D. discoideum possesses two different life cycles: a vegetative cycle, responsible for cell division, and a developmental cycle initiates upon nutrient depletion that culminates in the formation of dormant spores. During the developmental cycle D. discoideum cells aggregate to form a multicellular mobile organism, called a slug, which is able to move in the direction of light and heat, a process called photo-and thermotaxis, respectively, till it settles down to form the spore stalk and head. Roco4 expression is elevated during the later slug phase (> 12 h) and consistently, roco4- cells have previously been found to display severe development phenotypes, including delayed development and abnormal spore morphology (Figures 1A–E; van Egmond and van Haastert, 2010). The amino acid that is most frequently mutated in PD (G2019S) is conserved in Roco4 and corresponds to G1179S. Previously we have shown that both LRRK2G2019S and Roco4G1179S showed increased kinase activity in vitro (Kortholt et al., 2012). Interestingly, expression of Roco4 G1179S in vivo, results in a similar phenotype compared to roco4- cells (Figures 1A–C). Wild-type (WT) cells start forming slugs after 12 h, whereas both roco4- and Roco4 G1179S cells still remain in the mound stage after 12 h (Figure 1A). The two mutant cell lines remain in the mound stage for an excessive time period followed by multiple attempts to culminate. They eventually generate slugs and form fruiting bodies only after > 50 h (Roco4 G1179S) and > 70 h (roco4-) of starvation, respectively (Figure 1A). Furthermore, roco4- cells hardly make stalks, while Roco4 G1179S make much smaller stalks compared to WT (Figure 1B). Consistent with previous studies, we found that re-expression of the Roco4 WT protein (Roco4 rescue) completely restored the developmental defects (Figures 1A–C; van Egmond and van Haastert, 2010; Kortholt et al., 2012).

We next addressed whether the impaired development might be accompanied by a defect in phototaxis. Therefore, cells were washed, placed and developed on a non-nutrient agar (NNA) plate with a given light source in front. After 3 days the tracks of the migrated slug were blotted onto a plastic sheet, stained with Coomassie dye and the trails were quantified (Figures 1D,E). Wild-type cells showed highly efficient phototaxis toward the light source, with a phototactic index of 0.96 and a track length of 3.64 cm. In contrast both roco4- and G1179S cells showed severely impaired phototaxis. The roco4- slugs hardly move (Supplementary Movie 1) and therefore the accuracy of directional movement is indeterminate. The G1179S slugs do displace although impaired (1.37 cm) and with very poor directional movement toward the light (Figure 1E).

Together our results show that both the absence of Roco4 as well as the hyperactivity of the Roco4 kinase lead to impaired D. discoideum development and phototaxis. Interestingly, delayed development, impaired stalk formation and dysregulated phototaxis have been strongly linked to impaired mitochondrial function (Francione et al., 2011).

To analyze the mitochondrial activity in the Roco4 mutants we determined the accumulation of reactive oxygen species (ROS) levels by analyzing the dye intensity of 2′,7′-Dichlorofluorescin diacetate (DCF-DA). This dye is colorless until it gets oxidized and generates the fluorescent product 2′,7′-dichlorofluorescein (DCF) (Brubacher and Bols, 2001). Accumulation of DCF signal thus gives an indication of the amount of redox-active substances produced in the cell. WT and roco4- cells were washed with phosphate buffer to limit background noise and subsequently DCF-DA was added in the presence and absence of the Carbonyl cyanide m-chlorophenylhydrazone (CCCP). Mitochondrial uncoupler, CCCP, induces depolarization of the mitochondrial membrane and triggers maximum mitochondrial activity. The fluorescent signal accompanying the oxidation of DCF-DA to DCF has been measured after addition of CCCP. Initially WT and roco4- cells have similar low levels of ROS (Figure 2A), however, following CCCP addition the ROS production in roco4- cells appears to increase more compared to WT. Interestingly, in several roco4- cells the DCF fluorescence was so high that the complete cell is fluorescent and no individual patches can be observed (Figure 2A). To quantitatively analyze the DCF fluorescence intensity we used a plate reader assay (Figure 2B) and the vegetative WT and mutant cells were measured over time in the presence or absence of CCCP (Figure 2B). Consistent with the microscopy data, both cell lines display a similarly low level of initial ROS concentrations and roco4- cells show a significant increase of ROS levels following 30 min of CCCP challenge (Figure 2B).

To determine if the observed increase in ROS production is due to altered mitochondrial functioning, we analyzed the mitochondrial respiration of the WT, roco4-, Roco4 G1179S, and Roco4 rescue strains using the Seahorse Bioscience XF24 Extracellular Flux Analyzer (Figure 3). Seahorse technique enables kinetic measurement of oxidative metabolism by measuring the rate of oxygen consumption [oxygen consumption rate (OCR)] in real time. This method can provide detailed information on the metabolic phenotype by the addition of mitochondrial substrates and modulators of mitochondrial electron transport chain (Annesley et al., 2016). Cells were added to the Seahorse analyzer and basal OCR levels were determined prior to port injections and represent energetic demand of the cell under baseline conditions (Figures 3A,E). The decrease in OCR following the injection of ATP synthase inhibitor (DCCD or oligomycin) indicates the ATP synthesis rate (Figure 3B) while the maximum OCR is attained by adding the uncoupler (CCCP or FCCP) (Figures 3C,F). Complex I activity can be calculated by the decrease in OCR after rotenone addition (complex I inhibitor) (Figure 3D). Our data revealed that the basal mitochondrial respiration, ATP synthesis rate and complex I activity in the roco4- strain are significantly increased, while the maximum OCR is slightly increased yet not statistically significant compared to the WT strain (Figures 3A–D). In contrast, Roco4 G1179S and the Roco4 rescue cells exhibit a significantly reduced maximal respiration compared to the WT strain, while Roco4 G1179S cells have significantly lower basal respiration as well (Figures 3E,F). These data indicate that D. discoideum Roco4 mutants have an impact on different steps in the mitochondrial respiratory activity.

LRRK2 is highly expressed in immune cells and several studies have linked LRRK2 signaling to inflammation (Thévenet et al., 2011; Brockmann et al., 2017; Dzamko, 2017; Lee et al., 2017). We therefore used murine LRRK2 parental and LRRK2 KO RAW 264.7 macrophages to analyze whether the absence of LRRK2 (Knocking out) also has an effect on mitochondrial activity. LRRK2 KO RAW 264.7 cells show a higher maximal respiration and increased spare capacity compared to LRRK2 parental RAW 264.7 cells, whereas basal respiration, ATP-production and coupling efficiency [(ATP production/basal respiration)^∗100] were not significantly altered (Figures 4A–E). Previous studies have shown that the majority of LRRK2 is cytosolic and, if at all, only a small fraction of LRRK2 is present in mitochondria (West et al., 2005; Papkovskaia et al., 2012). We used high-resolution respirometry (Oroboros) to measure the activity of isolated mitochondria. Mitochondrial respiration was measured in the presence of the mitochondrial substrates, pyruvate, and malate (state 2). ADP was added in a saturating concentration to determine phosphorylating respiration (state3). Subsequently, oligomycin was applied to inhibit ATP synthase (state 4) and FCCP provided the maximal uncoupled-stimulated respiration (state 3U). Consistent with the proposed cytosolic LRRK2 localization we did not observe any significant differences in any of the mitochondrial respiration states between LRRK2 parental and KO RAW 264.7 cells, and can conclude that the OXPHOS complexes and complex V are functionally normal (Figure 4F).

Together our data show that D. discoideum Roco4 and LRRK2 mutants affect the mitochondrial respiration activity in a similar way. Furthermore, the differences between the whole cell and mitochondrial fraction measurements, suggest that this effect is indirect and that LRRK2 most likely requires other cytosolic cofactors to elicit mitochondrial effects.

To gain information on the shape and number of mitochondria in the D. discoideum cells, we used a specific mitochondrial marker coupled to Alexa-594-conjugated streptavidin (Life Technologies). Alexa-594-streptavidin binds to the heavily biotinylated protein 3-methylcrotonyl-CoA carboxylase α, which is localized in the mitochondrial lumen (Davidson et al., 2013). WT and roco4- cells show a similar mitochondrial network both in vegetative cells and in starved cells. In both strains mitochondrial elongation is enhanced upon starvation due to increasing energy demands (Figure 5A; Katz, 2002). The fixed cells show very clear elongated and interconnected mitochondria with no visible fragmentation in either WT or roco4- cells. Similar observations have been made in living cells using the previously published marker fluorescent marker GFP-gemA (Supplementary Figure 1; Vlahou et al., 2011; Davidson et al., 2013). Furthermore, we observed no obvious differences in mitochondrial morphology in LRRK2 parental and KO RAW 264.7 cells (Figure 5B). The ratio (percentage) of mitochondria to overall cell size was calculated using area measurements on images with ImageJ software; no differences were observed for both the roco4- and LRRK2 KO RAW 264.7 strains compared to the respective parental strains (Table 1). Consistently, manual counting of the mitochondria of the D. discoideum cells did not reveal significant differences between WT and roco4- cells (Table 1). The amount of mitochondria are in accordance with the previous defined number of 50 mitochondria per cell and observer focal lane in healthy WT cells (Gilson et al., 2003).

To visualize qualitative ultrastructural details of the mitochondria within cells, we made use of Electron Microscopy (EM). Due to the high resolution it is possible to visualize the overall mitochondrial shape and organization and also details of the inner lumen with regards to size, folding, and organization of the cristae (Figures 5C,D and Table 1). D. discoideum WT and roco4- and RAW 264.7 WT and KO cells were fixed and visualized under the EM. The mitochondria of both D. discoideum cell lines are neither aggregated nor clustered in close proximity to a specific organelle (Figure 5C). Shape, size and connectivity of the mitochondria are in agreement with previous literature analyzing wild-type-like mitochondrial membranes (Arnoult et al., 2001). In accordance, EM images of LRRK2 parental and KO RAW 264.7 cells do not indicate an abnormal distribution of the mitochondria within the cell or in clusters around an identifiable organelle (Figure 5D). The mitochondria appear similar in size, shape, and connectivity compared to the overall cell volume in both cell lines (Figure 5D and Table 1). The cristae are clearly visible and display proper folding and organization as normal for mitochondria in eukaryotic cells. The differences in mitochondrial physiology as observed between parental and Roco mutants are thus not expressed in their (ultra)structural organization.

The D. discoideum cells were cultured in HL5 (Formedium) media. For starvation the medium was washed away and the cells were incubated in phosphate buffer (PB; 10 mM phosphate buffer, pH 6.5) for 6 h prior to the experiment. Roco4 mutants have been created and described previously. The generation of the roco4- cell line (DBS0350095) and the Roco4 constructs (DDB_G0288251)/G1179S (DBS0350100) has been described previously (van Egmond and van Haastert, 2010; Gilsbach et al., 2012). Transformation of the cells has been carried out according to standard protocol and the cells were cultured in the presence of the according selection marker (van Egmond and van Haastert, 2010). Expression of the constructs was verified via Western Blot. For this, vegetative cells were washed in PB and lysed in 1:2 volume ratio in SDS buffer (200 mM Tris, pH 6.8, 400 mM DTT, 8% SDS, 0.05 Bromophenol Blue, 20% Glycerol) at 95°C for 10 min. Cell lysate was loaded on a SDS-PAGE gel, subsequently blotted on a nitrocellulose membrane and incubated with α-GFP according to the standard procedure (Gilsbach et al., 2012) [“GFP Antibody (B-2)” and “m-IgGκ BP-HRP,” Santa Cruz Biotechnology].

The murine cells were obtained from ATCC: parental RAW 264.7 and the LRRK2-KO (knockout) RAW 264.7. The cells were cultured in DMEM (LGC Standards GMBH) supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin at 37°C in 5% CO₂.

Starvation and onset of the developmental cycle have been induced by washing D. discoideum cells with PB. 2 × 10⁷ cells were spread on ϕ5 cm 1.5% non-nutrient agar. Developmental time and stalk formation was monitored and documented using a Stemi SV 11 (Zeiss) microscope. For phototaxis, 5 × 10⁵ cells were washed, resuspended in 10 μL and placed in a trench that was cut out on the far end of an agar plate. The plate was placed in a dark ϕ6 cm tube with a 1 mm wide open light slit opposite the cells. Cells were allowed to absolve the developmental cycle (24–72 h). Slugs were formed that migrated toward the light source over the agar surface. Slug tracks, consisting of slime and cell debris were blotted to a PVC sheet, stained with Coomassie Blue and scanned via a CanonScan 9000F (Canon). The tracks were digitized and measured via ImageJ and the phototaxis index is expressed as the ratio of the absolute distance traveled to the light divided by the total track length.

For mitochondrial staining, D. discoideum cells were settled on glass plates, then fixed with ultracold (–80°C) Ethanol, washed with PB and incubated with 50 ng/mL Alexa-594 conjugated streptavidin (Life Technologies) for 30 min with no subsequent washing steps according to protocol (Davidson et al., 2013). Cell images were taken via a Zeiss Axio Observer microscope. The number of mitochondria was manually counted in 25 cells per cell line. The percentage area of the cell occupied by mitochondria was calculated using ImageJ as the ratio of mitochondrial area to the whole cell area.

Mitochondria in the macrophage cell lines were stained with a mitotracker dye: cells (250,000 cells/well) were grown overnight on coverslips. Then the growth medium was replaced with a fresh medium containing 200 nM MitoTracker^TM Deep Red FM (Invitrogen) and incubated for 30 min at 37°C, followed by washing twice with PBS and fixation with 4% PFA for 25 min. After washing with PBS the cover slips were mounted on glass slides using Floroshield^TM with DAPI (Sigma) as a mounting medium. To study mitochondrial morphology, images were taken via Zeiss LSM800 confocal microscope and analyzed with ImageJ. The percentage area of the cell occupied by mitochondria was calculated as described above.

For transmission electron microscopy (TEM), D. discoideum and macrophage cells were grown to a confluent monolayer on Thermanox coverslips (Thermo Fisher Scientific). The coverslips with cells were fixed by immersion in ice-cold glutaraldehyde (6% in 0.1 M cacodylate-buffer pH 7.2) overnight, washed thoroughly with water and subsequently fixed and contrasted with 1% OsO₄ in 0.1 M cacodylate-buffer pH 7.2 (1 h at RT). For D. discoideum KMnO₄ (1.5% in double distilled water) was used as additional fixative. After thorough washing with water, the cells were incubated overnight in 1% uranyl-acetate. The coverslips were dehydrated in a rising ethanol series and embedded in Epon with the following steps: infiltration with increasing ratio of Epoxy-resin in ethanol (1:1, 3:1, pure resin, 15 min each step, pure resin overnight). The coverslips were cut into smaller pieces and transferred to beem capsules and covered with fresh resin which was polymerized at 60^oC for 48 h. Ultrathin sections were cut and the samples were examined with a transmission electron microscope (Philips CM12) at 80 kV.

For the detection of intracellular ROS levels, 5 × 10⁵ vegetative D. discoideum cells were settled on glass plates for 30 min. A 10 μM 2′,7′-Dichlorofluorescin diacetate (DCF-DA, Sigma-Aldrich) was added to the cells and allowed to incubate for 30 min. Images (1 frame/min) of cells have been taken with a Zeiss LSM800 microscope with and without the addition of 1 μM Carbonyl cyanide m-chlorophenylhydrazone (CCCP; Sigma- Aldrich) over a time period of 30 min. Alternatively, 5 × 10⁵ vegetative WT and mutant cells, washed and resuspended in PB, were seeded into wells of a black 96-wells-plate. A 10 μM DCF-DA was added to the wells and left to incubate for 30 min. Measurements (every minute) were taken immediately after the addition of 1 μM CCCP to the respective wells for 30 min via the microplate reader Synergy HTX (BioTek). Background measurements were taken of HL5 media with 10 μM DCF-DA and subtracted from the data. Measurements have been conducted five times.

Oxygen consumption measurements were performed on intact D. discoideum cells using the Seahorse XFe24 Flux Analyzer (Seahorse Biosciences) as previously described (Lay et al., 2016). Seahorse measurements were performed in two groups; one for the WT and roco4- strains and the other for the WT, Roco4 G1179S and Roco4 rescue. Briefly, D. discoideum cells were collected, washed and resuspended in SIH assay medium (SIH0102, ForMedium^TM, United Kingdom) supplemented with 20 mM sodium pyruvate and 5 mM sodium malate (pH 7.4). To ensure cell adherence during the assay, cells were seeded in XF24 cell culture plates coated with Matrigel^® (growth factor reduced basement membrane matrix; Corning, MA, United States) at 1 × 10⁵ cells/well and allowed to settle for 1–2 h. Bioenergetic profiling was performed by monitoring basal oxygen consumption rate (OCR) for 48 min followed by the sequential injection of the following inhibitors; N,N’-dicyclohexylcarbodiimide (DCCD; 10 μM; Sigma- Aldrich; Port A), carbonyl cyanide 3-chlorophenylhydrazone (CCCP; 10 μM; Sigma- Aldrich; Port B), rotenone (20 μM; Sigma- Aldrich; Port C), and antimycin A (10 μM; Sigma- Aldrich; Port D) at 32 min interval. Mitochondrial responses were monitored every 8 min following a cycle of 3 min mixing/2 min delay/3 min measurement. The following respiratory parameters were utilized to analyze the bioenergetic function: basal OCR, ATP synthesis rate (with DCCD), maximum OCR (with CCCP), complex I activity (with rotenone) and mitochondrial spare capacity (with antimycin A).

For bioenergetics assessment of the effect of LRRK2 on murine macrophage cells, adherent LRRK2 parental RAW 264.7 and LRRK2 KO RAW 264.7 cells were used. Cells were transferred to XF microplates at a seeding density of 75,000 cells/well 24 h before OCR measurement. After replacing the growth medium with 180 μL of assay medium supplemented with 25 mM glucose, 2 mM L-glutamine and 1 mM sodium pyruvate (pH 7.35), cells were preincubated for 1 h in a CO₂ free incubator at 37°C before starting the assay procedure. Following basal respiration measurement, various compounds such as: oligomycin (2.5 μM; Sigma-Aldrich), Carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP; 2.5 μM; Sigma-Aldrich), rotenone (1 μM) and antimycin A (1 μM) were sequentially injected to assess mitochondrial coupling of respiratory chain, maximal and non-mitochondrial oxygen consumption. OCR values were normalized to protein content in each well, determined with a BCA assay.

The function of the respiratory chain on isolated mitochondria in LRRK2 parental and KO RAW 264.7 cells was assessed via high resolution respirometry at 37°C, Oxygraph 2K (Oroboros systems, Innsbruck, Austria). Mitochondrial isolation was performed using a pump-controlled cell rupture system as formerly described (Schmitt et al., 2013). Isolated mitochondria (200 μg) were injected to an air tight chamber containing Mir05 (0.5 mM EGTA, 3 mM MgCl₂, 60 mM lactobionic acid, 20 mM taurine, 10 mM KH₂PO₄, 20 mM HEPES, 110 mM D-Sucrose and 1 mg/mL fatty acid-free BSA). Sequential addition of the following substances were done to measure mitochondrial respiration states; pyruvate (5 mM) and malate (2 mM) to assess non-phosphorylating respiration (mitochondrial state 2), saturating concentration of ADP (500 μM) to quantify the OXPHOS capacity of complex I-linked respiration (state 3), oligomycin (2 μg/mL) to inhibit the ATP-synthase (state 4), and stepwise titration of FCCP (1 μM) to calculate the maximal respiration (state 3U). Oxygen flux were recorded in real-time using the DatLab software and the data was analyzed with DatLab5 (version 5.1.1.91).