their help with acquiring platelet concentrate units for this study.

Footnotes

FUNDING AND RESOURCES

This work was supported by FDA intramural research funds. PG is a recipient of Oak Ridge Institute for Science and Education postdoctoral trainee fellowship.

AUTHORSHIP CONTRIBUTIONS

PG performed experiments, analysed data and wrote the manuscript. SDP assisted with design of microscopy experiments and performed confocal microscopy and analysis of microscopy images. PG, TG, SW, and CA designed experiments, interpreted data and edited the manuscript.

The Authors declare no conflicts of interest.

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