Cell lines were grown and maintained on SM/2 agar with Klebsiella aerogenes at 21°C (Fey et al., 2007). Cells were also grown axenically in HL5 medium at 21°C and 150 rpm. For all experiments, cells were harvested in the mid-log phase of growth (1–5 × 10⁶ cells/ml). Cultures were supplemented with 100 μg/ml ampicillin and 300 μg/ml streptomycin sulfate to prevent bacterial growth. AX4, hereafter referred to as WT, was the parental cell line for mfsd8 ⁻ , which was purchased from the Genome Wide Dictyostelium Insertion (GWDI) bank via the Dicty Stock Center (https://remi-seq.org) (Fey et al., 2019; Gruenheit et al., 2021). Blasticidin S hydrochloride (10 μg/ml) was used to select mfsd8 ⁻ cells. HL5 and low-fluorescence HL5 were purchased from Formedium (Hunstanton, Norfolk, United Kingdom). 2-N-morpholinoethanesulfonic acid (MES) was purchased from Fisher Scientific Company (Ottawa, ON, Canada). Rabbit polyclonal antibodies against autocrine proliferation repressor (AprA) and countin (CtnA) were provided as gifts by Dr. Richard Gomer (Brock and Gomer, 1999; Brock and Gomer, 2005). Rabbit polyclonal antibody against calcium dependent cell adhesion protein (CadA) was generated and validated in a previous study (McLaren et al., 2021). Mouse monoclonal anti-α-actinin (47-18-9) and mouse monoclonal anti-discoidin (DscA) (80-52-13) were purchased from the Developmental Studies Hybridoma Bank (University of Iowa, Iowa City, IA, United States) (Stadler et al., 1984). Mouse monoclonal anti-β-actin was purchased from Santa Cruz Biotechnology Incorporated (Dallas, TX, United States). Horseradish peroxidase (HRP)-conjugated secondary antibodies were purchased from New England Biolabs (Whitby, ON, Canada). p-Nitrophenyl-β-D-glucopyranoside (487507) (substrate for β-glucosidase), p-Nitrophenyl-a-D-glucopyranoside (487506) (substrate for α-glucosidase), 4-Nitrophenyl-α-D-mannopyranoside (N2127) (substrate for α-mannosidase), o-Nitrophenyl-β-D-galactopyranoside (48712-M) (substrate for β-galactosidase), 4-Nitrophenyl-α-D-galactopyranoside (N0877) (substrate for α-galactosidase), 4-Nitrophenyl N-acetyl-β-D-glucosaminide (N9376) (substrate for N-acetylglucosaminidase, NAG), Ala-Ala-Phe-7-amido-4-methylcoumarin (A3401) (substrate for TPP1), Fluorogenic Cathepsin B Substrate III (219392) (substrate for cathepsin B, CTSB), and β-glucosidase (49290) were purchased from Sigma Aldrich Canada (Oakville, ON, Canada). 4-Methylumbelliferyl 6-thio-Palmitate-β-D-Glucopyranoside (19524) (substrate for palmitoyl-protein thioesterase 1, PPT1) and Fluorogenic Cathepsin F Substrate (80350-BP) (substrate for cathepsin F, CTSF) were purchased from Cedarlane Laboratories (Burlington, ON, Canada). The Cathepsin D Activity Assay Kit (10013-596) was purchased from VWR International (Mississauga, ON, Canada).

To assess cell proliferation, cells in the mid-log phase of growth were washed thrice with fresh HL5. Cells were then diluted to 1–2 × 10⁵ cells/ml in HL5 and incubated at 21°C and 150 rpm. Cell concentrations were measured every 24 h over a 96-h growth period using a hemocytometer. The pinocytosis assay was conducted using a method described elsewhere (Rivero and Maniak, 2006; Huber et al., 2014). Briefly, cells in the mid-log phase of growth were placed in 5 ml of HL5 at a density of 5 × 10⁶ cells/ml. 100 μl of a 20 mg/ml fluorescein isothiocyanate (FITC)-dextran (70,000 M_r) stock solution was added to the 5 ml suspension, which was then incubated for 120 min at 21°C and 150 rpm. 500 μl were collected every 15 min, washed twice with ice-cold Sorenson’s buffer (2 mM Na₂HPO₄, 14.6 mM KH₂PO₄, pH 6.0) and lysed with 1 ml of buffer containing 50 mM Na₂HPO₄ (pH 9.3) and 0.2% Triton-X. Lysates were added to separate wells of black bottom 96-well plates and fluorescence was measured using a BioTek Synergy HTX plate reader and the following filters (460/40 nm for excitation, 528/20 nm for emission) (BioTek Instruments Incorporated, Winooski, VT, United States). To measure cell area, cells in the mid-log phase of growth were deposited onto a hemocytometer. Cells were imaged with a Nikon Ts2R-FL inverted microscope equipped with a Nikon Digital Sight Qi2 monochrome camera (Nikon Canada Incorporated Instruments Division, Mississauga, ON, Canada). The area of each individual cell was quantified using Fiji/ImageJ (Schindelin et al., 2012). To assess cytokinesis, cells (5 × 10⁵ total) in the mid-log phase of growth were collected and deposited onto coverslips placed inside separate wells of a 12-well dish. Coverslips were then submerged in low-fluorescence HL5 and incubated overnight at 21°C. The following day, cells were fixed in −80°C methanol for 45 min and mounted onto slides with Prolong Gold Anti-Fade Reagent containing DAPI (Fisher Scientific Company, Ottawa, ON, Canada). Cells were then imaged using a Nikon Ts2R-FL inverted microscope equipped with a Nikon Digital Sight Qi2 monochrome camera. For each independent experiment, the number of nuclei within each cell (at least 100) was scored and expressed as a percentage of the total number of cells analyzed.

Three full inoculation loops of K. aerogenes were collected and resuspended in KK2 buffer (0.7 g/L K₂HPO₄ and 2.2 g/L KH₂PO₄, pH 6.5). 25 μl of the suspension were then deposited onto SM/2 agar and incubated at 21°C. Two days later, D. discoideum cells in the mid-log phase of growth were harvested, washed thrice with KK2 buffer, and resuspended in KK2 buffer to obtain a final concentration of 0.4 × 10⁶ cells/ml. Cells (1 × 10² total) were deposited onto the center of the K. aerogenes lawns. Plaques were captured at the indicated time points using a Leica EZ4W stereomicroscope equipped with an internal 5MP CMOS camera (Leica Microsystems Incorporated, Concord, ON, Canada). Plaque diameters were quantified using Fiji/ImageJ.

Cells from the proliferation assay described above were harvested after 24, 48, and 72 h of growth (Huber et al., 2014). Cells were lysed with buffer containing 50 mM Tris–HCl (pH 8.0), 150 mM NaCl, 0.5% NP40, and a protease inhibitor tablet (PIA32965) (Fisher Scientific Company, Ottawa, ON, Canada). Samples of conditioned media (CM) were collected and standardized based on cell number (1 × 10⁶ total). Whole cell (WC) lysates and equal volumes of CM were separated by SDS-PAGE and analyzed by western blotting. The following primary and secondary antibodies were used: anti-AprA (1∶1000), anti-β-actin (1∶1000), and HRP-conjugated secondary antibodies (1:2000). Protein bands were imaged using the ChemiDoc Imaging System (Bio-Rad Laboratories Limited, Mississauga, ON, Canada) and quantified using Fiji/ImageJ.

Aggregation was examined using a method described previously with minor modifications (Huber, 2017). Briefly, cells (6 × 10⁶ total) harvested from the mid-log phase of growth were deposited into separate wells of a 6-well dish. Cells were allowed to adhere to the surface of the dish for 1 h after which time they were washed two times with KK2 buffer, and then starved in 1 ml of KK2 buffer. Cells were imaged at the indicated times with a Nikon Ts2R-FL inverted microscope equipped with a Nikon 10 Digital Sight Qi2 monochrome camera. A conditioned buffer (CB) swap experiment was conducted to test the effect of proteins secreted by WT cells on the aggregation of mfsd8 ⁻ cells (Huber et al., 2017). Briefly, the CB from WT cells starved for 2 h in 1.5 ml of KK2 buffer was collected and spun down to remove any cells present in the buffer. mfsd8 ⁻ cells were submerged in either 1 ml of KK2 buffer (control) or CB collected from starving WT cells. Cells were imaged at the indicated times with a Nikon Ts2R-FL inverted microscope equipped with a Nikon 10 Digital Sight Qi2 monochrome camera. Aggregation was also examined on 0.5% agar/KK2 (Huber and Mathavarajah, 2018). Briefly, cells in the mid-log phase of growth were harvested from HL5, washed two times with KK2 buffer, and plated (1.5 × 10⁸ cells/ml) in 0.5 μl volumes on 0.5% agar/KK2. Cell spots were imaged at 0 and 5 h using a Nikon Ts2R-FL inverted microscope equipped with a Nikon Digital Sight Qi2 monochrome camera. Images were viewed using NIS Elements Basic Research and analyzed using Fiji/ImageJ. The area after 5 h was expressed as a percentage of the area at 0 h to provide a measure of the amount of aggregation.

Cells (8 × 10⁶ total) in the mid-log phase of growth were deposited into 60 mm × 15 mm Petri dishes and allowed to adhere for 1 h, after which time the HL5 was removed. Adherent cells were washed two times with KK2 buffer and then submerged in 4 ml of KK2 buffer for 4 h, after which time the CB was collected and concentrated using an Amicon Ultra-4 centrifugal filter unit (UFC801024) (Fisher Scientific Company, Ottawa, ON, Canada) according to the manufacturer’s instructions. Adherent cells were lysed with NP40 lysis buffer (recipe noted above). SDS-PAGE and western blotting were then used to determine the effect of mfsd8 loss on the intracellular and extracellular amounts of CtnA, CadA, and DscA. The following primary and secondary antibodies were used: anti-CtnA (1∶1000), anti-CadA (1∶1000), anti-DscA (1:1000), anti-β-actin (1∶1000), anti-α-actinin (1∶1000), and HRP-conjugated secondary antibodies (1:2000). Protein bands were imaged using the ChemiDoc Imaging System (Bio-Rad Laboratories Limited, Mississauga, ON, Canada) and quantified using Fiji/ImageJ.

Cell-substrate adhesion was assessed using a previously described method with minor modifications (Huber et al., 2017; Huber and Mathavarajah, 2018). Briefly, cells (6 × 10⁶ total) were deposited into separate wells of a 6-well dish and allowed to adhere for 1 h after which time they were washed twice with KK2 buffer and starved in 1.5 ml of KK2 buffer for 4 h. After 4 h, cells were shaken at 150 rpm for 30 min. Samples of CB were collected and cells in CB were counted using a hemocytometer to determine cell dissociation. Cells remaining on the dish were also lysed with NP40 lysis buffer (recipe noted above) and protein concentrations of the lysates were quantified using the Qubit Protein Assay Kit (Q33211) and a Qubit 2.0 Fluorometer (Fisher Scientific Company, Ottawa, ON, Canada).

To assess various enzyme activities, cells were grown in HL5 overnight to confluency in 60 mm × 15 mm Petri dishes (8 × 10⁶ total). Growth-phase cells and cells starved for 4 h in KK2 buffer were then lysed with buffer containing 50 mM MES pH 6.53 and 0.1% NP40, unless stated otherwise (Phillips and Gomer, 2015). Protein concentrations of lysates were quantified using the Qubit Protein Assay Kit and a Qubit 2.0 Fluorometer, and 150 μg of protein was used for the enzyme assays described below. For all enzyme assays, the absorbance and fluorescence values for the experimental samples were corrected using a lysis buffer control.

Lysosomal hydrolases: To assay α-galactosidase activity, lysates were added to 18 μl of 2 mM of 4-Nitrophenyl-α-D-galactopyranoside in sodium citrate/phosphate buffer (pH 4.5) (Kilpatrick and Stirling, 1976). Samples were then incubated for 45 min at 37°C and quenched with equal volume of 1 M sodium glycinate buffer (pH 10.4). The reactions were then deposited into 96-well black clear bottom plates and the absorbance (405 nm) was measured using a BioTek Synergy HTX plate reader. To assay β-galactosidase activity, lysates were added to 36 μl of 25 mM of o-Nitrophenyl-β-D-galactopyranoside in 100 mM citrate buffer (pH 4.0) (Maruhn, 1976). The reaction mixtures were incubated for 45 min at 37°C, after which time an equal volume of 2-amino-2-methyl-1-propnanol/HCl solution was added. The reactions were then deposited into 96-well black clear bottom plates and the absorbance (405 nm) was measured using a BioTek Synergy HTX plate reader. To assay α-glucosidase activity, WC lysates were incubated in 2 mM p-Nitrophenyl-a-D-glucopyranoside within 0.1 M sodium succinate (pH 6.0) in a total volume of 150 μl (Wimmer et al., 1997). The reaction solution was incubated for 1 h at 65°C followed by the addition of 300 μl of 1 M sodium carbonate to quench the reaction. The quenched reactions were then deposited into 96-well black clear bottom plates and the absorbance (395 nm) was measured using a BioTek Synergy HTX plate reader. To assay β-glucosidase activity, lysates were added to 18 μl of 10 mM of p-Nitrophenyl-β-D-glucopyranoside in 50 mM acetate buffer (pH 5.0) (Coston and Loomis, 1969). The reactions were incubated at 35°C for 45 min. The reactions were then quenched with an equal volume of 1 M sodium carbonate and deposited into 96-well black clear bottom plates and the absorbance (405 nm) was measured using a BioTek Synergy HTX plate reader. To assay α-mannosidase activity, lysates were added to 18 μl of 5 mM of 4-Nitrophenyl-α-D-mannopyranoside in 5 mM acetate buffer (pH 5.0) (Loomis, 1970). Samples were incubated for 45 min at 35°C after which time an equal volume of 1 M Na₂CO₃ was added to quench the reactions. The quenched reactions were then deposited into 96-well black clear bottom plates and the absorbance (405 nm) was measured using a BioTek Synergy HTX plate reader. To assay NAG activity, WC lysates were added to 75 µl of 7 mM of 4-Nitrophenyl N-acetyl-β-D-glucosaminide in 100 mM acetate buffer (pH 5.0). Samples were then incubated for 5 min at 35°C, after which time an equal volume of 1 M Na₂CO₃ was added to quench the reactions (Loomis, 1969; Huber and Mathavarajah, 2018). The quenched reactions were then deposited into 96-well black clear bottom plates and the absorbance (405 nm) was measured using a BioTek Synergy HTX plate reader.

Ppt1 and Tpp1: To assay Ppt1 activity, lysates were added to 5 µl of 9 mM of 4-Methylumbelliferyl 6-thio-Palmitate-β-D-Glucopyranoside dissolved in McIlvaine phosphate/citric-acid buffer (pH 4) supplemented with 15 mM dithiothreitol and 0.375% Triton X-100. The reactions were incubated at 37°C for 1 h after which time they were boiled for 3 min at 95°C (van Diggelen et al., 1999; Brand et al., 2018). Once the reactions were cooled, 2.75 µl of 2.5 M NaOH and 4 µl of 0.025 U/μl β-glucosidase were added to the reactions and incubated for another hour at 37°C. The reaction was quenched by adding 0.5 M sodium carbonate-bicarbonate buffer containing 0.025% Triton X-100 (pH 10.7). The quenched reactions were then deposited into 96-well opaque black bottom plates and the fluorescence was measured using a BioTek Synergy HTX plate reader and the following filters (360/40 nm for excitation, 460/40 nm for emission). To assay Tpp1 activity, cells were lysed with buffer containing 50 mM sodium phosphate (pH 6.5) and 0.5% NP40, and then added to 80 µl of 200 μM Ala-Ala-Phe-7-amido-4-methylcoumarin dissolved in reaction buffer (150 mM NaCl, 100 mM sodium acetate, pH 4.5, 0.1% Triton X-100) (Stumpf et al., 2017). Reactions were incubated in the dark at 37°C for 1 h and then quenched by adding stop solution (150 mM NaCl, 100 mM sodium acetate, pH 4.3). The quenched reactions were then deposited into 96-well opaque black bottom plates and the fluorescence was measured using a BioTek Synergy HTX plate reader and the following filters (360/40 nm for excitation, 460/40 nm for emission).

Cathepsins: To assay CtsB activity, methods were adapted with minor revisions (Barrett, 1980). Briefly, WC lysates were incubated in reaction buffer (352 mM KH₂PO₄, 48 mM Na₂HPO₄, 4 mM EDTA, pH 6.0) with 8 mM cysteine in a total volume of 150 μl and incubated at 40°C for 5 min. A final concentration of 5 μM of substrate was added to the reaction solution, which was then incubated at 40°C for 30 min. The reaction solutions were quenched with 200 μl of 100 mM sodium chloroacetate (30 mM NaC₂H₃O₂, 70 mM HC₂H₃O₂, pH 4.3). The quenched reactions were then deposited into 96-well opaque black bottom plates and the fluorescence was measured using a BioTek Synergy HTX plate reader and the following filters (360/40 nm for excitation, 460/40 nm for emission). CtsD activity was measured following the manufacturer’s instructions specified in the Cathepsin D Activity Assay Kit. Briefly, cells were lysed with 100 µl of chilled cell lysis buffer (provided in kit) and incubated on ice for 10 min. The solution was then centrifuged for 5 min. Following centrifugation, 27.5 µg of clear cell lysate was added to 100 µl of reaction buffer (provided in kit) mixed with 2 µl of substrate. Reactions were incubated at 37°C for 1 h and 30 min. The reactions were then deposited into 96-well opaque black bottom plates and the fluorescence was measured using a BioTek Synergy HTX plate reader and the following filters (360/40 nm for excitation, 460/40 nm for emission). To assay CtsF activity, WC lysates were added to 0.5 µl of 25% HCl and 1 µl of 250 μg/ml pepsin. 50 µl reactions were incubated for 1 h at 37°C after which time 0.24 µl of 0.5 mM Fluorogenic Cathepsin F Substrate followed by 5.76 µl dimethyl sulfoxide and 70 µl of 0.1 M sodium phosphate buffer containing 1 mM EDTA and 0.1% (v/v) PEG 6000 (pH 6.5) were added to the reaction mixture (total volume of 120 µl). Reactions were incubated at 27°C for 1 h (Fonovic et al., 2004). The reactions were then deposited into 96-well opaque black bottom plates and the fluorescence was measured using a BioTek Synergy HTX plate reader and the following filters (360/40 nm for excitation, 460/40 nm for emission).

Numeric data are reported as the means ± SEM. Statistical analyses for all experiments were performed using GraphPad Prism 8 (GraphPad Software Incorporated, La Jolla, CA, United States). A one-sample t-test was used for evaluating data where the raw data between biological replicates was inherently variable (e.g., quantifying pixel intensity from a western blot). A p-value < 0.05 was considered significant for all analyses and n represents the number of independent experiments that were performed. Details on the specific statistical analyses performed are found in the figure captions.

A previous study in D. discoideum showed that Mfsd8 localizes to endocytic compartments, linked the function of Mfsd8 to protein secretion, and revealed the Mfsd8 interactome during growth and starvation (Huber et al., 2020b). To gain further insight into the function of Mfsd8 in D. discoideum, here, we performed an in-depth characterization of an mfsd8 ⁻ cell line. The expression of mfsd8 increases during the early stages of development, which involves the chemotactic aggregation of cells into multicellular mounds (Rot et al., 2009; Mathavarajah et al., 2017). Following mound formation, mfsd8 expression decreases dramatically reaching its lowest level just prior to terminal differentiation of pre-spore and pre-stalk cells. Expression then rises slightly during fruiting body formation. This expression profile suggested to us that Mfsd8 primarily functions during growth and the early stages of development. As a result, we focused our analysis on the effects of mfsd8-deficiency on processes that occur during these stages of the life cycle.

When cultured in liquid growth medium, mfsd8 ⁻ cells proliferated at a significantly increased rate compared to WT cells (Figure 1A). Since D. discoideum cells ingest extracellular liquid nutrients through macropinocytosis, we assessed whether the rate of pinocytosis was affected by mfsd8-deficiency (Hacker et al., 1997). Cells were incubated in liquid growth medium containing FITC-dextran for a 120-min period and the amount of intracellular fluorescence was measured every 15 min. Like proliferation, the rate of FITC-dextran accumulation was significantly increased in mfsd8 ⁻ cells relative to WT cells (Figure 1B).

Since loss of mfsd8 impacted cell proliferation and the accumulation of FITC-dextran in liquid growth medium, we tested whether mfsd8 ⁻ cells would display the same phenotype when grown on bacteria lawns. Cell lines were plated on lawns of K. aerogenes and plaque formation was monitored. In general, we observed that mfsd8 ⁻ cells formed plaques earlier than WT cells and the plaques were larger than WT plaques at all time points examined (Figure 1C). Of note, 32 h after depositing cells onto lawns, mfsd8 ⁻ plaques were translucent, while WT plaques were semi-translucent, indicating that mfsd8 ⁻ cells cleared bacterial lawns earlier than WT cells. When feeding on bacterial lawns, D. discoideum amoebae chemotactically respond to folic acid that is secreted by bacteria (Pan et al., 1972). However, we observed no significant effect of mfsd8-deficiency on folic acid-mediated chemotaxis (Supplementary Figure S1). Together, these results suggest that Mfsd8 regulates cell proliferation, pinocytosis, and growth on bacterial lawns in D. discoideum.

When examining the proliferation of mfsd8 ⁻ cells, we observed that mfsd8 ⁻ cells appeared larger than WT cells. Indeed, when we quantified the area of cells cultured in liquid growth medium, we found that mfsd8 ⁻ cultures contained a significantly higher proportion of cells > 200 μm² compared to WT cultures, and a correlated lower proportion of cells 100–200 μm² (Figure 2A).

When cultured in liquid growth medium, the majority of D. discoideum cells are mononucleated. However, it is well established that axenically grown cultures of D. discoideum can also contain polynucleated cells, which form due to defects in cytokinesis and tend to be larger than mononucleated cells (Waddell et al., 1987). Based on these findings, we examined the proportions of mononucleated and polynucleated cells in mfsd8 ⁻ cultures. We observed that loss of mfsd8 significantly decreased the proportion of mononucleated cells in culture and increased the proportion of polynucleated cells (Figure 2B). Together, these findings showed that mfsd8-deficiency increases cell size and reduces cytokinesis during the growth phase of the D. discoideum life cycle.

To gain insight into the possible mechanisms underlying the increased proliferation of mfsd8 ⁻ cells, the intracellular and extracellular levels of AprA were examined. AprA functions extracellularly to repress cell proliferation in D. discoideum and has previously been shown to be aberrantly secreted by cln3 ⁻ cells, which like mfsd8 ⁻ cells, also display increased rates of proliferation (Brock and Gomer, 2005; Huber et al., 2014). WC lysates and CM from WT and mfsd8 ⁻ cells were collected after 24, 48, and 72 h of axenic growth (Figure 1A). When analyzed by western blotting, anti-AprA detected two protein bands; one at 60 kDa and the other at 55 kDa, which is consistent with the banding pattern reported in previous studies (Brock and Gomer, 2005; Huber et al., 2014) (Figure 3). Intracellularly, the amounts of 60 kDa AprA remained constant during all stages of axenic growth for both WT and mfsd8 ⁻ cultures (Figure 3A). However, the amounts of 60 kDa AprA in mfsd8 ⁻ cells were lower at all time points analyzed compared to WT cells. The amounts of intracellular 55 kDa AprA decreased in WT and mfsd8 ⁻ cells during axenic growth but there were no differences between cell lines. In samples of CM, loss of mfsd8 reduced the extracellular amount of 60 kDa AprA after 24 and 48 h of axenic growth, compared to WT cells, but there was no effect at the 72-h time point (Figure 3B). In addition, CM collected from mfsd8 ⁻ cells after 72 h of axenic growth contained more 55 kDa AprA than CM collected from WT cells. Blots containing samples of CM were also probed with anti-β-actin to validate that CM did not contain intracellular proteins due to cell lysis (data not shown). Since AprA functions as a proliferation repressor, these findings suggest that altered levels of intracellular and extracellular AprA may have contributed to the increased rate of proliferation of mfsd8 ⁻ cells.

Lysosomal enzymes play an essential role in degrading material internalized by amoebae during the growth phase of the D. discoideum life cycle to provide cells with nutrients (Ashworth and Quance, 1972). Interestingly, in mice, loss of Mfsd8 has been shown to affect the levels of several lysosomal enzymes (Danyukova et al., 2018). Thus, we were interested in assessing lysosomal enzyme activity in mfsd8 ⁻ cells during growth. Loss of mfsd8 significantly increased the intracellular activities of α-galactosidase, α-glucosidase, β-glucosidase, α-mannosidase, and N-acetylglucosaminidase (Figure 4). The activity of β-galactosidase was also slightly elevated but not statistically significant (p = 0.08). mfsd8-deficiency also elevated the activities of Ppt1 and CtsF. In humans, mutations in PPT1 and CTSF cause the CLN1 and CLN13 subtypes of NCL, respectively (Mole and Cotman, 2015). Finally, there was no effect of mfsd8 loss on the activities of Tpp1, CtsB, or CtsD. In humans, mutations in TPP1 and CTSD cause the CLN2 and CLN10 subtypes of NCL, respectively, and CTSB has been identified as a potential biomarker for CLN6 disease (Huber, 2021). Together, these data suggest that mfsd8 ⁻ cells increase the activities of some, but not all, lysosomal enzymes to support their increased rates of proliferation and pinocytosis.

Based on the increased expression of mfsd8 during the early stages of development (Rot et al., 2009), we suspected that loss of mfsd8 would affect cellular processes during this stage of the life cycle. Therefore, we performed two assays to examine the impact of mfsd8-deficiency on aggregation. In the first assay, cells adhered to Petri dishes were submerged in KK2 buffer to initiate the developmental program. After 8 h, there was a noticeable delay in the aggregation of mfsd8 ⁻ cells compared to WT cells (Figure 5A). A delay in mfsd8 ⁻ aggregation was also observed when cells were deposited on KK2-buffered agar (Figure 5B). In D. discoideum, mound formation occurs through the chemotactic aggregation of cells towards 3′,5′-cyclic adenosine monophosphate (cAMP) (Konijn et al., 1967). However, we did not observe a significant effect of mfsd8-deficiency on cAMP-mediated chemotaxis (Supplementary Figure S1).

A previous study showed that mfsd8-deficiency alters the secretion of two other CLN protein homologs in D. discoideum, Cln5 and CtsD, during the early stages of development (Huber et al., 2020b). Thus, we performed a CB swap experiment to examine the possible role of altered secretion in the delayed aggregation of mfsd8 ⁻ cells. For this assay, we starved WT cells for 2 h and then collected the CB. In nine independent experiments, we observed that incubating mfsd8 ⁻ cells in CB collected from WT cells partially restored the delayed aggregation of mfsd8 ⁻ cells relative to WT cells (Figure 6). We also examined the effect of mfsd8-deficiency on cell-substrate adhesion, which plays an essential role in aggregation (Tarantola et al., 2014). During the early stages of development, loss of mfsd8 caused more cells to de-adhere from Petri dishes relative to WT cells (Figure 7). Combined, these data suggest that reduced cell-substrate adhesion along with aberrant protein secretion likely contribute to the delayed aggregation of mfsd8 ⁻ cells during the early stages of D. discoideum development.

To gain further insight into the cellular mechanisms affected by mfsd8-deficiency, we starved WT and mfsd8 ⁻ cells for 4 h and analyzed the intracellular and extracellular amounts of CtnA, CadA, and DscA. CtnA is a component of a secreted 450 kDa protein complex that regulates group size during aggregation by repressing cell-cell adhesion (Roisin-Bouffay et al., 2000). CadA and DscA, which were previously shown to interact with Mfsd8 (Huber et al., 2020b), both play important roles in regulating cell adhesion during the early stages of development (Cano and Pestaña, 1984; Springer et al., 1984; Knecht et al., 1987; Brar and Siu, 1993). In addition, loss of dscA impairs cell-substrate adhesion during aggregation (Springer et al., 1984; Bastounis et al., 2016). In this study, loss of mfsd8 significantly reduced the intracellular level of CtnA but increased the amount of the protein in CB, suggesting that secretion of CtnA was increased due to mfsd8-deficiency (Figure 8). Loss of mfsd8 had no effect on the amount of CadA inside cells but significantly increased its level extracellularly. Finally, loss of mfsd8 reduced the intracellular and extracellular levels of DscA. Combined, these data indicate that mfsd8-deficiency affects the intracellular and extracellular amounts of proteins involved in aggregation and cell adhesion, which could explain the delayed aggregation and reduced adhesion of mfsd8 ⁻ cells during the early stages of development.

During multicellular development, D. discoideum amoebae rely on autophagy and the actions of several lysosomal enzymes to provide cells with energy and building blocks required for fueling aggregation and fruiting body formation (Loomis, 1969; Loomis, 1970; Dimond et al., 1973; Kilpatrick and Stirling, 1976; Knecht et al., 1985; Otto et al., 2003; Kiel, 2010). As noted above, previous work in mice showed that loss of Mfsd8 affects the levels of several lysosomal enzymes (Danyukova et al., 2018). Thus, we assessed the effect of mfsd8-deficiency on lysosomal enzyme activity during the early stages of D. discoideum development. After 4 h of starvation, we observed increased intracellular activity of α-mannosidase and decreased activity of Ppt1 and CtsF (Figure 9). There was no significant effect of mfsd8-deficiency on the activities of α-galactosidase, β-galactosidase, α-glucosidase, β-glucosidase, N-acetylglucosaminidase, Tpp1, CtsB, or CtsD. Combined, these findings show that loss of mfsd8 alters the activities of some, but not all, lysosomal enzymes during aggregation.