This study was reviewed and approved by the University of Florida Institutional Review Board (IRB201500437) and the University of Sydney Human Research Ethics Committee (2018/1017). Participants provided written informed consent to participate. Blood was initially collected from healthy volunteers to establish and optimize assay parameters, and then 15 subjects with PD and 13 age-matched, neurologically normal control subjects were recruited through the University of Florida for this validation study. Subjects were excluded based on age (younger than 50 and over 85 years of age), known familial PD mutations and/or other known neurological, chronic or recent infections, or autoimmune comorbidities. Subjects were genotyped for the G2019S LRRK2 mutation (Life Technologies #4351378, Grand Island, NY) and excluded from this study if they were shown to be mutation carriers.

During recruitment, a family history and environmental questionnaire was used to assess history of disease and inflammation/immune-relevant environmental exposures and comorbidities. Caffeine, non-steroidal anti-inflammatory drug, and nicotine exposure was calculated as milligram-years, milligram-years, and pack-years, respectively. The study populations were balanced with respect to risk factors for PD, including age, smoking, non-steroidal anti-inflammatory drug use, caffeine intake, and rs3129882 (HLA-DRA SNP) genotype (Table 1).

For the cell isolation by SepMate, 10 mL of blood was collected from healthy volunteers per BD Vacutainer^® spray coated K2EDTA Tube (BD Vacutainer^®, 367863). 15 mL of Ficoll-Paque PREMIUM (GE Healthcare, 17-5442-02) was added to each SepMate™ tube (STEMCELL, 85450, 50 mL capacity) through the central hole of the SepMate™ tube insert. Blood samples were diluted with an equal volume of Dulbecco’s phosphate-buffered saline (PBS) (ThermoFisher, 14190094) containing 2% fetal bovine serum (FBS; Atlanta Biological, S11150) and 30 mL of diluted sample was added to each SepMate™ tube. Loaded SepMate™ tubes were centrifuged at 1200 x g for 10 min at room temperature, after which the resulting plasma layer was aspirated off and enriched PBMCs transferred into a 15-mL Falcon tube. PBMCs were washed x 2 with 12 mL PBS containing 2% FBS and centrifuged at 1000 x g for 2 min at 4°C. The PBMCs were gently resuspended in 10.5 mL PBS with 2% FBS and counted using a hemocytometer with Trypan blue (ThermoFisher, T10282) at a 1:20 dilution.

For the cell isolation by CPT vacutainer tubes, 8 mL of blood was collected from healthy volunteers per BD Vacutainer CPT Cell Preparation Tube with Sodium Citrate (BD Biosciences, 362761). CPT tubes were inverted 8–10 times and centrifuged at room temperature at 1500 x g for 20 min at room temperature. The PBMC enriched layer was transferred to a new 50-mL conical tube and MACS buffer (PBS, 0.5% bovine serum albumin, 20 mM EDTA, pH 7.2) was added to a final volume of 50 mL, followed by centrifugation at 1800 x g for 10 min at room temperature. Following removal of the supernatant, PBMCs were resuspended in 10 mL MACS buffer and counted using a hemocytometer with Trypan blue at a 1:20 dilution.

To isolate monocytes from PBMCs, a CD14 MicroBeads kit (Miltenyi Biotech, 130-050-201) was used. PBMCs were resuspended in 80 μL MACS buffer per 1 × 10⁷ cells in a 1.5-mL Eppendorf tube. 20 μL of human CD14 Microbeads per 1 × 10⁷ cells was added to the cell suspension. Samples were mixed well and incubated at 4°C in the dark for 15 min. Cells were washed by adding 1 mL MACS buffer and centrifuged at 300 x g for 10 min at 4°C. Supernatant was aspirated and cells resuspended in 500 μL of MACS buffer. MS columns (Miltenyi Biotech, 130-042-201) were placed on the MACS MultiStand (Miltenyi Biotech, 130-042-303) and prepped by adding 500 μL MACS buffer to wet. Cell suspension was added to the MS column and unlabeled cells that passed through were collected. Columns were washed x 3 with 500 μL MACS buffer. Columns were removed from the magnetic stand and 500 μL MACS buffer was added to the column reservoir. Magnetically labeled monocytes were immediately flushed out using the column plunger and collected in a 1.5-mL Eppendorf tube. Monocytes were counted using a hemocytometer with Trypan blue at a 1:20 dilution.

Peripheral blood mononuclear cells or monocytes were centrifuged for 10 min at 300 x g at 4°C. Supernatant was aspirated and cell pellets were gently resuspended in cryopreservation media (RPMI 1640, with either 6, 40, or 90% FBS) at a final concentration of 1 × 10⁷ cells/mL in cryovials (Simport, T311-2). DMSO was slowly added to each cryovial to a final concentration of 10% and cryovials placed in a room-temperature Mr. Frosty freezing container with isopropanol as per manufacturer’s instructions and stored at −80°C overnight. Cryovials were removed from freezing containers and immediately placed into liquid nitrogen for long-term storage. For cryorecovery, PBMCs or monocytes were retrieved from liquid nitrogen, thawed at 37°C, slowly added to 37°C filter sterilized complete culture media (RPMI 1640 media, 10% low endotoxin heat-inactivated FBS, 1 mM Penicillin-Streptomycin) and pelleted via centrifugation at 300 x g for 10 min at room temperature. Supernatant was removed and cells were resuspended in 37°C complete culture media for cell counting using Trypan blue.

Peripheral blood mononuclear cells were diluted to a final concentration of 1 × 10⁶ per mL in complete culture media and allowed to rest for 2 h at 37°C, 5% CO2, 95% relative humidity. After resting, cells were treated with either vehicle, 100 nM MLi-2 (Tocris, 5756), or 50 μM Conduritol B epoxide (CBE; Sigma, C5424), each in the presence and absence of 100U human IFN-γ (Peprotech, 50813413), for 18 h at 37°C, 5% CO₂, 95% relative humidity.

After the 18-h stimulation, cells were harvested and centrifuged at 300 x g for 5 min at 4°C. Cell pellets were resuspended in 100 μL of media containing the same inhibitors as treated with overnight and transferred to a v-bottom 96-well plate (Sigma, CLS3896-48EA). PFB-FDGlu [5-(Pentafluorobenzoylamino)-Fluorescein Di-β-D-Glucopyranoside] (Thermo Fisher, P11947) was added to each well to a final concentration of 0.375 mM and BMV109 Pan Cathepsin probe (Vergent Bioscience, 40200-200) was added to each well to a final concentration of 1 μM for 1 h at 37°C in the dark. Samples were centrifuged at 300 x g for 5 min at 4°C. Cell pellets were resuspended in PBS and washed x 2 by centrifuging at 300 x g for 5 min at 4°C. Cells were resuspended in 100 μL of Live/Dead stain (diluted 1:2000 in PBS, Invitrogen, L34965) and incubated in the dark at room temperature for 30 min. Cells were centrifuged at 300 x g for 5 min at 4°C washed in PBS x 2. Cells were resuspended in 50 μL of PBS containing diluted antibodies (see Table 2) including inhibitors (MLi-2 or CBE) and incubated in the dark at 4°C for 20 min. Cells were centrifuged at 300 x g for 5 min at 4°C washed in FACS buffer (PBS, 0.5 mM EDTA, 0.1% sodium azide) x 3. At UF, cells were taken for flow cytometry on a Macs Quant Analyzer (Miltenyi); at USYD, cells were taken for flow cytometry on a Cytek Aurora spectral analyser (Cytek Biosciences). A minimum of 100,000 events were captured per sample and data were analyzed using FlowJo version 10.6.2 software (BD Biosciences).

After the 18-h stimulation, cells were harvested and centrifuged at 300 x g for 5 min at 4°C. Cell pellets were resuspended in 100 μL of PBS containing either LRRK2 or GCase inhibitors (MLi-2 or CBE, respectively) according to the 18-h treatments and transferred to a v-bottom 96-well plate (Sigma, CLS3896-48EA). Samples were centrifuged at 300 x g for 5 min at 4°C. Cells were resuspended in 100 μL of Live/Dead stain (diluted 1:2000, Invitrogen, L34965) and incubated in the dark at room temperature for 30 min. Cells were centrifuged at 300 x g for 5 min at 4°C, and then washed in PBS x 2. Cells were resuspended in 50 μL of PBS containing diluted fluorophore-conjugated antibodies (see Table 2) including LRRK2 or GCase inhibitors (MLi-2 or CBE) and incubated in the dark at 4°C for 20 min. Cells were centrifuged at 300 x g for 5 min at 4°C and washed in PBS x 2. Cells were fixed in 50 μL of 4% paraformaldehyde (PFA) at room temperature in the dark for 10 min. Cells were centrifuged at 300 x g for 5 min at 4°C washed in PBS x 2. Cells were resuspended in 100 μL of permeabilization buffer (eBiosciences, 00-8333-56) and incubated on ice for 15 min. Anti-pT73 Rab10 antibody (Abcam, ab241060) was added to each well at 0.55 μG per well and incubated at 4°C for 45 min with rocking. Cells were centrifuged at 300 x g for 5 min at 4°C washed in PBS x 2. Cells were resuspended in 100 μL of PBS containing 1% normal goat/donkey serum, 2% BSA and 1:1000 AF488 donkey anti-rabbit secondary (Thermo Fisher, A-21206) and incubated at 4°C for 30 min with rocking. Cells were centrifuged at 300 x g for 5 min at 4°C washed in PBS x 2. Cells were resuspended in 100 μL of PBS containing 1% normal goat/donkey serum, 2% BSA 1:100 anti-LRRK2 AF700 antibody and incubated at 4°C covered for 20 min. Cells were centrifuged at 300 x g for 5 min at 4°C, and then washed in FACS buffer × 3. At UF, cells were taken for flow cytometry on a Macs Quant Analyzer (Miltenyi); at USYD, cells were taken for flow cytometry on a Cytek Aurora spectral analyser (Cytek Biosciences). A minimum of 100,000 events were captured per sample and data were analyzed using FlowJo version 10.6.2 software (BD Biosciences). When validating flow cytometry panels and antibodies, fluorescence minus one controls (FMOCs) were used to set gates and isotype controls were used to ensure antibody-specific binding.

V-PLEX human pro-inflammatory panel 1 kit (MSD; K15049D) was used to quantify cytokines in conditioned media from PBMCs. Media was diluted 1:1 with MSD kit diluent and incubated at room temperature in the provided MSD plate with capture antibodies for 2 h as per manufacturer’s instructions. Plates were then washed x 3 with PBS with 0.1% Tween-20 and detection antibodies conjugated with electrochemiluminescent labels were added and incubated at room temperature for another 2 h. After 3 x washes with PBS containing 0.1% Tween-20, MSD buffer was added and the plates were loaded into the QuickPlex MSD instrument for quantification.

Peripheral blood mononuclear cells were collected, pelleted via centrifugation at 300 x g at 4°C, washed in PBS and centrifuged at 300 x g at 4 degrees and lysed in Lysis buffer [50 mM Tris-HCl pH 7.5, 1% (v/v) Triton X-100, 1 mM EGTA, 1 mM Na3VO4, 50 mM NaF, 10 mM β-glycerophosphate, 5 mM sodium pyrophosphate, 0.27 M sucrose, 1X complete (EDTA-free) protease inhibitor cocktail (Roche), 1 μg/mL Microcystin-LR, 0.5 mM diisopropylfluorophosphate (DIFP)]. Cell lysates were then centrifuged at 14,000 rpm for 15 min at 4°C. 6X Laemmli sample buffer added (12% SDS, 30% β-mercaptoethanol, 60% Glycerol, 0.012% Bromophenol blue, and 375 mM Tris pH 6.8) and samples were reduced and denatured at 95°C for 5 min. Samples were loaded into 4–20% Criterion Tris-HCl polyacrylamide gels (BioRad) alongside Precision plus protein dual-color ladder (Biorad) to determine target protein molecular weight. Electrophoresis was performed at 100 V for ∼60 min and proteins transferred to a polyvinylidene difluoride (PVDF) membrane using a Trans-Blot Turbo Transfer System (BioRad) which utilizes Trans-Blot Turbo Midi PVDF transfer packs (BioRad) in accordance to manufacturer’s instructions. Prior to blocking, total protein was measured using Revert total protein stain (Licor) and imaged on the Odyssey FC imaging system (Licor). Membranes were then blocked in 5% non-fat milk in TBS/0.1% Tween-20 (TBS-T) for 1 h at room temperate and subsequently incubated with primary antibody in blocking solution overnight at 4°C. Membranes were washed with TBS-T (3 × 5 min) and incubated in horseradish peroxidase (HRP)-conjugated secondary antibody (1:1000) (BioRad) in blocking solution for 1 h. Membranes were washed in TBS-T (3 × 5 min) and developed using Super signal west femto/pico (Thermo). Membranes were imaged using the Odyssey FC imaging system and quantified using Image Studio Lite Version 5.2 (Licor).

Data and statistical analyses were performed using IBM SPSS statistics 27 or GraphPad Prism 9. For assessing differences between groups, data were analyzed by either 1-way or 2-way analysis of variance (ANOVA), or by t-test. In instances when data did not fit parametric assumptions, Kruskal–Wallis non-parametric ANOVA was used. Post-hoc tests following ANOVAs were conducted using Tukey HSD or Bonferroni correction. For assessing relationships between read-outs, data were analyzed by Pearson’s r. In instances when data did not fit parametric assumptions, Spearman’s rank was used to assess relationships between variables. Two-tailed levels of significance were used and p < 0.05 was considered statistically significant. Graphs are depicted by means ± standard error of the mean (SEM).

To define the optimal protocol to assess LRRK2 and GCase enzymatic activity simultaneously on a large number of samples in two separate laboratories across the globe, we started by assessing two different isolation methods of human PBMCs. To this end, we compared the cell count and viability of PBMCs collected and isolated from blood using CPT or SepMate tubes in order to determine the best cell isolation method. This was conducted across two institutes (University of Florida and University of Sydney) in order to assess the reproducibility of these outcomes. Using flow cytometry to compare PBMC isolation protocols it was found that monocyte viability (Figures 1A–C) and purity (Figures 1D,E) were superior when CPT tubes were used and site (operator) independent. Therefore, we recommend the use of CPT tubes in WHOPPA.

To determine optimal cryopreservation conditions for PBMCs and monocytes, we compared 6, 40, and 90% serum in the cryopreservation media to determine effects on cell viability and purity of cryopreserved PBMCs and monocytes. For PBMCs the levels of serum in the media did not significantly affect viability upon thawing at both research sites (Figures 1F,G). Based on these data, we conclude that serum in the range of 6–40% was best and thus proceeded with 20% FBS in the cryopreservation media in WHOPPA.

Cryopreserved cells have the added advantages of allowing for multisite collection followed by central analysis, as well as reducing batch effects previously seen using fresh cells and different reagents over time. However, the effects of cryopreservation on readouts such as LRRK2 and GCase enzymatic activity had to be determined. By using the optimized protocols to isolate, cryopreserve, thaw and stimulate PBMCs, we assessed the effects of cryopreservation on stimulation-dependent changes in LRRK2 levels and kinase activity, as assessed by levels of pRab10, in classical and non-classical monocytes gated from total PBMCs. A significant effect of cryopreservation on pRab10 was observed in both classical and non-classical monocytes, with an increased effect of IFN- γ stimulation observed in cryopreserved cells (Figures 2A,B). Despite this, a significant increase in pRab10 signal was observed in both fresh and cryopreserved monocytes, which was decreased with MLi-2 treatment. No significant changes in total Rab10 or pRab10 protein were observed biochemically in lysates from CD14⁺ monocytes (Figure 2C). In addition, no significant effects of cryopreservation on LRRK2 levels were observed in classical monocytes, with IFN-γ significantly increasing LRRK2 levels in both fresh and cryopreserved cells (Figure 2D). However, a significant effect of cryopreservation was observed in non-classical monocyte LRRK2 levels, with cryopreservation significantly decreasing LRRK2 levels (Figure 2E). This was confirmed biochemically in total CD14⁺ monocytes via Western blot (Figure 2F). These results indicate that, despite cryopreservation having a significant effect on some of the LRRK2-related readouts assessed here at baseline, changes could still be observed and are therefore an appropriate readout for cryopreserved monocytes.

To determine if cryopreservation alters GCase levels and enzymatic activity, and if stimulation-dependent changes could still be observed post-cryopreservation, we also assessed the effects of cryopreservation on stimulation-dependent changes in GCase activity. We assessed median fluorescence intensity (MFI) of the GCase activity probe, PFB-FDGlu, in classical and non-classical monocytes gated from total PBMCs. The GBA index was calculated to quantify GCase activity as previously described (Atashrazm et al., 2018). A significant effect of cryopreservation on GBA indices was observed in both classical and non-classical monocytes, with an increase and decrease observed, respectively (Figures 2G,H). Despite this, when stimulated with IFN-γ, an increase in GBA index was observed in both fresh and cryopreserved monocytes (Figure 2I). These results indicate that, despite cryopreservation having a significant effect on GCase activity levels at baseline, stimulation-dependent changes could still be observed and are therefore an appropriate readout for cryopreserved monocytes.

To assess the effects of cryopreservation on cytokine release from PBMCs, we plated PBMCs and collected conditioned media after an 18-h resting period. Using multiplexed immunoassays on conditioned media from PBMCs, we observed that cryopreservation altered the secretion of IL-1β and IL-13, however had no significant effect on other cytokines (Supplementary Figure 1), suggesting that cytokine release ex vivo is an appropriate readout to assess baseline inflammation in PBMCs after cryorecovery.

To ensure our ability to use WHOPPA to measure LRRK2 enzymatic activity, we first assessed the specificity of the pRab10 antibody for human pRab10 using flow cytometry applications. The MJF-R21 Abcam pRab10 antibody is a rabbit monoclonal antibody used in the studies herein to detect human Rab10 protein phosphorylated at T73 in human primary peripheral monocytes gated from total PBMCs (Supplementary Figure 2). Because the pRab10 antibody was not conjugated to a fluorophore, an Alexa Fluor 488 (AF488)-conjugated secondary anti-rabbit IgG antibody was used for the flow cytometry studies. We performed experiments in which total PBMCs were plated in the presence or absence of the LRRK2 kinase inhibitor, MLi-2. As Rab10 is a bona fide LRRK2 substrate, we expected a decrease in pRab10 signal via flow cytometry. A significant decrease in pRab10 signal by flow cytometry was observed with three different concentrations of MLi-2 in pRab10⁺ classical monocytes (Figure 3A), indicating that the pRab10 MFI is sensitive to LRRK2 kinase inhibition. Following stimulation with IFN-γ, a significant increase in pRab10 signal was observed, which was significantly decreased by MLi-2 treatment in a dose dependent manner (Figures 3A,B), indicating that the pRab10 MFI is sensitive to LRRK2 kinase inhibition and is a suitable readout for LRRK2 kinase activity in monocytes at both rest and after IFN-γ treatment.

In addition, we assessed the suitability of a LRRK2 antibody. The LRRK2 Alexa Fluor^®700 (AF700) Novus biologicals antibody is a rabbit polyclonal antibody conjugated to AF700 used here to detect human LRRK2 in monocytes gated from total human PBMCs. Monocytes gated from human PBMCs express detectable levels of human LRRK2 protein by flow cytometry with the LRRK2-AF700 antibody (Figure 3C). Importantly, the levels of LRRK2 in the primary monocytes could be increased by stimulation with IFN-γ, as measured by flow cytometry, in both classical and non-classical monocytes (Figures 3D,E), replicating previous reports (Cook et al., 2017).

Due to the fact that pRab10 and LRRK2 staining in PBMCs requires permeabilization to ensure intracellular staining, we assessed whether intracellular staining and permeabilization significantly altered the GBA MFI of these cells in order to determine if these readouts could be assessed via the same flow-cytometry panel. We assessed MFI of GBA as before in total PBMCs in the presence or absence of CBE. It was observed that fixation and permeabilization of cells for intracellular staining significantly reduced the GBA MFI quantified in PBMCs relative to cells that did not undergo fixation and permeabilization (Figure 3F). It was therefore concluded that the GBA index needed to be assessed in live cells, and that two flow cytometry panels, a live-cell panel and a separate fixed- and permeabilized-cell panel, would be used for WHOPPA.

To ensure the potential wide-spread use of our pipeline to assess large cohorts of human samples from multiple sites, we tested WHOPPA at multiple research institutes and compared the results. PBMCs from HCs were collected from 4 different sites (University of Florida, University of Sydney and two biorepositories). All four sites cryopreserved PBMCs from five HCs into six cryovials per volunteer, with three of these cryovials being shipped to the University of Florida and three to the University of Sydney, resulting in PBMCs from the same 20 volunteers being analyzed at both sites (Figure 4A). PBMCs were cryorecovered, plated, stimulated and assessed via flow cytometry as done in the previously described experiments. Data from all treatments from both University of Florida and University of Sydney were analyzed for correlations. A significantly positive correlation was observed between data from the two institutes on classical and non-classical monocyte frequency (Figures 4B,C), LRRK2 expression and enzymatic activity (Figures 4D–G) and HLA-DR expression (Figures 4H,I). A significantly positive correlation was observed between data from the two institutes on classical monocyte GBA index and BMV109 signal (Figures 4J,K). Although a high degree of correlation was observed between data from the two institutes on non-classical GBA index and BMV109 signal, these failed to reach statistical significance (Figures 4L,M). These results illustrate that WHOPPA provides reproducibility and consistency in LRRK2 and GCase site (operator) independent enzymatic readouts across research institutes and provides a reliable tool to be used in multi-center immunophenotyping studies.

To assess expression and activity levels of LRRK2 and GCase in human monocytes and the possible effects of PD on this, we applied the WHOPPA to a total of 15 samples from subjects with PD and 13 samples from age-matched, HCs.

To investigate whether LRRK2 enzymatic activity and levels differ between healthy individuals and those with PD, both at rest and in a stimulation-dependent manner, pRab10 and LRRK2 levels were assessed via flow-cytometry in cells from subjects with PD and age-matched HC subjects using the methods optimized in this study. Until recently, the most commonly used approach to ascertain LRRK2 expression levels had been western blotting and quantitative reverse transcriptase-polymerase chain reaction. However, we previously reported the use of the c41-2 Abcam LRRK2 antibody to assess LRRK2 levels in PBMCs via flow cytometry (Cook et al., 2017). Such a method allows researchers to specifically investigate particular immune cell subgroups; indeed, differences in LRRK2 expression were reported in different immune cell types, with increases in non-classical, CD16⁺ monocytes specifically observed in PD patients relative to HCs (Cook et al., 2017). However, to date, there is still no comparable method to quantify LRRK2 kinase activity via flow cytometry. The assessment of pT73-Rab10, a downstream substrate of LRRK2, was therefore used to provide a readout of LRRK2 kinase activity.

We found that pRab10 MFI was not significantly different between subjects with PD and HCs in classical or non-classical monocytes at rest (Figure 5A and Supplementary Figure 3A). However, upon IFN-γ stimulation, a treatment effect on pRab10 MFI was observed in classical monocytes, with a significant increase observed in PD classical monocytes relative to HCs. These increases in pRab10 MFI were significantly decreased upon LRRK2 kinase inhibition with MLi-2. Interestingly, in classical monocytes at rest, LRRK2 MFI was significantly increased in PD cells relative to HCs (Figure 5B), with both PD and HC classical monocytes upregulating LRRK2 MFI upon IFN-γ stimulation to comparable levels. These observations suggest that there is a stimulation-dependent increase in LRRK2 kinase activity levels in PD classical monocytes that is not driven by an overall increase in LRRK2 levels. Interestingly, there is a positive correlation between LRRK2 and pRab10 MFI in classical monocytes from both subjects with PD and HCs that is increased in PD cells (Figure 5C), suggesting an increased association between LRRK2 and pRab10 levels in these cells. No significant changes were observed between PD and HC non-classical monocytes (Supplementary Figures 3A,B).

Next, to assess whether alterations in stimulation-dependent LRRK2 enzymatic activity were associated with alterations in monocyte activation, classical monocytes were also assessed for their expression of human leukocyte antigen (HLA), which is the antigen-presenting molecule expressed on cells such as monocytes that activates CD4⁺ T cells and is upregulated following inflammatory stimuli. In humans, there are three different isotypes, HLA-DR, -DQ, and-DP, encoded by the MHC-II locus. To determine changes in antigen presentation in subjects with PD, we assessed expression levels of HLA-DR on monocytes. Both PD and HC groups displayed induction of HLA-DR proteins after IFN-γ stimulation (Supplementary Figure 3C) with no significant changes observed between groups. However, a positive correlation was observed between HLA-DR expression and both pRab10 and LRRK2 levels (Figures 5D,E), which was increased in PD cells. These observations were specific to classical monocytes and were not seen in non-classical monocytes, where a weaker correlation was observed between LRRK2 and HLA-DR (Supplementary Figure 3E), and LRRK2 and pRab10 (Supplementary Figures 3F,G).

To investigate whether WHOPPA could be used to assess potential GCase activity level differences at rest and in a stimulation-dependent manner between HCs and those with PD, the GBA index was calculated. Control and PD participant PBMCs were treated with and without CBE both at rest and with IFN-γ treatment, and the fluorescent signal resulting from PFB-FDGlu metabolism by lysosomal GCase was calculated. We found a significant decrease in GBA indices in PD classical monocytes at rest relative to HCs (Figure 5F). Interestingly, GBA indices were significantly increased with IFN-γ treatment in classical monocytes from both groups, indicating that an increase in GCase activity is associated with a pro-inflammatory state. Although no significant differences were seen between groups in non-classical monocytes, IFN-γ significantly increased GBA indices in these cells, with MLi-2 having no effect (Supplementary Figure 3H). These data suggest an association between GCase activity levels and pro-inflammatory responses in non-classical monocytes.

As both LRRK2 and GCase have been frequently implicated in lysosomal function, which in turn is instrumental in immune cell function as previously discussed, WHOPPA was used to assess lysosomal cathepsin activity levels in these human immune cells. Cysteine cathepsins are a group of 11 proteases which regulate degradation of endocytosed material and are important regulators of both health and disease (Reiser et al., 2010). BMV109 is a pan-cysteine cathepsin, quenched activity-based probe (qABP) which is intrinsically dark and emits fluorescence only when cleaved by proteases, allowing for simultaneous monitoring of cathepsin X, B, S and L activity (Verdoes et al., 2013). We found that BMV109 MFI was significantly upregulated when both classical and non-classical monocytes were treated with IFN-γ, suggesting increased cathepsin activity in response to pro-inflammatory stimulus (Supplementary Figures 3I,J). Interestingly, in classical monocytes at rest, an increase in BMV109 signal was observed in PD cells relative to HCs.

Finally, to assess whether alterations in LRRK2 and GCase activity levels in PD PBMCs were accompanied by alterations in cytokine release, we performed multiplexed immunoassays on conditioned media from plated PBMCs stimulated with IFN-γ and co-treated with MLi-2 for 18 h. When measuring cytokine secretion from PBMCs normalized to the percentage of live cells, taken from flow cytometry data previously described, few significant differences were detected between PD-subjects and HCs (Supplementary Figure 4). However, when these values were quantified as a stimulation-dependent fold-change from baseline, cytokine secretion was increased in the PD group with no significant effect of LRRK2 kinase inhibition (Figures 6A–H).