Abstract
Following biomaterial implantation, modulation of the acute immune response is essential for tissue regeneration. Polymorphonuclear leukocytes (PMNs) are critical effector cells in wound healing, and PMN dysfunction is mediated by mitochondrial dysfunction and can lead to prolonged inflammation and tissue damage. It was hypothesized that mitochondrial transplantation could be applied to PMNs in pro-inflammatory states as a means of upregulating regenerative proteins. Primary human PMNs were isolated from donor blood. Isolated PMNs and exogenous mitochondria were co-incubated to induce mitochondrial transplantation. Resulting interactions were assessed through microscopy to confirm initial uptake and mitochondria membrane potential retention, intracellular reactive oxygen species (ROS) analyses (n = 5), and PMN secretome quantification (n = 10) using multiplex protein analysis. Human PMNs were able to successfully uptake delivered mitochondria, and regenerative factors essential for tissue repair and immune cell recruitment including fibroblast growth factor-2 (FGF-2), interleukin (IL)-22, monocyte chemoattractant protein-1 (MCP-1), and granulocyte colony-stimulating factor (G-CSF) were significantly upregulated, indicating that exogenous mitochondria represent promising modulators of PMN function with broad clinical potential.
Keywords
Introduction
Following the implantation of biomaterials and tissue engineered constructs, the adequate facilitation of the acute immune response is critical to functional tissue regeneration and implant success 1 . Specifically, upon implantation of any biomaterial, the surgically induced tissue damage elicits an entire cascade of molecular events that influence the behavior of interfacing cells. The release of damage-associated molecular patterns (DAMPs) prompts the recruitment of immune cells, initiating the acute inflammatory response 2 . One of the first immune cells to interact with any implant is the polymorphonuclear leukocyte (PMN) 3 . PMNs are activated within seconds of making an incision, initiating the wound-healing cascade, and “priming” the cellular microenvironment for subsequent responses through the release of inflammatory mediators, consisting of various essential pro-inflammatory and regenerative proteins and signaling molecules 4 . PMNs have classically been considered as short-lived, stimulators of inflammation; however, this model neglects to acknowledge their impact as critical effector cells in the wound-healing continuum, as well as their complete range of functions and phenotypic presentations 4 . Particularly, PMNs are highly plastic and possess a wide-ranging span of essential, regenerative functions such as the attacking of extracellular pathogens through the release of lytic enzymes, stimulating matrix reprogramming and angiogenesis through the release of matrix metalloproteinases, and modulating the behavior of other immune cell populations such as monocytes and macrophages toward anti-inflammatory and regenerative phenotypes4,5. During wound healing, PMNs and macrophages engage in intercellular crosstalk, which can form positive feedback loops, amplifying inflammatory responses4,6,7. In addition, under the influence of inflammatory mediator dysregulation, this feedback loop can lead to impairment of the wound-healing cascade, ultimately generating tissue damage, prolonged inflammation, or tissue fibrosis4,6–8. Previous biomaterial implantation strategies have often resulted in these adverse reactions due to a failure to acknowledge and target the initial interactions between implants and effected immune cells. Therefore, recent research efforts in the field of biomaterials have been focused on the characterization and modulation of immune cells to allow for implant integration and functional tissue regeneration.
Mitochondria play significant roles in the regulation of cellular metabolism and homeostasis; however, historically, the role of the mitochondria in PMN function was often overlooked due to glycolysis being a major metabolic pathway for PMN energy production 9 . Recent studies have shown that although PMNs are primarily glycolytic, their mitochondria are widely involved in PMN function and disease pathogenesis, significantly contributing to their innate plasticity 9 . Specifically, PMN mitochondrial dysfunction can be associated with PMN activation, neutrophil extracellular trap (NET) formation, respiratory burst, differentiation/development, degranulation, and impaired chemotaxis 9 . In addition, PMN mitochondrial dysfunction has been correlated with several disease states characterized by immune dysregulation, including lupus, cancer, rheumatoid arthritis, and type 2 diabetes9,10.
Previously, cell-free mitochondria were associated with the release of pro-inflammatory signals in immune cells 11 . Mitochondrial molecules including mitochondrial DNA (mtDNA), mitochondrial RNA, N-formyl peptides, cardiolipin, and transcription factor A mitochondrial are able to bind to immune cell pattern-recognition receptors, eliciting a response comparable to that of other DAMPs 12 . However, a process called mitochondrial transfer has been found to induce modulatory effects on immune cells, situationally upregulating pro-inflammatory mediators and anti-inflammatory mediators 13 . In PMNs specifically, mitochondrial transfer studies have exhibited a range of potential benefits such as inhibited NET formation and increased respiratory capacity14,15.
Several pathways of mitochondrial transfer have been identified, including tunnel nanotubules, mitochondrial extrusion, cell fusion, gap junction channels, and extracellular vesicles (EVs) 16 . In vivo, mitochondria EVs provide protection for transferred mitochondria from the extracellular environment. Moreover, previous studies have identified that mitochondria transferred by EVs exhibit immunomodulatory potential. Specifically, EV transferred mitochondria have demonstrated regulatory functions in neutrophils as well as various other immune cell populations such as macrophages and T cells17–20. Although promising, EV-mediated mitochondrial transfer is currently limited by difficulties isolating and purifying EV populations, as well as understanding the mechanisms behind the encapsulation of intact mitochondria into EVs as opposed to mitochondrial fragments or proteins21,22.
Because mitochondrial dysfunction leads to a disruption of cellular homeostasis, several studies have attempted to address this issue through the process of mitochondrial transplantation, an artificially induced alternative to mitochondrial transfer 23 . Specifically, in mitochondrial transplantation, mitochondria are isolated from a healthy cell source. The exogenous mitochondria are then delivered to target cells with the intention of being incorporated into a recipient cell’s endogenous mitochondrial network, initiating metabolic reprogramming 24 . Multiple in vitro and in vivo studies have delivered these isolated, exogenous mitochondria to recipient cells via direct injection, intravenous injection/infusion, intranasal instillation, and co-culture, targeting several tissue and cell types including but not limited to myocardial cells, human oocytes, foam cells, pediatric heart tissue, intracerebral tissue, and alveolar epithelia25–30. Furthermore, this therapeutic delivery of mitochondria has shown significant potential in the resolution of dysregulated cellular metabolism and the mitigation of various disease states 24 . However, across these studies, the precise mechanistic behavior driving these outcomes has yet to be fully elucidated. For example, many of the published findings on mitochondrial transplantation have exhibited a disproportionate amount of uptaken mitochondria in respect to the administered dosages and the observed impacts, suggesting additional yet-to-be-realized interactions within the tissue microenvironment23,31. Particularly, the effects of this promising therapy on multiple critical interfacing cell types and signaling cascades essential to tissue homeostasis still need to be determined.
PMNs are the first innate immune cells to interact with any agent introduced to the body, shaping the host response and immune homeostasis4,32. Therefore, the interplay between exogenous mitochondria and PMNs has potentially significant effects on the success or failure of the mitochondrial transplantation process for any target application. Previous studies have begun to investigate these interactions. Specifically, the interactions between neutrophils and mitochondria-containing EVs have been explored, exhibiting potential immunomodulatory and uptake capabilities15,33,34. However, EVs have also been noted to have immunomodulatory capabilities due to their lipid composition 35 . Therefore, in these studies, it is difficult to distinguish the impacts of the delivered mitochondria from the EVs. In addition, EVs have the ability to encapsulate mitochondrial fragments and components, which are known to stimulate pro-inflammatory responses in PMNs and other innate immune cell populations11,36. Studies have also analyzed the impacts of naked, cell-free mitochondria on neutrophil function; however, these have been limited to exogenous mitochondria released by activated platelets17,37.
It has been observed that all mitochondria possess the core capacity for oxidative phosphorylation; however, they also possess numerous tissue-specific morphological and ultrastructural features that ultimately determine their functional capacity 38 . Specifically, mitochondria possess substantial genetic and non-genetic heterogeneity between and within different cell types, tissue types, and microenvironmental conditions38–40. Therefore, different mitochondrial sources have the potential to uniquely impact individual cell types. In addition, the previously mentioned studies have not holistically investigated the impacts of mitochondrial transplantation on PMN function, primarily attributing cell-free mitochondria to PMN pro-inflammatory inducing responses. Particularly, there are few to no studies analyzing the potential impacts of intact viable mitochondria on PMN cytokine production, surface receptor expression, or specific metabolic pathways 37 .
Due to PMNs’ complex roles in wound healing, as well as the existing mechanistic unknowns of the mitochondrial transplantation process regarding this critical cell type, two questions were assessed in this work. First, can PMNs undergo mitochondrial transplantation from non-platelet sources? And second, what unexplored impacts does mitochondrial transplantation have on PMN inflammatory mediator production? An ideal immunomodulatory strategy for implantable constructs would promote regenerative mediators while maintaining tissue homeostasis by allowing for the critical maintenance of necessary levels of pro-inflammatory mediators. The hypothesis of this study is that PMNs will incorporate transplanted mitochondria into their endogenous mitochondrial networks, initiating an upregulation of regenerative and anti-inflammatory proteins. This study is the first to explicitly investigate the impacts of mitochondrial transplantation, from a non-activated platelet source, on PMNs. Results will allow for more effective applications of mitochondrial transplantation, as well as potentially identifying a new target of its perceived benefits.
Materials and methods
PMN isolation and cell seeding
PMNs were obtained from healthy adult donors of varying sex and race. Individuals with autoimmune, endocrine, cardiovascular, or inflammatory disorders, or who reported tobacco use, were excluded to minimize confounding effects on PMN function. Donors abstained from alcohol and non-steroidal anti-inflammatory drugs for 48 h and fasted for ≥12 h prior to blood collection41–43. Donor screening, blood collection, and data-management procedures were approved by the University of Memphis Institutional Review Board (IRB ID: PRO-FY2020-230); written informed consent was obtained from all donors.
PMNs were isolated from whole blood using a well-established, density-based separation protocol validated in previous literature41,44,45. Using this method, the purity of neutrophils specifically has been reported at >96%45–47. Peripheral blood was collected into EDTA vacutainers (BD, Franklin Lakes, NJ, USA; 366643), and autologous serum was collected in untreated serum tubes (BD, 366668). Following gravitational sedimentation to separate leukocyte and erythrocyte fractions, the leukocyte layer was aspirated and centrifuged at 200 × g for 10 min at room temperature (RT) using a Sorvall ST8 centrifuge (Thermo Scientific, Waltham, MA, USA; rotor 75005701). The pellet was resuspended in 1X phosphate buffered saline (PBS) and layered onto 3 mL of Isolymph (CTL, 1.077 ± 0.001 g/mL, 759050), followed by centrifugation at 300 × g for 40 min at RT with the brake disabled. The peripheral blood mononuclear cell layer and supernatant were removed, and residual erythrocytes were lysed by treatment with 0.2% NaCl for 30 s at 4°C. Isotonicity was restored with 1.6% NaCl prepared using American Chemical Society (ACS)-grade sodium chloride (MP Biomedicals, Santa Ana, CA, USA; 194738) in sterile, endotoxin-free water (Cytiva, Marlborough, MA, USA; SH30529.02). Cells were centrifuged at 200 × g for 7 min at 4°C and washed in ice-cold 1X PBS. PMNs were resuspended in HBSS (Hank’s Balanced Salt Solution; Gibco, Waltham, MA, USA; 14175-095) supplemented with 0.2% autologous serum and 10 mM N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid (HEPES; Corning, Corning, NY, USA; 25-060-CI), hereafter referred to as HBSS+. Cell concentration and viability were assessed by 0.4% trypan blue (Gibco, 15250-061) exclusion using a Countess II FL automated cell counter.
PMNs (1 × 106 cells/mL) were seeded at 100 µL per well in a BioLite 96-well plate (Fisher). To standardize the final volume to 150 µL, negative controls received 40 µL of HBSS+, and positive controls received 30 µL of HBSS+ prior to cell seeding. To dissociate NET-associated myeloperoxidase (MPO), 10 µL of 150-U/mL Heparin (Sigma-Aldrich, H3393) was added to each well 48 . Negative control wells were left untreated, and positive controls were stimulated with 10 µL of 1.5-µM phorbol 12-myristate 13-acetate (PMA; Sigma-Aldrich, P8139).
Mitochondria isolation
Mitochondria were isolated from adult normal human dermal fibroblasts (HDF; American Type Culture Collection, Manassas, VA, USA; PCS-201-012, Manassas, Virginia, US) using a well-established protocol, validated in previous literature49,50. Using this method, the resulting mitochondrial purity has been reported as <0.01% being fractured or damaged, with non-mitochondrial particle contamination being <0.001% 50 . Initially, four T-150 flasks of HDFs were grown to ~80%–85% confluency, resulting in cell counts ranging from ~3.43×106 to 5.93×106 cells/ml (n = 4, T-150 HDF flasks), in Dulbecco’s Modified Eagle Medium (Gibco, 11965-092) supplemented with 10% heat-inactivated fetal bovine serum (Gibco, A56708-01) and 10% penicillin streptomycin (Corning, 30-002-Cl), hereafter referred to as complete medium. The complete medium was decanted from culture flasks followed by a 1X PBS wash. Each flask of cells was then homogenized by incubating cells at RT with 5 mL of homogenization buffer (300 mmol/L sucrose, 1 mmol/L EGTA (Ethylenediaminetetraacetic acid; Sigma-Aldrich, E4378-25G), 10 mmol/L HEPES (Corning, 25-060-CI), 2 mg protease from bacillus licheniformis (Subtilisin A, Sigma-Aldrich, 9014-01-1), in sterile, endotoxin-free cell culture–grade water (Cytiva, SH30529.02)). Homogenate was equally distributed between two pre-chilled 15-mL tubes and allowed to chill on ice for 15 min. The chilled homogenate was centrifuged at 500 × g for 5 min. The supernatant was filtered into a pre-chilled 50-mL tube through a 40-µm sterile nylon mesh cell strainer (Fisherbrand, 22363547) twice, using a new strainer each time. Supernatant was filtered a final time through a 10-µm ff mesh strainer (PluriSelect, El Cajon, CA, USA; 43-10010-50) and equally split between two new pre-chilled 15-mL tubes. Tubes were centrifuged at 800 × g for 5 min. Supernatants were poured into two new pre-chilled 15-mL tubes and centrifuged at 3500 × g for 10 min. Supernatant was aspirated, and the resulting pellets were reconstituted in 50-µL cold 1X PBS lacking calcium chloride and magnesium chloride, combined, and brought up to a final volume of 300 µL with additional cold 1X PBS.
Mitochondria ATP and protein quantification
Mitochondrial ATP content in each dose was assessed using CellTiter-Glo Luminescent Cell Viability Assay (Promega, Madison, WI, USA; G7570). Following the manufacturer protocol, 100 µL of mitochondria suspension and 100 µL of CellTiter-Glo Buffer were added to a white-walled 96-well plate (Thermo Scientific, 236105). Plate contents were mixed on an orbital shaker for 2 min, followed by a 10-min incubation at RT. Luminescence was recorded using a microplate reader, and resulting values (n = 3) were compared against ATP standards. Mitochondrial protein content in each dose was assessed using a Bicinchoninic (BCA) protein assay (Thermo Scientific, 23227). Following manufacturer protocol, 25 µL of mitochondria suspension and 200 µL of working reagent were added to a clear, flat-bottom 96-well plate (Thermo Scientific, 130188) and placed on an orbital shaker for 30 s. The plate was then incubated at RT for 30 min, and sample absorbance (n = 3) was read at 562 nm on a microplate reader.
Mitochondria uptake
Following mitochondria isolation, resuspended and combined mitochondria were centrifuged at 3500 × g to pellet. Supernatant was removed, and mitochondria pellet was resuspended in 250 µL of cold PBS. Isolated mitochondria were stained prior to delivery to distinguish between endogenous and exogenous PMN mitochondria. Reconstituted MitoTracker Orange CMTMRos (Invitrogen, Waltham, MA, USA; M7510) was added to PBS-resuspended mitochondria to achieve a working concentration of 190 µM. Exogenous mitochondria were incubated with stain for 30 min. Following incubation, stained mitochondria were centrifuged at 3500 × g to pellet. The supernatant was aspirated, and mitochondria were resuspended in cold PBS to achieve a final working volume of 300 µL.
PMNs (n = 5) were seeded in 96-well plates and activated using PMA as previously described. Negative controls included unstimulated PMNs without exogenous mitochondria, and positive controls included stimulated PMNs with exogenous mitochondria. PMNs were co-incubated with 10 µL of isolated mitochondria, resuspended as described in “Mitochondria isolation” section, for 1 h at 37°C following PMA activation. PMNs were then placed on ice for 10 min to inhibit activity. Following this, plates were centrifuged at 400 × g for 5 min, and supernatants were aspirated. PMNs were washed three times with 1X PBS, centrifuging plates at 400 × g for 5 min after each PBS addition to prevent unintentional disposal of cells. PBS was removed, and 10% formalin was added to wells at RT for 15 min to fix cells. Following fixation, wells were washed twice with 1X PBS, as previously described. PBS was removed, and PMNs were permeabilized in 0.1% Triton X-100/1X PBS for 15 min. Plates were then centrifuged at 400 × g for 5 min, Triton X-100 was removed, and wells were washed twice with 1X PBS, centrifuging as previously described. PMN nuclei and F-actin filaments were stained using a double fluorescence stain consisting of the NucBlue Fixed Cell Stain ReadyProbes reagent (Invitrogen, R37606) and Alexa Fluor 488 Phalloidin (Invitrogen, A12379), prepared according to manufacturer protocols. Cells were stained at RT for 1 h and protected from direct light exposure. Following the incubation period, PMNs were centrifuged at 400 × g for 5 min, staining solution was aspirated, and cells were washed twice with 1X PBS as previously described. Following the final PBS wash, each well was resuspended in 50 µL of 1X PBS.
Uptake imaging procedure
Cells were imaged at 20X magnification using a BioTek Cytation 1 imaging reader (Agilent, Santa Clara, CA, USA), and 60X magnification images were obtained using a Nikon A1 confocal scanning laser fluorescence microscope (Nikon, Tokyo, Japan). Confocal image acquisition was performed using a 1 AU pinhole and a laser intensity and PMT gain appropriate for the intensity of fluorescence staining. The same settings were used for all observations. Well plates containing 50 µL of resuspended PBS cell solution were used for 20X magnification images, and images were taken using the TRITC color filter at 15 min, 30 min, and 1 h. For 60X magnification images, cells were mounted by pipetting 30 µL from each resuspended well onto a glass slide (Fisher Scientific, cat. no. 1255007). Slides were left in the dark at RT until PBS evaporated, leaving only the stained cells. Once slides were dry, 10 µL of ProLong Gold antifade reagent (ref. no. P36930) was added on top of dried sample and covered with a glass cover slip (Fisher Scientific, cat. no. 2-545-87). The mountant was allowed to cure overnight at RT in the dark. Slides were imaged using the 4’,6-diamidino-2-phenylindole (DAPI), Fluorescein isothiocyanate (FITC), and Tetramethylrhodamine isothiocyanate (TRITC) color filters for NucBlue, Alexa Fluor 488 Phalloidin, and MitoTracker Orange CMTMRos, respectively.
Z-stack staining and imaging procedure
Cells were imaged at 60X magnification using a Nikon A1 confocal scanning laser fluorescence microscope (Nikon, Tokyo, Japan). Confocal image acquisition was performed using a 1 AU pinhole and a laser intensity and PMT gain appropriate for the intensity of fluorescent staining. The same settings were used for all observations. PMNs were isolated as previously described and stained for endogenous mitochondria using MitoTracker Green (Invitrogen, M7514) and for nuclei using NucBlue Live Cell Stain ReadyProbes reagent (Invitrogen, R37605), followed by centrifugation at 200 × g for 7 min at RT using a Sorvall ST8 centrifuge (Thermo Scientific, rotor 75005701). The supernatant was removed, and cells were washed by resuspension in 15 mL of cold PBS (pH 7.4) and centrifuged at 200 × g for 7 min at RT. The pellet was resuspended in HBSS+, and stained PMNs (1 × 106 cells/mL) were seeded in HBSS+ in glass bottom six-well plates (Matek, P06G-1.5-10-F). Cells were activated for 1 h using PMA to better simulate an in vivo wound-healing microenvironment that exogenous mitochondria would encounter. HDF mitochondria were isolated and stained with MitoTracker Orange CMTROS as previously described. PMNs and exogenous mitochondria were co-incubated for 15 min at 37°C. This was followed by the addition of the ProLong Live Antifade Reagent (Invitrogen, P36975) for an additional 15 min at 37°C. Representative volume snapshots and Z-stack images were then acquired.
Mitochondrial membrane potential retention
Membrane potential staining procedure
To determine if mitochondria remained metabolically intact following uptake, PMNs (n = 4) were seeded in HBSS+ in 96-well plates as previously described. Cells were activated for 1 h using PMA. Following PMA activation, exogenous mitochondria were stained with tetramethylrhodamine, methyl esther (TMRM) (Invitrogen, I34361, T668), an indicator of mitochondrial membrane potential, and 10 µl of stained mitochondria was co-incubated with stimulated PMNs. Controls were as previously described. Plates were incubated at 37°C.
Membrane Potential Imaging Procedure
Plates were imaged at 30 min and 3 h following addition of mitochondria. Images were taken at 20X magnification using a BioTek Cytation 1 imaging reader (Agilent, Santa Clara, CA, USA). Representative brightfield and fluorescence images using the TRITC filter were acquired.
DCFDA-based intracellular ROS assay
Intracellular reactive oxygen species (ROS) production was quantified using 2’,7’-dichlorofluorescein diacetate (DCFDA; Abcam, ab113851), a cell-permeable probe that is converted to fluorescent DCF upon oxidation. The assay was adapted from the manufacturer’s protocol and the method described by Nadesalingam et al. 51 . Isolated PMNs were resuspended at 1.5 × 106 cells/mL in HBSS+ containing 5-µM DCFDA. Cells were incubated for 30 min at 37°C in the dark, followed by centrifugation at 200 × g for 7 min at RT using a Sorvall ST8 centrifuge (Thermo Scientific, rotor 75005701). The supernatant was removed, and cells were washed by resuspension in 15 mL of cold PBS (pH 7.4) and centrifuged at 200 × g for 7 min at RT. The pellet was resuspended in HBSS+ and adjusted to 1 × 106 cells/mL. PMNs were seeded at 100,000 cells per well in appropriate culture plates. Isolated mitochondria were loaded into wells at volumes of 10 µl using dilutions of 1:3, 1:6, and 1:12 in cold 1X PBS. Controls were as previously described. Samples were incubated at 37°C with 5% CO2 for 1 and 3 h. Following incubation, fluorescence was measured using a microplate spectrophotometer with excitation at 485 nm and emission at 535 nm, in accordance with the manufacturer’s protocol. Data were normalized to the positive control for each donor to account for inter-donor variability in ROS responses and reported as the mean fold change of the positive control ± standard deviation for five independent experiments (n = 5).
Cytokine and protein analysis of PMN supernatants
PMNs were plated as previously described and incubated with the highest tested concentration of mitochondria (1:3) at 37°C with 5% CO2 for 1 and 3 h. Following incubation, PMN activity was halted by placing plates on ice for 10 min prior to supernatant collection. Supernatants (100 µL) were transferred to microcentrifuge tubes and centrifuged at 500 × g for 5 min at RT using a Sorvall Legend XTR centrifuge (Thermo Scientific, rotor 6133415). Clarified supernatants (50 µL) were transferred to new tubes and stored at −20°C until analysis 52 . PMN supernatants were analyzed using a multiplexed magnetic bead immunoassay (Thermo Fisher, Minneapolis, MA, USA) on a MAGPIX® reader (Luminex, Austin, TX, USA) according to the manufacturer’s instructions along with the Procartaplex (Thermo Scientific, MA, USA) Analysis App. Matrix metalloproteinase-9 (MMP-9) and MPO were assayed using a 1:50 dilution in HBSS+ (Catalog Number: PPX-02-MXT2CHW, Lot: 463553-000). All other analytes, including angiopoietin-1, fibroblast growth factor-2 (FGF-2), granulocyte colony-stimulating factor (G-CSF), hepatocyte growth factor (HGF), IL-1β, IL-10, Il-1RA, IL-22, IL-6, IL-8, monocyte chemoattractant protein-1 (MCP-1), tumor necrosis factor alpha (TNF-α), and vascular endothelial growth factor-A (VEGF-A), were assayed using a 1:2 dilution in HBSS+ (Catalog Number: PPX-13-MXRWGXY, Lot: 450895-000). Of the 13 analytes, IL-10 was below the detection limit of the assay. All analytes released at a sufficient level are reported as the mean fold change of the positive control ± standard deviation (n = 10).
Statistics
A priori power analyses were utilized to ensure that the selected sample sizes allowed for the study power to be at least 80% using a significance level of 0.05 for all PMN experiments. A minimum sample size of n = 5 healthy, adult donors was used for all quantitative studies, and all samples were collected in quadruplicate. To account for high analyte variability as found in prior literature, data were normalized to donor-specific positive controls using Microsoft Excel53–55. Statistical analysis was performed using GraphPad Prism. Specifically, normality was assessed for all data groups using Shapiro–Wilk tests. Multiple data groups across this study exhibited strong skewedness or outlying values. Therefore, non-parametric methods were used. Particularly, Kruskal–Wallis tests and Dunn’s multiple comparison tests were used to determine statistical significance in these studies.
Results
Mitochondria ATP and protein quantification
To confirm mitochondria isolation and desired dosage, ATP and mitochondrial protein content were quantified. Isolated HDF-derived mitochondria from four T-150 flasks (n = 3) at ~80%–85% confluency (~3.43 × 106–5.93 × 106 cells/ml) resulted in an average ATP concentration of 3.08 µM. The same number and confluency of T-150 flasks resulted in an average mitochondrial protein concentration of 560 µg/mL.
Mitochondria uptake
Fluorescence microscopy confirmed isolation and delivery of exogenous mitochondria into recipient PMNs. In initial 20X images, PMN nuclei appeared colocalized with delivered mitochondria after 15 min of co-incubation in both PMA-stimulated and PMA-unstimulated mitochondria-treated groups (Figure 1). Representative, z-stack images at 60X were then taken to further confirm colocalization of delivered mitochondria and endogenous PMN structures that would confirm successful transplantation. PMA-stimulated mitochondria-treated groups saw significant colocalization between delivered mitochondria and endogenous PMN mitochondria (Figure 2). Unstimulated mitochondria-treated groups saw significant colocalization between delivered mitochondria and PMN nuclei (Figure 3). Additional non-z-stack 60X images were obtained at 1 h of co-incubation to further assess changes in nuclear morphology (Figure 4). A lobed nuclear morphology was exhibited in mitochondria-treated groups and HBSS+ control groups for both stimulated and unstimulated PMNs.
Representative fluorescent microscopy images display PMA and non-PMA-activated human peripheral blood PMNs (20X) (n = 5) that were co-incubated with exogenous mitochondria from HDFs for 15 min. Phalloidin (Alexa Fluor 488) fluorescent stain indicates F-actin, DAPI (NucBlue) fluorescent stain indicates nuclei, and MitoTracker Orange CMTMRos fluorescent stain indicates delivered mitochondria. Scale bar = 100 µm.
Images display representative z-tack images of a non-PMA-activated human PMN (60X) co-incubated with delivered mitochondria from HDFs for 15 min prior to imaging preparation. Color channels indicate (a) merged, (b) DAPI (NucBlue) staining nuclei, (c) FITC (MitoTracker Green) staining endogenous PMN mitochondria, and (d) TRITC (MitoTracker Orange CMTMRos) staining delivered mitochondria. Scale bar = 15 µm.
Images display representative z-tack images of a PMA-activated human PMN (60X) co-incubated with delivered mitochondria from HDFs for 15 min prior to imaging preparation. Color channels indicate (a) merged, (b) DAPI (NucBlue) staining nuclei, (c) FITC (MitoTracker Green) staining endogenous PMN mitochondria, and (d) TRITC (MitoTracker Orange CMTMRos) staining delivered mitochondria. Scale bar = 25 µm.
Representative confocal images display PMA and non-PMA-activated human peripheral blood PMNs (60X) (n = 5) that were co-incubated with delivered mitochondria from HDFs for 1 h. Phalloidin (Alexa Fluor 488) fluorescent stain indicates F-actin, DAPI (NucBlue) fluorescent stain indicates nuclei, and MitoTracker Orange CMTMRos fluorescent stain indicates exogenous mitochondria. Scale bar = 5 µm for PMA-stimulated HBSS+ control and 10 µm for all other columns.
Membrane potential retention
The retention of delivered mitochondrial membrane potential following PMN uptake was assessed using TMRM, a fluorescent stain that only fluoresces when mitochondrial membrane potential is intact. Representative images of varying donors and well locations were used to assess general ability of transplanted mitochondria to retain membrane potential following uptake (Figure 5). TMRM-stained transplanted mitochondria produced fluorescent signals at all timepoints, 30 min, 90 min, and 180 min, for both PMA-stimulated and PMA-unstimulated PMN groups. Fluorescent signal decreased as time progressed, indicating a slight loss in exogenous mitochondrial membrane potential over time.
Representative images display PMA and non-PMA-activated human peripheral blood PMNs (20X) that were co-incubated with HDF-derived mitochondria stained with TMRM. Mitochondrial membrane potential following uptake was monitored at 30 min, 90 min, and 180 min. All selected images use PMNs from the same donor. Scale bar = 100 µm.
Intracellular ROS activity
The impacts of transplanted mitochondria on PMN intracellular ROS were used to initially determine whether mitochondria transplantation initiated or further facilitated the production of PMN pro-inflammatory mediators in unstimulated or stimulated PMNs, respectively. Mitochondrial transplantation caused no significant differences in either stimulated or unstimulated PMN intracellular ROS activity relative to the respective untreated controls (Figure 6). This was reflected at both the 1- and 3-h timepoints for all mitochondria doses. However, significant differences were observed between positive (100 nM PMA HBSS+ Ctrl) and negative control (unstimulated HBSS+ Ctrl) groups at both timepoints, validating experimental methods.
Graphs display intracellular ROS levels relative to positive controls per donor (n = 5) at (a) 1 and (b) 3 h. Asterisks indicate statistically significant differences between compared groups, and degrees of significance are indicated as **P < 0.01.
Cytokine and protein analysis of PMN supernatants
Baseline positive control (100 nM PMA HBSS+ Ctrl) values for each inflammatory mediator of interest, donor sample, and timepoint were used to normalize all cytokine and protein analysis data to account for sample variability (Tables 1–5). All cytokine and protein analysis graphs are reported in terms of normalized relative fold change. Non-normalized data for all donors and proteins of interest are included in Supplemental Data.
MPO production values for baseline, non-normalized, positive control (100 nM PMA HBSS+), negative control (unstimulated HBSS+), and normalized unstimulated control values per donor and timepoint.
Bolded font indicates excluded donors due to elevated baseline MPO levels. (lower limit of quantification [LLOQ] = 101 pg/ml) (upper limit of quantification [ULOQ] = 103,100 pg/ml).
Table contains baseline, non-normalized, positive control (100 nM PMA HBSS+) values per donor and timepoint for pro-inflammatory cytokines.
Bolded font indicates excluded donors due to elevated baseline MPO levels.
Table contains baseline, non-normalized, positive control (100-nM PMA HBSS+) values per donor and timepoint for regenerative growth factors.
Bolded font indicates excluded donors due to elevated baseline MPO levels.
Table contains baseline, non-normalized, positive control (100 nM PMA HBSS+) values per donor and timepoint for regenerative and anti-inflammatory cytokines.
Bolded font indicates excluded donors due to elevated baseline MPO levels.
Table contains baseline, non-normalized, positive control (100 nM PMA HBSS+) values per donor and timepoint for immune cell recruitment and production cytokines.
Bolded font indicates excluded donors due to elevated baseline MPO levels.
Significant differences in MPO production were observed between the PMA-stimulated HBSS+ control group and the unstimulated HBSS+ control group, confirming activation. In addition, MPO production in baseline unstimulated, negative controls were used as donor selection criteria. Donors with normalized, baseline negative control values higher than 70% (0.7) of the respective positive control, per timepoint, were removed from all subsequent cytokine analyses to limit confounding variables due to elevated baseline MPO levels (Table 1). Excluded donors are indicated in all data tables by bolded font. After removing donors with baseline MPO values that did not meet the selection criteria, a minimum sample size of n = 5 was maintained. Power analysis was repeated and confirmed that the resulting sample sizes retained a minimum of 80% power due to their large effect sizes. The lower limit of quantification (LLOQ) and upper limit of quantification (ULOQ) are indicated in each table or caption for each analyte. The analysis software provided proprietary low-end extrapolation, allowing for extension of standard curves past the LLOQ, limited by the limit of detection. However, due to the proprietary nature of the company-provided data extrapolation, the LLOQ and ULOQ were used as non-zero extremes for all out-of-range (OOR) data analyses. All obtained OOR protein values were assigned the ULOQ.
Significant differences in MMP-9 production were observed between positive and negative controls at both 1 (***P < 0.001) and 3 (*P < 0.05) h (Figure 7). However, no differences were observed in response to mitochondria treatment at either timepoint (Figure 7). Significant differences were also not observed in pro-inflammatory cytokines IL-1β or IL-6 at either timepoints (Figure 8) relative to positive controls. IL-8 exhibited significant differences (**P < 0.01) between positive and negative controls at 3 h (Figure 8). TNF-α data exhibits statistically significant differences between PMA-stimulated mitochondria-treated groups and PMA-stimulated HBSS+ control groups at both 1 (***P < 0.001) and 3 h (***P < 0.01) (Figure 8). Significant differences were observed in regenerative growth factor FGF-2 at 1 (***P < 0.001) and 3 h (***P < 0.01) between the PMA-stimulated mitochondria-treated group and the HBSS+ control group (Figure 9). Significant differences in FGF-2 production were also observed between unstimulated mitochondria-treated groups and HBSS+ control groups at 1 (**P < 0.01) and 3 h (*P < 0.05) (Figure 9). Significant differences in HGF production were observed between control groups at 1 (*P < 0.05) and 3 (**P < 0.01) h.
Graphs display fold change of MPO and MMP-9 relative to positive controls per donor (n = 10) (a) 1 h and (b) 3 h after the addition of exogenous mitochondria. Asterisks indicate statistically significant differences between compared groups, and degrees of significance are indicated as *P < 0.05, **P < 0.01, or ***P < 0.001.
Graphs display fold change of pro-inflammatory cytokines relative to positive controls per donor (n = 10) (a) 1 h and (b) 3 h after the addition of exogenous mitochondria. Asterisks indicate statistically significant differences between compared groups, and degrees of significance are indicated as ***P < 0.001.
Graphs display regenerative growth factor fold change relative to positive controls per donor (n = 10) (a) 1 h and (b) 3 h after the addition of exogenous mitochondria. Asterisks indicate statistically significant differences between compared groups, and degrees of significance are indicated as *P < 0.05, **P < 0.01, or ***P < 0.001.
Significant differences were not observed between data groups for regenerative cytokines angiopoietin-1 or IL-1RA (Figure 10). Significant differences were observed for VEGF-A between control groups at both 1 (***P < 0.001) and 3 h (***P < 0.001) (Figure 10). PMA-stimulated mitochondria-treated groups significantly increased in IL-22 production at 3 h (*P < 0.05) (Figure 10). Significant increases in cytokines associated with immune cell recruitment and activation, G-CSF and MCP-1, were observed. G-CSF had significant upregulation between the PMA-stimulated mitochondria-treated group in comparison to the HBSS+ control at 1 (****P < 0.0001) and 3 (**P < 0.01) h and unstimulated mitochondria-treated groups in comparison to the HBSS+ control at 1 (*P < 0.05) and 3 (*P < 0.05) h (Figure 11). MCP-1 had significant upregulation between the PMA-stimulated mitochondria-treated group in comparison to the HBSS+ control at 1 (***P < 0.001) and 3 (*P < 0.05) h and unstimulated mitochondria-treated groups in comparison to the HBSS+ control at 1 (**P < 0.01) and 3 (***P < 0.001) h (Figure 11).
Graphs display regenerative and anti-inflammatory cytokine fold change relative to positive controls per donor (n = 10) (a) 1 h and (b) 3 h after the addition of exogenous mitochondria. Asterisks indicate statistically significant differences between compared groups, and degrees of significance are indicated as ***P < 0.001.
Graphs display fold change of cytokines responsible for immune cell recruitment and production relative to positive controls per donor (n = 10) (a) 1 h and (b) 3 h after the addition of exogenous mitochondria. Asterisks indicate statistically significant differences between compared groups, and degrees of significance are indicated as *P < 0.05, **P < 0.01, ***P < 0.001, or ****P < 0.0001.
Discussion
This work investigated the impacts of mitochondrial transplantation on PMN behavior. While other in vitro and in vivo studies have examined the impacts of mitochondrial transplantation on various target cells, tissues, and diseases, few studies address the interactions of this promising therapy with PMNs, which are orchestrators of the inflammatory response, ultimately dictating the fate of the tissue regenerative process4,25–30. This work confirmed the ability of human PMNs to uptake metabolically intact exogenous mitochondria. It was also confirmed that mitochondrial transplantation has immunomodulatory impacts on human PMNs through the upregulation and maintenance of various critical inflammatory mediators.
Results from fluorescence images indicated that exogenous mitochondria were taken in by human peripheral blood PMNs as early as 15 min following co-incubation (Figures 1 to 4). Colocalization of transplanted mitochondria with unstimulated PMN nuclei and stimulated PMN endogenous mitochondria also indicated potential integration of transplanted mitochondria into the endogenous mitochondrial network of recipient PMNs, which is typically located near the nucleus topographically and functionally (Figures 2 and 3) 56 . The impact of this uptake on cell morphology was further analyzed through additional confocal microscopy. Specifically, following 1 h of incubation of PMNs with exogenous mitochondria, it was observed that cells retained a lobed nuclear morphology (Figure 4). Upon extracellular trap formation, PMN nuclei shift to a rounder shape, losing their lobed structure 57 . Therefore, the retention of this lobed structure served as an early indicator that exogenous mitochondria did not stimulate NETs.
This prompted further investigation into the interactions between delivered mitochondria and recipient PMNs. Particularly, the functional status of transplantation mitochondria was assessed following uptake through live-cell mitochondrial membrane potential analysis. TMRM staining indicated that although some depolarization was observed, several transplanted mitochondria retained their membrane potential until the maximum observed time point of 3 h following uptake (Figure 5). Mitochondrial membrane potential maintenance is critical to overall mitochondrial function, as well as the resulting impacts on PMN function9,58. Therefore, the retention of membrane potential can be associated with maximum capacity for mitochondrial impact on recipient PMNs.
The specific impacts of metabolically intact transplanted mitochondria on PMN function were assessed through the analysis of inflammatory mediator production. Results demonstrated that intracellular ROS production of mitochondria-treated groups displayed no significant differences from positive controls in response to all mitochondria doses including dilutions of 1:3, 1:6, and 1:12 (Figure 6). This indicated that transplanted mitochondria neither stimulated nor inhibited the production of intracellular ROS, although mitochondria are known to be key contributors to the production of cellular ROS. In addition, this opposes previous studies on other, non-PMN cell types, indicating that mitochondrial transplantation can reduce ROS activity and enhance antioxidant capacity59,60. However, the lack of significant increases in intracellular ROS production builds upon the assessment of recipient PMN nuclear morphology, further indicating that transplanted mitochondria did not upregulate a pro-inflammatory response.
Inflammatory mediator analysis was continued through the assessment of multiple cytokines and proteins of interest using a multiplex analysis (n = 10). Initially, two proteins critical to PMN function, MPO and MMP-9, were assessed. MPO is an enzyme that coats chromatin released during NETosis 61 . In addition, previous studies have indicated that MPO contributes to the occurrence of PMA-mediated NETosis 61 . Therefore, MPO production was used as a baseline indicator for both induced and innate PMN activation. At both 1 and 3 h, there were significant differences in MPO production between control groups, indicating proper activation of stimulated groups. No significant differences were observed between treatment groups in respect to positive controls, indicating that mitochondrial transplantation neither promotes nor downregulates NETosis. In addition, the lack of significant increases in MPO production further supports the lobed nuclear morphology observed in Figure 4. Human PMNs also have the ability to produce tissue inhibitors of metalloproteinases (TIMP)-free MMP-9, a potent stimulator of extracellular matrix remodeling and angiogenesis 62 . Differences in MMP-9 production were observed between control groups at 1 h; however, mitochondrial transplantation had no impact on MMP-9, preserving but not upregulating the production of this critical, PMN-specific, matrix remodeling enzyme.
Assessed, detectable, pro-inflammatory cytokines included IL-1β, IL-6, TNF-α, and IL-8 (Figure 8). Pro-inflammatory cytokines IL-β and IL-6 saw no significant increases after exposure to mitochondria. IL-8, a critical regulator of PMN infiltration during the acute immune response, had significant differences between control groups, further validating the PMN activation model used 63 . However, no significant differences were present between mitochondria-treated groups and controls. These findings further indicate that transplanted mitochondria are not inducing damage-associated pro-inflammatory responses in recipient PMNs.
There was significant upregulation in the pro-inflammatory cytokine TNF-α between the mitochondria-treated PMA-stimulated group and the PMA-stimulated HBSS+ control at both timepoints. TNF-α is associated with increases in ROS production and apoptosis in PMNs 64 . However, it has been proven that these effects are dependent on the concentration of TNF-α present in the microenvironment. Previous studies have indicated that TNF-α has two potential effects on PMN function. It has been found that at lower concentrations, TNF-α possess anti-apoptotic qualities, over a broad range of concentrations 65 . Alternatively, at higher concentrations, TNF-α production has been associated with significant pro-apoptotic effects that are directly linked to the production of reactive oxidative species 65 . Therefore, it has been suggested that reactive oxidative species production in PMNs can be used as an assessment for the functional state of TNF-α. If respiratory burst does not take place, TNF-α possesses protective effects, decreasing PMN apoptosis 65 . Therefore, for the assessed data, due to the lack of significant increases in intracellular ROS following mitochondria co-incubation (Figure 6), it can be assessed that although statistically significant, the TNF-α produced was in the protective, anti-apoptotic range as opposed to the pro-apoptotic range traditionally associated with TNF-α upregulation.
The regenerative growth factors FGF-2 and HGF were also assessed (Figure 9). The median relative fold change for FGF-2 production in both stimulated and unstimulated mitochondria-treated groups in comparison to controls ranged between ~200X and 500X. FGF-2 plays a key role in tissue regeneration, and its release by PMNs has emerged as an important element of early wound-healing responses. As first responders in innate immunity, PMNs rapidly infiltrate injured tissue and secrete mediators that influence angiogenesis, cell proliferation, and matrix remodeling. Among these factors, FGF-2 is particularly significant due to its potent pro-angiogenic and pro-proliferative activities. PMNs contribute to FGF-2 availability through both direct release and enzymatic mechanisms, thereby enhancing endothelial and fibroblast proliferation and supporting neovascularization, consistent with evidence demonstrating FGF-2’s central role in orchestrating the early and late phases of tissue repair42,66,67. FGF-2 acts in concert with MMPs to remodel ECM components and facilitate the transition from inflammation to regeneration. FGF-2 supports fibroblast and keratinocyte migration and proliferation, facilitating matrix deposition and restoration of tissue integrity. These coordinated PMN-FGF-2 interactions are associated with accelerated wound closure and improved regenerative outcomes across multiple tissue types68–71. In prior studies, FGF-2 has also been associated with regenerative PMN phenotypes68,72. Therapeutically, FGF-2 has shown promise in promoting the regeneration of periodontal and other connective tissues, enhancing cell proliferation and ECM production, and ameliorating inflammation. Its consistent upregulation following injury further supports its role as an active mediator of repair rather than a passive byproduct of inflammation73,74. In Figure 9(b), a sizable effect was observed at the 3-h timepoint for a single donor for both PMA-stimulated and PMA-unstimulated mitochondria-treated groups; however, when removed, significant FGF-2 upregulation remained as indicated in Supplemental Materials. As a result, the substantial upregulation of FGF-2 at both timepoints indicates significant promise for the promotion of PMN-mediated tissue regeneration via mitochondrial transplantation. HGF, another potent stimulator of angiogenesis, was not significantly affected by mitochondrial transplantation, as the only observed significant differences were between control groups 75 .
Assessed, detectable, anti-inflammatory, and regenerative cytokines included angiopoietin-1, IL-22, VEGF-A, and IL-1RA (Figure 10). No significant differences were observed in angiopoietin-1 and IL-1RA production between any experimental groups. IL-22 had significant upregulation between the PMA-stimulated mitochondria-treated group and the PMA-stimulated HBSS+ control group at 3 h. PMN-derived IL-22 also contributes to functional wound resolution by promoting the recruitment and activation of additional immune cells, thereby amplifying regenerative processes. IL-22 influences fibroblast behavior, driving their proliferation and activation during matrix remodeling and tissue repair 76 . Moreover, PMNs interact with a broader cytokine network, including IL-6 and TNF-α, to coordinate and sustain regenerative signaling, underscoring the collaborative roles of multiple cytokines during wound healing77,78. The role of IL-22 in tissue regeneration is closely tied to its ability to strengthen epithelial barrier integrity and stimulate cellular proliferation. PMNs have emerged as important sources of IL-22 in both skin and intestinal wound-healing contexts76,79. IL-22 acts through its receptor on epithelial and other target cells, initiating downstream signaling pathways that promote repair, regeneration, and restoration of tissue homeostasis79,80. In epithelial tissues, IL-22 enhances mucus and antimicrobial peptide production, supporting barrier function and creating a microenvironment conducive to healing 79 . Moreover, IL-22 appears to have a role beyond just promoting cell survival; it helps orchestrate the transition from inflammation to resolution. Elevated IL-22 levels have been associated with increased PMN apoptosis and subsequent phagocytic clearance of dead cells, which is vital for avoiding chronic inflammation 81 . This process is reflected in the modulation of PMN function, where IL-22 influences the expression of various surface markers that are pivotal in mediating PMN interactions with other immune cells, helping in the resolution of inflammation82,83. Another dimension of IL-22’s role in inflammation resolution is its ability to counteract pro-inflammatory signals. Specifically, while IL-22 does not diminish the initial immune response, it modulates the inflammatory environment by promoting tissue repair over prolonged inflammation 84 . This is particularly important in contexts where unresolved inflammation can lead to further tissue damage and sustained inflammatory disorders, such as autoimmune diseases or host-graft rejection 81 . Due to these regenerative benefits, upregulation of this cytokine further indicates the regenerative potential of mitochondrial transplantation on PMN function and the acute inflammatory response overall. In addition, VEGF-A, a critical stimulator of angiogenesis, production had significant differences between control groups but not between treatment groups (Figure 10), indicating that mitochondrial transplantation had no observed negative impacts on angiogenic factors 85 .
Analyzed cytokines responsible for immune cell recruitment and production included MCP-1 and G-CSF (Figure 11). Results indicated a significant upregulation in MCP-1, a cytokine associated with monocyte recruitment, between mitochondria-treated groups and control groups for both stimulated and unstimulated conditions at both timepoints 86 . PMNs and macrophages engage in a dynamic interplay that is central to the host-immune response and tissue regenerative process 87 . Specifically, PMNs produce inflammatory mediators such as MCP-1 that are directly involved in the recruitment of not only monocytes but also tissue resident macrophages 88 . Recruited and activated macrophages are then able to release their own unique inflammatory mediators into the healing microenvironment that impact PMN effector functions, migratory behavior, and cell death 87 . PMNs have also been associated with the polarization of macrophages toward regenerative, M2 phenotypes 4 . Therefore, this cyclic communication between PMNs and macrophages is a key determinant of host-biomaterial outcomes. In a balanced inflammatory response, PMN and macrophage crosstalk is critical to the promotion of inflammatory resolution and tissue regeneration.
In addition, there was a similar significant increase in G-CSF between stimulated and unstimulated mitochondria-treated groups and control groups at both timepoints. G-CSF is a key regulator of PMN biology. It is associated with the production, differentiation, and release of neutrophil precursors in the bone marrow 89 . However, G-CSF also contributed to the function of mature neutrophils at sites of inflammation after leaving the bone marrow 89 . This mediator has also shown been shown to interact with other facets of the tissue-regenerative process such as mesenchymal stem cell-immune cell crosstalk and macrophage polarization toward regenerative phenotypes90,91. In addition, prior literature has associated G-CSF production with the promotion of regenerative neutrophil phenotypes, as well as preventing inflated neutrophil mobilization, leading to excess tissue damage5,92.
This study began to explore the influences of mitochondrial transplantation on human PMNs. However, additional studies are needed to fully understand the implications of applying mitochondrial transplantation to human PMNs. Limiting factors included mitochondria dosage, mitochondrial source, the assessed inflammatory mediators, donor variability, and the heterogeneous PMN population used. Particularly, this study determined mitochondria dosage based on prior lab experiments; however, further dose-dependent optimization is needed. In addition, mitochondria were isolated from non-donor-specific HDF cells. Therefore, due to the diversity of mtDNA, it would be beneficial to explore the impacts of mitochondrial source on overall transplantation outcomes, as well as any resulting immunomodulation 93 . For example, HDF-derived mitochondria caused sizable increases in FGF-2, a growth factor secreted by fibroblast cells 94 . Therefore, due to mitochondrial heterogeneity, these increases are potentially due to mitochondrial source 38 . In addition, although several inflammatory mediators were assessed in this study, several key PMN-produced inflammatory mediators remain to be explored. Specifically, the impacts of mitochondrial transplantation on the production of PMN extracellular ROS and additional PMN-specific cytokines, chemokines, surface receptors, and lipid mediators need to be further assessed. In addition, at the 1-h timepoint, multiple analytes in the multiplex analyses were at, below, or near the LLOQ. For future studies, it would be beneficial to analyze timepoints past the observed 3 h, as well as optimize sample dilution factors used in the assays. Due to high sample variability between donors, clinical studies assessing the impacts of this therapy on a larger number of subjects would also significantly contribute to the clarity of findings. Particularly, baseline cytokine and protein release between donors was highly variable. Prior literature has established that this is to be expected due to PMN heterogeneity53–55. Therefore, although data trends were consistent across donors, normalization to donor-specific positive controls was necessary, and additional further study using larger sample sizes would further verify observations 95 . In addition, studies that separate individual PMN populations would be beneficial. Although neutrophils comprise the majority of PMNs, eosinophils that are also present in this population have the ability to produce inflammatory mediators as well 96 . Therefore, to identify potential modulatory behavior in specific PMN populations, cell sorting would also be beneficial.
Previous studies have demonstrated that PMNs prime the wound-healing microenvironment to orchestrate the coming cellular interactions during the implantation of biomaterials. However, there has been a lack of effective strategies to guide the behavior of PMNs toward states that are conducive to non-fibrotic, regenerative implant environments. In addition, mitochondrial transplantation is an emerging research strategy for targeting mitochondria-dependent cellular processes and disease states; however, this reparative approach currently has unaddressed determinants of its success, including the lack of understanding of the interactions between this therapy and the acute immune response. This study provides foundational contributions toward tackling both research gaps. Next steps include identifying specific endocytic and signaling pathways involved in the observed immunomodulatory benefits, characterizing the impacts of mitochondrial transplantation on intercellular crosstalk between PMNs and other cell populations in the acute inflammatory response, and beginning to explore the capacity of mitochondrial transplantation to benefit or to adversely affect other innate immune cells.
Conclusion
Mitochondrial transplantation exhibits significant potential for PMN-mediated acute immune modulation due to its upregulation of regenerative proteins and its retention of critical, PMN-produced mediators. Overall, this study indicates the need to investigate the interactions between mitochondrial transplantation and the acute immune response to tailor this therapy most effectively for clinical applications. Additional studies are needed to elucidate the endocytic and signaling pathways involved in the exhibited PMN modulation.
Supplemental Material
sj-docx-1-cll-10.1177_09636897261453307 – Supplemental material for Mitochondrial transplantation as an immunomodulatory strategy: Modulating polymorphonuclear leukocytes for functional tissue regeneration
Supplemental material, sj-docx-1-cll-10.1177_09636897261453307 for Mitochondrial transplantation as an immunomodulatory strategy: Modulating polymorphonuclear leukocytes for functional tissue regeneration by Samantha C. Hall, Evan N. Main and Gary L. Bowlin in Cell Transplantation
Footnotes
Acknowledgements
The authors thank Drs. Marie van der Merwe and Jacqueline Pence for their assistance in multiplex assay preparation and protocols. The authors would also like to thank the University of Memphis Integrated Microscopy Center for their assistance in confocal imaging. The authors would also like to thank Audrey Alberson for her work and diligence as an undergraduate trainee, as well as for her assistance with sample collection and donor recruitment.
Ethical considerations
The study was conducted in accordance with the Declaration of Helsinki and was approved by the institutional review board of the University of Memphis (protocol code #PRO-FY2020-230, dated 8 November 2022). Informed consent was obtained from all subjects involved in the study.
Consent to participate
All donors provided written informed consent prior to participation. Written informed consent was obtained for anonymized patient information to be published in this article.
Consent for publication
Not applicable.
Author contributions
Funding
The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: Support was provided by the Herbert Herff Chair of Excellence Biomedical Engineering, University of Memphis, held by Gary L. Bowlin.
Declaration of conflicting interests
The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.
Data availability statements
All raw data supporting the conclusions of this article will be made available by the authors upon request.
Supplemental material
Supplemental material for this article is available online.
Statement of human and animal rights
This article does not contain any studies with human or animal subjects.
Statement of informed consent
Informed consent was obtained from all subjects involved in the study.
References
Supplementary Material
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