PTP1B phosphatase dampens iPSC-derived neutrophil motility and antimicrobial function

Abstract

Neutrophils are rapidly recruited to sites of infection and are critical for pathogen clearance. Therapeutic use of primary neutrophils has been limited, as they have a short lifespan and are not amenable to genetic manipulation. Human induced pluripotent stem cells (iPSCs) can provide a robust source of neutrophils for infusion and are genetically tractable. However, current work has indicated that dampened intracellular signaling limits iPSC-derived neutrophil (iNeutrophil) cellular activation and antimicrobial response. Here, we show that protein tyrosine phosphatase 1B (PTP1B) inhibits intracellular signaling and dampens iNeutrophil effector function. Deletion of the PTP1B phosphatase increased PI3K and ERK signaling and was associated with increased F-actin polymerization, cell migration, and phagocytosis. In contrast, other effector functions like NETosis and reactive oxygen species production were reduced. PTP1B-deficient neutrophils were more responsive to Aspergillus fumigatus and displayed rapid recruitment and control of hyphal growth. Accordingly, depletion of PTP1B increased production of inflammatory factors including the neutrophil chemokine interleukin-8. Taken together, these findings suggest that PTP1B limits iNeutrophil motility and antimicrobial function.

Deletion of the phosphatase PTP1B increases induced pluripotent stem cell–derived neutrophil intracellular signaling to improve motility, phagocytosis, and antifungal response.

1. Introduction

Rapid progress has been made in the development of infusible cell and gene therapies in the past 2 decades.^1–3 Much of this work has focused on adoptive transfer of genetically modified host immune cells, such as chimeric antigen receptor T cells, for cancer treatment.^3–5 However, this approach can be limiting, as genetic modification of primary immune cells is a challenging and lengthy process, and the quality of patient-derived cells can be variable. Induced pluripotent stem cells (iPSCs) have emerged as a promising therapeutic option, as they can be genetically modified and provide a robust source of differentiated cells.⁶ Furthermore, iPSCs can generate all immune cell types, thus allowing for development of novel cell therapies including those utilizing neutrophils.^7–9

Neutrophils are highly motile cells and as first responders of innate immunity are critical for clearing bacterial and fungal infections. They exhibit an arsenal of antimicrobial functions, including production of reactive oxygen species (ROS), release of neutrophil extracellular traps (NETs), and phagocytosis. Primary neutrophils have a short lifespan and are not genetically malleable, and thus their therapeutic use has been limited. iPSC-derived neutrophils (iNeutrophils) are currently being explored as an adoptive cell therapy for neutropenic patients with bacterial infections^10,11 or as an anticancer therapy.^12–15 iNeutrophils have been shown to exert classic functions including cell motility, phagocytosis, and production of ROS and NETs.¹⁶ However, as shown by many groups, their functional capacity is often reduced compared with primary human neutrophils.^10,12,16,17 Thus, further study of the molecular pathways that regulate iNeutrophil function is necessary for any future clinical use.

We hypothesized that the relative decrease in iNeutrophil motility and phagocytosis could be due to dampened activation of intracellular signaling pathways, such as pAKT and pERK, which mediate actin polarization.^17–19 Protein tyrosine phosphatases such as PTP1B negatively regulate PI3 K and MAPK signaling pathways that are involved in many neutrophil functions.²⁰ In murine models, PTP1B inhibits neutrophil phagocytosis and bacterial killing^21,22 and thus may similarly dampen human iNeutrophil function.

Here, we show that we can genetically engineer iNeutrophils to dissect mechanisms that alter cellular activation, motility, and antimicrobial response. Deletion of PTP1B increased iNeutrophil PI3 K and ERK signaling associated with increased F-actin polymerization, cell migration, and phagocytosis. PTP1B knockout (PTP1B-KO) iNeutrophils produced higher levels of inflammatory chemokines, including the neutrophil chemoattractant interleukin (IL)-8. Furthermore, PTP1B-KO iNeutrophils displayed a highly activated morphology in the presence of the fungal pathogen Aspergillus fumigatus, efficiently migrated to hyphae, and limited fungal growth. Thus, deletion of the PTP1B phosphatase increases iNeutrophil intracellular signaling and function.

2. Methods

2.1. Stem cell culture and neutrophil differentiation

Neutrophils were differentiated from bone marrow–derived human iPSCs (hiPSCs) as previously described.⁷ Briefly, bone marrow–derived IISH2i-BM9 were obtained from WiCell. hiPSCs were cultured on Matrigel-coated tissue culture plates in mTeSR-Plus medium (STEMCELL Technologies).

To induce hemogenic endothelium, hiPSCs are transfected with ETV2 modified messenger RNA (mRNA) (TriLink Biotechnologies) in TeSR-E8 media (STEMCELL Technologies) using TransIT reagent and mRNA boost (Mirus Bio). One hour prior to transfection, cells were detached by TrypLE Select (Life Technologies). Cells were replated onto collagen (2.4 µg/mL)-coated plates in TeSR-E8 media with 10 µM ROCK inhibitor Y-27632 (ROCKi; Tocris). One day following transfection, media was changed to StemLineII media with VEGF-165 (20 ng/mL; PeproTech) and FGF2 (10 ng/mL; PeproTech) to induce differentiation into hemogenic endothelial cells. We refer to this media cocktail as media A. After 2 d, the media was changed to differentiate the cells into granulocyte-monocyte progenitors with StemLineII media supplemented with FGF2 (20 ng/mL), granulocyte-macrophage colony-stimulating factor (25 ng/mL; PeproTech), and UM171 (50 nM; Xcess Biosciences). We refer to this media cocktail as media B. On days 8 to 10, floating cells were gently harvested and used for terminal neutrophil differentiation. These cells were cultured in StemSpan SFEM II medium (STEMCELL Technologies), supplemented with GlutaMAX 100 × (1×; Thermo Fisher Scientific), ExCyte 0.2% (Merck Millipore), human granulocyte colony-stimulating factor (150 ng/mL; PeproTech), and Am580 retinoic acid agonist (2.5 μM; STEMCELL Technologies) at 1 × 10⁶ cells/mL density. We refer to this media cocktail as media C. After 4 d, fresh media C was added on the top of the cells. Neutrophils were harvested from the supernatant 8 to 10 d after plating in media C.

2.2. Generation of PTP1B^−/− BM9iPSCs

To generate PTP1B-KO BM9 iPSCs, 2 single guide RNAs were designed using the Synthego CRISPR design tool to target PTPN1 exon 3: 5′-GATGTAGTTTAATCCGACTA-3′ (sgRNA1) and 5′-TAAAAATGGAAGAAGCCCAA-3′ (sgRNA2). Prior to nucleofection, BM9 iPSCs were detached by TrypLE Select (Life Technologies) and singularized by pipetting. A total of 5 µg of both sgNRAs and 5 µg of Cas9 protein (PNA Bio) were incubated together for 10 min, then the cells were nucleofected using the Human Stem Cell Nucleofector Kit 2 (#VPH-5022; Lonza). Cells were plated at 25 cell/cm² on a Matrigel-coated plate in mTeSR-Plus supplemented with 1× CloneR supplement (STEMCELL Technologies; #05888). After 2 d, media was changed to mTeSR-Plus only. Individual colonies were picked after 7 d and further expanded. To confirm biallelic mutation in PTPN1, genomic DNA was extracted from individual clones, then screened by polymerase chain reaction (PCR) for the acquisition of a 67 bp deletion using primer1 (5′-TGCATCAGAGAACAGATCCT-3′) and primer2 (5′-CTGGGTAAGAATGTAACTCC-3′). After Western blot verification and chemotaxis analysis of multiple PTP1B^−/− clones, we selected a single clone (#6) for further experimentation.

2.3. Human neutrophil isolation

All blood samples were obtained from healthy donors and were drawn according to the University of Wisconsin–Madison Minimal Risk Research Institutional Review Board–approved protocol (ID: 2017–0032) per the Declaration of Helsinki. Human blood was obtained from volunteering donors with informed written consent through a protocol that was approved by the Internal Review Board of the University of Wisconsin–Madison. Neutrophils were isolated using MACSxpress negative antibody selection kit and purified with the MACSxpress erythrocyte depletion kit (Miltenyi Biotec), following manufacturer's instructions. Isolated neutrophils were resuspended in modified Hank’s Balanced Salt Solution (mHBSS) (+0.1% human serum albumin [HSA] + 10 mM HEPES) and utilized for various experiments. Incubations involving neutrophils were performed at 37 °C with 5% CO₂.

2.4. Cytospins

To confirm neutrophil morphology, 90,000 cells were spun onto a glass slide for 5 min at 1200 rpm using a Shandon Cytospin 3 centrifuge (Thermo Fisher Scientific). Cytospin slides were then stained with Differential Quick III Stain Kit (Electron Microscopy Sciences; #26096-25) following manufacturer's instructions. Slides were imaged at 20 × on an Olympus IX70 microscope and evaluated for neutrophils by the presence of hypersegmented nuclei.

2.5. Flow cytometry

Flow cytometry analysis was used to evaluate expression of cell lineage markers on iNeutrophils. Neutrophils were stained in phosphate-buffered saline (PBS) + 1% HSA media + Brilliant Buffer (Thermo Fisher Scientific; #00-4409-42) and Human TruStain FcX Fc Receptor Blocking Solution (BioLegend; #422302), then fixed with 2% PFA. Data acquisition was performed on an Aurora Cytometer (CytekBio). Antibodies used in this study can be found in Supplementary Table 1. Forward and side scatter parameters identified single cells and live cells were identified with Ghost Dye Red 780 or Zombie NIR dye. Myeloid cells were identified by CD11b+ expression and mature neutrophils were identified by CD15+ or CD15+ CD16+ expression. Monocytes were identified as CD14+ cells. Data were analyzed using FlowJo software (v10.8.1; TreeStar).

2.5.1. Flow cytometry analysis in R using Cytofworkflow pipeline

Flow cytometry (.fcs) files were read into FlowJo after spectral unmixing. Initial gates using forward and side scatter parameters isolated single cells, and a viability dye was used to exclude dead or dying cells. For each sample, live, single cells were exported, selecting the “all compensated parameters” option to generate new .fcs files in FlowJo for further analysis using R (v4.0.3; R Foundaiton for Statistical Computing) with the prepData function from the Cytofworkflow (v1.14.0) pipeline.²³ Data transformation was calculated for all fluorescent parameters using arcsinh normalization with a cofactor of 3,000. The distribution of marker expression for transformed data was plotted using a smoothed density function using ggplot2 to compare wild-type (WT) and KO samples.

2.6. PrestoBlue viability

Primary human neutrophils and iNeutrophils were resuspended in RPMI media supplemented with 10% fetal bovine serum (FBS). A total of 100,000 cells were plated into each well of a black clear-bottom 96-well plate. Separate plates were prepared for each time point (days 0, 1, 3, and 5). On day 0, cells were allowed to rest for at least 1 h prior to addition of PrestoBlue HS (Thermo Fisher Scientific; #P50200). For each time point, PrestoBlue HS was added and incubated for 30 min at 37 °C before reading fluorescence at 560/590 nm in a microplate reader (Synergy H1; Bio-Tek Instruments). Background fluorescence of media-only wells was subtracted from each sample, and the fold change was calculated compared with day 0, respective to each cell line.

2.7. Western blot cell signaling

Western blotting was conducted to quantify PTP1B protein expression and evaluate phospho-signaling in our iNeutrophils. Cells were stimulated with 1 μM fMLP for 3 min. Cell pellets were collected in lysis buffer (25 mM HEPES, pH 7.5, 150 mM NaCl2, 1% Nonidet P-40, 10 mM MgCl₂, 1 mM EDTA, 10% glycerol supplemented with 1 g/mL pepstatin A, 2 g/mL aprotinin, 1 g/mL leupeptin, 200 nM phenylmethanesulfonyl fluoride, 1 mM sodium orthovanadate, 1% protease inhibitor mixture 2 [#p5726, Sigma-Aldrich]). Cells were incubated on ice for 10 min and then sonicated with 20% amplitude for 3 × 5 s. Cells were then clarified by centrifugation at 15,000 g, 4 °C for 15 min. Protein concentrations were determined using the Pierce BCA Protein Assay (Thermo Fisher Scientific; 23225) and samples were stored at −80 °C. Immunoblotting of cell lysates was performed and blots was imaged with an infrared imaging system (Odyssey; LI-COR Biosciences). Primary and secondary antibodies used can be found in Supplementary Table 2.

2.8. Chemotaxis

Chemotaxis was assessed using 2 separate microfluidic devices as described previously.^24,25 Regarding the former, PDMS devices were plasma treated and adhered to glass coverslips. Devices were coated with 10 μg/mL fibrinogen (Sigma-Aldrich) in PBS for 30 min at 37 °C, 5% CO₂. The devices were blocked with 2% bovine serum albumin (BSA)–PBS for 30 min at 37 °C, 5% CO₂, and then washed twice with mHBSS (+0.1% HSA + 10 mM HEPES). Cells were stained with Calcein AM (Molecular Probes) in PBS for 10 min at room temperature followed by resuspension in mHBSS (+0.1% HSA + 10 mM HEPES). Cells were seeded at 5 × 10⁶/mL to allow adherence for 30 min before addition of chemoattractant. Then, 3 µL of 1 μM fMLP (Sigma-Aldrich) chemoattractant was loaded into the input port of the microfluidic device. Cells were imaged every 30 s for 45 to 90 min on a Nikon Eclipse TE300 inverted fluorescent microscope with a 10× objective and an automated stage using MetaMorph software v7.8 (Molecular Devices). Automated cell tracking analysis was done using JEX software²⁶ to calculate chemotactic index and velocity and to generate rose plots.

For analysis of confined migration, a second device was utilized. Channels bound by a liquid-liquid (oil-media) interface were constructed as previously described.^25,27 Briefly, PDMS-silane was grafted onto a glass slide by chemical vapor deposition. The slides were then exposed to O₂ plasma following overlay with a PDMS mask, allowing for differential surface patterning of hydrophilic and exclusively liquid-repellent regions.^28,29 Channels of 100 µm width and 500 µm length were constructed by sweeping culture media (RPMI + 2% FBS) with 10 μg/mL fibrinogen (Sigma-Aldrich). After a 30-min incubation period at 37 °C, channels were washed by sweeping PBS across the channels. A 1 µL droplet of cells (40k cells/µL) was added to the inlet droplet across from either 100 nM fMLP or 11.25 µM IL-8 in the outlet droplet. Cells were imaged every 30 s for 45 to 90 min on a Nikon Eclipse TE300 inverted fluorescent microscope with a 20 × objective and an automated stage using MetaMorph software. Cells were tracked manually using the FIJI/ImageJ Manual Tracking plugin. A minimum of 5 cells per device were tracked and 3 devices were tracked per condition.

2.9. Immunofluorescent imaging

iNeutrophils were stimulated and prepared for immunofluorescent imaging as previously described.³⁰ Acid-washed 22-mm glass circle coverslips were coated with 10 µg/mL fibronectin for at least 1 h at 37 °C and then blocked for 30 min with 2% BSA-PBS. Cells (3 × 10⁵) in 500 µL 0.5%HSA-RPMI were seeded per coverslip in a 24-well plate and allowed to rest for 30 min. fMLP was added for a final concentration of 100 nM fMLP, and cells were allowed to adhere for 30 min at 37 °C. Media was aspirated and fixation was performed as follows: 1 mL of 37 °C preheated 4% paraformaldehyde in PEM buffer (80 mM PIPTES, pH 6.8; 5 mM EGTA, pH 8.0; 2 mM MgCl₂) for 15 min at room temperature (RT). Fixative was aspirated and 0.25% Triton X-100 in PEM buffer was added for 10 min at 37 °C. Coverslips were washed 3 times with PBS, blocked in 5% BSA-PBS for 60 min at RT, washed, and incubated in rhodamine-phalloidin (Thermo Fisher Scientific; #R415) overnight at 4 °C. Coverslips were washed, counterstained with Hoechst 33342 (Thermo Fisher Scientific; #H3570; 1:500) for 5 to 10 min, washed with ddH₂O, and mounted on Rite-On Frosted Slides (Thermo Fisher Scientific; 3050-002) with ProLong Gold Antifade Mountant (P36930; Invitrogen). Slides were imaged on an upright Zeiss LSM 800 Laser Scanning Confocal Microscope equipped with a motorized stage and Airyscan module. Images were acquired with a 60× oil, NA 1.40 objective using the Airyscan acquisition mode. Images were processed in ZenBlue software (Zeiss) using the Airyscan processing method and maximum intensity projections generated for further analysis. Integrated density of the actin channel was quantified using ImageJ (v1.52a) (National Institutes of Health).

2.10. Phagocytosis

Phagocytosis was quantified using pHrodo Green Escherichia coli BioParticles (Invitrogen; #P35366) following manufacturer's instructions. Briefly, the E. coli BioParticles were opsonized with 30% pooled human serum (#MP092930149; MP Biomedicals) for 30 min at 37 °C then washed 3 times in PBS. One million cells were resuspended in 80 µL of media C and 20 µL of opsonized beads were added at 100:1 multiplicity of infection. Cells and beads were incubated in an Eppendorf tube for 1 h at 37 °C, then stopped by addition of ice-cold PBS. While keeping tubes on ice, cells were stained with neutrophil lineage markers (Supplementary Table 1), then fixed with 2% PFA before flow cytometry analysis on the Aurora Cytometer (CytekBio).

2.11. Intracellular ROS

Intracellular peroxynitrite production by iPSC-differentiated neutrophils was quantified using the peroxynitrite indicator DHR123 (Invitrogen; #D23806). A black-walled clear-bottom 96-well plate was precoated with 10 µg/mL fibronectin. 100,000 cells in Phenol Red Free-RPMI containing 2% FBS and 5 µg/mL DHR123 reagent were plated into each well. Cells were allowed to rest for 30 min at 37 °C. PMA at a final concentration of 50 ng/mL was added to appropriate wells. Wells were plated in quadruplicate to account for technical error. Reads were taken every 15 min for 2 h at 485/535 nm using the Victor3V microplate reader (PerkinElmer). Background signal of unstimulated cells was subtracted from the corresponding PMA stimulated cell line and plotted over time. Samples were normalized to the percent of maximal ROS production by WT iNeutrophils to account for daily fluctuations in RFUs.

2.12. NETosis assay

A black-walled clear-bottom 96-well plate was precoated with 10 µg/mL fibronectin. A total of 200,000 cells in 100 µL Phenol Red Free RPMI + 2% FBS were plated into each well. Cells were allowed to rest for 30 min at 37 °C. A final concentration of 100 ng/mL PMA was added to appropriate wells and the plate was incubated for 4 h at 37 °C. Sytox Green (Invitrogen; #S7020) was added at 375 nM final concentration. The microplate was incubated for 10 min and an endpoint reading was taken using a Victor3V microplate reader (PerkinElmer) to quantify extracellular DNA by fluorescence (500/528 nm). Background signal of Sytox Green unstimulated cells was subtracted from the corresponding PMA stimulated cell line. Fold change of fluorescence was calculated compared with WT.

2.13. CD15 positive selection

hiPSC-derived neutrophils were positively selected for mature neutrophils using surface expression of CD15. CD15 microbeads (Miltenyi; #130-046-601) were incubated with neutrophils following the manufacturer's protocol and then positively selected using LD columns (Miltenyi; #130-042-901). Cells were allowed to rest overnight in media C with added 10% FBS before use in further experiments.

2.14. Real time quantitative PCR

iPSC-differentiated neutrophils were stimulated with lipopolysaccharide (LPS) to evaluate expression of inflammatory cytokines. A 6-well TC-treated plate was precoated with 10 µg/mL fibronectin. Approximately 6 million cells resuspended in 2% FBS-RPMI were plated per well. Cells were allowed to rest for 30 min at 37 °C before stimulation. Final concentration of 200 ng/mL E. coli LPS (Sigma-Aldrich; #L2755) was added to appropriate wells and the plate was incubated for 2 h at 37 °C. Floating cells were collected and spun down at 300 g. To collect RNA, 1 mL of Trizol (Invitrogen) was added to adherent cells and then combined with the pelleted floating cell sample. Samples were pipetted to mix and then stored at −80 °C until RNA extraction. RNA was isolated by Trizol extraction following manufacturer's instructions (Invitrogen; DNA preparation. cDNA was synthesized using oligo-dT primers and the Superscript III First Strand Synthesis kit (Invitrogen #18080-051) following manufacturer's instructions. Complementary DNA was used as the template for quantitative PCR (qPCR) using FastStart Essential Green DNA Master (Roche) and a LightCycler96 (Roche). Data were normalized to ef1a within each sample using the ΔΔCq method.³¹ Fold change represents the change in cytokine expression over the unstimulated WT sample. All qPCR primers are listed in Supplementary Table 3.

2.15. Inflammatory cytokine and LTB4 enzyme-linked immunosorbent assay

iNeutrophil secretion of inflammatory cytokines IL-8, IL-1β, IL-6, and tumor necrosis factor α (TNFα) protein was quantified by enzyme-linked immunosorbent assay (ELISA). A 12-well TC-treated plate was coated with 10 µg/mL fibronectin. iNeutrophils were resuspended in 2% FBS supplemented RPMI and 1.25 to 1.5 million cells were plated per well. Cells were allowed to rest for 30 min at 37 °C before stimulation with 200 ng/mL E. coli LPS or 10 µg/mL Zymosan for 4 h at 37 °C. Wells were harvested and the media was spun down at 300 g to pellet the cells. Supernatants were collected, aliquoted, and frozen at −80 °C until use. Cytokines were quantified with the Human IL-8/CXCL8 DuoSet ELISA (#DY208; Bio-Techne), Human IL-6 DuoSet ELISA (Bio-Techne; DY206), Human TNFα DuoSet ELISA (Bio-Techne; DY210), and Human IL-1 beta/IL-1F2 DuoSet ELISA (Bio-Techne; DY201) following manufacturer's instructions. Samples were diluted to fit in the range of the standard curve and normalized to the input cell number.

LTB4 release was quantified following stimulation with fMLP. A 24-well TC-treated plate was coated with 10 µg/mL fibronectin. iNeutrophils were resuspended in 0.5% HSA supplemented RPMI and 1 million cells were plated per well. Cells were allowed to rest for 30 min at 37 °C before stimulation with 1 μM fMLP for 5 min or 30 min. Wells were harvested and the media was spun down at 300 g to pellet the cells. Supernatants were collected, aliquoted, and frozen at −80 °C until use. LTB4 release was quantified with the LTB4 Parameter Assay Kit (Bio-Techne; KGE006B) following manufacturer's instructions.

2.16. Fungal co-culture and imaging

A. fumigatus (CEA10) expressing RFP was grown as previously described.³² Briefly, A. fumigatus (CEA10) was grown on glucose minimal medium (GMM) plates at 37 °C in the dark to promote asexual conidiation. Aspergillus was plated at 1 × 10⁶ conidia/10 cm plate for 3 to 4 d. Conidia were harvested in 0.01% Tween water by scraping with an L-spreader and then passed through sterile Miracloth into a 50 mL conical tube. The spore suspension was centrifuged at 900 g for 10 min at room temperature and resuspended in 50 mL 1× PBS. The spore suspension was then vacuum filtrated using a Buchner filter funnel with a glass disc containing 10- to 15-µm-diameter pores. The filtered suspension was centrifuged at 900 g for 10 min and resuspended in 1 mL 1× PBS. Conidia were counted using a hemacytometer and the concentration was adjusted to 1.5 × 10⁸ spores/mL. Conidial stocks were stored at 4 °C and used up to 1 mo after harvesting.

Live imaging was conducted to visualize iNeutrophil interactions with fungal hyphae. A total of 2 × 10³  A. fumigatus spores/well were plated in 100 µL GMM media in a black 24-well plate coated with fibronectin (Corning). The plate was incubated at 37 °C for 8 h, or until germling stage. Spore germination was confirmed by microscopy prior to adding neutrophils. iNeutrophils were resuspended in RPMI + 2% FBS at 6 × 10⁵ cells/mL. GMM media was removed from the wells and replaced with 100 µL of neutrophil suspension to yield a neutrophil-to-spore ratio of 150:1. For MK886 treatment, peripheral blood (PB) neutrophils or iNeutrophils were preincubated with 1 μM MK886 (Calbiochem; #475889) for 30 min at 37 °C prior to addition to the plate. Neutrophil-fungal interactions were imaged every 3 min on an inverted fluorescent microscope Nikon Eclipse TE300 (Nikon) with a 20× objective and an automated stage (Ludl Electronic Products) with a Prime BSI Express camera (Teledyne Photometrics). Environmental controls were set to 37 °C with 5%C CO₂. Videos were compiled using ImageJ software.

2.16.1. Image analysis

iNeutrophil circularity was quantified after 0 and 4 h of co-incubation with RFP-expressing A. fumigatus. Individual cells were outlined using the Polygon selection tool in ImageJ and circularity (4π*area/perimeter²) was calculated and is reported in Fig. 5C. The hyphal length for individual RFP-expressing germlings was measured using fluorescent images after 0 and 4 h of co-incubation with iNeutrophils. Germlings and hyphae were traced using the segmented line feature in ImageJ and the cumulative lengths of individual germlings are reported in Fig. 5G. The percent of germlings with clustered iNeutrophils and cluster size was quantified after 0, 1, 2, and 4 h of co-incubation with A. fumigatus. A cluster was identified as a tightly formed group of at least 5 neutrophils in contact with a germling or hyphal branch. For cluster formation, an initiating neutrophil is identified and upon contact with Aspergillus results in rapid neutrophil recruitment in the following frames. The first frame to show rapid recruitment of >4 neutrophils was counted as the first time point of cluster formation. The cluster area was quantified by outlining the perimeter using the Polygon selection tool (ImageJ). Only iNeutrophils that stayed in contact with the cluster for at least 3 frames were included in the quantification. Cluster size and percent of clustered germlings are reported in Fig. 5E, F.

Fig. 5.

Deletion of PTP1B increases iNeutrophil clustering and inhibition of fungal growth. (A) Representative bright-field images of Aspergillus fumigatus coculture with WT or PTP1B-KO iNeutrophils over the course of 8 h. (B) Higher magnification bright-field images of WT or PTP1B-KO iNeutrophil cell morphology and interaction with A. fumigatus hyphae. Individual cells are outlined. (C) Quantification of iNeutrophil cell shape (circularity) at 0 and 4 h of co-incubation. Small dots represent individual cells. WT: n = 414; KO: n = 396. (D) Representative time-lapse images of PTP1B-KO iNeutrophils phagocytosing and clustering around A. fumigatus hyphae. Clusters are outlined. (E) Quantification of percent of A. fumigatus germlings surrounded by iNeutrophil clusters at 0, 1, 2, and 4 h of co-incubation. Dotted lines indicate individual biological replicates. Number of quantified germlings: WT, n = 155; KO, n = 99. (F) Quantification of iNeutrophil cluster size at 0, 1, 2, and 4 h of co-incubation. Dotted lines indicate individual biological replicates. Number of quantified clusters: WT, n = 39; KO, n = 80. (G) Quantification of hyph­al length at 0 and 4 h of coculture with WT or PTP1B-KO iNeutrophils. Small dots represent individual germlings. WT: n = 138; KO: n = 110. In (A, D) Black arrows indicate germlings, white arrows indicate a phagocytosed germling, and asterisks indicate iNeutrophil swarming and cluster formation. Scale bars represent 50 µm. Experiments were conducted at least 3 times, or as indicated on the plot. Bars in panels C and G represent least-squares adjusted means ± SEM. Black dots in panels C and G represent the averages for independent biological replicates. Dots in panels E and F represent the mean ± SEM for 4 independent biological replicates. P values were calculated by analysis of variance with (C, G) Tukey's multiple comparisons, (E) linear regression, or (F) unpaired Student's t test on the calculated area under the curve for each replicate. ****P < 0.0001.

2.17. Statistical analysis

All experiments and statistical analyses represent at least 3 independent biological replicates (N), meaning 3 independent stem cell differentiations. Technical replicates are completed with the same cell differentiation sample by measuring multiple wells or imaging positions. The replicate number of cells or germlings (n) for experiments in Figs. 2I and 5C, G is indicated in the figure legend. Statistical significance was set to 0.05. Experiments in which cell populations were unresponsive or unhealthy looking were not included in analysis.

Fig. 2.

Deletion of PTP1B promotes intracellular signaling, motility and actin polymerization. (A) Representative Western blots and (B) quantification of ERK, HS1, and AKT phospho-signaling after stimulation with 1 μM fMLP for 3 min. (C) iNeutrophil chemotactic index and (D) mean velocity in response to an fMLP gradient over 45 min of imaging. (E) Representative track plots of cells migrating in response to an fMLP gradient. Blue tracks indicate cells that traveled toward the fMLP source, whereas red tracks indicate cells that moved away. (F) PB neutrophil and iNeutrophil mean velocity under confinement in response to an fMLP gradient. (G) iNeutrophil mean velocity under confinement in response to an IL-8 gradient. (H) Representative immunofluorescence images of F-actin staining after 100 nM fMLP stim. (I) Quantified integrated density of F-actin staining. Small dots represent individual cells. WT: n = 47; KO: n = 54. Scale bar represents 20 µm. Experiments were conducted at least 3 times, or as indicated on the plot. Black dots in panels B, C, D, F, G, and I represent independent biological replicates. Small colored dots in panels C, D, F, and G represent technical replicates. Bars in panel B, C, D, F, G, and I represent means ± SEM. P values were calculated by (B) 1-sample t test, (C, D, G) unpaired Student's t test, (F) analysis of variance with Tukey's multiple comparisons, or (I) Welch's t test. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Analysis of receptor expression by flow cytometry, chemotactic index and cell velocity, phagocytosis, qPCR gene expression, and ELISA was performed using unpaired Student's t test. Technical replicates were included for motility data by imaging at least 4 positions on 2 separate microfluidic devices per sample and tracking all the cells in the field of view. Cell velocity of primary neutrophils and iNeutrophils combined was analyzed by 1-way analysis of variance. Analysis of PrestoBlue viability, intracellular ROS production, and percent of clustered germlings was performed using simple linear regression, and the P value was calculated by comparing the slope of each line. Technical replicates comprised 3 to 4 wells for the viability and ROS assays, which were conducted on a plate reader. Analysis of cluster size over time was performed by calculating the area under the curve for each replicate and then determining the P value using a unpaired Student's t test. For percent clustering and cluster size, 4 positions were imaged over time, and each cluster was quantified and averaged to represent 4 technical replicates. Analysis of pHrodo median fluorescence intensity, NETosis, and Western blot phospho-signaling was performed using 1-sample t test. Technical replicates for NETosis comprised measuring fluorescence in 3 wells. For Western blotting, a single stimulated sample was collected and used as a single technical replicate. The previous analyses were conducted using GraphPad Prism (v9; GraphPad Software). For analysis of the F-actin integrated density data, outliers were identified using the robust regression and outlier removal method.³³ Equivariance was checked using an F test and it was determined that the samples did not meet the requirement; thus, a Welch's unequal variances t test was used. iNeutrophil circularity and hyphal length analyses represent least-squares adjusted means ± SEM and were compared using analysis of variance with Tukey's multiple comparisons (R v4.3.3). For fungal analysis, 4 positions were imaged over time, and each fungal hyphae was quantified and averaged to represent 4 technical replicates. For MK886 treatment, the time to cluster for each fungal germling was determined and then averaged for each imaging position. The P value was determined by unpaired Student's t test. All graphical representations of data were created in GraphPad Prism (v9), and figures were ultimately assembled using Adobe Illustrator (version 23.0.6).

3. Results

3.1. Generation of PTP1B-KO iNeutrophils

We hypothesized that dampened intracellular signaling may be responsible for the reduced function of iNeutrophils compared with primary human neutrophils. We differentiated hiPSCs into neutrophils using serum- and feeder-free conditions, following published protocols (Fig. 1A).^7,16 Upon fMLP stimulation, iNeutrophils were found to have significantly lower levels of pERK1/2 compared with primary human neutrophils (Fig. 1B). To increase activation of intracellular signaling pathways in iNeutrophils, we targeted the phosphatase PTP1B. PTP1B has been shown to limit murine neutrophil effector functions^21,22,34; however, its role in human neutrophils is unclear.

Fig. 1.

Generation of PTP1B-KO iNeutrophils. (A) Timeline for neutrophil differentiation from bone marrow–derived iPSCs. (B) Quantification of phosphorylated-ERK signaling in fMLP stimulated human PB neutrophils and WT iNeutrophils. (C) Diagram illustrates sgRNAs targeting exon 3 of PTPN1 for CRISPR/Cas9-mediated deletion of a 67 bp region at the stem cell stage. (D) Representative blot for Western blot confirmation of PTP1B CRISPR/Cas9-mediated deletion in differentiated neutrophils. (E) Representative cytospins showing morphological confirmation of neutrophil differentiation. (F) Flow cytometry staining of differentiated neutrophils and myeloid cells. Cells were gated on live cells. (G) Histogram plots of normalized flow cytometry surface receptor expression data in panel F. Data were normalized individually to each fluorescent marker and presents the range of expression. (H) Cell viability of iNeutrophils compared with 3 independent human PB neutrophil donors. The diagram in panel A was created with BioRender.com. Experiments were conducted at least 3 times, or as indicated on the plot. Dots in panel F represent 4 independent biological replicates and dots in panel H represent the average of 3 replicates. Data in panel G are the average of all data in panel F. Means ± SEM are shown. P values were calculated (F) by unpaired Student's t test or by (H) simple linear regression. *P < 0.05; ***P < 0.001. G-CSF = granulocyte colony-stimulating factor; GM-CSF = granulocyte-macrophage colony-stimulating factor; mmRNA = modified messenger RNA; ns = not significant.

To generate PTP1B-KO iNeutrophils, we mutated PTPN1, which encodes PTP1B, using CRISPR/Cas9 at the iPSC stage (Fig. 1C) and generated clonal cell lines with biallelic mutation (Supplementary Fig. 1A). Loss of PTP1B protein expression was confirmed for multiple clones both at the stem cell stage and after neutrophil differentiation by Western blot (Fig. 1D, Supplementary Fig. 1B). Cytospin preparations confirmed that PTP1B KO iNeutrophils display the hypersegmented nuclei characteristic of differentiated neutrophils (Fig. 1E). Multiple PTP1B-KO clones were tested with similar results, and we continued all further analysis with clone 6.

Further validation of neutrophil differentiation was completed by staining for neutrophil surface receptors. As previously reported, almost all iNeutrophils expressed the common myeloid marker CD11b, but <20% expressed the canonical human neutrophil marker CD66b or mature neutrophil marker CD10. However, the majority of iNeutrophils expressed primary blood and mature neutrophil markers CD15 and CD16 (Fig. 1F, G).^12,16 While deletion of PTP1B resulted in a lower proportion of fully mature CD16+ neutrophils, the majority of PTP1B-KO cells still expressed CD15 (Fig. 1F). PTP1B has been shown to modify murine myelopoiesis by negatively regulating monocyte differentiation³⁵; however, we did not find a significant increase in CD14+ monocytes upon PTP1B deletion in human iPSCs (Fig. 1F). Our data indicate that deletion of PTP1B at the stem cell stage does not prevent neutrophil differentiation.

A limitation of primary human PB neutrophils is their short lifespan ex vivo. To determine if the lifespan of iNeutrophils is longer than that of PB neutrophils and if PTP1B impacts viability, we evaluated the longevity of these cells in culture. iNeutrophils had a significantly longer lifespan than PB neutrophils, as PB neutrophils exhibited less than 30% viability at 3 d, and iNeutrophils maintained 50% viability after 5 d. Loss of PTP1B did not impact iNeutrophil viability (Fig. 1H). The lower maturation level of our INeutrophils may explain the longer lifespan of these cells compared with PB neutrophils.

3.2. Deletion of PTP1B promotes intracellular signaling, motility, and actin polymerization

Previously, we and others have reported that PTP1B regulates the actin cytoskeleton and promotes migration of cancer cells.^36,37 To determine if PTP1B also positively regulates signaling and motility of human iNeutrophils, we first quantified MAPK and PI3 K phospho-signaling associated with neutrophil migration. Following stimulation with the bacterial formylated peptide fMLP, PTP1B-KO iNeutrophils showed increased phosphorylation of ERK1/2, AKT, and the actin binding protein HS1 compared with WT iNeutrophils (Fig. 2A, B).

To investigate motility of PTP1B-KO cells, we conducted short-term live imaging of neutrophil migration in response to the chemoattractant fMLP, using a previously published microfluidic device (Video 1).³⁸ PTP1B-KO iNeutrophils displayed enhanced motility with higher chemotactic index and velocity compared with WT cells (Fig. 2C, D). Representative cell tracks show increased directed migration and track length indicative of increased persistence and speed of PTP1B-KO iNeutrophils (Fig. 2E). We also observed increased velocity of PTP1B-KO iNeutrophils under mechanical confinement with both fMLP and IL-8 (Fig. 2F, G).²⁵ While expression of the fMLP receptor FPR1 was elevated on KO cells, the IL-8 receptor CXCR1/2 was not (Supplementary Fig. 2), indicating that increased receptor expression alone is not responsible for enhanced migration of PTP1B-KO iNeutrophils.

During migration, actin is polymerized at the leading edge to drive pseudopod formation.³⁹ As elevated p-HS1 may increase actin polymerization to promote PTP1B-KO iNeutrophil chemotaxis, we next imaged F-actin polymerization in iNeutrophils after stimulation in an fMLP bath. PTP1B-KO cells displayed increased density of total F-actin compared with WT cells by immunofluorescent staining (Fig. 2H, I). Thus, deletion of the PTP1B phosphatase likely promotes neutrophil motility via increased intracellular ERK and AKT signaling and actin polymerization.

3.3. Deletion of PTP1B improves neutrophil phagocytosis but decreases production of ROS and NETs

Increased intracellular phospho-signaling may also promote neutrophil antimicrobial functions, including the ability to phagocytose microbes.²⁰ Furthermore, phagocytosis is mediated by actin polymerization to form the phagosome,⁴⁰ and thus we hypothesized that this function may be enhanced in PTP1B-KO iNeutrophils. To test this, we quantified phagocytosis of E. coli–coated beads by flow cytometry and found a significant increase in phagocytosis by CD15+ PTP1B-KO iNeutrophils compared with WT cells (Fig. 3A). This effect was heightened when gating on CD15+ CD16+ mature neutrophils with 80% of PTP1B-KO iNeutrophils phagocytosing and acidifying E. coli–coated beads, compared with fewer than 50% of WT cells. Furthermore, PTP1B-KO cells took up twice as many E. coli beads as WT cells (Fig. 3B).

Fig. 3.

Deletion of PTP1B improves neutrophil phagocytosis but decreases production of ROS and NETs. (A) iNeutrophil phagocytosis of acidified pHrodo Escherichia coli beads quantified by flow cytometry. Percent pHrodo+ cells of CD11b+ CD15+ neutrophils or CD11b+ CD15+ CD16+ mature neutrophils. (B) Quantification of median fluorescence intensity (MFI) of pHrodo+ neutrophils. Fold change calculated over WT. (C) Quantification of iNeutrophil intracellular ROS production over time using DHR123 indicator following stimulation with 50 ng/mL PMA. (D) Real-time qPCR of NADPH oxidase gene expression in unstimulated iNeutrophils or after stimulation with 200 ng/mL LPS for 2 h. Real-time qPCR samples were normalized to the WT control. (E) NETosis quantified with Sytox Green DNA indicator after 4 h stimulation with 100 ng/mL PMA. Experiments were conducted at least 3 times, or as indicated on the plot. Dots in panels A, B, D, and E represent independent biological replicates and dots in panel C represent the average of 3 replicates. Mean ± SEM are shown. P values were calculated by (A, D) unpaired Student's t test, (C) simple linear regression, or (B, E) 1-sample t test. *P < 0.05; **P < 0.001; ****P < 0.0001. ns = not significant.

Following phagocytosis, neutrophils can kill pathogens by production of intracellular ROS. iNeutrophils were previously shown to produce ROS and NETs with a similar capacity to primary human neutrophils.¹⁶ To determine the impact of PTP1B on ROS production, we stimulated cells with PMA and measured intracellular ROS production over time. PTP1B-KO iNeutrophils are capable of producing ROS, although at lower levels than WT cells (Fig. 3C). As ROS are generated by the NADPH oxidase complex,⁴¹ we quantified mRNA expression of NADPH oxidase genes and found decreased levels in PTP1B-KO neutrophils both at basal state and with LPS stimulation (Fig. 3D). Thus, decreased expression of the NADPH oxidase complex may limit ROS production by PTP1B-KO iNeutrophils.

Another antimicrobial mechanism utilized by neutrophils is the release of NETs. Upon PMA stimulation, we found that PTP1B-KO iNeutrophils make fewer NETs (Fig. 3E). ROS production regulates NET release⁴²; thus, the decreased production of NETs correlates with the decrease in ROS we observed in PTP1B-KO iNeutrophils. Thus, PTP1B affects multiple pathways involved in the neutrophil antimicrobial response.

3.4. Deletion of PTP1B increases inflammatory cytokine production

Upon migrating to sites of infection, neutrophils can produce inflammatory cytokines to further recruit and promote the innate and adaptive immune response. IL-8 is a strong neutrophil chemoattractant and activating factor. We found increased IL-8 production with both LPS and Zymosan stimulation (Fig. 4A). Increased production of IL-8 may contribute to enhanced iNeutrophil activation and migration during infection. We also found differential production of the inflammatory cytokines IL-6 and TNFα, with elevated secretion by PTP1B-KO iNeutrophils after stimulation (Fig. 4B, C). IL-1β secretion was very low and showed no difference between WT and KO iNeutrophils (Fig. 4D).

Fig. 4.

Deletion of PTP1B increases inflammatory cytokine production. (A–D) Inflammatory cytokines produced by unstimulated iNeutrophils or after stimulation with 200 ng/mL LPS or 10 µg/mL Zymosan for 4 h. Experiments were conducted 3 times. Dots represent independent biological replicates. (E) Flow cytometry staining for pathogen recognition receptors. Percent was quantified of CD15+ CD16+ iNeutrophils. Dots represent independent biological replicates. (F) Quantification of median fluorescence intensity (MFI) for pathogen recognition receptors on CD15+ CD16+ mature neutrophils. (A–F) Mean ± SEM are shown. P values were calculated by unpaired Student's t test. *P < 0.05; ***P < 0.001. ns = not significant.

To determine if increased IL-8 production by PTP1B-KO iNeutrophils is associated with altered expression of pathogen recognition receptors, we quantified expression of Dectin-1, TLR2, and TLR4 on WT and KO iNeutrophils. Zymosan is recognized by Dectin-1 and TLR2, whereas LPS is primarily recognized by TLR4.⁴³ While we found similar levels of expression of Dectin-1 and TLR4, the proportion of TLR2+ iNeutrophils was elevated in the PTP1B-KO cells (Fig. 4E). Furthermore, TLR2 expression was significantly enhanced indicating that increased inflammatory cytokine expression in response to Zymosan may be due to increased recognition (Fig. 4F). Taken together, our findings show that PTP1B-KO cells have increased production of inflammatory mediators compared with WT iNeutrophils.

3.5. Deletion of PTP1B increases iNeutrophil clustering and inhibition of fungal growth

After evaluating the effect of PTP1B deletion on specific neutrophil functions, we next wanted to determine if PTP1B expression affects iNeutrophil response to the fungal pathogen A. fumigatus as neutrophils are the primary phagocytic cell defense against this pathogen.⁴⁴ We cocultured iNeutrophils with A. fumigatus at the germling stage and then live imaged iNeutrophil-fungal interactions over the course of 8 h (Fig. 5A). We noticed a stark difference in the morphology of WT compared with PTP1B-KO iNeutrophils in the presence of A. fumigatus. During co-incubation, KO iNeutrophils displayed an elongated cell shape indicating cell activation (Fig. 5B). Specifically, PTP1B-KO iNeutrophils showed significantly decreased circularity in the presence of A. fumigatus (Fig. 5C). In contrast, the majority of WT cells maintained a rounded morphology, even after 4 h (Fig. 5B, C).

We next analyzed iNeutrophil motility in response to A. fumigatus. We found that PTP1B-KO iNeutrophils were highly migratory and rapidly recruited and clustered A. fumigatus (Video 2). In contrast, WT iNeutrophils showed a delayed response with fewer cells migrating and physically interacting with germlings or hyphae (Videos 3 and 4). Furthermore, we observed many instances of PTP1B-KO iNeutrophils phagocytosing germlings that led to enhanced neutrophil recruitment and clustering (Fig. 5D, Video 5). PTP1B-KO iNeutrophils rapidly formed tight clusters around A. fumigatus germlings and hyphae that were larger in size compared with WT cells (Fig. 5E, F). Neutrophils can amplify their recruitment by swarming, which is mediated by self-generated gradients of LTB4.⁴⁵ To determine if the clustering we observed was a swarming response, we quantified expression of the LTB4 receptor BLT1R and LTB4 release after fMLP stimulation. WT and KO iNeutrophils both expressed high levels of BLT1R and we found no difference in the levels of LTB4 produced (Supplementary Fig. 3A–C). Furthermore, addition of the LTB4 inhibitor MK886 did not prevent cluster formation (Supplementary Fig. 3D, E).

Lastly, we wanted to determine if heightened PTP1B-KO iNeutrophil recruitment to A. fumigatus limited fungal growth. We quantified the hyphal length for each germling at 0 and 4 h of co-incubation (Supplementary Fig. 3F). After 4 h, hyphal growth was significantly decreased when cultured with PTP1B-KO neutrophils, compared with WT cells (Fig. 5G). Taken together, our findings demonstrate that PTP1B-KO iNeutrophils are more responsive to A. fumigatus, resulting in enhanced recruitment and inhibition of fungal growth.

4. Discussion

iNeutrophils can exert many classic primary neutrophil functions. In vivo mouse studies using infusible iNeutrophils show promise for treating diseases ranging from bacterial infection to cancer.^{10,13–15,17} However, many of these iNeutrophils display inhibited or lower functional capacity compared with primary human neutrophils.^10,12,16,17 Therefore, there is a need to improve our understanding of the molecular signaling pathways that regulate iNeutrophil function to enhance their use as a clinical therapy. iNeutrophils can be genetically modified and thus are a valuable tool for dissecting pathways regulating cell migration and antimicrobial response. In this study, we found that the phosphatase PTP1B dampens iNeutrophil intracellular signaling resulting in inhibited motility and antimicrobial function.

PTP1B is a nonreceptor tyrosine phosphatase that targets a variety of signaling pathways including JAK/STAT, PI3 K, and Ras/MAPK.⁴⁶ Indeed, PTP1B acts as an intracellular checkpoint to negatively regulate cell responses. Specific neutrophil effector functions including migration, phagocytosis, NETosis, and cytokine production have been shown to be affected by PTP1B phosphatase activity.^21,22,34 Accordingly, we found that deletion of PTP1B increased PI3 K and MAPK signaling and improved key neutrophil functions including cell motility and phagocytosis.

Neutrophils are critical for clearance of fungal infections, including the opportunistic pathogen A. fumigatus, which commonly infects immunocompromised patients.⁴⁷ We found that PTP1B-KO iNeutrophils are significantly better at inhibiting fungal growth over WT cells, due to enhanced recruitment to A. fumigatus. Prior work suggested that PTP1B is required for migration of many cell types, including cancer cells.^34,36,37 Here we discovered a new function for PTP1B in limiting the migration of neutrophils, as deletion of PTP1B resulted in increased chemotaxis. PTP1B directly targets p38 to inhibit MAPK signaling involved in cell migration toward fMLP.⁴⁸ Indeed, MAPK signaling was increased in stimulated PTP1B-KO iNeutrophils and may promote cell motility toward other stimuli including A. fumigatus. Additionally, overexpression of PTP1B results in disorganized distribution of F-actin and focal adhesions.⁴⁹ We found that PTP1B-KO iNeutrophils had increased phosphorylated HS1 and actin polymerization after fMLP stimulation, as well as increased cell polarization in response to A. fumigatus. HS1 activates Rac-GTPase signaling and Arp2/3-mediated actin polymerization necessary for cell polarity during chemotaxis.^50,51 Thus, PTP1B may limit iNeutrophil motility through inhibition of MAPK signaling and actin organization.

During tissue damage and pathogen clearance, neutrophils produce chemotactic factors such as IL-8 and LTB4 to amplify recruitment.^19,52 Here we show that PTP1B-KO iNeutrophils are significantly better at clustering fungal hyphae than WT cells. WT iNeutrophils are capable of clustering but do so infrequently due to limited recruitment to A. fumigatus. The clustering that we observed was reminiscent of neutrophil swarming, which is regulated by self-generated production of LTB4.^19,52 However, we found no difference in the expression of the LTB4 receptor BLT1R or LTB4 release between WT and KO cells. Furthermore, the LTB4 inhibitor MK886 had no effect on PB neutrophil or iNeutrophil clustering of fungal germlings, suggesting that neutrophil response to this fungal stage is independent of LTB4. IL-8 can promote primary neutrophil clustering and inhibition of A. fumigatus growth.⁵³ Indeed, IL-8 production was increased by KO cells with Zymosan stimulation. Thus, the increased ability of PTP1B-KO iNeutrophils to cluster is likely due to elevated inflammatory signaling initiated upon interaction with fungal components.

We identified differences in receptor expression on PTP1B-KO iNeutrophils, which may contribute to increased pathogen recognition and cellular activation. Increased expression of FRP1 and TLR2 correlated to enhanced chemotaxis and antifungal response. However, PTP1B-KO iNeutrophils still exhibited greater chemotaxis toward IL-8 compared with WT despite similar CXCR1/2 levels. Additionally, LPS stimulation increased IL-8 and IL-6 production by KO iNeutrophils even with comparable TLR4 expression to WT cells. Thus, altered receptor expression cannot fully explain the increased responsiveness of PTP1B-KO iNeutrophils. Studies with murine neutrophils and macrophages have shown that PTP1B suppresses MyD88- and TRIF-dependent inflammatory signaling pathways,^21,54 supporting our data showing increased intracellular signaling in PTP1B-KO iNeutrophils.

We found that PTP1B-KO iNeutrophils are significantly better at inhibiting fungal growth over WT cells. Neutrophils limit fungal growth via phagocytosis, production of NETs and ROS, or degranulation, the release of antimicrobial proteins such as myeloperoxidase or lactoferrin.^55–58 PTP1B-KO iNeutrophils readily phagocytosed A. fumigatus germlings, correlating with the increased uptake of E. coli coated beads. Other functions less dependent on the dynamic actin cytoskeleton such as ROS and NETosis were reduced in PTP1B-KO iNeutrophils. KO cells expressed lower transcript levels of NADPH oxidase complex components which may contribute to limited ROS production and ROS-dependent NETosis. Although, WT iNeutrophils were more responsive to PMA than KO iNeutrophils, the majority of WT cells remained inactive in the presence of A. fumigatus and were not recruited to hyphae. Limited activation of WT iNeutrophils is likely due to decreased expression of pathogen recognition receptors such as TLR2 and dampened signaling mediated by PTP1B. We predict that PTP1B-KO iNeutrophils produce ROS and NETs in response to A. fumigatus due to increased interaction with hyphae. Taken together, PTP1B-KO iNeutrophils are better able to control Aspergillus growth due to enhanced recruitment and activation of antimicrobial functions.

An intriguing question is whether iNeutrophils can be used to control infection in neutropenic hosts with invasive fungal disease. Alongside administration of antifungals or antibiotics, granulocyte transfusion therapy can be utilized to improve patient outcomes. However, success of this therapy has suffered from low donor yield, as well as the short lifespan and functional capacity of primary human neutrophils ex vivo.⁵⁹ We hypothesize that iNeutrophils could serve as an alternative source of infusible neutrophils.

Further applications of iNeutrophils include use as a model system to study human neutrophil biology. iNeutrophils are derived from healthy tissue and can provide a robust source of genetically tractable human neutrophils. Additionally, iNeutrophils were shown to more closely recapitulate motility defects found in primary patient neutrophils than PLB-985 cells.⁶⁰

A caveat of our work is the heterogeneity of differentiated iNeutrophils. Our flow cytometry analysis of lineage marker expression showed that PTP1B-KO iNeutrophils are more immature and heterogeneous, with lower expression of neutrophil markers CD15 and CD16. Immature neutrophils have reduced capacity for some effector functions including NETosis and phagocytosis.^61,62 It is interesting that PTP1B-KO cells exhibit an enhanced antifungal response despite lower maturation. A recent report suggests that deletion of the transcriptional regulator GATA1 can improve the maturation and NETosis function of iNeutrophils.⁶³ Additionally, while multiple clones were tested, this work did not include a full genetic rescue of PTP1B. Re-expression of PTP1B would be informative for confirming reversal of the phenotypes observed in the KO.

In summary, we identified PTP1B as a negative regulator of iNeutrophil cellular activation and motility. PTP1B phosphatase activity is likely only one mechanism that inhibits iNeutrophil intracellular signaling and dampens effector function. Further study of the mechanisms that regulate iNeutrophil function will improve their use as a clinical cellular therapy.

Author contributions

M.A.G., D.A.B., T.J.S., A.N.P., J.H.S., and H.S.J., performed experiments. M.A.G. D.A.B., T.J.S., A.N.P., J.H.S., J.B, D.J.B., H.Q.D., I.I.S., N.P.K., and A.H. analyzed and interpreted data. M.A.G. and A.H. designed the research and wrote the article.

Supplementary material

Supplementary materials are available at Journal of Leukocyte Biology online.

Funding

The research reported in this publication was supported by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health under Award Numbers RO1 AI134749-05 and T32AI055397. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Conflict-of-interest disclosure

I.I.S. serves on Scientific Advisory Board of Umoja Biopharma.

Data sharing statement

All relevant data are included in the manuscript.