To form water-dispersible nanoparticles, the iNC must be coated with a PEGylated amphiphilic block co-polymer. To achieve this, we employed the well-established technique of Flash Nanoprecipitation (FNP),²³ which requires the dispersion of the nanocarrier in a water miscible organic solvent (such as 1,4-dioxane). As the formation of the initial iNC results in nanoparticles dispersed in chloroform along with excess spermine and triethylamine, we first performed an aqueous extraction with 150 mM sodium chloride to remove the residual 1,4-dioxane, spermine and triethylamine, and any unincorporated DNase 1. Secondly, we performed a solvent exchange with a rotary-evaporator to disperse the nanoparticles in 1,4-dioxane (Scheme 1B, Step 3 & 4). Representative intensity distributions of the iNC prior to extraction, following extraction, and the solvent exchange steps are shown in Figure 2B. Minor changes in the size distribution are observed, which may be due to the changes in the solvent quality for the polymer corona. The monodispersed particle size distribution is maintained.

To coat the iNC with PEG, the iNC was dispersed in 1,4-dioxane alongside dissolved methoxy-PEG-PLA (20 mg). This polymer/nanocarrier mixture was rapidly mixed with an antisolvent water, in a CIJ to generate PEG-coated DNase 1 nanoparticles (DNPs). A representative, monomodal trace of the final PEGylated nanoparticles is shown in Figure 2C. We show that the nanoparticles detected by DLS is not afforded by simple mixing of the free PEG-PLA micelles and iNCs as shown in Figure 2C. When mixed, the free micelle population (~20 nm) is distinguishable from the DNP particles on DLS. To make fluorescent particles, we added ~1 weight% of rubrene relative to the PEG BCP (0.2 mg), which did not significantly alter size or polydispersity of the PEG-coated DNase 1 nanoparticles, as shown in Figure 2D. Stability of the particles was measured using the correlation function via DLS over 48 hrs of incubation with the nanoparticles in 10 mM Tris-HCl buffer (Figure 2E). A summary of the sizes and polydispersity indices and physiochemical characterization of DNPs at each processing step of iFNP is shown in Table 4 and Table 5, respectively. A release assay was then performed employing Tris-HCl buffer at 37°C (Figure 2F). This assay demonstrated an initial release of ~10% of the DNase 1 from the DNPs within the first 10 h, followed by a sustained release of DNase 1 over 8 days resulting in a release of up to 80% of the content of the DNPs. Finally, to confirm particle morphology, transmission electron microscopy (TEM) images demonstrated production of spherical nanoparticles (Figure 3A).

The aqueous extraction step in the processing of the iNC allows for the extraction of unencapsulated DNase 1 to be solubilized in the aqueous phase. Thus, we can measure DNase 1 content in the aqueous phase to quantify encapsulation efficiency.²⁴ We performed control experiments where DNase 1 was mixed in the MIVM in the absence of stabilizing BCP, such that no nanoparticles were formed. Thus, we confirmed that DNase 1 could be quantitatively extracted into the aqueous phase. Overall, we employed two protein quantification methods (micro-BCA and protein absorbance at 280nm) to quantify that the encapsulation efficiency was 94 ± 1%, and 96 ± 5%, respectively. These results are summarized in Table 6. Combining the micro-BCA results with measurements by thermogravimetric analysis, we were able to measure the loading capacity of the iNC. This was determined to be 3.7 ± 0.3%. Thermogravimetric analysis also allowed us to confirm that there was no detectable loss of nanoparticle mass during the extraction step, as masses stayed the same before and post extraction (data not shown). This result, combined with the stability in the size and polydispersity of the particle through the processing steps, support that DNase 1 that is encapsulated at the iNC stage was carried through to the PEG-coated nanoparticle.

To quantify DNase 1 protein content in water-dispersible PEGylated DNPs we performed denaturing gel electrophoresis, and subsequently stained the gel with the highly sensitive protein stain, Silver Stain (Figure 3B). We found that the Silver Stain process allows for the quantitation of DNase 1 down to ~20 ng. Employing a combination of Silver Staining and thermogravimetric analysis of the total mass content of the DNP we determined that the final loading capacity was 1.09 ± 0.29%, with an average total amount of DNase 1 yield in a formulation being 169 ± 69 μg of DNase 1. To demonstrate that DNase 1 protein is present in the final preparation of aqueous stabilized DNase 1 nanoparticles, we employed a zymography assay to measure DNase activity, with a denatured DNA-containing SDS-gel. In this assay, DNA degradation will appear as a dark band and can only be achieved in the presence of an active DNA degrading enzyme. Briefly, the nanoparticles were degraded by incubating them with 0.1M NaOH, and 5% SDS over 48 hours, and then loaded into the denatured DNA-containing SDS-gel. Following protein refolding, we showed that DNase 1 activity could be demonstrated in aqueous stabilized DNase 1 nanoparticle samples (Figure 3C).

As a release assay demonstrated that DNase 1 is released by DNP (Figure 2F), we then demonstrated that the released DNase 1 is capable of degrading NETs in vitro. We generated NETs ex vivo from bone-marrow derived murine neutrophils, stimulated with PMA (Figure 4A, left panel). The NETs were then incubated with DNPs (1 U/mL of DNase 1), and NET degradation was quantified over 6 hours (Figure 4) compared to time-matched buffer only conditions. SYTOX green nucleic acid stain was used to visualize NETs and measure their degradation (Figure 4A). Impressively, we found that DNPs could achieve NET degradation within 1 hr of incubation (Figure 4B). Collectively, this provides compelling evidence demonstrating that DNase 1 is not only released from the DNPs, but that the protein has preserved functionality and can perform a therapeutic function. This finding underscores the dual achievement that DNase 1 is successfully released and retains enzymatic activity, demonstrating a potent and timely therapeutic response, holding significant implications for the therapeutic potential of this delivery system.

We next explored the potential of neutrophils as carriers for depositing DNP cargo and NPs in in vitro thrombi. Here we used both fluorescent DNP to confirm internalization by neutrophils, and rubrene/quantum-dot containing nanoparticles (QD-NP, and Ru-QD-NPs) with equivalent size and surface chemistry to DNPs for NP visualization by immunofluorescence. The synthesis and characterization of these quantum-dot containing NP are detailed in the supplementary (Supplementary Table 2, Supplementary Figures 3 & 4). Representative characterization of murine bone marrow derived neutrophils employed for these studies is shown in Supplementary Figure 6. Our findings revealed that neutrophils effectively internalize fluorescent DNPs, with greater than 80% of neutrophils being positive for fluorescent DNPs at 1.5 h post-administration (Figure 5A). Notably, the viability of neutrophils remained unaffected, indicating the non-toxic nature of the DNPs at the tested concentration and exposure time (Figure 5A). To confirm the observed signal as internalization, we compared the percentage of cells positive for fluorescent DNP at low temperatures, where active processes like internalization are hindered (Figure 5A). Confocal microscopy further validated internalization, as the DNP signal localized within the neutrophils, which bounds were delineated by the cell membrane marker CD45 (Figure 5B, 5C). This can also be visualized in Supplementary Movie 1. Similarly, Ru-QD-NP are efficiently and rapidly internalized by both murine and human neutrophils (Supplementary 7 & 8, respectively). We next showed that internalization of DNPs did not induce activation of the respiratory burst, indicated by a lack of reactive oxygen species production, measured with the de-hydro ethidium fluorescence assay (Figure 5D). Moreover, the internalization of DNPs by neutrophils did not hinder their ability to perform the oxidative burst following stimulation with either PMA or ionomycin and PMA (Figure 5D). Finally, CD11b expression in murine neutrophils remained unchanged over 90 minutes of incubation with Ru-QD-NPs (Supplementary Figure 7). Together, these data indicate that internalization of DNPs, and parameter-matched Ru-QD-NP, does not perturb important neutrophil features: viability and activation.

Next, we assessed whether DNP internalization would affect neutrophils’ capacity to bind and penetrate in vitro thrombi (Figure 6A). BMDNs, labeled with either Cell Tracker Deep Red or Green CMFDA dyes to distinguish DNP-loaded from empty cells, respectively, exhibited equivalent clot association, as quantified by flow cytometry of single cell suspensions of clot (Figure 6B) and observed with immunofluorescence (Figure 6C). To more easily visualize thrombus margins and tissue morphology, directly adjacent tissue sections of the representative thrombi in Figure 6C were stained with H&E (Supplementary Figure 9). The gating strategy and representative controls for flow cytometry can be found in the Appendix (Supplementary Figure 10). To validate whether the Cell Tracker dye itself influenced the number of neutrophils detected by flow cytometry in the in vitro clot, we performed the experiment two ways by crossing over the BMDN dyes and DNP treatment, thus: (1) Green CMFDA/DNP-loaded, Deep Red/empty; and (2) Deep Red/DNP-loaded and Green CMFDA/empty. In Supplementary Figure 11, we observe that Cell Tracker Deep Red labelling alone can result in a significant overrepresentation of the number of neutrophils detected via flow cytometry, whereas DNP internalization has no significant influence on the number of neutrophils in the in vitro clot. Next, these in vitro thrombi were stained with an anti-bovine DNase 1 antibody and imaged with both wide-field and confocal fluorescence microscopy (Figure 6D & E). These representative images demonstrated positive signal adjacent to the DNP-loaded neutrophils, indicating delivery of DNase 1 cargo to in vitro thrombi and exocytosis of DNP from the DNP-loaded neutrophil, as the signal was outside the cellular membrane boundaries (Figure 6D & E, Supplementary Figure 12).

To visualize nanoparticle delivery to in vitro clots, quantum dot-containing nanoparticles (QD-NPs) with equivalent size and surface chemistry to DNPs were employed (see Supplementary Table 2). Human peripheral neutrophils (characterized in Supplementary Figure 13) internalized QD-NPs within one hour (Supplementary Fig. 8). In vitro human clots incubated with QD-NP-loaded neutrophils demonstrated significant QD delivery compared to clots without QD-NP-loaded neutrophils (Figure 7A, C, D), confirming neutrophil-mediated nanoparticle transport to in vitro thrombi. We also corroborated that QD-NP internalization did not impact neutrophils’ association with in vitro thrombi compared to empty neutrophils (Figure 7B). Further representative fluorescent images of thrombus sections and directly adjacent thrombus sections stained with H&E, to more easily visualize tissue morphology and thrombus margins, are shown in Supplementary Figure 14. Thus, our results demonstrate that neutrophils loaded with DNPs retain their clot-binding capabilities and neutrophils can serve as efficient vehicles for nanoparticle delivery into in vitro thrombi.

Poly(D,L-lactide-block-acrylic acid) (sigma Aldrich, 802190–1G), Methoxy poly(ethylene glycol)-b-poly(D,L-lactide) (5kDa-5kDa, Sigma Aldrich, 900658–500MG), Poly(ethylene oxide)-block-poly(D,L lactide) (5kDa-5kDa, Evonik Industries), Poly acrylic acid homopolymer (Sigma Aldrich, 9003-01-04), Poly(L-lactide)-PEG-Maleimide (PLLA-PEG-Mal, 5k-5k, Nanosoft Polymers), Rubrene (Sigma Aldrich, 554073–100MG), Quantum dots (580nm emission, stabilized with oleic acid, suspended in toluene were obtained from NNCrystal US), Triethylamine (Thermo Fisher, O4884100), DNase 1 (Sigma Aldrich, 10104159001), spermine (Sigma Aldrich, AAL11956203), Chloroform (CHCl₃, Thermo Fisher, AC423550025), 1,4-dioxane (Fisher Scientific, AC408820025), DMSO (Fisher Scientific, D128–1), Methanol (HPLC Grade, VWR, EM-MX0475–1), THF (Fisher Scientific, T425–4), Glacial Acetic Acid (Fisher Scientific, A38S-500), Molecular biology grade nuclease free water (VWR, 97062–794), RPMI 1640 media (11875135, Gibco), Heat-inactivated fetal bovine serum (FBS) (16140071; Gibco), Penicillin-Streptomycin 10,000U/mL (15140122, Gibco), Phorbol 12-myristate 13-acetate (PMA, Sigma Aldrich P8139–5MG), Paraformaldehyde (158127; Sigma-Aldrich), 10X PBS (20–134; Apex Bioresearch Products, San Diego, CA), Calcium Chloride dihydrate (Fisher Scientific, C79–500), Magnesium Chloride Hexahydrate (Fisher Scientific, M33–500), Tris-HCl (Sigma Aldrich, T3253–1KG), Citric Acid (Sigma Aldrich, 251275–500G), Bovine Serum Albumin (Sigma Aldrich, A9418–5G), HBSS, no calcium, no magnesium, no phenol red (Thermo Fisher, Gibco, 14175095), Deoxyribonucleic acid sodium salt from salmon testes (Sigma-Aldrich, D1626–5G), 40% Acrylamide (Sigma Aldrich, A2792), TEMED (Sigma Aldrich, 50-197-8483), Ammonium persulfate (Sigma Aldrich, A3678).

All animal handling and experimental procedures were approved by both the Institutional Animal Care and Use Committee (IACUC) at the University of North Carolina – Chapel Hill (IACUC ID: 20–148 and 23–116) and the IACUC at Wake Forest University Health Sciences (IACUC ID: A23–148). C57Bl/6 mice (strain number:000664, age: 8–12 wk old, male) were purchased from Jackson Laboratory and were fed standard chow. Mice were allowed ad libitum access to food and water throughout the study.

This study was approved by the Institutional Review Board (IRB) of Wake Forest University Health Sciences (IRB protocol number: IRB00104255). All procedures were conducted in accordance with the 1964 Declaration of Helsinki and its later amendments. All participants provided written informed consent prior to participation. Inclusion criteria were healthy adults over the age of 18 with a weight > 110lb or 50 kg. Exclusion criteria included a history of cardiovascular disease, cancer, liver cirrhosis, diabetes, current pregnancy, chronic inflammatory conditions, or the use of anti-inflammatory or anticoagulant medications and smoking.

Venipuncture was performed by a certified nurse in the Teammate Clinic at the Biotech Place (Wake Forest University Health Sciences) using a sterile technique under standardized conditions. For each participant, approximately 10 mL of venous blood was collected from the antecubital vein using a 21-gauge butterfly needle (BD Vacutainer, Becton Dickinson, USA) and a standard vacuum collection system. Blood was drawn into an EDTA-coated tube (BD Vacutainer K2 EDTA, USA). Tubes were inverted gently 8–10 times immediately after collection to ensure proper mixing of anticoagulant with the blood.

DNase1 nanoparticles (DNPs) were synthesized via inverse flash nanoprecipitation (iFNP) with methods adapted from previous literature^{23, 26}.

DNase 1 activity in solution was measured with the commercially available fluorometric plate-reader assay from Abcam (ab23406). Briefly, solutions containing 5mg/mL of DNase 1 were diluted to 0.1–0.5 μg/mL in molecular biology grade water to be used as samples. In the case where organic solvents were used for the original suspension, a control was run containing a similar percentage of final organic solvent with control DNase 1 to ensure that this did not adversely affect the plate reader assay. Preparation of the samples and a standard curve were performed as per the protocol instructions in the product insert. Fluorescence was measured (Ex/Em = 651/681 nm) in kinetic mode, every 30 seconds, for 90 minutes at 37°C. DNase 1 activity from DNase 1 dissolved in organic solvent mixtures, or poly acrylic acid / spermine / triethylamine mixtures, was recorded as a percentage of the activity for DNase 1 from the same lot, dissolved in water.

Measurements for nanoparticle size with intensity distribution and correlation function were performed with Dynamic Light Scattering (DLS) and were carried out with a Zetasizer NanoZS (Malvern Panalytical, Ltd, UK). Size measurements were conducted at a 173-degree scattering angle, with the appropriate settings for the solvent. Measurements were conducted at least 3 times with 30 s to 240 s of equilibration time. For measurements in organic solvents, we employed the Malvern Panalytical Sq Cell W Cap 12 mm O.D. glass cuvette (Fisher scientific, NC0631452), whilst measurements in aqueous solvents were recorded in Malvern Panalytical Inc 40μL Cuvettes (Fisher Scientific, NC0628994). Samples were typically diluted 10-fold for measurements into appropriate solvents (Chloroform, 1,4-dioxane or 0.02 μm filtered water or 1X PBS).

Conventional TEM images were captured on a JEOL JEM 1230 TEM at 80 kV, tungsten filament, with a Gatan Orius SC1000 CCD camera and Gatan Microscopy Suite 3.0 software. Rubrene-DNase1-NPs in water were prepared for TEM by pipetting 8 μL atop Formvar on carbon mesh TEM grids that were treated with glow discharge (PELCO easiGlowTM Glow Discharge Cleaning System). After two minutes, samples were rinsed with deionized water and stained with 2% uranyl acetate negative stain for two minutes prior to imaging.

Encapsulation Efficiency was broadly determined by measuring the amount of DNase 1 protein recovered into the aqueous extract phase following formation of the iNC. INCs containing DNase 1 were synthesized as described, but in the absence of spermine and triethylamine, and then extracted with 0.15M NaCl solution. Controls included: iNCs without DNase 1 added, DNase 1 alone with no stabilizing polymer added, aqueous extract buffer alone. The Encapsulation Efficiency was calculated using the following equation: EncapsulationEfficiency=100-(100×DNase1inverseDNase1max)

DNase 1_inverse = the amount of DNase 1 recovered into the aqueous extract phase following formation of DNase 1 containing iNCs

DNase 1_max = the amount of DNase 1 recovered into the aqueous extract phase following DNase 1 passing through the MIVM with no stabilizing BCP added (no iNC formed).

The aqueous extraction phase was measured for protein using the commercially available micro BCA assay (Thermo Fisher, 23235). Instructions as per the user’s manual were followed and sample absorbances were measured at 562 nm. Measurements were made in triplicate with three independent batches of nanoparticles synthesized.

A second, orthogonal, measure was utilized to confirm Encapsulation Efficiency. We measured protein absorbance at 280 nm using the Duetta Fluorescence and Absorbance Spectrometer (Horiba Scientific). Measurements were made in triplicate with three independent batches of nanoparticles synthesized.

To determine the Loading Capacity and loss to extraction of the iNC we used a Thermogravimetric Analyzer (TGA, TA instruments). INCs containing DNase 1 were synthesized as described, but in the absence of spermine and triethylamine, and then extracted with 0.15M NaCl solution. 90 μL of the iNC solution in chloroform was added to the TGA pan prior to and following the extraction step, and the dry mass recorded. Loading capacity was determined by employing the following equation: LoadingCapacity(inversenanocarrier)=DNase1max×EENanoparticleMass×100

DNase 1_max = the amount of DNase 1 recovered into the aqueous extract phase following DNase 1 passing through the MIVM with no stabilizing BCP added (no iNC formed).

EE = Encapsulation Efficiency

Nanoparticle Mass = total dry mass of the nanoparticle as calculated from measurements with the TGA. Mass in 90μL was extrapolated to calculate the total mass based on the total volume of the nanoparticle suspension.

Loading of the final aqueous stabilized PEG-coated nanoparticle was confirmed by gel electrophoresis followed by Silver Stain. Briefly, following dialysis, nanoparticles were diluted with 6X SDS loading buffer (Fisher Scientific, 50-196-785) and boiled (10 min, 90°C). A series of DNase 1 standards were similarly treated. These were loaded into a Sodium dodecyl sulfate (SDS)—polyacrylamide gel (4% (v/v) stacking gels and 10% (v/v) running gels). Electrophoresis was carried out at 120 V using Tris/glycine electrophoresis buffer (25 mM Tris, 192 mM glycine, 0.1% (w/v) SDS, pH 8.7), just until the protein band had surpassed the stacking gel. To stain for the protein content, we used the commercially available Silver Stain Plus Kit, and followed the protocol instructions supplied (Biorad, 1610449). Gels were imaged using an Azure Biosystems Gel Doc imaging system.

DNase 1_{(silver stain)}= the amount of DNase 1 as measured by silver stain of a dispersion of DNPs in water. Nanoparticle mass = total solid mass of a dispersion of DNPs in water via thermogravimetric analysis

We performed this assay as previously described in Jimenez-Alcázar and co-authors⁸³, and Rosenthal and Lacks⁸⁴, with adaptations. Briefly, nanoparticles were degraded by diluting them 2-fold into a solution with a final concentration of 5% SDS and 0.1M NaOH, and then stirred rapidly at room temperature for 48 hrs. This sample was then diluted in water to represent ~100 pg of DNase 1. DNase 1 standards were prepared in water representing 51, 102 and 204 pg of DNase 1. The sample and standards were diluted with 6X SDS sample loading buffer, and then boiled for 10 minutes before loading onto the gels.

Sodium dodecyl sulfate (SDS)—polyacrylamide gels were prepared with 4% (v/v) stacking gels without DNA and 10% (v/v) running gels containing 100 μg/ml of denatured salmon testes DNA. DNA was first dissolved in water (30–60’ sonication) then boiled (10 min, 90°C), and placed on ice before addition to gel reagents. Electrophoresis was carried out at 120 V using Tris-glycine electrophoresis buffer (25 mM Tris, 192 mM glycine, 0.1% (w/v) SDS, pH 8.7). After electrophoresis, SDS was removed by washing the gel twice with 10 mM Tris/HCl pH 7.8 for 30 min at 50°C. Proteins were refolded by incubating the gels overnight at 37°C in a re-folding buffer containing 5% (w/v) milk powder, 10 mM Tris/HCl pH 7.8, 3 mM CaCl2, 3 mM MgCl2, 100 U/ml penicillin, and 100 μg/ml streptomycin. The gels were transferred to a refolding buffer without milk powder (10 mM Tris/HCl pH 7.8, 3 mM CaCl2, 3 mM MgCl2, 100 U/ml penicillin, and 100 μg/ml streptomycin) and incubated for additional 24 hrs at 37°C. The gels were then stained with for DNA with SYBR^™ Safe DNA Gel Stain in 1X TAE (Life Technologies, S33111). Fluorescent images of the gels were recorded using a molecular gel imager.

DNPs were dispersed into 10 mM Tris-HCl buffer containing 3 mM MgCl₂ and CaCl₂ and 2–3 mL placed into Tube-O-Dialyzer (G Biosciences, VWR, Cat#: 95057–606, MWCO 50K). They were then dialyzed while shaking at 37°C for up to 8 days against a volume of 30 mL of buffer. The dialysate was then reduced to 0.5mL using a passivated Pall Centrifugal Filter (MWCO 3K). The centrifugal filters were passivated overnight with 5% Tween-20 in water (0.22um filtered) and then washed copiously (10x volume) with 0.22um filtered water, then left in 1X PBS until use. The dialysate volume was measured with a mass balance to determine an accurate volume, and the protein content measured with the microBCA assay. The protein content in the dialysate was calculated as a % of the initial amount of protein in the DNPs prior to the release assay.

Bone marrow derived neutrophils were isolated as previously described with minor alterations. Briefly, the femur and tibia of 10–14-week-old C57bl/6 mice were excised, and the bone marrow collected into cold HBSS prior to red blood cell lysis with hypotonic sodium chloride (0.2%), followed by addition of NaCl solution (1.6%). Cells were strained (40 μm nylon cell strainer) and then pelleted (300 g, 5min) and resuspending in HBSS. Cells were counted and then neutrophils were isolated by magnetic separation with a mouse neutrophil isolation kit (Miltenyi, 130-097-658). The instructions were followed as per the kit protocol. A Midi Macs separator (Miltenyi, 130-042-302) with LS columns (Miltenyi, 130-042-401) was employed. Neutrophils were finally resuspended at ~1×10⁶ cells/mL in 1X RPMI (no serum or antibiotics added). The typical yield from 1 mouse was ~30×10⁶ bone marrow cells and ~5×10⁶ neutrophils, with 98% viability. Cells were checked for initial cell count and viability with the Muse Count and Viability kit (Sigma Aldrich, Luminex MCH100104). Neutrophils were always used fresh and within 2 hrs of isolation, they were kept at 4°C until use.

Human neutrophils were isolated directly from venous blood by magnetic separation with the EasySep^™ Direct Human Neutrophil Isolation Kit. The instructions were followed as per the kit protocol. An Easy 50 magnet (Catalog #18002) with 50 ml tube was employed. Cell count and viability were measured with the Nexcelom ViaStain AO/PI Solution (CS2-0101-5ml). Following isolation, neutrophils were maintained in 1X RPMI media. Neutrophils were used fresh, within 2 hr of isolation. They were kept 37°C until use.

Flow Cytometry was performed on the Thermo Fisher Attune NxT instrument and data acquired on the Attune NxT 2.6 software. Data was analyzed using FlowJo software. For staining we employed Anti mouse CD45-PeCy5 (Fisher Scientific, 15-0451-81, 1:3000, final ~0.07ug/mL) and the Violet live dead stain (Thermo Fisher, L34963, 1:5,000). Cells were gated in the order of cell granularity/size, singlet events, live cells, CD45⁺, rubrene⁺.

For single cell suspensions of blood clots, cells were gated in the order of singlet events, DAPI+ve events, Ly6G positivity (PE), and finally either Green-CMFDA fluorescence or Deep Red Fluorescence. Exogenous neutrophils had been stained with Anti-ly6G PE (Thermo Fisher, Clone 1RB6–8C5, Cat#: 12-5931-85, 1:1500 dilution, 0.13μg/mL).

This method was adapted from previous literature⁵⁰. Briefly, 96 well U-bottom microplates (08-772-54, Fisher scientific) were treated overnight with 1% BSA/1XPBS. On the day of the experiment, 100,000 murine neutrophils in RPMI media (no serum) at 1×10⁶ cells/mL were added to the treated wells. 10μL of rubrene-DNase 1-NPs (final concentration of ~0.2μg/mL rubrene) were added to the neutrophils and incubated either at 37°C with 5% CO2 and humidity or 4°C or up to 2 hours. Neutrophils were pelleted (300 g, 5min), and washed with flow cytometry staining buffer (0.5% BSA in 1X PBS). Cells were pelleted and resuspended in 1X PBS and stained with the violet live/dead stain for 30 minutes, protected from light at room temperature. Cells were washed and then stained with CD45 antibody for 30 minutes at 4°C. Cells were washed and then fixed with 1% PFA in 1X PBS for 15 minutes at 4°C. Cells were stored overnight in 1X PBS at 4°C and analyzed by flow cytometry within 48 hours.

Oxidative burst of bone marrow derived neutrophils was measured using the Abcam DHE ROS kit (Cat#Ab236206). Neutrophils (1×10⁶ cells/mL, RPMI) were incubated in the presence or absence of DNase1-nanoparticles (3.4U/mL) for 2 hrs at 37⁰C, 5% CO₂, and then pelleted and washed to remove unincorporated nanoparticles. These neutrophils were subsequently stimulated with PMA (100 nM) or PMA (100 nM) and ionomycin (1 μM) for 30 minutes at 37⁰C, 5% CO₂. The instructions for the Abcam kit were then followed to stain these cells for ROS production.

NET degradation was analyzed as previously described^{83, 85}, with minor alterations. 50,000 purified murine neutrophils (250,000 cells/mL) in serum-free F12 media were seeded onto sterile 96 well glass bottom plates (VWR, 655891). To induce NET formation, neutrophils were treated with 100 nM PMA for 4 hrs at 37°C with 5% CO2 and humidity. Wells were then washed three times with 1X PBS and stored in 1X PBS overnight at 4°C. DNase 1 or DNase 1-NPs at ~1U/mL of DNase 1 were added to wells in Tris-HCl, 10mM pH 7.8, containing 3mM CaCl₂ and 3 mM MgCl₂. Control wells were incubated with buffer alone. NET degradation was allowed to occur for up to 4 hrs. To stop NET degradation, wells were washed and treated with cold 2mM EDTA for 15 min at 4°C. Well were washed, and then NETs fixed with 2% PFA in 1X PBS for 15 min in the fridge. Wells were washed and DNA stained with 2μM Sytox Green (Thermo Fisher, S7020) for 15 min at room temperature. Wells were washed and stored until imaging in 1X PBS, protected from light at 4°C. Entire wells were imaged for GFP fluorescence with High Content Analysis Nikon Elements software using an inverted fluorescence microscope (Nikon Ti2 eclipse, Objective: Plan Apochromat Lambda 10X, NA = 0.45, WD = 4mm). Analysis was performed in Image J. Firstly, a standardized well area was drawn that excluded the very edges of the well, and this region was duplicated. The same area was applied to all wells. The fluorescence was thresholded and the area of the thresholded fluorescence was measured. To restrict measurements to the filamentous NET structures, circular particles were counted, and the area of the particles measured. Area from spherical particles was deducted from the total initial area. The remaining area was considered to be that of filamentous NET structures.

To generate the pre-formed blood clot, the method of Kattula et al. was modified slightly⁸⁶. Briefly, venous blood was drawn from healthy male C57bl/6 mice (8–12 wk old) into 4% sodium citrate. Blood was then recalcified (final concentration 30mM) and treated with tissue factor (TF; Innovin, diluted 1:9000 [1 pM TF]) and clot formation proceeded in siliconized wells for 1 h at 37°C. Bone marrow derived neutrophils from C57bl/6 mice (8–12 wk old) were isolated and labelled with 1μM Cell Tracker Deep Red (Thermo Fisher, Catalog #: C34565) or Green-CMFDA (Thermo Fisher, Catalog #: C7025) according to manufacturer’s instructions. Green or Deep Red labelled neutrophils were then loaded with DNPs (5U DNase 1/mL) for 2h at 37°C, 5% CO₂. Following incubation and washing, neutrophil count was standardized so that empty and DNP-loaded neutrophils were at 12 million cells/mL. Subsequently both empty and DNP-loaded neutrophils were added to the same pre-formed clot at equal concentrations and allowed to associate with the clot overnight (18 h). Preparation of single cell suspension: The clot was washed with 1X cold HBSS, and then ground into 10 mL of 1X cold HBSS and pelleted. The pellet was resuspended in warm DNase 1 digest solution (60U/mL DNase 1, 1X PBS 3mM MgCl₂) and incubated for 30 min at 37°C, 5% CO₂. The digest of free extracellular DNA was halted by the addition of EDTA to a final concentration of 2 mM in PBS. Following pelleting and washing with flow cytometry staining buffer, the cells were then stained with Anti-ly6G PE (Thermo Fisher, Clone 1RB6–8C5, Cat#: 12-5931-85, 1:1500 dilution, 0.13μg/mL). for 30 min at 4°C. Red blood cells were then lysed and the remaining single cell suspension cells were fixed using RBC Lyse-&-Fix (Thermo Fisher, Catalog #: 00-5333-54). The single cell suspension was then stained with DAPI (300 nM) for 15 min, room temperature. Histological sections: The clot was washed with 1X cold HBSS, and then fixed with 4%PFA/1X PBS for 1 h at 4°C. The clot was then placed in 30% sucrose/1XPBS for 1–2 h at 4°C, and then frozen in OCT freezing medium over liquid N₂ (Fisher Scientific, Catalog #: 23-730-625). 10μm sections were taken with a cryostat at -20°C. Tissue sections were washed with 1X PBS and stained with DAPI (300 nM) for 5 min, room temperature, and subsequently mounted in ProLong Gold AntiFade Mounting Medium (ThermoFisher Scientific, Catalog #: P36930), with #1 thickness cover slips (VWR, Catalog #: 16004–098). Fluorescent microscopy images were acquired on a Nikon Ti2 eclipse with Nikon NIS Elements AR software (Objective: Plan Apochromat Lambda 10X, NA = 0.45, WD = 4mm; and Plan Fluor 40X, NA = 0.75, WD = 0.66mm, Camera: pco.edge 4.2Q High QE sCMOS). Fluorescence was detected on the DAPI, GFP and Narrow Cy5 channels. Image analysis was performed on Image J (FIJI).

Data was graphed using Graphpad Prism 10 software (v10.2.2, Graphpad Software LLC). Data was analyzed with Graphpad Prism 10 software or SAS (9.4). To compare the means between multiple groups we employed factorial ANOVA. Ratios, which cannot be normally distributed, were analyzed using a beta-distributed generalized linear model. Percentages were log transformed and then analyzed via factorial ANOVA followed by Sidak Multiple Comparisons post-hoc test.