DPTIP, a newly identified potent brain penetrant neutral sphingomyelinase 2 inhibitor, regulates astrocyte-peripheral immune communication following brain inflammation
DPTIP, a newly identified
Ajit G. Thomas
Amrita Datta Chaudhuri
Sarah C. Zimmermann
Alexandra G. Gadiano
Barbara S. Slusher
Brain injury and inflammation induces a local release of extracellular vesicles (EVs) from astrocytes carrying proteins, RNAs, and microRNAs into the circulation. When these vesicles reach the liver, they stimulate the secretion of cytokines that mobilize peripheral immune cell infiltration into the brain, which can cause secondary tissue damage and impair recovery. Recent studies suggest that suppression of EV biosynthesis through neutral sphingomyelinase 2 (nSMase2) inhibition may represent a new therapeutic strategy. Unfortunately, currently available nSMase2 inhibitors exhibit low potency (IC50 ? 1 ?M), poor solubility and/or limited brain penetration. Through a high throughput screening campaign of >365,000 compounds against human nSMase2 we identified 2,6-Dimethoxy-4-(5-Phenyl4-Thiophen-2-yl-1H-Imidazol-2-yl)-Phenol (DPTIP), a potent (IC50 30 nM), selective, metabolically stable, and brain penetrable (AUCbrain/AUCplasma = 0.26) nSMase2 inhibitor. DPTIP dose-dependently inhibited EV release in primary astrocyte cultures. In a mouse model of brain injury conducted in GFAP-GFP mice, DPTIP potently (10 mg/kg IP) inhibited IL-1?-induced astrocyte-derived EV release (51 ? 13%; p < 0.001). This inhibition led to a reduction of cytokine upregulation in liver and attenuation of the infiltration of immune cells into the brain (80 ? 23%; p < 0.01). A structurally similar but inactive analog had no effect in vitro or in vivo.
nSMase25. Upregulation of nSMase2 activity is associated with cognitive impairment in HIV infection6, and with
plaque deposition in AD7,8. Moreover, astrocyte-derived EVs (ADEVs) isolated from the plasma of AD patients
contain increased amounts of complement proteins, implying that glial activation leads to the release of EVs
that may play some role in regulating innate immunity9. Our group has shown that brain inflammation, a
common theme in many neurodegenerative disorders10, can trigger the release of EVs from astrocytes which primes
the infiltration of immune cells into the brain via upregulation of cytokines in the periphery11. Taken together,
inhibition of EV secretion through inhibition of nSMAse2 is emerging as a novel avenue for the treatment of
diseases associated with aberrant exosomal intercellular communication11?13. Unfortunately, limitations of
currently available nSMase2 inhibitors have prevented a detailed evaluation of the role of nSMase2 in disease
models and the advancement of drug-like nSMase2 inhibitors to the clinic. Currently available nSMase2 inhibitors
have low potency (IC50?s in ? M level), poor aqueous solubility, and/or limited brain penetration. GW486914,
the most widely used inhibitor, has low inhibitory potency (IC50 = 1 ? M) in biochemical assays and very poor
solubility (practically insoluble in water with poor solubility in organic solvents such as DMSO (0.2 mg/ml).
These attributes have hampered GW4869?s clinical development. Cambinol, an inhibitor our group identified
from a pilot screen of commercially available small chemical libraries15 showed better solubility, but it was
metabolically unstable and exhibited a poor in vivo pharmacokinetic profile. Chemistry efforts by our laboratory to
improve cambinol?s potency (IC50 = 5 ?M) and stability were unsuccessful. Herein, we report on a high
throughput screening (HTS) campaign of over 365,000 compounds that identified a potent inhibitor of nSMase2 termed
DPTIP, with an excellent pharmacokinetic profile including significant brain penetration, which was capable of
dose-dependently blocking EV release from primary astrocytes. Moreover, in a mouse model of brain
inflammation that recapitulates common features of neurodegenerative diseases, DPTIP potently inhibited IL-1?-induced
ADEV release, peripheral cytokine upregulation and neutrophil migration into the brain.
Results and Discussion
Development of a 1536-well cell-free human recombinant nSMase2 enzyme activity
assay. Human nSMase2 catalyzes the hydrolysis of sphingomyelin (SM) to phosphorylcholine and ceramide.
As we reported previously, we used the Amplex Red system to monitor nSMase2 activity15. In this reaction, one
of the enzymatic products, phosphorylcholine, is stoichiometrically converted through a series of
enzyme-coupled reactions to fluorescent resorufin, so that fluorescence signal is directly proportional to nSMase2
activity (Fig.?1A). An enzymatic assay protocol was developed in 1536-well format for implementation for HTS.
Several parameters were first optimized through the measurement of the fluorescence signal. Fluorescence
signal increased with longer times of incubation (15?150 min) and increasing nSMase2 concentrations (0.03 to
0.5 ? g protein/mL) at a constant SM concentration (20 ? M) (Fig.?1B). Similarly, fluorescence signal increased with
longer time of incubation (30?150 min) and increasing SM concentrations (5?40 ?M) at a constant enzyme
concentration (0.063 ? g protein/ml) (Fig.?1C). Based on these results, we chose 0.1 ? g protein/mL human nSMase2
cell lysate, 20 ? M SM in a total volume of 4 ? L and 2 h incubation at 37 ?C to assess assay performance in HTS
format. Under these conditions, reaction rate was linear with a robust fluorescence signal of approximately 2500
relative fluorescent units (RFU). Cambinol was used as the positive inhibitor control15; it was pre-incubated with
human nSMase2 for 15 min prior to addition of SM. Final DMSO concentration was 0.57%. The assay exhibited
signal/background = 21 and Z? = 0.8 (Fig.?1D). We also evaluated the dose response of inhibition by cambinol
and GW4869 to determine variability in the IC50 values from plate to plate. GW4869 was insoluble in DMSO and
appeared as a yellow pellet at the 3 highest concentrations so it was excluded as a positive control. Cambinol?s
average IC50 from 4 independent determinations was 27 ? 1 ? M (Fig.?1E). The final stage of validation of the
assay for HTS was the screening of the Library of Pharmacologically Active Compounds (LOPAC) in 1536-well
plates using the same assay conditions at four different inhibitor concentrations (0.4, 2, 11 and 57 ? M). Overall,
the sample field was even, there were no plate positional effects and the number of active hits increased as the
HTS campaign and data analysis of hits led to the identification of seven potent nSMase2
inhibitors. Following assay validation, we screened 365,000 compounds from the Molecular Libraries Small
Molecule Repository (MLSMR) and 2816 compounds from the NCGC pharmaceutical collection (NPC) library
for human nSMase2 inhibitors. Compounds were screened at 4 concentrations: 1.1, 11, 57 and 114 ? M. Cambinol
(full dose response in each plate) was used as positive control. After eliminating promiscuous compounds, 1990
compounds that had maximal inhibitory responses >50% at the highest concentration tested and robust curve
response classes (CRC)16 were selected for re-testing in the same human nSMase2 activity assay and counter
screen. The purpose of the counter screen was to identify false positives, i.e., compounds that inhibited the
enzyme-coupled reactions of the assay system; it was carried out in the absence of human nSMase2 and SM
and using phosphorylcholine as substrate. Out of the 1990 compounds, 1782 (90%) were confirmed in the 7
dose-response hnSMase2 confirmatory assay, but most (1718; 86%) were found to be false positives in the
counter screen, resulting in 64 bona fide nSMase2 inhibitors. We also considered the difference between potency and
response in the counter screen to select 156 additional hits that showed robust inhibition of the overall reaction,
but were weakly active in the counter screen. There were a total of 220 compounds for follow-up confirmation
(Fig.?2A). Out of the 220 compounds tested, 7 compounds exhibited dose responses with IC50 ? 10 ?M that were
also inactive in the counter assay (Fig.?2B).
DPTIP is the most potent nSMase2 inhibitor reported to date. Filtering of the HTS hits as
outlined above resulted in the identification of MLS000523327 or DPTIP
(2,6-Dimethoxy-4-(5-Phenyl-4-Thiophen2-yl-1H-Imidazol-2-yl)-Phenol) as the most promising compound based on potency and chemical optimization
feasibility. The IC50 for DPTIP using an extended inhibitor concentration range (10 pM ? 100 ? M) was 30 nM
(Fig.?3A). This IC50 is 30- and 160-fold more potent than the prototype inhibitors GW4869 (1 ? M)14 and
cambinol (5 ? M)15. To our knowledge, this is the first nSMase2 inhibitor described with nanomolar potency. Because
DPTIP contains a hydroxyl group which could be a metabolic liability in vivo (Fig.?3A), we determined the
importance of this group for inhibitory activity. We synthesized the des-hydroxyl analog of DPTIP (Fig.?3B) and showed
that it was inactive against human nSMase2 (IC50 > 100 ?M) (Fig.? 3B). These results demonstrate the importance
of the hydroxyl group for inhibition, and also provide a structurally similar inactive DPTIP analog for use as a
comparison compound in subsequent pharmacological assays.
DPTIP exhibited non-competitive mode of inhibition and showed selectivity for nSMase2
versus related enzymes. DPTIP exhibited the hallmarks of noncompetitive inhibition; when the rate of
reaction with respect to SM concentrations was monitored at increasing inhibitor concentrations, there was a decrease
in maximal rate (Vmax) while the Michaelis constant (Km) was unchanged (Fig.?3C). Vmax and Km for each data set
at a given inhibitor concentration were obtained from non-linear regression fits to Michaelis-Menten kinetics
DPTIP did not inhibit members of two related enzyme families including alkaline phosphatase (IC50 > 100 ? M
in counter screen), a phosphomonoesterase, or acid sphingomyelinase (IC50 > 100 ? M), a phosphodiesterase
closely related to nSMase2 (results not shown). Inhibitor selectivity with respect to enzymes from related families
is consistent with a noncompetitive mode of inhibition, as DPTIP is likely acting at a site other than the catalytic
site. Additional data also indicate that DPTIP exhibits specificity for nSMase2; DPTIP has been screened in 759
bioassays assays at NCATS and only weak activity (2?50 ?M) was observed in 19 (2.5%) of these assays. (https://
DPTIP showed metabolic stability in mouse and human liver microsomes. One potential liability
when using chemical probes in vivo is lack of metabolic stability which structurally inactivates the compound
before it can reach its molecular target. We evaluated DPTIP for metabolic stability using human and mouse
liver microsomes as we have previously described17. Percent of drug remaining over time was determined by
liquid chromatography?tandem mass spectrometry analysis (LC/MS/MS). In the presence of NADPH, DPTIP
remained intact (100% remaining at 1 h) in both mouse and human liver microsomes (Fig.?4A) indicating that the
compound is not affected by CYP-450-mediated metabolism. These in vitro results indicate DPTIP does not have
major liver metabolic liabilities that would preclude its use as an in vivo probe.
DPTIP exhibited plasma exposure and brain penetration after systemic dosing in mice. In the
next set of experiments, we evaluated the in vivo pharmacokinetic profile of systemically administered DPTIP.
Mice were given DPTIP (10 mg/kg IP) and plasma and brain levels of DPTIP were measured at 0.25, 0.50, 1, 2, 4
and 6 h post dose (n = 3 per time point). DPTIP peak concentration in both plasma and brain was at 0.5 h (Cmax
plasma = 11.6 ? 0.5 ? M; Cmax brain = 2.5 ? M) (Fig.?4B). The AUC0-? of DPTIP in plasma and brain was 10 ? 1
and 2.6 ? 0.5 ? M*h, respectively, resulting in an AUCbrain/AUCplasma = 0.26. Brain levels of DPTIP exceeded its
IC50 for inhibition of nSMase2 up to 4 h following 10 mg/kg systemic dosing (Fig.?4B).
DPTIP inhibited EV release from primary astrocytes whereas its inactive analog had no
effect. Independent laboratories have shown that pharmacological and genetic inhibition of nSMase2 blocks
EV secretion from glial cells12. Consequently, we evaluated DPTIP for its ability to inhibit EV release from
primary glial cells in vitro. Mouse primary astrocytes were activated by FBS withdrawal as we have previously
described11 and treated with DPTIP or its closely related inactive des-hydroxyl analog (Fig.?5A) at a dose range
of concentrations (0.03?30 ? M) using DMSO (0.02%) as vehicle control. Two hours after treatment, EVs were
isolated from the media and quantified by nanoparticle tracking analysis. DPTIP inhibited EV release from
astrocytes in a dose dependent manner (Fig.?5A). In contrast, its closely related inactive analog had no effect on EV
release suggesting DPTIP inhibits EV release via inhibition of nSMase2. We also determined the activation status
of (+/?) serum-deprived astrocytes after DPTIP treatment. Rat primary astrocytes were treated with DPTIP (10
?M) or inactive analog for two hours along with (+/?) serum deprivation-induced stress. Cells were fixed and
immunofluorescence labeling for GFAP was performed. DPTIP and inactive analog without serum starvation
did not change GFAP levels (Fig.?5B,C). Serum deprivation resulted in activation of astrocytes as evidenced by
increase in GFAP fluorescence intensity compared to non-treated controls. Treatment with DPTIP prevented
astrocyte activation in response to serum starvation, while the inactive analog failed to prevent astrocyte
DPTIP inhibited biomarkers of brain inflammation in vivo whereas its inactive analog had no
effect. Given DPTIP?s brain penetration in mice and its ability to inhibit EV release in vitro, we next
evaluated the ability of DPTIP to ameliorate EV release from astrocytes, cytokine upregulation in liver and neutrophil
migration into brain in an in vivo mouse model of brain inflammation. As we have previously shown11,18, striatal
injection of IL-1? in mice expressing GFP-GFAP in astroglia triggers a release of GFP-labelled EVs that rapidly
enter into plasma, resulting in cytokine upregulation in liver and peripheral immune cell migration into brain11.
Mice were dosed (10 mg/kg IP, DPTIP or inactive analog) 0.5 h prior to IL-1? striatal injection. At this dose, brain
concentrations of DPTIP are above its IC50 for nSMase2 inhibition for at least 4 h after compound administration
(Fig.?4C). There were two groups of mice: the first group was sacrificed 2h after IL-1? administration by heart
puncture, and GFP-labeled circulating EVs were measured with liver cytokines. Mice in the second group were
dosed a second time with DPTIP or inactive analog at 12 h and sacrificed at 24 h after IL-1? administration to
measure brain neutrophils (Fig.?6A). Counting of astrocyte-released EV (GFP+) from blood and liver cytokine
analysis was conducted by single injection of DPTIP. Although release of EVs from astrocytes can be initiated
immediately after intracranial injection of IL-1?, infiltration of neutrophils in brain parenchyma occurred
12h24h after the IL-1? injection. Since the pharmacokinetic profiles of DPTIP in plasma and brain following 10mg/
kg IP dose showed that brain levels of DPTIP exceeded its IC50 for nSMase2 for only 4 h post dose, we
administered DPTIP twice after IL-1? injection to ensure inhibition of nSMase2 was sustained during the experiment.
When mice were dosed with DPTIP, number of astrocyte-derived EVs was reduced by 51 ? 13% 2 h post IL-1?
administration (Fig.?6B). Western analysis using the isolated exosomal fraction confirmed the presence of CD63
(transmembrane protein), TSG101 (cytosolic protein) and Flotilin-1 (lipid raft associated protein), commonly
used EV markers19,20. The GFP signal was an indication that these EVs originated in brain11 while lack of
mitofilin and ?-actinin signals indicated the vesicles were not of mitochondrial21 or cytoskeletal22 origin respectively
Upregulation of liver cytokines upon IL-1? treatment was inhibited by DPTIP (Fig.?6C). Neutrophils, as
measured by immunohistochemistry of coronal brain sections using LY6b antibody, showed reduced
staining in sections from animals treated with DPTIP compared to IL-1?-treated animals (Fig.?6D);
corresponding quantification showed neutrophil migration into brain was reduced by 80 ? 23% compared to IL-1?-treated
animals (Fig.?6E). Administration of the closely related inactive analog, had no statistically significant effect on
IL-1?-induced EV release (Fig.?6B). The effects of the inactive des-hydroxyl DPTIP on production of TNF-? and
IL-6 were marginal and not statistically significant. Although the magnitude of reduction in CCL2 production
by the inactive analog was high, the data were variable and also not statistically significant (Fig.?6C). Finally,
des-hydroxyl DPTIP had no effect on neutrophil migration (Fig.?6D,E). Results with the inactive analog were
consistent with the suggestion that DPTIP effects occur through nSMase2 inhibition. Importantly, these results
are in agreement with our previous findings that co-injection of IL-1? with nSMase2 inhibition (either GW4869,
altenusin, lentivirus targeting astrocytic nSMase2, or using nSMase2 KO mice) suppress neutrophil infiltration
into brain parenchyma11. The same studies also indicated that nSMase2 inhibition suppressed activation of
astrocytes and microglia11.
Within this study, we focused our efforts on astrocytes because of the intimate association of these cells with
the blood-brain barrier (BBB), and because in our previous study we knocked down nSMase2 expression
selectively in astrocytes and showed that this inhibited the release of astrocyte-derived EVs (ADEVs) and prevented
the liver cytokine response, and leukocyte trafficking into brain following parenchymal injection of IL-1beta11.
Although it remains possible that neuronal or microglial- derived EV are also affected by nSMase2 inhibition,
these earlier findings suggest that ADEVs are a major source of brain EVs that regulate the peripheral response to
CNS injury. Future studies will include the use of neuronal and microglial derived EVs.
The exact mechanism of serum deprivation-induced EV release is not known. Serum deprivation is known to
produce a stress response that stimulates secretory pathways in astrocytes23. Additionally, nutrient deprivation
has been shown to cause accumulation of ceramides in astrocytes, likely due to a stress response activation of
nSMase224. Nutrient starvation has been reported to increase nSMase2 activity and induce its expression in other
cell types25. Serum deprivation induced EV release observed in our experiments may therefore be the result of
nSMase2 activation in response to nutrient deprivation stress.
A schematic illustration of the in vivo experiment are shown in Fig.?7 which are consistent with the data
detailed above as well as previous literature. In brief, striatal IL-1? injection activates the IL-1? receptor on the
plasma membrane of astrocytes that in turn activates nSMase2 enzymatic activity to catalyze the hydrolysis of
sphingomyelin to produce ceramide26. Ceramide is used to manufacture intracellular vesicles (IVs)1 that are
released from astrocytes as EVs and migrate into plasma where they induce a peripheral acute cytokine response,
mainly in liver, and prime immune cells to transmigrate to the brain11. In the presence of DPTIP, inhibition of
nSMase2 prevents ceramide production, EV formation and secretion (Fig.?6B) cytokine upregulation (Fig.?6C)
and neutrophil migration (Fig.?6D).
In summary, DPTIP is the most potent nSMase2 inhibitor identified to date (IC50 30 nM), exhibits selectivity,
is metabolically stable and brain penetrant. DPTIP is an inhibitor of EV release in primary glial cells and in vivo.
In addition, biomarkers that have been associated with EV release from brain, including cytokine upregulation
and immune cell migration to brain, were also inhibited by DPTIP. The des hydroxyl inactive analog of DPTIP
did not inhibit EV release in vitro and had no effect on IL-1?-induced cytokine regulation or neutrophil migration
to brain in vivo. DPTIP is a considerable improvement over other nSMase2 inhibitors identified to date, it can be
used as a probe in animal models of disease associated with EV dysregulation and it contains a structural scaffold
that is actively being optimized for clinical translation.
Expression of human nSMase2. Full length human nSMase2 cDNA with a C-terminal Flag tag cloned
into a pCMV6-Entry expression vector (Origene) was transfected into HEK293 cells using lipofectamine 2000
(Life Technologies). Selection of transfected cells was carried out for two weeks with 500 ? g/ml G418 in EMEM
containing 10% FBS (ATCC) and 2 mM glutamine (Life Technologies). Expression of human nSMase2 was
confirmed by Western-blot analysis using an antibody specific against nSMase2 (R&D) diluted to 0.4? g/ml in
Tris-buffered saline with 0.1% Tween 20 and 5% bovine serum albumin. Cells expressing human nSMase2 were
grown to confluency in 150 mm dishes, washed twice with cold PBS and harvested using a cell scraper in lysis
buffer pH 7.5, Tris-HCl 100 mM, 1 mM EDTA, 100 mM sucrose, 100 ?M PMSF, 1X protease inhibitor cocktail III
(Calbiochem), 1 ml per dish. Cell lysis was achieved by sonicating 3 times on ice for 30 sec. Protein concentration
was determined using the bicinchoninic acid (BCA) assay. Aliquots of cell lysate were snap frozen and stored at
?80 ?C. Activity of recombinant human nSMase2 from cell lysates remained stable for at least six months.
Fluorescence-based nSMase2 activity assay in 1536-well format. Measurements of nSMase2
activity using fluorescence as readout was optimized for dose response quantitative HTS (qHTS). The assay was
carried out in black solid bottom, medium binding, 1536-well plates (Greiner, 789176-F). Fluorescence response
was optimized with respect to nSMase2 concentration, incubation time and SM concentration. Recombinant
human nSMase2 preparations (2 ? L) at various concentrations (0.03 to 0.5 ? g protein/? L solution) were
incubated with substrate/detection reaction mixture (2 ? L) containing various concentrations of SM (5 to 40 ?M),
coupling enzymes (alkaline phosphatase 4 U/mL, choline oxidase 0.1 U/mL and horseradish peroxidase 0.1 U/
mL) and Amplex red? (50 ?M). Hydrolysis of SM was carried out for different incubation times (15?160 min) at
37 ?C in pH 7.4 Tris-HCl buffer 100 mM, containing 10 mM MgCl2 and 0.2% Triton X-100. Phosphorylcholine
made during the nSMase2-catalyzed reaction is dephosphorylated by alkaline phosphatase to produce choline,
which in turn undergoes oxidation in the presence of choline oxidase to produce betaine and peroxide. Peroxide
in the presence of horseradish peroxidase and Amplex Red generates fluorescent resorufin (Ex 525/Em 598).
Resorufin was monitored with Viewlux ?HTS Microplate Imagers (Perkin Elmer) at energy levels 1,000 or 3,000
and exposure times of 1 or 2 sec. Fluorescence readings varied when using Viewlux offline (assay
characterization) vs. Viewlux online (HTS); in order to account for differences in fluorescence efficiency, assay performance
was monitored from machine to machine based on assay dynamic range and cambinol IC50 reproducibility. Based
on results of the different conditions outlined above, the HTS campaign was carried out using 0.1 ? g protein/
? L nSMase2 preparation, 20 ?M SM and 2 h time of incubation. Control inhibitors or test compounds (23 nL)
were added from various concentrations in DMSO solution into to the nSMase2 preparation and incubated for
15 min prior to the addition of substrate and enzyme-coupling detection reagents. Compounds were screened in
4 doses, starting at 57 ?M, and doing 5-fold dilutions. A customized screening robot (Kalypsys) was used for the
primary screen. A step-by step HTS assay protocol is given in the Supplementary Data (Table?S1). Inhibitors of
nSMase2 were selected using compound dose response curve algorithms developed at NCGC to score actives,
which assigns each tested compound a compound response class (CRC) number16. This method classifies primary
hits into different categories according to their potency (IC50), magnitude of response (efficacy), quality of curve
fitting (r2), and number of asymptotes. For example, CRC of ?1.1 represents complete curve and high efficacy;
CRC of ?1.2 represents complete curve but partial efficacy. Compounds with CRCs of ?1.1, ?1.2, ?2.1 and
?2.2 were generally selected for confirmation and validation. Structural analysis of selected compounds was
performed and promiscuous compounds were filtered out. A counter-assay to rule out compounds that inhibited
the detection reaction was carried out in the absence of human nSMase2. The reaction was initiated with addition
of phosphorylcholine (alkaline phosphatase substrate), added at a final concentration of 2 ?M. Compounds that
showed inhibitory activity in the counter-assay were removed from further validation.
IC50 determination of selected compounds. Human nSMase2 (0.1 ?g protein/?L) was added to a
reaction mixture containing SM (20 ?M), and detection reagents as indicated above and different compound
concentrations in the 10 pM ? 100 ? M range in a total volume of 100 ? L (96-well format). Kinetic measurements were
obtained from 2 h traces when the reaction was linear. Percent inhibition was obtained using the formula [(rate
of change of fluorescence in the presence of inhibitor divided by rate of change of fluorescence in the absence of
inhibitor) ? 100].
Synthesis and characterization of DPTIP and des-hydroxyl DPTIP. Detailed descriptions of the
synthesis of DPTIP and its des-hydroxyl analog along with corresponding authentication information are given
in Supplementary Data.
Metabolic stability. Metabolic stability assay was conducted in mouse or human liver microsomes as we
have described previously with minor modifications17. Briefly, the reaction was carried out using potassium
phosphate buffer (100 mM, pH 7.4), in the presence of an NADPH regenerating system (compound final
concentration was 1 ?M; 0.2 mg/mL microsomes). Compound disappearance was monitored over time using a
liquid chromatography and tandem mass spectrometry (LC/MS/MS) method. Chromatographic analysis was
performed using an Accela ultra high-performance system consisting of an analytical pump and an autosampler
coupled with a TSQ Vantage mass spectrometer (Thermo Fisher Scientific Inc., Waltham, MA). Separation of
analyte was achieved at ambient temperature using Agilent Eclipse Plus column (100 ? 2.1 mm i.d.) packed with
a 1.8 ?m C18 stationary phase. The mobile phase consisted of 0.1% formic acid in acetonitrile and 0.1% formic
acid in water with gradient elution. The [M+ H]+ ion transition of DPTIP (m/z 378.956 ? 363.073, 200.055) and
losartan (IS) (m/z 423.200 ? 207.107, 180.880).
In vivo pharmacokinetics. Pharmacokinetic studies in mice were approved by the Animal Care and Use
Committee at Johns Hopkins University. Male CD1 mice between 25 and 30 g were obtained from Harlan and
maintained on a 12 h light?dark cycle with ad libitum access to food and water. Test compounds were dosed at
10 mg/kg IP at a dosing volume of 10 mL/kg. Blood and brain tissue were collected at 0.25, 0.5, 1, 2, 4 and 6 h post
dose (n = 3 per time point). Blood was obtained via cardiac puncture and plasma was harvested from blood by
centrifugation at 3000 ? g for 15 min and stored at ?80 ?C. Brain tissues were harvested following blood
collection and immediately snap frozen in liquid nitrogen and stored at ?80 ?C until LC?MS analysis. Calibration
standards were prepared using na?ve mouse plasma or brain spiked with DPTIP. DPTIP standards and samples
were extracted from plasma and brain by a one-step protein precipitation using acetonitrile (100% v/v) containing
internal standard (losartan: 0.5 ?M). The samples were vortex mixed for 30 secs and centrifuged at 10000 ? g for
10 min at 4 ?C. Fifty microliter of the supernatant was diluted with 50 ? L water and transferred to a 250 ? L
polypropylene vial sealed with a Teflon cap and analyzed via LC/MS/MS as described above. Plasma concentrations
(pmol/mL) as well as tissue concentrations (pmol/g) were determined and plots of mean plasma concentration
versus time were constructed for PK analysis. Non-compartmental-analysis modules in Phoenix WinNonlin
version 7.0 (Certara USA, Inc., Princeton, NJ) were used to assess pharmacokinetic parameters including maximal
concentration (Cmax), time to Cmax (Tmax), and area under the curve extrapolated to infinity (AUC0-?).
Inhibition of EV release from primary glial cells. Potential inhibition of test compounds on EV release
from primary astrocytes was carried out as previously described
(Dickens et al., 2017)
. Briefly, rat primary
astrocytes were seeded onto 6-well plates at a density of 20,000 cells/well. Twenty-four hours after seeding, astrocytes
were washed with PBS and the medium changed to media without FBS. Absence of FBS mimics a trophic factor
withdrawal stimulus causing EVs to be released from astrocytes via an nSMase2-dependent pathway. Astrocytes
were then treated with test compounds at different concentrations: 0.03, 0.1, 0.3, 1, 3, and 10 ?M. DMSO (0.02%)
was used as control. Two hours after treatment, media was collected and centrifuged at 2700 ? g for 15 min at 4 ?C.
The supernatant was collected and the number of EVs quantified using ZetaView Nanoparticle Tracker (Particle
Metrix GmBH, Meerbusch, Germany) and the corresponding ZetaVeiw software (8.03.04.01). Nanosphere size
standard 100 nm (Thermo Scientific) was used to calibrate the instrument prior to sample readings. Instrument
pre-acquisition parameters were set to 23 ?C, a sensitivity of 65, a frame rate of 30 frames per second (fps), a
shutter speed of 100, and laser pulse duration equal to that of shutter duration. Post-acquisition parameters were set
to a minimum brightness of 25, a maximum size of 200 pixels, and a minimum size of 10 pixels. For each sample
1 mL of the supernatant was injected into the sample-carrier cell and the particle count measured at 5 positions,
with 2 cycles of reading per position. The cell was washed with PBS after every sample. Mean concentration of
EVs/mL (?SEM) was calculated from 4 replicates.
Inhibition of EV release in vivo. All experimental protocols using vertebrate animals were reviewed by
the Institutional Animal Care and Use Committee at Johns Hopkins University and are in accordance with the
guidelines of the NIH guide for the care and use of laboratory animals. Striatal injections and EV measurements
were performed as previously described by our group in adult (2?3 month) male GFAP-GFP mice (Jackson
Laboratories)11,18. Mice were anesthetized with 3% Isoflourane (Baxter) in oxygen (Airgas), and placed in a
stereotaxic frame (Stoelting Co.). A small burr hole was drilled in the skull over the left striatum using a dental drill
(Fine Scientific Tools). IL-1? (0.1 ng/3 ? L) was injected (total volume of 3 ?L) at the rate 0.5 ? L/min via a pulled
glass capillary tip diameter <50 ? m18; using the stereotaxic coordinates: A/P + 0.5; M/L ?2; ?3 D/V. Saline was
used as a control. When DPTIP or its des-hydroxyl analog were used, they were given IP (10 mg/kg, 5% DMSO,
5% Tween-80 in saline) 30 min before IL-1? injection. Following infusion, the capillary was held in place for 5 min
to allow for solution to diffuse into the tissue. Animals were sacrificed at 2h by an overdose of anesthetic, and
transcardially perfused with ice-cold saline containing heparin (20 ? L per 100 ml, Sigma). Blood was collected
via cardiac puncture using a heparin (Sigma Aldrich) coated syringe and EDTA tubes (BD) 2 h following
striatal injections. Blood was immediately centrifuged at 2,700 x g for 15 min (20 ?C) to obtain plasma. Plasma was
further centrifuged at 10,000 g for 15 min (4 ?C) to generate platelet free plasma. This procedure removes large
particles such as apoptotic bodies.
Quantitation of Plasma EVs. Dynabeads M-450 Epoxy (Invitrogen) were coupled with anti-GFP antibody
(Thermo Fisher) at a ratio of 200?g antibody per 4 ? 108 beads. Plasma from GFAP-GFP mouse (50 ?L) was
incubated with 2 ? 107 anti-GFP antibody-coupled Dynabeads at 4 ?C overnight. The beads were washed and
placed on a magnet to separate EVs bound to anti-GFP antibody-coupled Dynabeads. The precipitated EVs were
eluted using 0.1 M glycine (pH 3.0). The concentration of immunoprecipitated GFP+ EVs was quantified using
ZetaView nanoparticle tracking analysis (Particle Metrix) as described above.
Western analysis. Proteins were resolved by 10% SDS?polyacrylamide gel electrophoresis and transferred to
polyvinylidene difluoride membranes (Bio-Rad). Nonspecific binding sites were blocked with 5% (w/v) milk in
TBS containing 0.1% Tween 20 (TBS-T). After blocking, blots were incubated overnight with the primary
polyclonal antibodies to GFP (1:1000; Thermo Fisher) CD63 (1:200; Santa Cruz Biotechnology), flotillin1 (1:1000;
Abcam), TSG101 (1:1000; BD Biosciences), mitofilin (1:5000 Thermo Fisher Scientific) and ?-actinin (1:1000;
Gentex). After washes with TBS-T, blots were incubated for 2 h with the corresponding IgG horseradish
peroxidase?linked secondary antibody (1:1000; Cell Signaling Technology) and developed by enhanced
chemiluminescence. Image analysis was performed using a G: BOX imaging system (Syngene).
Cytokine measurements. RNA was isolated from fresh frozen tissues (10 to 50 mg) using the RNeasy Mini Kit
(Qiagen). Total RNA was reverse-transcribed and quantified using previously published methods27. For
quantitative real-time PCR (qRT-PCR), each reaction contained SYBR Green Master Mix (12.5 ml; Life Technologies),
diethyl pyrocarbonate H2O (10.5 ml), forward and reverse primers to CCL2, TNF?, IL-6, IL-1b, IL-17a, IL-10,
IGFR1, and CXCL1 (0.5 ml each; Sigma-Aldrich), and cDNA (1 ml). Each 96-well plate included a nontemplate
control, and samples were analyzed in triplicate on an Applied Biosystems 7300 (Life Technologies). Cycling
parameters were as follows: one cycle for 2 min at 50 ?C, one cycle for 10 min at 95 ?C, and 40 cycles for 15 s at
95 ?C and for 1 min at 60 ?C. The change in threshold cycle (?Ct) for each sample was normalized to ?-actin, and
??Ct was calculated by comparing ?Ct for the treatment group to the average ?Ct of the control group28.
Immunohistochemistry. Coronal brain sections (30 ? m) were prepared using a cryostat microtome (Leica).
Endogenous peroxidase activity was quenched using a 1% solution of H2O2 in methanol, and primary antibody
Ly6b (1:1000, AbD Serotec), was incubated at 4 ?C overnight. Sections were washed (3 ? PBS), and biotinylated
secondary antibody (1:100, Vector Laboratories) was added at room temperature for 2 hours. Staining was
visualized using an avidin-biotin complex (1:100 of A and B, Vector Laboratories) and DAB-HCl using a microscope
to monitor staining progression. Stereological quantitation was performed using a one-in-five series (270-? m
spacing), from the rostral point of bregma +1.10 mm to the caudal point of bregma ?0.58 mm as previously
Ethical approval. All experimental protocols using vertebrate animals were reviewed by the Institutional
Animal Care and Use Committee at Johns Hopkins University and are in accordance with the guidelines of the
NIH guide for the care and use of laboratory animals. Johns Hopkins Medical Institution is fully accredited by the
American Association for Accreditation in Laboratory Animal Care (AAALAC).
Data Availability Statement
Experimental data used to generate the results reported in this manuscript are available upon request.
This research was supported by NIH grants RO1 MH107659 (CR) and P30 MH075673 (BSS and NH).
C.R., J.M., R.R., T.T., Norman H., M.F. and B.S.S. were responsible for experimental design and writing of
manuscript. E.B., X.H., N.S. were responsible for H.T.S. and data analysis. Nyada H. and O.S. were responsible
for chemical synthesis and authentication. A.G.T. carried out IC50 determinations and mechanism of inhibition
studies. Y.W., S.C.Z. and A.G.G. performed metabolic stability and PK studies. ADC carried out in vitro inhibition
of EV secretion in glial cells. S.-W.Y. was responsible for in vivo studies using GFAP-GFP mice and for the
illustration in Figure?7.
Supplementary information accompanies this paper at https://doi.org/10.1038/s41598-018-36144-2.
Competing Interests: The authors declare no competing interests. Publisher?s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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