Scholarly article on topic ' Preferential Extracellular Generation of the Active Parkinsonian Toxin MPP + by Transporter-Independent Export of the Intermediate MPDP + '

Preferential Extracellular Generation of the Active Parkinsonian Toxin MPP + by Transporter-Independent Export of the Intermediate MPDP + Academic research paper on "Biological sciences"

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Antioxidants & Redox Signaling
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Academic research paper on topic " Preferential Extracellular Generation of the Active Parkinsonian Toxin MPP + by Transporter-Independent Export of the Intermediate MPDP + "

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Original Research Communication Preferential extracellular generation of the active parkinsonian toxin MPP+ by transporter-independent export of the

<3 [£ Stefan Schildknecht,1* Regina Pape,1* Johannes Meiser,2 Christiaan Karreman,1 Tobias

_l_ ^ 3 3 3 3 4

Ph13 Strittmatter, Meike Odermatt, Erica Cirri, Anke Friemel, Markus Ringwald, Noemi

Pasquarelli,5 Boris Ferger,5 Thomas Brunner,6 Andreas Marx,3 Heiko M. Möller,7

2 1 Karsten Hiller, Marcel Leist

ja h 1In vitro Toxicology and Biomedicine, Department of Biology, University of Konstanz, D-

¿J § 78457 Konstanz, Germany;

o | Metabolomics Junior Research Group, Luxembourg Centre for Systems Biomedicine,

o g University of Luxembourg, L-4362 Esch-Belval, Luxembourg;

§*o 3Department of Chemistry and Konstanz Research School Chemical Biology, University of

Konstanz, D-78457 Konstanz, Germany;

4MCAT GmbH, Hermann-von-Vicari-Str. 23, D-78464 Konstanz, Germany; 5CNS Disease Research, Boehringer Ingelheim Pharma GmbH & Co. KG,

D-88397 Biberach an der Riss, Germany; 1 ^ 6Biochemical Pharmacology, Department of Biology, University of Konstanz, D-78457

^ Konstanz, Germany;

42 Ja Is 7Institute of Chemistry, University of Potsdam, D-14476 Potsdam, Germany

S These authors contributed equally

" Running head: MPDP export from astrocytes

■ — ¡a

•l"^ Word count:

§•.§ Reference numbers: 56

^ Number of greyscale illustrations: 4

Number of color illustrations: 2


J To whom correspondence should be addressed:

■M (U t+H O

g| Stefan Schildknecht, PhD

id ^ University of Konstanz

§ jg Department of Biology

£ In vitro Toxicology and Biomedicine

1 § PO Box M657

Universitätsstr. 10

£ § 78457 Konstanz, Germany

J Fax: +49-7531-88-5039

13 Tel: +49 7531-88-5053

f E-mail:

2 Key words: astrocytes, autoxidation, parkinsons disease, MPDP , MPP


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£ Aims: 1-methy-4-phenyl-tetrahydropyridine (MPTP) is among the most widely used

oo ^ neurotoxins for inducing experimental Parkinsonism. MPTP causes Parkinsonian symptoms

"H § in mice, primates, and humans by killing a subpopulation of dopaminergic neurons.

Extrapolations of data obtained using MPTP-based Parkinsonism models to human disease are common; however, the precise mechanism by which MPTP is converted into its active neurotoxic metabolite 1-methyl-4-phenyl-pyridinium (MPP+) has not been fully elucidated. In


' Z rf this study, we aimed to address two unanswered questions related to MPTP toxicology: 1)

is why are MPTP-converting astrocytes largely spared from toxicity; and 2) how does MPP+

^ 13 reach the extracellular space? Results: In MPTP treated astrocytes, we discovered that the

'■3 § +

g fxM membrane-impermeable MPP , which is generally assumed to be formed inside astrocytes, is


almost exclusively detected outside of these cells. Instead of a transporter-mediated export, we found that the intermediate MPDP+ and/or its uncharged conjugate base passively diffused across cell membranes and that MPP+ was formed predominately by the extracellular ^ s « oxidation of MPDP+ into MPP+. This non-enzymatic extracellular conversion of MPDP+ was

g promoted by O2, a more alkaline pH, and dopamine autoxidation products. Innovation and

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converting astrocytes, and provide a rationale for the preferential formation of MPP+ in the

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extracellular space. The mechanism of transporter-independent extracellular MPP formation

S Conclusion: Our data indicate that MPTP metabolism is compartmentalized between

intracellular and extracellular environments, explain the absence of toxicity in MPTP-

described here indicates that extracellular genesis of MPP+ from MPDP is a necessary

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prerequisite for the selective uptake of this toxin by catecholaminergic neurons.


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£ The experimental study of dopamine (DA) neuron degeneration (a hallmark of Parkinson's

oo ^ disease) relies on toxicants that specifically induce pathological states reminiscent of human

§ Parkinsonism by inducing the selective death of DA neurons. Among the chemical tools

+-T3 currently used to induce DA neuron degeneration, MPTP (1-methyl-4-phenyl-1,2,3,6-

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tetrahydropyridine) most closely reproduces human Parkinsonian pathology in mice and primates (3,31). MPTP is a lipophilic and non-cytotoxic prodrug that crosses the blood brain

J rf barrier and is then converted into the active toxicant MPP+ (1-methyl-4-phenylpyridinium)

tt 13 (23,32,54). Once MPP+ reaches

a critical concentration in the cerebrospinal fluid, it acts as a

2 molecular 'Trojan Horse' in DA neurons as result of its selective uptake by the DA

_ a M transporter (DAT) (26). Unlike DA, MPP accumulates in mitochondria, where it acts as an


inhibitor of complex I of the respiratory chain (38), and thereby kills its 'host cell' (10). However, a vexing enigma pertaining to the neurotoxic mechanisms of MPTP remains unsolved. MPTP is converted to MPP+ in astrocytes by the enzyme monoamine oxidase-B 2 s « (MAO-B), which is localized on the outer mitochondrial membrane of astrocytes (44). How

MPP exits astrocytes rather than accumulating in their mitochondria is unknown.

gs Due to its charge, the active toxicant MPP cannot cross membranes. Recent reports have

suggested that passive transporters, such as the family of organic cation transporters (OCT) or

the plasma membrane monoamine transporter (PMAT), are necessary for MPP+-efflux from

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g,^ astrocytes (9,37). OCT3 was observed by Cui et al. to be preferentially expressed in glial cells

> in the vicinity of midbrain DA neurons that degenerated in response to MPTP treatment, and

it was hypothesized that a preferential export of intracellular astrocytic MPP+ via OCT3 might

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^« contribute to the loss of adjacent DA neurons (9). However, a recent study clearly

f demonstrated the global formation of MPP in the whole brain only minutes after MPTP

^ administration, and a subsequent rapid clearance from most regions (28). This indicates that

2 glial cells devoid of OCT3 expression must also contribute to MPTP conversion and MPP

2 ~ release. Moreover, conventions in membrane biophysics suggest that passive transporters like

¡« S3 OCT3 would contribute to an uptake of MPP+ from the cerebrospinal fluid into cells (49). A

2 & plasma membrane potential of -70 mV would drive a 15-fold increase in the accumulation of

'3'g a positively charged molecule inside cells according to the Nernst equation. Hence, the

presence of OCT transporters in glial cells would rather lead to a reduction of extracellular ,

» MPP and its toxicity in neurons as it was observed with monovalent paraquat (42).

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§h MPTP can form several metabolites other than MPP in the brain (52), and their biochemical

and physical properties need to be taken into account in order to understand the possible

¡Its mechanisms of MPP+ transport across the membrane. The first metabolic step in MPP+

.Sgc formation is always an MAO-dependent two-electron oxidation yielding the unstable

^•j?^ intermediate MPDP (5). A non-enzymatic autoxidation of MPDP has been suggested as the

most likely second step of MPP+ formation (30). In addition, a disproportionation reaction yielding one molecule of MPTP and MPP+ from two molecules of MPDP+ has been described

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(4,39). However, this reaction takes place only at concentrations of MPDP+ (millimolar range)

unlikely to occur in biological systems. The complex chemistry of MPDP+ involves further

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spontaneous reactions; as an a,P-unsaturated iminium ion, MPDP can exist in tautomeric

forms and/or in its conjugate base form, 1-methy-4-phenyl-1,2-dihydropyridine (1,2-MPDP)

(4,5). The latter non-charged metabolite is capable of crossing biological membranes.

In the present work, we report two main findings: 1) the MAO-catalyzed MPTP intermediate

metabolite MPDP+/1.2-MPDP undergoes transporter-independent export from the cell, and 2)

MPDP+/1.2-MPDP is responsible for non-enzymatic MPP+ formation in the extracellular space. These mechanisms explain why MPTP-converting glial cells are spared from MPP+ toxicity and how extracellular MPP+ is generated, which is a necessary prerequisite for the

accumulation of MPP in catecholaminergic cells.


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^ g Extracellular MPP generation by MPTP-converting astrocytes predominates.

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We studied the conversion of MPTP to MPP by astrocytes in the mouse astrocytic cell line

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2 g IMA 2.1, which displays MPTP metabolism properties comparable to primary astrocytes (Fig.

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1A,B (45)). We observed an accumulation of the biologically active toxicant MPP in the

IMA culture medium, but not within the cells (Fig. 1C, D). Addition of the OCT3 inhibitor

tij^ D22 to the culture medium had no influence on MPP+ release or uptake (data not shown),

S.B suggesting that the previously suggested OCT3 transport mechanism (9) was not obligatory

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f for extracellular MPTP/MPP accumulation. However, the MPTP converting enzyme

monoamine oxidase-B (MAO-B) is located exclusively within the astrocytic cell body, and

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MPP+, as a charged molecule, cannot penetrate biological membranes. This raises the

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question: how can MPP be efficiently generated extracellularly in the absence of a system for its transport from the cell (Fig. 1C)?

To resolve this issue, we generated IMA sub-clones that stably express OCT3 (a passive 3§ R transporter of MPP+), DAT (accumulates MPP+ against a concentration gradient), or both

^ 3 DAT and OCT3; these cells lines are referred to as OCT3-IMA, DAT-IMA, and OCT3/DAT-

IMA, respectively. Expression levels of OCT3 in the OCT3-IMA and OCT3/DAT-IMA lines ^ a were identical (Suppl. Fig. S1A), and the transport capacity and pharmacological features

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endogenously expresses OCT3 (22). We next assessed MPP transport in these cell lines. In

§ g DAT-IMAs, uptake of MPP+ was prevented by the DAT-blocker GBR12909, while in OCT3-

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were comparable to those of the human kidney carcinoma cell line Caki-1 which

3 g IMAs MPP+ uptake was prevented by the OCT3 blocker D22 (Suppl. Fig. S1A-D).

We next assessed the response of wild type and the transporter-expressing cells lines to MPP treatment. All cells were exposed to a fixed concentration (20 |iM) of extracellular MPP

jS (Fig. 1E). The toxicant did not enter wild type IMAs, which was as expected given the

2 absence of functional carriers in the plasma membrane in these cells and the inability of MPP

~ to cross lipid bilayers. Strong intracellular accumulation was however observed in DAT-

g jj IMAs. Ectopic OCT-3 expression in OCT3-IMAs and OCT3/DAT-IMAs enabled influx of

2 £ MPP+ into cells. In OCT-3/DAT-IMAs, MPP+ efflux from cells that had accumulated very

'S'g high concentrations of MPP+ due to DAT activity was observed (Fig. 1E). Steady state levels

were reached within 2 h. At this time point, the absolute amount of accumulated MPP+ was ^ ft measured, and the cellular volume of IMA was determined (2.5 pl/cell) so that average

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° 8 found in mice exposed to MPTP (19). Within 24 h, these concentrations were sufficient to

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tJ3ft diminish the cellular capacity to reduce resazurin or to produce ATP (Suppl. Fig. S2). By

contrast, IMAs neither accumulated MPP+ nor displayed impaired viability in response to up to a 120 |iM concentration of extracellular MPP+. Indeed, the degree of MPP+-induced reductions in viability across the four cell lines correlated with their respective capacity to accumulate MPP+ (Fig. 1G). These data support the notion that astrocytes are not resistant to

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MPP+ toxicity, as reduced viability was observed as soon MPP+ accumulated within the

transporter-expressing cells. Our findings also confirm that MPP cannot cross the

5-2 m .p

intracellular MPP+ concentrations could be calculated (Fig. 1F). The transporter-expressing cells reached levels of around 180-680 |iM, which is highly similar to tissue concentrations

membranes of IMAs unless they ectopically express MPP+ transporters. This led us to posit

course of MPTP metabolism.

'•g'2 that MPP crossed the IMA membrane through an OCT-3 independent mechanism during the

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go MPDP+ is a membrane-crossing intermediate generated from MPTP by MAO-B.

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B Since the pioneering work of Castagnoli and colleagues, it has been known that MAO-B

| g converts MPTP to intermediates such as MPDP+, and that further oxidation to MPP+ proceeds

'•gj non-enzymatically (4,30,39). Some of these initial publications speculated that these

^ & intermediates might play a role in transport processes, but further investigation of their

2 biological formation and properties over the last 30 years has been highly limited. We thus

~ focused here on the transport of MPDP+ as a potential missing link that might explain

g ö extracellular MPP+ formation. Analysis of the products formed in the conversion of MPTP by

^ purified MAO-B enzyme confirmed that MPDP is a major intermediate in the generation of

■¡3'i« MPP+ (Fig. 2A). Similar observations were made in analysis of wildtype IMA and OCT3-

qJö IMA cultures treated with MPTP, as MPDP+ was detected in the supernatant before MPP+.

^ a We further found that MPDP+ export was not dependent on OCT3 (Fig. 2B). To determine if MPDP+ formation in this in vitro model also occurs in vivo, mice were injected with MPTP

5-2 « .p

and the striatum and cerebellum was collected from injected mice at different time intervals.

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° 8 These experiments revealed a rapid (30 min) rise of MPDP in both brain areas that preceded

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^ the formation of MPP+. MPP+ remained elevated in the striatum, but not the cerebellum (Fig.

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ijf 2C). C-5 carbons of tetrahydropyridine structures such as MPDP+ are acidic (i.e. they possess

a a,ß-unsaturated iminium ion structure). However, their conjugate free base form 1,2-MPDP was also observed (4,30,39) (Fig. 2D) and this molecule is expected to be able to freely diffuse across biological membranes. To more carefully examine the formation of

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'S j3 intermediates that are generated in the MAO-dependent conversion of MPTP to MPP+, we

S a conducted time-resolved H-NMR analysis (Suppl. Fig. S3). These data confirmed a

significant time lag between the enzymatic formation of MPDP+ and its subsequent

'•g^ autoxidation to MPP . Based on these findings, we hypothesized that the acid-base pair

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g exported from astrocytes, and would thereby explain the presence of extracellular of MPP .

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1 Free bidirectional flux of MPDP+/1.2-MPDP across biological membranes.

f In order to test the hypothesis that MPDP+/1.2-MPDP possesses transporter-independent

h membrane permeability, IMAs were treated with 50 |iM fresh MPDP or decomposed MPDP

MPDP71.2-MPDP serves as membrane-permeable intermediate in MPP production that is

2 (stored in an open tube at room temperature prior to use). Both the resazurin reduction assay

2 ~ and measurements of intracellular ATP (Fig. 2E) indicated that only fresh MPDP+ led to cell

g £ damage. For an appropriate interpretation of the data, it is important to note that the

S3 ^ extracellular volume in these experimental setups is substantially greater than the intracellular

'3'g volume. Treatment of the cells with MPTP hence results in a constant and steep concentration

qJ2 gradient of freshly generated MPDP+ that supports its efflux into the extracellular space. In

+a ft contrast, extracellular addition of MPDP+ generates conditions in which the intracellular

gjy volume is rapidly saturated (i.e. intracellular volume <<< extracellular volume).

% To confirm a direct influence of intracellular MPP on mitochondria, metabolic flux

1*3 measurements were obtained as a more sensitive readout. MPP+ specifically inhibits complex

.5 g c I of the electron transport chain. As a result, NADH cannot be oxidized via complex I and

^•5,3 accumulates in the cell. An increase in the NADH/NAD ratio results in an inhibition of

NAD+-dependent reactions of the TCA cycle, such as pyruvate dehydrogenase activity. In the case of complex I inhibition, glucose carbon contribution to the TCA cycle strongly decreases, while pyruvate reduction to lactate increases. By increasing the rate of glycolysis, the cell compensates for the lack of ATP, similar to the Warburg effect. To assess metabolic

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flux, we labeled cells with the tracer [U- C6] glucose, which can be used to monitor relative

decarboxylating pyruvate in the mitochondrion during the production of acetyl-CoA, (Fig.

ft|| pyruvate oxidation that results from the NAD+-dependent pyruvate dehydrogenase reaction

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e % 2F). The activity of pyruvate dehydrogenase and of ATP dependent pyruvate carboxylase,

were both inhibited by MPDP+, but not by MPTP or MPP+ (Fig. 2F, Suppl. Fig. S4).

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To directly compare MPP+ and MPDP+ uptake, intracellular MPP+ levels in IMAs exposed to either MPP+ or MPDP+ were investigated. To exclude any potential involvement of cellular transporter systems, all experiments were conducted in parallel with artificial lipid vesicles

(Fig. 3A). As a third model system, purified human erythrocytes were used as they possess no

2 MPP transporter activity, but do allow the conversion of MPDP to MPP due to the

2 ~ presence of iron-containing hemoglobin. In all three model systems, extracellular

¡« £ concentrations of MPP+ as high as 200 |M did not result in significant accumulation of

2 ^ intracellular MPP+, while the addition of MPDP+ immediately led to the formation of

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intracellular/intravesicular MPP+. Uptake of MPDP+ and MPP+ into erythrocytes was then

qJ tested in vivo by intravenous injection of these compounds into mice and collection of blood

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in the intracellular accumulation of MPP+, providing further evidence of transporter-

4a ft after 3 min. Similar to the ex vivo treatment of erythrocytes, the addition of MPDP+ resulted

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■li^ independent membrane passage ofMPDP+ (Suppl. Fig. S5).

^ To further test the kinetics of cell loading and unloading of MPDP+, IMAs were exposed to

¿fll MPDP+ (50 |M) for

a 30 min loading phase. Analysis of the cellular compartment showed a

^•j?^ rapid increase

of MPDP+, followed by a rise in MPP concentrations. MPP was apparently generated within these cells, as extracellular MPP+ cannot cross the cell membrane (Fig. 3B, Left). After the loading phase, cells were washed, new medium was added, and metabolite levels were detected over time in both the intracellular (Fig. 3B, Middle) and extracellular compartments (Fig. 3B, Right). Accumulated MPDP+ vanished rapidly from the cells into the

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gp extracellular media, whereas intracellular MPP was not able to exit the cells (Fig. 3B, Middle

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ft¡^ and Right). Considering these data together, we concluded that extracellular MPP+ originated

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e % MPDP concentration gradient is the predominant driving force underlying its transport across

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membranes, which explains the apparent export of MPP+. These observations indicate that

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■3 8 MPDP+/1.2-MPDP

is membrane permeable and suggest that MPP+ will remain in a given

from the autoxidation of extracellular MPDP+. In addition, these findings indicate that the

i, biological compartment, unless transporters are present.

§ The role of extracellular MPDP+ autoxidation in dopamine neuron toxicity.

2 ^ Our data here-to-fore have suggested that upon MPTP intoxication,

MPDP+/1.2-MPDP would

<« k have the potential to freely diffuse across the membranes of several cells, and eventually

® ^ convert to MPP+ in any cell type adjacent to MAO-containing glial cells. However, MPP+

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accumulation is largely limited to catecholaminergic neurons that take up the molecule via

their catecholamine transporter (e.g. DAT). To compare the respective contribution of DAT-

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via DAT (46,47,48) were treated with MPP+ or MPDP+ (20 |M) in the presence or absence of

2 8 the DAT blocker GBR12909 (GBR). This compound blocks the uptake of MPP+ via DAT

& mediated MPP accumulation and passive MPDP /1.2-MPDP-dependent influx (Fig. 4A),

human dopaminergic neuronal cells (LUHMES), which can accumulate extracellular MPP+

Js & inhibition, but not MPDP+/1.2-MPDP as it is not a DAT substrate. Metabolic flux was

measured with labeled [U-13C6] glucose, which provides a readout of the inhibition of oxidative phosphorylation. Both MPP+ and fresh MPDP+ led to mitochondrial inhibition, and GBR blocked toxicity in both cases (Fig. 4B). These observations were confirmed by cellular staining with an anti-P-III-tubulin antibody, which demonstrated that pronounced neurite

U degeneration was triggered by both MPP+ and fresh MPDP+, and these effects were

•ga completely prevented by GBR (Fig. 4C). Taken together these data suggest that at the

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j^g moderate MPDP+/1.2-MPDP concentrations (low micromolar range) that exist in vivo,

••g<8 intracellular autoxidation of MPDP would not be sufficient to generate damaging

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intracellular MPP+ concentrations.

To explore whether toxic MPP+ can be formed within neurons from freely diffusing and

| 2 autoxidizing MPDP+, LUHMES cells were exposed to high (200 |M) or low (20 |M) MPDP+

| g concentrations in the presence or absence of the DAT blocker GBR. High extracellular

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MPDP+ levels led to a DAT-independent accumulation of intracellular MPP+ doses sufficient

^ & to inhibit mitochondria, whereas low concentrations did not (Fig. 4D). Thus, toxicity evoked

2 by free diffusion of freshly generated MPDP in cells adjacent to astrocytes may play only a

2 ~ marginal role. To test this assumption, a co-culture model containing IMA (astrocytes) and

¡« S3 LUHMES (neurons) was exposed to MPTP. This led to neurotoxicity via glial-dependent

S3 ^ generation of MPP . The addition of GBR completely inhibited neurotoxicity in this co-

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culture system. These data confirm that the passive diffusion of freshly generated MPDP /1.2-

qJ MPDP itself from glia immediately adjacent to LUHMES was not able to evoke significant

£ ft toxicity (Suppl. Fig. S6).

§ h Identification of physiological factors that promote the autoxidation of MPDP+ in the

~ •.§ extracellular space

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The findings described thus far indicate that MPP needs to be formed extracellularly in order

Mg^ to become a substrate for DAT, and thus induce neurotoxicity. We next explored how a

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i^B'-S preferential extracellular generation of MPP might proceed in structures such as the brain

with a high ratio between the intra- and the extracellular volume. As oxygen levels are likely to play a role in MPDP+ autoxidation, we studied this environmental factor. High-resolution

measurements of oxygen concentrations in cells have provided evidence for steep oxygen

'g,^ concentration gradients between the cytosol and mitochondria (27,36). This may result in

3 g average intracellular oxygen concentrations that are significantly lower than the interstitial

go levels in a tissue or cell culture media (17,51). We found that the speed of MPP formation

^ strongly depended on oxygen levels, and that MPDP+ is highly stable under low oxygen

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astrocytes (Fig. 5A, B). Lower intracellular oxygen levels would hence stabilize intracellular

g MPDP while at the higher extracellular oxygen tension, the non-enzymatic conversion into

conditions such as those that are typically found near the respiring mitochondria (27,36,51) of

MPP+ would be favored.

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We posited that another important factor in determining the rate of MPDP autoxidation

pS § might be the local pH. The cytosolic pH of most cells in the body is typically lower than in

g extracellular fluids and the blood. Local acidification is largely driven by respiratory chain

~ proton pump activity, particularly around mitochondria. We observed an accelerated

g is autoxidation of MPDP+/1.2-MPDP under more alkaline conditions (Fig. 5C, D), while more

2 ^ acidic conditions (e.g. close to mitochondria) stabilized MPDP . The combination of a more

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acidic pH and lower oxygen tensions (e.g. the conditions typically encountered inside cells)

qJ strongly prevented MPDP+ autoxidation (Fig. 5 E, F).

^ Based on these findings, we hypothesized that, in addition to DAT activity, the conditions in

ja h brain regions enriched in dopaminergic neurons might promote greater autoxidation than in

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other areas. The nigrostriatal system of the basal ganglia has a particularly high iron content, 1*3 and also contains various autoxidation products of dopamine (12,35). All of these compounds

« g c promote the formation of radicals, which is required for autoxidation of MPDP . While free

radicals have very short half-lives inside cells due to several efficient free-radical buffering mechanisms (e.g. glutathione system, iron buffering), they can persist for longer times extracellularly. Taking this into consideration, several candidates were tested for their influence on MPDP+ autoxidation. A striking acceleration of MPP+ genesis from MPDP+ was observed in response to the treatment with autoxidized dopamine, melanin, iron containing

'§3 p

ferritin, or with free ferrous iron (Fig. 5G, H). Addition of the iron-chelator deferoxamine

a ¡^ prevented the iron-mediated acceleration of MPDP+ autoxidation (Suppl. Fig. S7). This is in

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e ^ (11,29), although these reports did not demonstrate whether the conversion of MPTP was

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affected by treatment with iron chelators or other oxidative stress related events. Enhanced

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accordance with reports indicating a protective role of iron chelators in MPTP PD models

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^ + 3 2 autoxidation in the extracellular milieu would not only explain preferential MPP generation

g outside cells, but also ensures there is a constant, steep concentration gradient between

cytosolic MPDP+ and extracellular MPDP+, which is the main driving force for its transporter-

^ Î independent efflux (Fig. 6).


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g Among the known human neurotoxicants, MPTP is exceptional in that it triggers relatively

selective death of nigrostriatal DA neurons upon systemic administration (3,25,31). A

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necessary prerequisite for the selective DAT-dependent uptake of the active toxin MPP+ is the

presence of MPP+ in the extracellular space, as was elegantly confirmed by several

gia microdialysis studies (9,21,33). The mechanism by which MPP , a membrane-impermeable

molecule generated intracellularly by the enzymatic conversion of MPTP, reaches the

extracellular space has been elusive. The present work aimed at addressing this question, and

P was inspired by the observation of efficient extracellular MPP+ generation in MPTP-

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converting astrocytes in absence of significant intracellular MPP+ accumulation and toxicity.

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— _______ .... :___________ ..

l3 § We first determined that astrocytes (IMAs) were not immune to MPP+, as these cells died

when cytosolic accumulation of extracellular MPP+ was enabled by the ectopic expression of the MPP+ transporters DAT and/or OCT3. Although it can be concluded from these results that astrocytes are not per se resistant to MPP+, they do have the capacity to accelerate

3S " glycolytic turnover and hence are able to at least partially compensate for inhibited

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p mitochondrial ATP synthesis caused by MPP . This conclusion is corroborated by several

a'S experiments in the literature showing that any cell tested thus far, e.g. HEK293, hepatocytes,

^ ft COS, or HeLa cell, is killed by MPP+ if the toxicant is allowed to enter the cytosol.

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g MPTP with endogenous MAO, such as astrocytes, platelets, and hepatocytes, are not

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J3 cells, MPP accumulation would therefore be transient and minimal, as


Moreover, the findings of the present study provide a rationale for why cells that metabolize

necessarily killed by the final reaction product MPP+. We found that the intermediate MPDP+ (or its neutral base 1.2-MPDP) exits cells along a concentration gradient via membrane diffusion. MPP+ is mainly formed extracellularly, and in most cases it would be diluted and carried away by body fluids or taken up by passive transporters in neighboring cells. In most

2 would exit the cell as soon as the extracellular concentration drops. Only cells that accumulate

2. ~ MPP+ through an active intracellular transport mechanism, e.g. DA neurons in the central

g £ nervous system, would therefore be able to retain the toxicant for a sufficient period of time

2 ^ and at high enough concentrations to cause damage (Fig. 6). While accumulation of MPP in

'3'g cells is a necessary condition for the toxicity of this molecule, the degree of susceptibility of

particular cell types to intracellular MPP+ may vary due to the presence or absence of ^ a resistance mechanisms, e.g. by sequestration as in adrenomedullary cells (43) or other more

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g, 8 suggested to directly contribute to the conversion of MPTP in neuronal cells (1). By contrast,

Mg^ in LUHMES cells, MPTP treatment resulted in neither the formation of MPP+ nor cellular

.5 c c

-S—S-ji degeneration (not shown).

The concentration of MPP+ required to cause toxicity in different cell types may therefore vary. Studies of isolated mitochondria and submitochondrial particles have shown that MPP+ concentrations of about 5-20 millimolar need to be reached in the mitochondrial matrix for inhibition of complex I (40,41), which would in turn require cytosolic levels in the high

(U ¡5

c3 a ^

poorly understood cellular factors (7,46). Alternatively, certain cells may possess factors that promote MPP+ formation and toxicity; for instance, cytochrome P-450 2D6 was recently

'§3 li

micromolar range (> 100 |iM). This is corroborated by our measurements showing a ä mitochondrial effects and toxicity within a period of 24-48 h in IMAs expressing OCT3 or

<u £ a

■M (U <+H O

e ^ measurements of wildtype IMAs showed that concentrations were approximately 10 |iM (~ 1

'•§ £

§1 nmol/mg protein), which is in close accordance with the data of Di Monte et al. obtained from

DAT if intracellular MPP+ concentrations were larger than 100 |iM (Fig. 1F). By contrast, our

3 2 primary mouse astrocyte cultures (1 nmol/mg, after 48 h exposure to 250 |iM MPTP)

g (15,16,55).

Here we have demonstrated that MPDP+ is the pivotal MPTP metabolite responsible for the

jg extracellular genesis of MPP , which minimizes its toxicity to most cell types. MPDP was

2 described and synthesized by Neil Castagnoli and colleagues in the 1980s (4,5). Their

2 ~ pioneering discovery of this metabolite and the seminal work of Di Monte et al. (13-16,55)

¡« £ were quickly incorporated into all major toxicology textbooks, but experimental study of this

S3 ^ compound has been virtually non-existent for the past 25 years, despite its wide use in PD

'S'g research during this time. Here, we have advanced this early work by providing direct

evidence by NMR and MS of MPDP+ formation during MPTP to MPP+ conversion by MAO-

42 ft B. It was originally postulated that the disproportionation of two molecules of MPDP+ into

MPP+ and MPTP would be the dominating mechanism of MPP+ formation, however, our

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Рч Я

previous studies on the autoxidation of resynthesized MPDP+ suggest that direct oxidation of

S u ui

t+ч О

о S3 MPDP+ into MPP+ dominates (4,5). Although the mechanism of MPDP+ autoxidation has not

tJ^ft yet fully been elucidated, kinetic and thermodynamic principles observed in structurally

<u сз 5 Л вд

.Пз-S related compounds allow us to postulate that the reaction sequence is as follows: in the initial

step, a stabilized allyl radical is formed by the abstraction of a proton at C5 as a result of the interaction of MPDP+ with molecular oxygen (O2), transition metals, or free radical species (e.g. dopamine autoxidation products) (Suppl. Fig. S8); in a second reaction, the allyl radical

iU fj 2 ^

'Я j3 interacts with a second free radical species or O2; when there is an interaction with O2, C-5

s3 si would carry a peroxide group, which would render the carbon 6 acidic and hence trigger the

Й rt •¡2 cj

■§3 release of a proton from C-6 (peroxometabolite); energetically, the elimination of the C-5

(D ^ ^ О

'•g« hydroperoxide substituent from the base form would strongly favor aromatization to the stable


<u 42 — a

■M (U t+ч О

Й ^ о^З

^ favored under alkaline conditions. This sequence of events is strongly supported by our

% I? experimental observations indicating accelerated MPDP+ autoxidation in the presence of high

§ & oxygen levels and transition metals, and a relative stabilization under slightly acidic

toxin MPP+ (Suppl. Fig. S8); the final step of proton abstraction from the peroxometabolite is

та^ conditions (Fig. 5). Importantly, autoxidation can efficiently proceed under conditions of

f relatively low MPDP+ concentrations in contrast to disproportionation reactions.

2 We have identified several factors that increase MPP formation in the extracellular space,

2 ~ most notably higher oxygen tension and pH. Indeed, the low oxygen levels within cells

¡« S3 compared to the extracellular space may be reduced even further in astrocytes by MAO-B

2 & metabolism of MPTP, which is an oxygen-consuming reaction (6). Thus, low oxygen tensions

'3'g and lowered pH around respiring astrocytes would prevent MPP+ formation within cells,

while this process would proceed faster in the extracellular space. Moreover, our data show ^ ft that MPTP conversion proceeds more than 10-times faster in the presence of biological

oxidation enhancers, like melanin, oxidized dopamine, or ferrous iron. Some of these factors have particularly high levels not only in the nigrostriatal system in which DA

o w tt £

° 8 neurodegeneration prevails, but also in hemoglobin-carrying erythrocytes (Fig. 3A and Suppl.

j3ft Fig. S5) (50,53,56). These factors are neutralized and scavenged to a large extent inside cells

5 Gh ££

.—2.5 due to powerful redox buffers, lysosomal removal, or iron binding by ferritin, while they may

exert their full activity in the extracellular space. Apart from oxygen tension or pH, other factors might indeed influence the autoxidation of MPDP+. Our observation of accelerated MPDP+ autoxidation, induced by ferrous iron and its inhibition by the iron chelator

U deferoxamine (Suppl. Fig. S7), provides insight into the molecular mechanisms that might

"3 g underlie previous observations of protection against MPTP-toxicity in vivo by iron chelator

"fis administration to model animals (11, 29). However, we tested ascorbic acid both in a cell-free

'•gmodel of MPDP autoxidation, as well as in IMAs that were exposed to MPTP. In both cases,

o £ a

■M (U

t+H O 0 &

|s These findings indicate that a more detailed analysis of the molecular mechanisms involved is

<D £ W)<D

the autoxidation of MPDP+ and the formation of MPP+ was not affected (Suppl. Fig. S7).


cs a ^

^ 1 brains. This suggested that preferential MPP release from astrocytes via OCT3 would be a

g A recent study reported that there is selective expression of OCT3 in glial cells located in

close spatial vicinity to nigrostriatal neurons that were primarily affected in MPTP-challenged

g major contributing factor in the selective loss of DA neurons in the substantia nigra (9). p a

2 ~ However, we observed that MPDP+ generation in the brains of MPTP-treated mice preceded

¡« ö the formation of MPP+ not only in the substantia nigra, but also in the cerebellum which is a

S3 ^ prerequisite for the transporter-independent efflux of MPDP into the extracellular space (Fig.

'3'g 2). A recent publication was in full accordance with our findings in demonstrating almost

qJö uniform MPP+ generation in MPTP exposed brains only minutes after MPTP administration.

4| ft This was followed by rapid clearance from the brain, except in the basal ganglia, ventral

mesencephalon, and olfactory bulb (28). Given the findings that MPP is generated globally

ö Ö c3 ft

in the extracellular space and taken up by specific cell types due to selective transporter

■: 5 o

tt £ +

° 8 expression, the tissue distribution of MPP may be determined by complex dynamics that owe

js^ to the differential expression of MPP+ transporters across cell types and brain regions. For

.wjg.^ instance, MPP+ would be partially sequestered into cells expressing OCT transporters (Fig. 6),

as indicated by our experiments with OCT-3-expressing IMAs (Fig. 1). Indeed, such a mechanism has been reported in vivo for the MPP+-related compound paraquat, which resulted in the protection of DA neurons in brains of paraquat-treated mice by lowering

'S J3 extracellular concentrations of the toxin (42).

The basic principles identified in the present work are not limited to the experimental toxin

ft ^ MPTP, but could also apply to endogenously formed small molecules such as quinolinic acid

<u £ ft ■M (U t+H O

e ^ are suspected to play key roles in neurological and psychiatric disorders.

'•§ £

<D £ W)<D

(34), isoquinolinates, beta-carbolines (8), or dopamine autoxidation products (2), all of which


13 p ft

The well-studied Parkinsonian toxicant MPTP requires astrocytic metabolism to generate

^ MPP+. Subsequently, MPP+ is transported into dopaminergic neurons by dopamine

J <D C3

< e »

m p Ö C3


<u £ ¿s a

■M (U

t+H O 0 &

O y ce a

§ transporters and it thereby accumulates and eventually kills these cells. Why MPP+ spares the

astrocytes in which it is supposedly generated and how MPP+ leaves these cells has been a vexing enigma for decades. We found here that MPP+ is generated extracellularly, and that the release of its membrane permeable precursor MPDP+/1,2-MPDP from astrocytes is

' Z rf transporter independent. Differences in O2 and pH between the intra- and extra-cellular

O <+h O

£ is environment favor the stabilization of the labile MPTP intermediate MPDP+/1,2-MPDP

within astrocytes and promote its nonenzymatic conversion in the extracellular space.

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Materials and Methods


p a (Nj3 wjS Sj3

<N <+H

00 ^ +

2g MPP (1-methyl-4-phenylpyridinium-iodide), purified monoamine oxidase-B (MAO-B),

• • o

GBR12909 (DAT blocker), dopamine, ferritin, and tyrosine-melanin were purchased from

q^ Sigma (St. Louis, MO, USA).

S d Synthesis of MPTP-HCI and MPDP+ bromide.

All reagents are commercially available and were used without further purification. Solvents

were dried over molecular sieves and used directly without further purification. All reactions

were conducted under exclusion of air and moisture. H- and C-NMR spectra were recorded on an Avance III 400 MHz spectrometer from Bruker at room temperature. The time course

£ — of enzymatic MPTP conversion was followed on a Bruker AV III 600 MHz spectrometer

^ i §

•2& £ equipped with a TCI-cytoprobe. Spectra were processed with the software MestReNova 6.1.1

9 2 £ ^ Ö «1

from MestRelab Research and the 1H and 13C chemical shifts are reported relative to the

go residual solvent peak. High resolution mass spectrometry (HRMS) was performed with

gÜ micrOTOF-Q II ESI-Qq-TOF from Bruker Daltonics.

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0 E^ Ö ^

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cd ft tt 0

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Ph cd w

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1= äs

5 <3 (U <3

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Synthesis of MPTP-HCI (4) and MPDP+ bromide (6)

1 2 3 4 5 6

Scheme 1. a) THF, -78°C^ rt, over night; (18) b) aq HCl, reflux, 5h, quantitative; (18) c) (i) NaEtO, EtOH, 0°C (ii) aq H2O2, EtOH, 50°C, quantitative; (20) d) DCM, TFAA, aq HBr,

0°C, 53%. (20)

Step a): 1-Methyl-4-phenyl-4-piperidinol 3 (18)

A solution of 24.6 ml phenyllithium 2 (1.8 M in dibuthylether) was added to a solution of 1-metyl-4-piperidone 1 (5 g, 44.19 mmol) in 20 ml of dry tetrahydrofuran (THF) at -78°C. The reaction was allowed to warm up to room temperature and was stirred over night. The solvents were removed in vacuo to give crude 3 as yellowish solid.

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■M (U t+H O

cs a « 0

Step b): MPTP-HCI 4 (18)

Refluxing of crude 3 in 50 ml 10% aq HCl for 5 h furnished compound 4. The solvent was removed in vacuo and crude 4 was re-crystallized from ethanol to give white crystalline 4 in a quantitative yield.

1H-NMR (400 MHz, CD3OD) Ö 7.53 - 7.47 (m, 2H), 7.44 - 7.31 (m, 3H), 6.15 (ddd, J = 5.1, 3.4, 1.7 Hz, 1H), 4.08 (d, J = 15.3 Hz, 1H), 3.92 - 3.79 (m, 1H), 3.79 - 3.70 (m, 1H), 3.45 -3.35 (m, 1H), 3.03 (s, 3H), 3.00 - 2.92 (m, 1H), 2.87 (d, J = 21.4 Hz, 1H). 1H-NMR (600 MHz, D2O) Ö 7.55 - 7.52 (m, 2H), 7.46 (dd, J = 10.2, 4.8 Hz, 2H), 7.43 - 7.39 (m, 1H), 6.15 - 6.13 (m, 1H), 4.05 (d, J = 16.4 Hz, 1H), 3.82 - 3.78 (m, 1H), 3.74 - 3.69 (m, 1H), 3.43 - 3.34 (m, 1H), 3.00 (s, 3H), 2.96 - 2.90 (m, 1H), 2.86 (d, J = 19.0 Hz, 1H).

§ 13C-NMR (101 MHz, CD3OD) S 139.9, 136.8, 129.7, 129.4, 126.2, 116.6, 53.5, 52.0, 42.9,

<N ^ n

13C-NMR (150 MHz, D2O) S 138.3, 134.6, 128.7, 128.4, 125.0, 115.5, 52.0, 50.6, 41.7, 23.8.

2| HRMS (ESI, positive ion mode): [M+H]+; calculated: C12H15N: 174.1277 m/z; measured:

^ 174.1288 m/z

Step c): 1 -Methyl -4-phenyl - 1,2,3,6-tetrahydropyri dine #-Oxide 5 (20)

(i) The solution of 4 (2.31 g, 11.01 mmol) in 60 ml ethanol was cooled to 0°C and 9.5 ml of a

£ h sodium ethoxide solution (1.17 M in ethanol, 11.12 mmol) was added. The mixture was

£ '-g filtered over celite and the filter cake was washed with 30 mL of ethanol.

o tt £

s<g (ii) Next, 2.6 ml of 30%

aq H2O2 was added and the mixture was stirred at 50°C. After 2 h, an

^ additional portion of 2 mL 30% aq H2O2 was added, and it was stirred over night. Afterwards


gft m the excess of H2O2 was quenched with 10% Pd/C. The resulting mixture was filtered over

celite and the filter cake was washed with 30 ml of ethanol. The solvent was removed in vacuo to give 5 2 H2O as yellow oil in a quantitative yield.

1H-NMR (400 MHz, CDO3) S 7.42 - 7.26 (m, 5H), 5.96 (s, 1H), 4.32 - 4.10 (m, 2H), 3.76 -

3.65 (m, 2H), 3.37 (s, 3H), 3.06 - 2.95 (m, 1H), 2.76 - 2.62 (m, 1H).

J-5 Step d): MPDP+ bromide 6 (20)

Sg The suspension of 5 2 H2O (2.51 g, 11.01 mmol) in 90 ml dichloromethane (DCM) was

■M (U t+H O

(2.06 g, 12.23 mmol). The mixture was evaporated to dryness under reduced pressure at room § is temperature. The resulting crude product was re-crystallized from acetone to yield 53 % 6

cooled to 0°C and trifluoroacetic anhydride (TFAA) (4.49 g; 21.38 mmol) was added dropwise within 20 min. It was stirred at 0°C for 2 h, followed by the addition of 48% aq HBr

(1.48 g, 5.87 mmol) as a yellow hygroscopic solid.

o u c3 ft

— 1H-NMR (400 MHz, CDCl3) S 9.23 (d, J = 4.1 Hz, 1H), 7.69 - 7.60 (m, 2H), 7.52 - 7.40 (m,

J| 3H), 6.89 (d, J = 4.6 Hz, 1H), 4.14 (t, J = 9.4 Hz, 2H), 3.93 (s, 3H), 3.32 (t, J = 9.4 Hz, 2H).

§ 1H-NMR (600 MHz, D2O) S 8.38 (d, J = 4.8 Hz, 1H), 7.82 (d, J = 7.9 Hz, 2H), 7.63 - 7.54

Sf (m, 3H), 6.90 (d, J = 5.0 Hz, 1H), 4.04 (t, J = 10.1 Hz, 2H), 3.69 (s, 3H), 3.28 (t, J = 10.1 Hz,

n 00 ^

©• S3 13C-NMR (101 MHz, CDCl3) S 164.1, 158.0, 134.5, 133.0, 129.5, 127.3, 113.9, 49.0, 47.7,

£ £ + -


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ft"B Q-5

13C-NMR (150 MHz, D2O) S 163.5, 158.8, 134.6, 132.5, 129.0, 126.7, 112.9, 47.6, 46.0, 24.5.

HRMS (ESI, positive ion mode): [M]; calculated C12H14N+: 172.1121 m/z; measured:

C a s3H

' £ a 172.1124 m/z

o w IS >3

Analysis of 1-Methyl-4-phenylpyridinium Iodide (MPP+ T) (Sigma Aldrich)

1H-NMR (400 MHz, DMSO) S 9.02 (d, J = 6.9 Hz, 2H), 8.51 (d, J = 7.0 Hz, 2H), 8.11 - 8.04 (m, 2H), 7.70 - 7.61 (m, 3H), 4.34 (s, 3H).

1H-NMR (600 MHz, D2O) S 8.74 (d, J = 6.7 Hz, 2H), 8.28 (d, J = 6.7 Hz, 2H), 7.97 - 7.92 (m, 2H), 7.70 - 7.62 (m, 3H), 4.36 (s, 3H).

• Ö in

C-NMR (101 MHz, DMSO) S 154.3, 145.6, 133.5, 132.0, 129.7, 128.0, 124.1, 47.1.

J| 13C-NMR (150 MHz, D2O) S 156.3, 144.8, 133.8, 131.9, 129.5, 127.7, 124.6, 46.9.

•§<8 HRMS (ESI, positive ion mode): [M]; calculated C12H12N+ 170.0964 m/z; measured:

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<D £ W)<D

170.0968 m/z

LUHMES cell culture.

Methods for the generation, handling, and properties of LUHMES cells have been described

in detail previously (47,48). The cells were grown in standard conditions (serum-free) at a

g density of 300.000 cells per well in 24-well plates. The cells were derived from human fetal

2 mesencephalon and were conditionally immortalized by expression of v-myc under a tet-off

~ promoter. Upon addition of tetracycline, LUHMES cells differentiate within 5 days to fully

¡« S3 post-mitotic human dopaminergic neurons with neurites of up to 1000 |im length (48). At this

stage, the cells express highly active dopamine transporters and are susceptible to low

© s ^ S3

'S'g micromolar levels of MPP+ (47,48). For uptake experiments, cells were differentiated for 6

Qja days and plated at a density of 300.000 cells/well in 24-well plates.

is Ö trt C o

£ d IMA 2.1 astrocytes (IMA).

o p tt £

Generation and properties of the IMA cell line have recently been described in detail (45).

ggg ^ These immortalized mouse astrocytes were cultured at 37 °C (5 % CO2) in DMEM (normal

^fl glucose) (GIBCO) with 5 % FCS (fetal calf serum, from PAA) and 1%

Penicillin/Streptomycin (10.000 U/ml stock). The cells were passaged by trypsinization for 2 min with 0.5% trypsin/DMEM every 2-3 days and reseeded at a ratio of 1:5 or 1:10.

Experiments were performed 2 days after cells reached confluency in DMEM containing 2 %

Overexpression of OCT3 and DAT.

¡¿-rt sfl

<u 4a a

o u Gene contracts for human solute carrier family 6, member 3 (dopamine transporter, DAT) and

2-T3 sls22a - solute carrier family 22, member 3 (organic cation transporter, OCT3) were

3 § purchased from SourceBioscience (Berlin, Germany). Both were obtained as inserts in the

=3 ft pCR vector (InVitroGene, Life Technologies Corporation USA) as clone number

| g IRCBp50050019Q (40147025 - IMAGE ID) and clone number IRCKp5014K038Q (8860680

¡3-p - IMAGE ID), respectively. The DAT gene was cloned into the phsCtR2AU-W expression

h construct. This HIV-based vector was used as a plasmid to transfect the IMA cell line using

2 Lipofectamine (InVitroGene, Life Technologies, Darmstadt, Germany). In this construct the

~ DAT gene was fused to the 3'-end of the tR2AU cassette (turbo red fluorescent protein from

¡« & Entacmaea quadricolor, the 2A sequence of Porcine teschovirus and human ubiquitin

2 ^ sequences). Translation of this gene produces free tRFP protein by ribosomal skipping at the

'S'g 2A site and free DAT protein by the action of cellular hydrolases which cut at the 3'-end of

the ubiquitin peptide. Cells that were successfully transfected and expressed the DAT gene ^ ft could therefore by be identified by their expression of red fluorescence. Cells were assayed 3

o p 1=

days after transfection.

The gene for OCT3 was cloned into the pBIGFP vector. This construct contains a

¡lis bidirectional inducible promoter allowing the expression of any gene cloned therein and the

«ce simultaneous expression of the linked marker gene eGFP (enhanced Green Fluorescent

Protein). Induction levels can be monitored by examination of eGFP intensities. Cells were transfected using Lipofectamine (InVitroGene, Life Technologies Corporation, Darmstadt, Germany) and subsequently selected with 200 |ig/ml G418 for 10 days.

S g Immunostaining.

Cells were grown in 24-well plastic cell culture plates (Nunclon™). Following treatment,

<u 4a ^ ft rt (U t+H O O tJ

1% BSA (Calbiochem, San Diego, CA) for 1 h, the primary antibody (anti-P-III-tubulin,

cells were fixed with 4% PFA for 20 min at 37°C and washed with PBS. After blocking with

g ^ Convance, mouse 1:1000) was added in PBS-Tween (0.1%) and the cells were incubated at 4

3 V °C overnight. Secondary antibody (anti rat IgG-Alexa 488, Invitrogen, Darmstadt, Germany)

O y c3 ft

was added for 45 min at RT. For visualization, an Olympus IX 81 microscope (Hamburg,

•S w

— Germany) equipped with an F-view CCD camera was used. Images were processed by Cell P

ft ^ software (Olympus).

§ Cell viability.

£ § Resazurin (Sigma) was added to the cell culture medium at a final concentration of 5 ug/ml,

Irs and was incubated at 37 °C for at least 30 min. The fluorescence intensity was measured at

530 nmex and 590 nmem with a Tecan Infinite M200 reader. Cell viability (in %) was calculated by normalizing the fluorescence values to the values obtained from untreated controls.

o № © S

• • O

s •y Q-5

ö tS C a

* -ü ATP assay.

o w 1=

§g Cells grown in 24-well plates were lysed in PBS-buffer containing 0.5% phosphatase inhibitor

s g § cocktail 2 (Sigma) and boiled at 95 °C for 10 min. Following centrifugation at 10.000 g for 5

^ • min to remove cell debris, protein content in the supernatant was measured and samples were

adjusted to have equal concentrations. Samples were then diluted 1:10 in PBS / 0.5% phosphatase-inhibitor buffer. For the detection of ATP levels, a commercially available ATP 3 § R assay reaction mixture (Sigma) containing luciferin and luciferase was used (50 |il of adjusted

~"3 sample plus 100 |il of assay-mix were added to a black 96-well plate). Standards were

prepared by serial dilutions of ATP disodium salt hydrate (Sigma) to obtain concentrations £ & ranging from 1000 nM to 7.8 nM.


o s ft ■M (U t+H O

a 8 Vesicle uptake assay.

For vesicle preparation, stock solutions of 100 mg/ml DOPC (1,2-dioleoyl-sn-glycero-3-

cd ft tt «

€S .a

phosphocholine) and 100 mg/ml DOPE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine) (Avanti) in CHCl3 were prepared. For the production of a 2 ml vesicle suspension, 102 |il of DOPC and 98 |il of DOPE stock solutions were mixed in a glass tube and evaporated for 30 min under vacuum while rotating to remove CHCl3. The lipid film was then resuspended in 1

2 ml of 20% Triton X-100 in PBS, and air was removed by N2-degassing. The mixture under N2

~ atmosphere was then briefly sonicated in a water bath and allowed to rest at room temperature

g £ until a clear solution was observed, indicating complete solubilization of the lipids. To

2 & remove the detergent, 600 mg Bio-Beads SM-2 (BioRad) preequilibrated in PBS were added

'3'g to the clear lipid/detergent mixture diluted 1:1 in PBS, and incubated at 4 °C on a rotator for

16 h. For removal of the beads, the vesicle suspension was collected with capillary tips and ^ ft transferred to a separate tube. For uptake assays, the vesicle suspension (10 mg lipids/ml) in

g <H C a

vesicles were separated by loading 100 |ul of the sample onto a NAPTM-5 Sephadex™ G-25

<+H CJ

O p tt &

° 8 column and a stepwise addition of 250 |ul volumes of PBS for elution. As control, vesicle-free

ipH ~ 2

solutions containing MPP+ or MPDP were added. The vesicle-containing fractions were

.5 a s

analyzed separately, MPP+ or MPDP+ values in the vesicle fractions were integrated.

PBS was incubated with MPP+ or MPDP+ for 2 h at 37 °C. To remove free MPP+/MPDP+,

Erythrocyte uptake assay

Human blood was collected in EDTA (1.6 mg/ml blood) containing tubes (Sarstedt S-

•rt J3

ox5 Monovette, Nuembrecht, Germany) and was centrifuged at 200 g for 4 min. The erythrocyte

•a d

o.o fraction was resuspended in a buffer containing 21 mM Tris, 4.7 mM KCl, 2 mM CaCl2, 140

mM NaCl, 1.2 mM MgSO4, 5.5 mM glucose, pH 7.4. The cells were washed twice with the buffer by centrifugation at 1000 g for 1 min. The cells were then treated with MPP or

■M (U

^ 9 MPDP+ as indicated. To assess intracellular MPP+ levels, the erythrocytes were again washed

•ja g

S by three consecutive centrifugation steps (1000 g, 1 min) to remove extracellular MPP+ or

% ju MPDP+. The erythrocytes were then lysed by resuspension in H2O for 20 min, supported by

2 ^ moderate sonication. Proteins were precipitated by the addition of 0.4 M perchloric acid, the

--jgJI supernatant was filtered through a 0.22 |im membrane and analyzed by HPLC.

Be ft c3

§ MPTP conversion in the brain

C57BL/6JRj male mice were kept under 12h light/dark cycles at a temperature of 23°C and

i-E 55% humidity. The animals had ad libitum access to food and water. All animal studies were

approved by the appropriate institutional governmental agency (Regierungspraesidium Tuebingen, Germany) and carried out in an Association for Assessment and Accreditation of Laboratory Animal Care International-accredited facility in accordance with the European tS^ Convention for Animal Care and Use of Laboratory Animals. The mice were intraperitoneally

o № © «

• • o

injected with MPTP (30 mg/kg; free base in 0.9% NaCl), cared for over the time intervals

% & indicated, and then killed by cervical dislocation. The striatum and cerebellum of each mouse

o ° ft °

^ was dissected after decapitation. The tissues were homogenized in 1 ml of 0.4 M perchloric

.5 g c acid by sonication. Following a centrifugation step at 2.000 g for 10 min at 4°C, the pellet was

resuspended in 250 |iM of1M NaOH. Total protein content was detected by a BCA assay kit. The supernatant was filtered through a 0.22 |im membrane and analyzed by HPLC.

Ö Ö c3 ft

HPLC analysis.


ö^o Detection of MPTP, MPP+, and MPDP+ was performed on a Kontron system (Goebel o.o

.5 8 Analytic, Au/Hallertau, Germany) comprising a model 520 pump, model 560 autosampler,

cCo <u £ ft rt (U t+H O

o sample and centrifuged at 10,000 x g for 15 min. The supernatant was filtered through a

Chromaphil PET-20/15MS-filter with 0.2 |im pore size from Macherey Nagel (Düren,

models 535 and 430 diode array detectors, set at 245 nm for MPTP, 295 nm for MPP+, and 345 nm for MPDP+. Samples were acidified with 9 |il perchloric acid (70%) per ml volume of

Germany). Separation was carried out on a C18 nucleosil column (250 x 4.6 mm; 5 |im particle size) from Macherey Nagel (Düren, Germany) at room temperature. The mobile phase consisted of acetonitrile : distilled water : triethylamine : sulfuric acid (12.50 : 86.18 :

^ „ 1.04 : 0.28, v/v, pH 2.3). The mobile phase was degassed with an online vacuum degasser and

2 delivered isocratically at a flow rate of 1 ml/min at an average pressure of 145 bar. Data

~ analysis was performed with Geminyx II software (Goebel Analytic). Peak areas of MPTP,

g is MPDP+, and MPP+ were evaluated with the Geminyx II software by the application of

2 ^ standard curves obtained for each of the three individual compounds.

+ ^ 13

feu Detection of C-labelled intracellular metabolites.

43 ft 13

;g73 For stable isotope labeling, cells were incubated with [U- C6] glucose for at least 18 h. To

4^ infer relative intracellular fluxes, intracellular metabolites were extracted and measured with

GC/MS. Cells grown in six-well plates were washed with 1 ml saline solution (0.9%) and

^ quenched with 0.4 ml of cold (- 20 °C) methanol. After adding an equal volume of cold water,

cells were collected with a cell scraper and transferred in tubes containing 0.4 ml of cold (-20

<D Cd .lit

' ö ¡S

°C) chloroform. The extracts were shaken at 1,400 rpm for 20 min at 4 °C and centrifuged at 16,000xg for 5 min at 4 °C. 0.3 ml of the upper aqueous phase was collected in specific GC glass vials and evaporated under vacuum at -4 °C using a refrigerated CentriVap Concentrator (Labconco). Metabolite derivatization was performed using an Agilent Autosampler. Dried

o ~ polar metabolites were dissolved in 15 pi of 2% methoxyamine hydrochloride in pyridine at

o.° 45 °C. After 60 min, an equal volume of 2,2,2-trifluoro-N-methyl-N-trimethylsilyl-acetamide

g^ +1% chloro-trimethyl-silane was added and metabolites were incubated for 30 min at 45 °C.

GC/MS analysis was performed using an Agilent 6890GC equipped with a 30m DB-35MS


o 43 a

■M (U t+H O

capillary column. The GC was connected to an Agilent 5975C MS operating under electron

§ is impact ionization at 70 eV. The MS source was held at 230 °C and the quadrupole at 150 °C.

The detector was operated in scan mode and 1 pi of derivatized sample was injected in splitless mode. Helium was used as carrier gas at a flow rate of 1 ml/min. The GC oven temperature was held at 80 °C for 6 min and increased to 300°C at a rate of 6°C/min. After 10

^ „ min, the temperature was increased to 325°C at a rate of 10 °C/min for 4 min. The run time

2 for each sample was 59 min. For determination of the mass isotopomer distribution (MID) of

2 ~ citrate, the mass spectrum was corrected for natural isotope abundance. Data processing from

raw spectra to MID correction and determination, was performed using MetaboliteDetector 2 & software (24).

• • o


This work was supported by RTG 1331, BMBF, the Doerenkamp-Zbinden-Foundation,

gja KoRS-Chemical Biology, and the Collaborative Research Center 969 "Chemical and

■5-S Biological Principles of Cellular Proteostasis", funded by the Deutsche

Forschungsgemeinschaft (DFG).

Author Disclosure Statement

No competing financial interests exist.

42 ££

If § List of Abbreviations

S 2 ^ « î

§ * 1,2-MPDP = 1 -methyl -4 -phenyl - 1,2-dihydropyridine

•S a DA = dopamine

.5 8 DAT = dopamine transporter

ftj^ DOPC = 1,2-dioleoyl-s«-glycero-3-phosphatidylcholine

DOPE = 1,2-dioleoyl-s«-glycero-3-phosphoethanolamine

■Ja & IMA = immortalized mouse astrocytes

<+h O O U

g,^ LUHMES = lund human mesencephalic neurons

MAO = monoamine oxidase

<u > +

4 MPDP = 1 -methyl -4-phenyl -2,3 -dihydropyridinium

MPP+ = 1-methyl-4-phenyl-pyridinium

£ g MPTP = 1-methyl-4-phenyl-tetrahydropyridine

, OCT = organic cation transporter

S3J2 PMAT = plasma membrane monoamine transporter

Be ft =3


13 p ft

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• • o

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cd ^ <D £

Ph cd m

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g Edwards RH, Greene LA, and Zecca L. Neuromelanin biosynthesis is driven by excess

• • o

s ^ Q-5

cytosolic catecholamines not accumulated by synaptic vesicles. Proc Natl Acad Sci USA 97(22): 11869-74, 2000.

^^ 51. Takahashi E, and Doi K. Impact of diffusional oxygen transport on oxidative metabolism in

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2 13 §

(MPTP). Toxicology 49(2-3): 513-9, 1988.

.5 Ö Ö

53. Tribl F, Asan E, Arzberger T, Tatschner T, Langenfeld E, Meyer HE, Bringmann G, Riederer

* ¿'S

P, Gerlach M, and Marcus K. Identification of L-ferritin in neuromelanin granules of the human substantia nigra: a targeted proteomics approach. Mol Cell Proteomics 8(8): 1832-8, 2009.

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55. Wu EY, Langston JW, and Di Monte DA. Toxicity of the 1-methyl-4-phenyl-2,3-^ a dihydropyridinium and 1-methyl-4-phenylpyridinium species in primary cultures of mouse

<u 4a a

■M (U <+H O

° £ 56. Zucca FA, Basso E, Cupaioli FA, Ferrari E, Sulzer D, Casella L, and Zecca L. Neuromelanin

<D £ W)<D

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Ph =d m

astrocytes. J Pharmacol Exp Ther 262(1): 225-30, 1992.

of the human substantia nigra: an update. Neurotox Res 25(1): 13-23, 2014.

Antioxidants & Redox Signaling

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Antioxidants & Redox Signaling

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§ FIG. 1. Formation and release of MPP+ by astrocytes. The mouse astrocyte cell line IMA

2 ~ 2.1 (A) and primary mouse glial cells (B) were incubated with MPTP (30 pM) for the

g S3 indicated time intervals. MPTP and MPP+ were measured in the supernatant by HPLC

2 ^ analysis. (C) A schematic overview of MPTP conversion by astrocytes. In the first step,

■¡3'i« MPTP is oxidized by mitochondrial monoamine oxidase B (MAO-B). In the second step, the

intermediate undergoes a nonenzymatic autoxidation to form MPP+. Intracellular formation of

42 & MPP+ would imply either an accumulation within mitochondria driven by their membrane

potential (AY) or a transporter-mediated passage across the membrane. The primary export mechanism for MPTP metabolites we propose herein assumes the formation of a membrane permeable intermediate that diffuses across the plasma membrane and ultimately forms MPP+

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in the extracellular space. (D) IMAs were exposed to MPTP (20 pM) for the time intervals

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indicated, and intracellular and extracellular (supernatant) MPP+ levels were analyzed. (E) IMA wildtype cells (wt) or transgenic cells stably expressing the organic cation transporter-3 (OCT3), the dopamine transporter (DAT), or a combination of both were exposed to MPP+

(20 pM) for the time periods indicated, and intracellular MPP+ was measured. (F) IMA clones

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й $ + + were exposed to MPP (20 pM) for 2 h to allow for saturation of uptake. Intracellular MPP

'3 g concentrations were calculated from measured MPP amounts in cell homogenates and cell

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ii^ OCT3, DAT, or a combination of both were exposed to varying MPP+ concentrations for 24

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Purified MAO-B enzyme (1 U/ml) was incubated with 30 pM MPTP for the time intervals indicated. In addition to MPP+, an intermediate was detected that was identified as MPDP+. (B) Wildtype (wt) IMAs or organic cation transporter 3 (OCT3)-expressing IMA were treated with MPTP (30 pM) for various incubation periods, MPP+ and MPDP+ levels in the

2 extracellular medium were assessed. (C) Mice received an intraperitoneal injection of MPTP

2 ~ (30 mg/kg) at t=0 min. After the time intervals indicated, samples from the striatum and the

¡« S3 cerebellum were collected and analyzed by HPLC. (D) Structures of MPTP and its derivatives

^ are shown. MPDP originates from the MAO-B catalyzed two-electron oxidation of MPTP.

'S'g Under physiological conditions, MPDP+ can also exist in its uncharged and membrane permeable base form 1,2-MPDP. This intermediate is converted by a non-enzymatic

^ & autoxidation into MPP+. (E) IMAs were incubated with 50 pM of MPTP, MPDP+, or

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decomposed MPDP+ for 18 h. Cell viability was detected by the resazurin reduction assay and by analysis of intracellular ATP. (F) For an analysis of intracellular fluxes of central carbon

° 8 metabolism, IMAs were incubated with [U- C6] glucose and treated with MPTP or MPDP

(50 pM) for 18 h. All data are means ± SD (n = 3). *p < 0.05

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FIG. 3. The intermediate MPDP+ crosses biological membranes. (A) Uptake velocities of MPP+ and MPDP+ were investigated by detecting intracellular MPP in IMAs incubated with 200 pM of compound for various time intervals. To exclude the involvement of cellular transporters or mitochondrial transmembrane potential, the same experimental setup was conducted with artificial vesicles composed of 50 mol % DOPC (1,2-dioleoyl-s«-glycero-3-phosphatidylcholine) and 50 mol % DOPE (1,2-dioleoyl-s«-glycero-3-phosphoethanolamine). Human erythrocytes were used as an alternative cellular model with high endogenous iron

2 content (hemoglobin), which facilitates the intracellular conversion of MPDP+. (B)

~ Bidirectional transport of MPDP+ across biological membranes was studied in IMAs loaded

g ö with 50 pM MPDP+ over the course of 30 min (left). Cells were then washed three times, new

2 medium was added, and the release of intracellular MPDP+ and MPP+ (middle) into the

'3'g medium (right) was detected. All data are means ± SD (n = 3). *p < 0.05

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FIG. 4. Transporter-independent uptake of MPDP into dopaminergic neurons. (A)

Neuronal LUHMES cells endogenously expressing the dopamine transporter (DAT) allow either a transporter-independent uptake of the MPDP+/1,2-MPDP intermediate followed by intracellular autoxidation into MPP+, or a DAT-dependent uptake of MPP+ originating from the extracellular autoxidation of MPDP+/1.2-MPDP. (B) As a sensitive readout for mitochondrial activity, glucose oxidation was monitored in LUHMES cells treated with MPP+ or MPDP (20 pM) by the addition of [U- C6] glucose in the presence or absence of the DAT blocker GBR (1 pM) for 18 h. (C) The morphology of LUHMES cells treated with MPP+ or MPDP+ (20 pM)

in the presence or absence of GBR (1 pM) for 24 h was visualized by staining with an anti-P-III-tubulin antibody. (D) To determine the respective contribution of DAT-dependent and independent uptake, LUHMES cells were incubated in low (20 pM) or high (200 pM) MPDP+ concentrations. Intracellular levels of ATP and MPP+ were detected after 24 h. All data are means ± SD (n = 3). *p < 0.05

Antioxidants & Redox Signaling

Preferential extracellular generation of the active parkinsonian toxin MPP+ by transporter-independent export of the intermediate MPDP+ (doi: 10.1089/ars.2015.6297) This article has been peer-reviewed and accepted for publication, but has yet to undergo copyediting and proof correction. The final published version may differ from this proof.

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FIG. 5. Modulation of the nonenzymatic conversion of MPDP+ into MPP+. (A) MPDP+ (100 pM) was kept under atmospheric conditions in DMEM medium at 37 °C, or under

g oxygen-free conditions (B) for the time intervals indicated. MPP , MPDP , and MPTP were

detected by HPLC. (C+D) MPDP+ (100 pM) was maintained in potassium phosphate buffer

g is with varying pH as indicated in each time interval. Autoxidation of MPDP+ into MPP+ was

monitored photospectrometrically. (E+F) MPDP (100 pM) was kept under atmospheric (E)

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or oxygen-free conditions (F) in potassium phosphate buffer of different pH at 37 °C for the

qJ indicated incubation intervals. (G,H) To identify the influence of metabolites of the dopamine

treated with the indicated compounds (100 pM each) under aerobic conditions with a pH of

4a a pathway and physiologically relevant iron-containing molecules, MPDP+ (100 pM) was

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FIG. 6. MPDP+/1,2-MPDP is the main exported form of MPTP metabolites. Monoamine oxidase-B (MAO-B)-dependent conversion of MPTP in astrocytes yields MPDP+ that partially exists in its conjugate base form 1,2-MPDP, and is able to freely diffuse across biological membranes. Conditions with an alkaline pH, elevated O2-tension, iron-containing proteins, or neuromelanin promote accelerated autoxidation of MPDP+ in the extracellular space. Due to its charge, stable MPP+ requires transporters such as the dopamine transporter (DAT) or the organic cation transporter (OCT) for cellular uptake.

Antioxidants & Redox Signaling

Preferential extracellular generation of the active parkinsonian toxin MPP+ by transporter-independent export of the intermediate MPDP+ (doi: 10.1089/ars.2015.6297) This article has been peer-reviewed and accepted for publication, but has yet to undergo copyediting and proof correction. The final published version may differ from this proof.

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Preferential extracellular generation of the active parkinsonian toxin MPP+ by transporter-independent export of the

intermediate MPDP+

Stefan Schildknecht, Regina Pape, Johannes Meiser, Christiaan Karreman, Tobias

Ts § Strittmatter, Meike Odermatt, Erica Cirri, Anke Friemel, Markus Ringwald, Noemi

lä h Pasquarelli, Boris Ferger, Thomas Brunner, Andreas Marx, Heiko M. Möller, Karsten

'S £ Hiller, Marcel Leist

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Supplementary information

J Supplementary Figures: 8

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FIG. S1. Characterization of DAT and OCT3 expressing IMA 2.1. IMA expressing either OCT3, DAT, or both transporters were analyzed for their respective OCT3 and DAT mRNA expression. RNA from the various IMA cell lines was isolated using the Purelink RNA mini kit of Ambion (Life Technologies GmbH, Darmstadt, Germany). The mRNA was reversed transcriped using the iScript Reverse Transcription Supermix for RT-qPCR and the PCR reactions were performed using the SsoAdvanced universal SYBR Green supermix both from Biorad (Bio-Rad Laboratories GmbH, Munich, Germany) . Reactions were carried out in a CFX96 Realtime Sytem/C1000Thermal cycler also purchased from BioRad.

Oligonucleotides for GAPDH, Dat1 and Oct3 were designed using the AiO program and manufactured by Eurofins MWG Operon (Ebersberg, Germany) The sequences are: GAPDH: 5'-



(A). To test the functionality of the transporters, DAT-IMA were treated with MPP+ (20 pM) in the presence or absence of the DAT-blocker GBR12909 (1 pM) (B). OCT3-IMA were treated in an identical manner but in the presence/absence of the OCT3 blocker D22 (2 pM) (C). Cells were then homogenized and analyzed for their intracellular content of MPP+. Interestingly to note: the OCT3 blocker efficiently blocked Oct3-dependent uptake of MPP+ in the range of 1-5 pM. At more extended incubation periods (> 2h), these concentrations however exhibited significant toxicity. (D) Corresponding to the intracellular detection of MPP+ illustrated in Fig. 1E, MPP+ levels in the extracellular compartment were assessed in wt-IMA, DAT-IMA, OCT3-IMA, and DAT/OCT3-IMA exposed to MPP+ (20 pM) for the time intervals as indicated.

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FIG. S2. MPP+ toxicity is determined by its uptake into the cytosol. Wildtype IMA (wt), respectively IMA expressing the organic cation transporter 3 (OCT3), the dopamine transporter (DAT), or both OCT3 and DAT (OCT3/DAT) were treated with MPP+ (5 pM) for the time intervals indicated. Viability was assessed (A) by the resazurin reduction assay, and by measuring intracellular levels of ATP (B). All data are means ± SD (n = 3). *p < 0.05

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FIG. S3. MAO-B-dependent conversion of MPTP. Purified monoamine oxidase-B (MAO-B) enzyme (10 U/ml) was added to PBS containing MPTP (100 pM). Consumption of MPTP and the concomitant generation of MPDP+ and MPP+ were detected by nuclear magnetic resonance analysis (NMR) over time. The numbers in the spectra indicate the position of the respective H-atom in the structures above.

Position of H atom MPTP [ppm] MPDP+ [ppm] MPP+ [ppm]

2 3.96+3.70 8.27 8.6

3 6.05 6.80 8.18

5 2.82+2.78 3.19 8.18

6 3.62+3.29 3.95 8.65

7 2.91 3.59 4.27

9 7.44 7,72 7.84

10 7.36 7.46 7.56

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FIG. S4. (A) Schematic overview on the incorporation of C-labeled carbon atoms in the

tricarboxylic acid (TCA) cycle. Uniformly labeled [U- C6] glucose (labeled carbons indicated in red) is added to cells and channeled into glycolysis and the TCA cycle. Input via

acetyl-CoA (M+2) leads to two C-labeled carbons while input via the pyruvate carboxylase pathway feeds in three 13C-labeled carbons (M+3). (B) IMA were treated with MPTP, MPP+, or MPDP (50 pM) together with [U- C6] glucose for 18 h. Pyruvate carboxylase activity was analyzed by determining fractional enrichment of (M+3) aspartate. (C) As an alternative

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indicator for TCA inhibition, extracellular lactate was detected. MPTP, MPP+ and MPDP+

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FIG. S5. In vivo transport of MPDP+ across membranes. Mice received an intravenous injection of either MPDP or MPP+ (60 mg/kg). After 3 min, the animals were sacrificed, blood was collected, erythrocytes were analyzed for intracellular MPP+. All data are means ± SD (n=3). *p < 0.05.

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FIG. S6. Comparison of dopamine neuron toxicity in mono- and in co-culture with IMA astrocytes. Low density IMA in mono-culture were treated with MPTP (20 pM), monocultures of LUHMES were treated with MPP+ (5 pM) for 48 h in the presence or absence of the MAO-B inhibitor deprenyl (10 pM), or the DAT-blocker GBR12909 (1 pM). Cell morphology of IMA was visualized by staining with an anti-S100ß antibody (green), LUHMES were stained with an anti-ß-III-tubulin antibody (orange). Co-cultures of confluent IMA (green) and LUHMES seeded on top (orange) were treated with MPTP (20 pM) for 5 days in the presence or absence of deprenyl (10 pM) or GBR12909 (1 pM).

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FIG. S7. Influence of antioxidants on the generation of MPP+. (A) IMA were incubated with MPTP (20 pM) for 6h in the presence of varying concentrations of ascorbic acid. MPTP, MPDP+, and MPP+ were assessed in the supernatant. (B) MPDP (100 pM) in a cell-free system was treated with Fe alone (100 pM), with a combination of Fe and desferoxamine (100 pM), or with a combination of Fe and ascorbic acid (100 pM) for the time intervals as indicated. The decrease of MPDP+ (B) and the corresponding formation of MPP+ (C) were photometrically detected.

FIG. S8. Autoxidation of MPDP+. Interaction of MPDP+ with molecular oxygen, transition metals, or reactive oxygen species (ROS) yields a stabilized allyl radical (M1). Reaction of

g the allyl radical with a second molecule of O2 leads to the formation of a peroxide group at

the C5 atom (M2), promoting the release of a proton from C6 to finally form MPP+ (M3).

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