Abstract

   Parkinson's disease (PD) is characterized by the progressive
   deterioration of dopamine (DA) neurons, and therapeutic endeavors are
   aimed at preventing DA loss. However, lack of effective brain delivery
   approaches limits this strategy. In this study, a “Trojan horse” system
   is used for substantia nigra‐targeted delivery of a blood brain
   barrier‐penetrating peptide (RVG29) conjugated to the surface of
   nanoparticles loaded with the natural autophagy inducer
   4,4′‐dimethoxychalcone (DMC) (designated as RVG‐nDMC). Here, the
   neuroprotective effects of DMC are demonstrated in PD. Specifically,
   RVG‐nDMC penetrates the blood brain barrier with enhanced
   brain‐targeted delivery efficiency and is internalized by DA neurons
   and microglia. In vivo studies demonstrate that RVG‐nDMC ameliorates
   motor deficits and nigral DA neuron death in PD mice without causing
   overt adverse effects in the brain or other major organs. Moreover,
   RVG‐nDMC reverses tyrosine hydroxylase ubiquitination and degradation,
   alleviates oxidative stress in DA neurons, and exerts antiinflammatory
   effects in microglia. The “Trojan horse” strategy for targeted delivery
   of DMC thus provides a potentially powerful and clinically feasible
   approach for PD intervention.

   Keywords: blood brain barrier, brain targeted delivery, DMC,
   neuroinflammation, oxidative stress, TH ubiquitination
     __________________________________________________________________

   A Trojan horse system (RVG‐nDMC) is constructed for blood brain
   barrier‐penetrated delivery of 4,4′‐dimethoxychalcone (DMC) to
   ameliorate motor deficits and nigral dopamine (DA) neuronal death in
   Parkinson's disease. Mechanistically, RVG‐nDMC reduces tyrosine
   hydroxylase ubiquitination and degradation, alleviates the oxidative
   stress in DA neurons, and exerts antiinflammation effects in microglia
   for synergistic Parkinsonian intervention.

   graphic file with name ADVS-8-2004555-g006.jpg

1. Introduction

   Parkinson's disease (PD) is the most prevalent neurodegenerative
   movement disorder, affecting ≈2–3% of the population ≥65 years of age
   worldwide.^[ [40]^1 , [41]^2 ^] Pathologically, PD is characterized by
   the loss of dopamine (DA) neurons in the pars compacta of the
   substantia nigra (SNpc) and the formation of intracytoplasmic Lewy
   bodies due to accumulation of misfolded α‐synuclein.^[ [42]^3 , [43]^4
   ^] The progressive deterioration of vulnerable DA neurons in the SNpc
   may be attributed to cellular disturbances characterized by
   mitochondrial dysfunction, disruption of the autophagy–lysosome system,
   misfolding and aggregation of misfolded proteins, and endoplasmic
   reticulum (ER) stress and neuroinflammation.^[ [44]^3 , [45]^5 ^]
   Notably, nearly 50% of DA neuron loss in the SNpc occurs when movement
   deficiencies appear; however, there is still a lack of efficient means
   for mediation to attenuate or prevent DA neuron death.^[ [46]^2 ,
   [47]^3 ^] Thus, mechanistic and therapeutic targets for rescuing DA
   neuron degeneration are strongly desirable.

   DA neuron replenishment represents a cardinal target for both
   idiopathic and symptomatic PD therapeutics; however, as the disease
   progresses, long‐term therapy may induce dyskinesia and other motor
   complications.^[ [48]^6 ^] Although several reprogrammable and
   gene‐editing approaches have yielded promising benefits for alleviating
   neurological deficits and DA neuron loss in PD animal models,
   investigations are still preclinical, and technical challenges and
   ethical considerations must be overcome before their clinical
   application.^[ [49]^7 ^]

   Tyrosine hydroxylase (TH) catalyzes the formation of
   L‐3,4‐dihydroxyphenylalanine (l‐DOPA), the rate‐limiting step in the
   biosynthesis of DA, and TH is, therefore, recognized to be a
   characteristic hallmark of the DA neuron. Nigrostriatal TH protein has
   been widely reported to be lost in PD patients,^[ [50]^8 , [51]^9 ^]
   and its loss correlates with the progression of DA neuron death.
   Emerging evidence reveals that TH is ubiquitinated in PD, which results
   in its degradation by the ubiquitin–proteasome system.^[ [52]^10 ,
   [53]^11 , [54]^12 ^] Importantly, pathological TH is a potential source
   and contributor to the generation of reactive oxygen species (ROS),
   which is a common feature of many neurodegenerative disorders,
   including PD.^[ [55]^13 , [56]^14 ^] Thus, TH may be a promising
   candidate target protein for developing new treatments of PD,^[ [57]^15
   ^] though the potential of targeting TH ubiquitination has not been
   extensively researched.

   Recently, 4,4′‐dimethoxychalcone (DMC) has been identified as a natural
   autophagy inducer with antiaging properties.^[ [58]^16 ^] Because DMC
   promotes cytoprotection and autophagy across species, we speculated
   that it may be neuroprotective in PD treatment. However, the blood
   brain barrier (BBB) would be likely to impede the potential
   intervention efficacy of DMC, and the proposed mechanisms of DMC for PD
   remain unexplored. Over the last decades, increasing efforts have
   focused on developing effective nanocarrier systems that cross the BBB
   to treat neurodegenerative diseases.^[ [59]^17 , [60]^18 ^]
   Appropriation of brain‐targeting ligands, such as transferrin receptors
   and glucose transporters, facilitates BBB penetration by nanodelivery
   systems for more effective neurodegenerative diseases therapy.^[
   [61]^19 , [62]^20 ^] Although several systems have been established to
   delivery nanomedicines that target α‐synuclein, as another pathological
   hallmark in PD progression,^[ [63]^20 , [64]^21 ^] targeted delivery
   systems to treat PD by restoring DA activity remain unexplored.

   In this study, a BBB‐penetrating peptide (RVG29)‐based “Trojan horse”
   system (designated as RVG‐nDMC) was prepared as a strategy for PD
   therapy by substantia nigra‐targeted delivery of DMC in a mouse model
   (Scheme [65] 1A). RVG‐nDMC penetrated the blood brain barrier and
   enhanced the brain‐targeted delivery efficiency by binding the
   nicotinic acetylcholine receptor (nAchR), which was subsequently
   internalized by DA neurons and microglia (Scheme [66]1B). Importantly,
   in vivo and in vitro studies demonstrated that RVG‐nDMC reduces TH
   ubiquitination and degradation independent of regulating TH
   phosphorylation and decreases subsequent ubiquitinated TH‐mediated
   oxidative stress in DA neurons. RVG‐nDMC also alleviates inflammatory
   cytokine release in microglia (Scheme [67]1C). Moreover, transcriptome
   sequencing and metabolomics analysis confirmed that RVG‐nDMC
   ameliorates motor deficits and nigral DA neuron death in PD mice
   without observable toxic effects in nontargeted organs, indicating
   great potential of RVG‐nDMC for clinical translation and
   neuroprotection in PD.

Scheme 1.

   Scheme 1
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   The preparation and proposed mechanism of RVG‐nDMC for PD intervention.
   A) Schematic illustration of the synthesis of RVG‐nDMC. RVG‐nDMC is
   composed of DMC modified with polymer nanoparticle linked with RVG
   peptide. B) Proposed mechanism by which RVG‐nDMC ameliorates motor
   deficits and nigral DA neuron death in the MPTP‐induced PD mouse model.
   C) Briefly, RVG‐nDMC penetrates the BBB and accumulates in DA neurons
   and microglia in the brain. Specifically, RVG‐nDMC reverses
   MPTP‐induced TH ubiquitination and degradation, as well as oxidative
   stress. Additionally, RVG‐nDMC exerts antiinflammatory effects in the
   PD mouse model.

2. Results and Discussion

2.1. Preparation and Characterization of RVG‐nDMC

   To obtain RVG‐nDMC, DMC was first physically encapsulated into the
   hydrophobic core of PEG‐b‐PLGA nanoparticles. PLGA20K‐b‐mPEG2K and
   PLGA20K‐PEG5K‐MAL were used to prepare nDMC, which was further modified
   with RVG29 peptide through a maleimide–thiol reaction (Figure [69] 1A).
   As shown in Figure [70]1B,[71]C, the nDMC and RVG‐nDMC appeared as
   uniform nanosized and sub‐200 nm nanoparticles. Moreover, the positive
   charge of nDMC reversed to a negative charge after RVG29 peptide
   modification, which confirms that RVG29 peptide was successfully
   conjugated onto nDMC (Figure [72]1D). Similar characteristic absorption
   peaks of DMC, nDMC, and RVG‐nDMC were observed at 350 nm, demonstrating
   that DMC was efficiently loaded into nDMC and RVG‐nDMC (Figure [73]1E).
   The drug encapsulation efficiency (EE), loading efficiency (LE), and
   release behaviors of nDMC and RVG‐nDMC were measured by using a
   standard curve of the UV–vis absorbance of DMC at 350 nm (Figure S1,
   Supporting Information), and the resulting EEs for nDMC and RVG‐nDMC
   were 94.2% and 91.1%, with LEs for nDMC and RVG‐nDMC of 15.8% and
   15.2%, respectively (Figure [74]1F). Furthermore, DMC underwent
   sustained released from nDMC and RVG‐nDMC under physiological
   conditions (Figure [75]1G), which has potential therapeutic advantages.
   Collectively, these results suggest successful preparation of RVG‐nDMC
   for assessment of the therapeutic benefit of DMC.

Figure 1.

   Figure 1
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   Characterization of nDMC and RVG‐nDMC. A) Schematic illustration of the
   synthesis process of RVG‐nDMC nanoparticles. Evaluation of nDMC and
   RVG‐nDMC by B) particle size analysis; C) TEM observation (Scale bars,
   200 nm); and D) zeta potential analysis. n = 3 per group. E) UV–vis
   absorbance of DMC, nDMC, and RVG‐nDMC. F) EE and LE of DMC in nDMC and
   RVG‐nDMC were measured with a spectrophotometer. G) The drug release
   kinetics of nDMC and RVG‐nDMC. n = 3 per group. Results are expressed
   as the mean ± SEM. Statistical significance was determined by Student's
   t‐tests.

2.2. RVG‐nDMC Displays Widespread Cellular Distribution and Restores
MPP^+‐Damaged Cell Viability in MN9D Cells

   RVG29 peptide has been shown to recognize neural nAchR or
   gamma‐aminobutyric acid (GABA) receptor, which may promote the
   intracellular delivery in nAchR or GABA‐overexpressed neural cells.^[
   [77]^22 ^] Therefore, we investigated the cellular uptake behavior of
   RVG29‐nDMC in DA neuron‐derived MN9D cells. As shown in Figure [78] 2A,
   RVG29‐nDMC was efficiently internalized by MN9D cells. Intriguingly,
   RVG‐nDMC promoted more cytoskeletal accumulation than RVG‐n did,
   suggesting that DMC conjugation may promote nanoparticle entry
   (Figure [79]2A–[80]C). To quantitatively analyze differences in the
   cellular uptake of nDMC and RVG‐nDMC, we employed flow cytometry. The
   cellular uptake of RVG‐nDMC was significantly increased as compared
   with the cellular uptake of nDMC (Figure S2A,B, Supporting
   Information), which is consistent with the microscopy results. These
   findings suggest that intracellularly released DMC may promote the
   membrane distribution or recycling of nAchR or GABA receptor, which may
   induce its sequential binding with RVG29‐nDMC and increase nanoparticle
   entry. Because DMC has been reported to be an autophagy inducer with
   antiaging properties,^[ [81]^16 ^] we examined the interactions of
   RVG‐nDMC with the mitochondria and lysosomes. RVG‐nDMC significantly
   accumulated in the mitochondria and lysosome as determined by
   costaining with mitochondrial tracker (Mito‐tracker), mitochondrial
   marker, pyruvate dehydrogenase E1α (PDH‐E1α), and lysosomal tracker
   (Lyso‐tracker) (Figure [82]2D–[83]G; Figure S3A,B, Supporting
   Information). In a previous study, a modified RVG peptide (RVG9R) was
   used to deliver short interfering RNA (siRNA) to the brain, and
   RVG9R‐loaded siRNA was found to specifically interact with the
   Mitochondrial Complex I.^[ [84]^23 ^] Thus, although there are no
   mitochondria‐targeting ligands on the surface of the nanoparticles, it
   would be of interest in the future to determine whether RVG29 can
   physically interact with mitochondria.

Figure 2.

   Figure 2
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   Biological effects of RVG‐nDMC on DA neuron‐derived MN9D cells. A)
   Representative images and B,C) quantitative fluorescence analysis of
   α‐Tubulin in MN9D cells after 6 h incubation with different
   Cy5.5‐labelled t nanoparticles. n = 6 per group. D) Representative
   images and E) quantitative fluorescence analysis of mitochondrial
   tracker (Mito‐tracker) in MN9D cells after 6 h incubation with
   Cy5.5‐labelled nDMC and RVG‐nDMC. n = 6 per group. F) Representative
   images and G) quantitative fluorescence analysis of lysosomal tracker
   (Lyso‐tracker) in MN9D cells after 6 h incubation with Cy5.5‐labelled
   nDMC and RVG‐nDMC. n = 6 per group. Scale bars, 50 µm for the original
   images and 10 µm for the magnified images. H) Cell viability of MN9D
   cells treated with increasing concentrations of DMC, nDMC, and
   RVG‐nDMC. n = 3 per group. I) Cell viability of MN9D cells treated with
   increasing concentrations of MPP^+. n = 3 per group. J) Cell viability
   of MPP^+ pretreated MN9D cells incubated with DMC, nDMC, and RVG‐nDMC.
   n = 3 per group. Results are expressed as the mean ± SEM. ^** p < 0.01.
   ^## p < 0.01, ^# p < 0.05 vs MPP^+ + RVG‐nDMC group. The Unpaired
   Student's t‐test was used for comparison between two groups. One‐way
   ANOVA with a Tukey post‐hoc analysis was used for comparison among
   multiple groups.

   Further analysis suggests that modification of DMC to nDMC and RVG‐nDMC
   significantly decreased the undesired cytotoxicity of DMC, especially
   at higher DMC concentrations (50 × 10^−6, 100 × 10^−6, and 200 × 10^−6
   m) (Figure [86]2H). We further constructed an in vitro PD model by
   using neurotoxin 1‐methyl‐4‐ phenylpyridinium (MPP^+) to treat MN9D
   cells with an optimized concentration of MPP^+ at 1000 × 10^−6 m
   (Figure [87]2I). RVG‐nDMC, but not free DMC or nDMC, extensively
   restored MPP^+‐decreased cell viability (Figure [88]2J). These results
   reveal that RVG‐nDMC targets mitochondria and lysosomes upon
   internalization, is relatively nontoxic, and shows beneficial effects
   in an in vitro model of PD.

2.3. Brain Targeting and Distribution of RVG‐nDMC in the SNpc

   The BBB impedes the efficiency of brain‐targeted drug delivery, which
   restricts the efficiency of PD interventions. Therefore, we determined
   whether RVG‐nDMC may have the ability to overcome this limitation.
   Indeed, our results indicate that RVG‐nDMC possesses favorable
   brain‐targeting ability. Compared to nDMC, RVG‐nDMC exhibited a greater
   efficiency in brain‐targeting delivery as evaluated by photoacoustic
   imaging (Figure [89] 3A,[90]B), fluorescence imaging
   (Figure [91]3C,[92]D), and ex vivo fluorescence imaging
   (Figure [93]3E,[94]F). Notably, the distribution of RVG‐nDMC‐Cy5.5 in
   the brain was earlier and prolonged relative to the distribution of
   nDMC‐Cy5.5. These results suggest that RVG‐nDMC can penetrate the BBB
   with more extensive distribution than for nDMC, which is of potential
   advantage for PD intervention (Figure [95]3G).

Figure 3.

   Figure 3
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   Brain Targeting Delivery of RVG‐nDMC. A) Real time photoacoustic
   imaging and B) corresponding photoacoustic signal analysis of mice
   after intravenously injection of nDMC or RVG‐nDMC. n = 3 per group. C)
   Real time fluorescence imaging and D) corresponding fluorescence
   analysis of mice after intravenously injection of Cy5.5‐labelled nDMC
   or RVG‐nDMC. n = 3 per group. E) Ex vivo imaging and F) corresponding
   fluorescence analysis of the sacrificed tissues after intravenously
   injection of Cy5.5‐labelled nDMC or RVG‐nDMC at 48 h. n = 3 per group.
   G) Schematic illustration of the proposed mechanism of RVG‐nDMC for
   brain targeting delivery. H–J) Representative images of TH, Iba1, and
   GFAP staining in SN derived from mice treated with Cy5.5, nDMC‐Cy5.5,
   or RVG‐nDMC‐Cy5.5 at 6 h postinjection. White arrows in the enlarged
   details of the right column show the presence of nanoparticles in DA
   neurons and microglia. Red arrows show the presence of nanoparticles
   outside of astrocytes and microglia. Scale bars, 50 µm for the original
   images and 10 µm for the magnified images. Results are expressed as the
   mean ± SEM. ^** p < 0.01, ^* p < 0.05. The Unpaired Student's t‐test
   was used for comparison between two groups.

   The SNpc is a well‐known vulnerable brain region in PD.^[ [97]^24 ^]
   Thus, to further evaluate the SNpc targeting ability of RVG‐nDMC, we
   examined the distribution of RVG‐nDMC in the SNpc at different time
   points (1, 6, and 24 h) using differential neural cell markers (TH for
   DA neuron, ionized calcium binding adapter protein 1 (Iba1) for
   microglia, and glial fibrillary acidic protein (GFAP) for astrocytes)
   after intravenous injection. RVG‐nDMC was localized in DA neurons at 1,
   6, and 24 h postinjection (Figure [98]3H; Figures S4A and S5A,
   Supporting Information), indicating favorable DA neuron targeting
   ability. Intriguingly, Cy5.5 signals were also observed in microglia at
   1, 6, and 24 h postinjection (Figure [99]3I; Figures S4B and S5B,
   Supporting Information). Microglia express much less nAchR and GABA
   receptor than neurons and function as resident professional phagocytes
   in the brain.^[ [100]^25 ^] Therefore, we speculate that RVG‐nDMC might
   be engulfed by microglia in the SNpc, which play a clear role in the
   inflammation of PD. In contrast, no RVG‐nDMC was observed in astrocytes
   (Figure [101]3J; Figures S4C and S5C, Supporting Information). Thus,
   these results reveal that RVG‐nDMC penetrates the SNpc and localizes
   primarily in DA neurons, with additional localization in microglia.

2.4. RVG‐nDMC Administration Improves Motor Deficits in PD Mice

   To evaluate whether RVG‐nDMC may provide in vivo PD intervention
   efficacy via DA neuron‐targeted delivery, we employed the
   1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP)‐induced PD mouse
   model.^[ [102]^26 ^] MPTP was administrated intraperitoneally
   continuously over 5 days to induce a subacute PD state (Figure [103]
   4A). Simultaneously, nDMC, RVG‐nDMC (equivalent dose of DMC), or the
   control vehicle were administered intravenously every other day for 12
   days. After 3 days, behavioral tests were carried out to evaluate the
   protective effects of RVG‐nDMC against motor dysfunction mediated by
   MPTP administration. No obvious significant differences were observed
   in the total travelled distance, time spent in the center, or movement
   speed in the open field test (Figure [104]4B; Figure S6A,B, Supporting
   Information). However, RVG‐nDMC reversed the MPTP‐induced increased
   pole‐climbing time and decreased the holding time and latency to fall
   in the rotarod test; and the improved behavior performance was not
   observed for nDMC or RVG‐n alone, suggesting that both the nanocarrier
   and DMC moieties of RVG‐nDMC are required for improved motor
   performance (Figure [105]4C–[106]E).

Figure 4.

   Figure 4
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   RVG‐nDMC attenuates motor deficits and DA neuron loss in PD mice. A)
   Experimental design for RVG‐nDMC administration in MPTP‐induced PD
   mice. B) Total distances travelled in the open‐field of PD mice treated
   with nDMC, RVG‐n, and RVG‐nDMC. C) The pole‐climbing test was used to
   examine the bradykinesia of PD mice. D) The grasping test was used to
   further examine the grip strength of PD mice. E) The rotarod test was
   used to examine the motor coordination of PD mice. n = 10, 9, 10, 10,
   and 9 for Control, MPTP, MPTP + nDMC, MPTP + RVG‐nDMC, and MPTP + RVG‐n
   groups, respectively. F,G) Quantification of the TH‐positive density in
   the striatum and TH‐positive neurons in the SN. n = 6 per group. H)
   Immunohistochemical staining of TH‐positive neurons in the striatum and
   SNpc in Control, MPTP, MPTP + nDMC, MPTP + RVG‐nDMC, and MPTP + RVG‐n
   groups. The ellipses in the middle column of panel H show the
   boundaries of the SNpc and the middle‐column boxes denote the areas
   that are expanded in the right‐hand columns. Scale bars, 1 mm for
   images in striatum; 800, 100, and 50 µm for the series of images in
   SNpc. Results are expressed as the mean ± SEM. ^** p < 0.01, ^* p <
   0.05 vs Control. ^## p < 0.01, ^# p < 0.05 vs MPTP group. ^&& p < 0.01,
   ^& p < 0.05 vs MPTP + RVG‐nDMC group. Statistical significance was
   determined by one‐way ANOVA and Tukey tests for post‐hoc comparisons.

   TH is the rate‐limiting enzyme in DA biosynthesis, and its dysfunction
   may underlie DA neuron loss in PD.^[ [108]^4 , [109]^8 , [110]^27 ^]
   Therefore, we evaluated the effect of RVG‐nDMC on TH levels after MPTP
   treatment. No obvious differences were observed in the striatal TH
   density between the different treatment groups (Figure [111]4F,[112]H).
   However, the MPTP + RVG‐nDMC group had greater amounts of TH‐positive
   neurons in the SNpc as compared with the MPTP, MPTP + nDMC, and MPTP +
   RVG‐n groups (Figure [113]4G,[114]H). Furthermore, RVG‐nDMC was
   verified to increase nigral TH and dopamine transporter (DAT)
   expression in PD mice by Western blot analysis (Figure [115]
   5A–[116]C). Consistently, RVG‐nDMC increased the number of DA synaptic
   vesicles in the substantia nigra (SN) in the PD model mice as evaluated
   by transmission electron microscopy (TEM) of the ultrastructural
   morphology of synaptic vesicles (Figure [117]5D,[118]E). These results
   suggested that RVG‐nDMC improves MPTP‐damaged DA synaptic plasticity.
   Notably, based on high‐performance liquid chromatography coupled with
   tandem mass spectrometry (HPLC/MS‐MS), RVG‐nDMC also reversed the
   MPTP‐induced decrease in DA, as well as 3,4‐dihydroxyphenylacetic acid
   (DOPAC), a product of the metabolism of DA (Figure [119]5F–[120]H). In
   contrast, RVG‐nDMC showed no obvious effect on the levels of other
   neurotransmitters, such as serotonin (5‐hydroxytryptamine, 5‐HT) and
   5‐hydroxyindoleacetic acid (5‐HIAA), the chief end‐product of serotonin
   metabolism (Figure [121]5F,[122]I,[123]J). Moreover, there was no
   obvious effect on the expression of several synaptic proteins after
   treatment with MPTP and RVG‐nDMC, such as Synapsin, Syntaxin,
   Synaptotagmin, and PSD‐95 in the SN (Figure [124]5K–[125]M), suggesting
   RVG‐nDMC may not alter nigral synaptic transmission in the PD model
   mice. Therefore, these results are consistent with specificity in the
   effect of RVG‐nDMC in ameliorating DA loss in the SNpc.

Figure 5.

   Figure 5
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   RVG‐nDMC increases TH expression and DA synthesis in PD mice. A–C)
   Representative blots and quantification showing TH and DAT expression
   levels in the SN of Control, MPTP, and MPTP + RVG‐nDMC groups. n = 6
   per group. D) Ultrastructural analysis of synaptic vesicles in the SN
   of the three groups. Scale bars, 500 nm. Black arrows show the
   morphological synaptic vesicles. E) Quantification of synaptic vesicles
   in the SN of the three groups. n = 10 per group. F) Typical HPLC
   chromatograms of DA, DOPAC, 5‐HT, and 5‐HIAA. G–J) The levels of DA,
   DOPAC, 5‐HT, and 5‐HIAA were assessed by HPLC‐MS/MS analysis. n = 4 per
   group. K–M) Representative blots and quantification showing Synapsin,
   Syntaxin, Synaptotagmin, and PSD‐95 expression levels in the SN of the
   three groups. n = 6 per group. Results are expressed as the mean ± SEM.
   ^** p < 0.01 vs Control. ^## p < 0.01, ^# p < 0.05 vs MPTP group.
   Statistical significance was determined by one‐way ANOVA and Tukey
   tests for post‐hoc comparisons.

   Given that RVG‐nDMC becomes localized, in part, in the liver, lung and
   kidney of mice after intravenous administration, we examined potential
   adverse effects of RVG‐nDMC on non‐targeted organs. After RVG‐nDMC
   administration, no detectable pathological changes were observed in the
   liver, heart, kidney, lung or spleen from normal mice as evaluated by
   H&E staining (Figure S7, Supporting Information). Consistently, no
   detectable alterations were observed in serum biochemical indicators of
   liver or kidney function following nDMC and RVG‐nDMC administration
   (Table S1, Supporting Information). However, nDMC resulted in
   observable renal tubular atrophy and interstitial loosening in the
   kidney without changes in serum biochemical indicators of kidney
   function (black arrows in Figure S7 and Table S1 in the Supporting
   Information). Substantially more RVG‐nDMC as compared with nDMC could
   be observed in the kidneys of the mice (Figure [127]3E,[128]F). This
   may be partially due to the expression of nAchR in the kidney,^[
   [129]^28 ^] which facilitates the binding of RVG29 peptide. The slight
   kidney damage upon nDMC treatment observed in this study may result
   from the interplay between MPTP and dysfunctional uptake of nDMC in the
   kidney. Moreover, the surface charge of nanoparticles dictates the
   suborgan distribution of nanoparticles in the kidney, which also may
   affect the metabolism of nanoparticles.^[ [130]^29 ^] As illustrated in
   Figure [131]1D, the modification of RVG29 peptide reverses the
   negatively charged nDMC to positively charged RVG‐nDMC, which might
   further affect drug metabolism and drug effects. Importantly, we did
   not find significant changes in biochemical indicators of kidney
   function. Thus, we hypothesize that this may be a temporary structural
   dysfunction, and further study is needed to evaluate the safety and
   toxicity of DMC in the future. However, these results indicate overall
   that RVG‐nDMC can efficiently exert neuroprotection without causing
   obvious side effects.

2.5. RVG‐nDMC Administration Inhibits TH Ubiquitination in PD Mice

   Next, we investigated the proposed mechanism of RVG‐nDMC for restoring
   TH positive neurons by RNA‐sequencing analysis. Volcano plots
   (Figure [132] 6A,[133]B) and hierarchical clustering (Figure [134]6C)
   identified 33 representative differentially expressed genes (DEGs) that
   were increased by MPTP and also were reversed by MPTP + RVG‐nDMC
   treatment. For further insight into the potential pathways involved in
   these DEGs, we performed Kyoto Encyclopedia of Genes and Genomes (KEGG)
   analysis (Figure [135]6D; Figure S8, Supporting Information).
   Intriguingly, the top 3 DEG‐enriched pathways were “Ribosome,”
   “Parkinson's disease,” and “Oxidative phosphorylation,” while there
   also was high enrichment for the “Dopaminergic synapse” pathway (see
   green arrows in Figure [136]6D). For further verification, we examined
   the number and p value of identified DEGs for the PD‐related KEGG
   pathways, including “Ribosome,” “Parkinson's disease,” “Oxidative
   phosphorylation,” “Dopaminergic synapse,” “Synaptic vesicle cycle,” and
   “Proteasome” (Figure [137]6E,[138]F). Additionally, the Log[2] fold
   change and p value of the 33 individual DEGs enriched in these pathways
   were measured. The results demonstrate that the overlapping
   dopaminergic synapse and PD‐related genes (Th, Ddc, Slc18a2, Slc6a3,
   and Ubb) were increased in the SN of PD mice as compared with control
   mice, and that RVG‐nDMC decreased the mRNA expression of these genes,
   with similar results for 18 ribosome related genes
   (Figure [139]6G,[140]H). Notably, the expression of specific genes
   encoding ribosomal proteins has been demonstrated to be decreased in
   the SN in the late stage of PD patients, with associated progressive
   neuronal loss in the SN.^[ [141]^30 ^] Therefore, the increased
   expression of ribosomal protein subunit genes might be compensatory to
   the DA neuron loss upon subacute MPTP damaged in SN. Consistently,
   further bioinformatic analysis verified that the altered genes have key
   roles in Ribosome, Parkinson's disease, Oxidative phosphorylation and
   Dopaminergic synapse pathways (Figure S9A–D, Supporting Information).

Figure 6.

   Figure 6
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   RNA‐sequencing analysis of RVG‐nDMC treatment in the MPTP‐induced PD
   mouse model. A,B) Volcano plots showing the DEGs between the MPTP and
   Control groups, as well as the MPTP + RVG‐nDMC and MPTP groups. C)
   Hierarchical clustering of 33 representative MPTP + RVG‐nDMC‐reversed
   DEGs that were increased in MPTP as compared with Control group. D)
   KEGG pathways enriched by DEGs among these three groups. Green arrows
   show the PD‐related pathways. E,F) The number and p value of identified
   DEGs in PD‐related KEGG pathways. G,H) The Log[2] fold change and p
   value of 33 individual DEGs enriched in the KEGG pathways in (E) and
   (F).

   Because we found the opposite expression pattern for TH mRNA and
   protein upon RVG‐nDMC treatment in the MPTP mouse model, we further
   explored the related mechanisms. Mechanistically, the activity of TH is
   subject to long‐term regulation of gene expression (enzyme stability,
   transcriptional and translational regulation, and RNA stability), as
   well as short‐term regulation of enzyme activity (allosteric
   regulation, feedback inhibition, and phosphorylation).^[ [143]^31 ^]
   Previous work has demonstrated that TH protein is ubiquitinated and
   degraded by the ubiquitin–proteasome system, dependent, in part, on its
   phosphorylation, and that this ubiquitination is linked DA neuron
   loss.^[ [144]^11 , [145]^12 , [146]^32 ^] Therefore, we sought to
   evaluate the effects of RVG‐nDMC on TH protein phosphorylation and
   ubiquitination. First, we evaluated three phosphorylation sites on TH
   located in the N‐terminal regulatory domain.^[ [147]^33 ^] RVG‐nDMC
   showed no obvious effects on TH phosphorylation at Ser19, Ser31, and
   Ser40 (Figure [148] 7A–[149]D). Next, we performed
   co‐immunoprecipitation (co‐IP) assays to determine whether TH is
   ubiquitinated upon MPTP treatment. A direct interaction between
   ubiquitin and TH was observed using capture antibody for either
   ubiquitin or TH, and RVG‐nDMC reduced this interaction, suggesting that
   RVG‐nDMC inhibits ubiquitination of TH (Figure [150]7E–[151]J). Similar
   results were also observed by costaining of TH and ubiquitin in the
   SNpc (Figure [152]7K,[153]L). Because RVG‐nDMC reverses the decreased
   TH expression induced by MPTP, these results are consistent with the
   possibility that RVG‐nDMC inhibits the degradation of ubiquitinated TH
   to restore its expression in the SN of PD mice. Generally,
   ubiquitination of TH appears to be triggered by enzyme phosphorylation;
   however, other factors also induce TH degradation via proteasomes
   independent of its phosphorylation. For example, the inflammatory
   cytokine, ciliary neurotrophic factor, leads to the ubiquitination of
   TH in a gp130‐dependent manner, and gp130 cytokines induce proteasomal
   degradation of TH through an extracellular signal‐regulated kinases1/2
   (ERK1/2)‐dependent pathway.^[ [154]^12 ^] ERK5 signaling is suggested
   to regulate ubiquitination of TH in DAergic PC12 cells independent of
   its phosphorylation.^[ [155]^34 ^] Additionally, the
   MgATP‐ubiquitin–proteasome‐dependent pathway is also indicated to
   regulate TH ubiquitination, suggesting that energy metabolism may play
   a specific role in the degradation of TH.^[ [156]^35 ^] Given that MPTP
   induces energy depletion by damaging mitochondrial function,^[ [157]^36
   ^] we hypothesize that abnormal energy metabolism mediated by MPTP
   administration may contribute to the ubiquitination of TH. Thus, these
   results suggest that potential signaling pathways may modulate the
   ubiquitination of TH independent of its phosphorylation.

Figure 7.

   Figure 7
   [158]Open in a new tab

   RVG‐nDMC reverses TH ubiquitination in the MPTP‐induced PD mouse model.
   A–D) Representative blots and quantification showing the expression
   levels of phosphorylated TH at Ser19, Ser31, and Ser40 in the SN of
   Control, MPTP, and MPTP + RVG‐nDMC groups. n = 6 per group. E–G)
   Representative blots and quantification of co‐IP assays showing the
   interaction between TH and Ub by using Ub as the capture antibody in
   the SN of the three groups. n = 3 per group. H–J) Representative blots
   and quantification of co‐IP assays showing the interaction between TH
   and Ub by using TH as the capture antibody in the SN of the three
   groups. n = 3 per group. K,L) Immunofluorescent staining and
   quantification of Ub within TH positive neurons. n = 6 per group. White
   arrows show the immunostaining of Ub in TH‐positive neurons. Scale
   bars, 50 and 10 µm for the original and magnification of images in (K).
   Results are expressed as the mean ± SEM. ^** p < 0.01 vs Control. ^## p
   < 0.01 vs MPTP group. Statistical significance was determined by
   one‐way ANOVA and Tukey tests for post‐hoc comparisons.

2.6. RVG‐nDMC Administration Reduces Oxidative Stress

   Pathological TH is a potential source and contributor of ROS
   generation, and ROS has been widely reported to induce DA neuron
   death.^[ [159]^13 ^] Therefore, we evaluated the potential effects of
   RVG‐nDMC on nigral metabolism in PD mice by performing GCTOF/MS
   (Figure [160] 8A; Figure S10A,B, Supporting Information). As shown in
   Figure [161]8B, a distinct separation of metabolites among the three
   groups of mice was observed in 2D boxplots and PCA score plots,
   indicating significant differences in the metabolic profiles. Z‐score
   plots showed relative variations of each individual metabolite across
   all groups, which is displayed in the form of a heatmap
   (Figure [162]8C). Next, metabolic pathway enrichment analysis (MPEA)
   was carried out to reveal potential pathways involved in the
   differential metabolites among these three groups (Figure [163]8D). Our
   results demonstrate that RVG‐nDMC improves glutathione metabolism by
   increasing Pyroglutamic acid and Ornithine (Figure S11A–C, Supporting
   Information). The polyamines putrescine, spermidine, and spermine are
   derived from ornithine under the catalyzation of ornithine
   decarboxylase.^[ [164]^37 ^] Because spermidine and spermine
   experimentally enhance longevity via autophagy induction,^[ [165]^38 ^]
   ample evidence supports their neuroprotective effects in PD. For
   example, spermidine protects PD from 6‐OHDA toxicity through its
   antioxidant and antiinflammatory properties, and it alleviates
   α‐synuclein neurotoxicity by inducing autophagy.^[ [166]^39 ^] It would
   therefore be of interest in the future to examine whether DMC exerts
   neuroprotective effects through the polyamine pathway.

Figure 8.

   Figure 8
   [167]Open in a new tab

   Metabolomic analysis of RVG‐nDMC treatment in the SN of PD mice. A) The
   relative abundances of representative metabolites in each group. B)
   Overall metabolic profiles of samples from the Control, MPTP, and MPTP
   + RVG‐nDMC groups using PCA plots. C) Z‐score plots showing the
   relative variations of each individual metabolite across all groups in
   the form of a heatmap. D) MPEA of potential pathways involved in the
   differential metabolite content between the indicated groups. E)
   Representative metabolites among the three groups. n = 6 per group. F)
   Schematic diagram showing RVG‐nDMC‐changed metabolites in the glucose
   metabolism pathway that affect oxidative stress in PD mice. Results are
   expressed as the mean ± SEM. ^** p < 0.01. Statistical significance was
   determined by one‐way ANOVA and Tukey tests for post‐hoc comparisons.

   RVG‐nDMC also reversed MPTP‐decreased d‐Glucose, Fructose, Glucose
   6‐phosphate, and Fructose 6‐phosphate, which are enriched in glucose
   metabolism (Figure [168]8C,[169]E). Previous studies have revealed an
   antioxidant‐like role of glucose, while glucose uptake inhibition has
   been shown to induce ROS accumulation.^[ [170]^40 ^] Thus, glucose
   metabolism may play an important role in regulating the balance of ROS
   production and scavenging in response to RVG‐nDMC (Figure [171]8F). On
   the one hand, glucose oxidation, through the pentose phosphate pathway
   (PPP), produces reducing equivalents in the form of nicotinamide
   adenine dinucleotide phosphate (NADPH), which are necessary to recycle
   antioxidant glutathione from its oxidized status.^[ [172]^41 ^] On the
   other hand, neuronal serine derived from glucose
   6‐phosphate/3‐phosphoglycerate (3PG) metabolism is the substrate for
   NADPH and glutathione (GSH) synthesis.^[ [173]^42 ^]

   Pyruvate derived from glucose metabolism mitigates oxidative stress,
   including ROS generation, by sustaining the citric acid cycle (CAC,
   also called TCA) cycle.^[ [174]^43 ^] Therefore, we postulated that
   RVG‐nDMC may cause changes in mitochondrial metabolism. TEM results
   suggest that RVG‐nDMC reversed the MPTP‐decreased number of
   mitochondria and ameliorated mitochondrial morphological changes caused
   by MPTP, such as mitochondrial swelling and mitochondrial cristae
   disappearance (Figure [175] 9A, red arrows; quantified in
   Figure [176]9B). Furthermore, RVG‐nDMC restored the MPTP‐decreased
   expression of the mitochondrial‐associated antiapoptotic protein Bcl‐2,
   though no obvious effects on the pro‐apoptotic protein Bax were
   observed (Figure [177]9C,[178]D). As further validation, we evaluated
   antioxidant‐like effects of RVG‐nDMC in MPP^+‐treated DA‐derived MN9D
   cells. Mito‐Sox appeared oxidized in MPP^+ treated cells, while
   RVG‐nDMC and N‐acetyl‐l‐cysteine (NAC) treatment significantly
   decreased oxidized Mito‐Sox and improved the mitochondrial damage
   (Figure [179]9E,[180]G). Consequently, RVG‐nDMC and NAC reversed the
   MPP^+‐induced ROS generation (Figure [181]9F,[182]H). Notably, RVG‐nDMC
   and NAC also increased the superoxide dismutase (SOD) activity, GSH
   concentration and GSH/glutathione disulfide (GSSG) ratio
   (Figure [183]9I–[184]K). Additionally, RVG‐nDMC, but not NAC, restored
   the reduced state of NADPH and its oxidized state (NADP^+) ratio
   (Figure [185]9L). These results suggest that RVG‐nDMC may prevent DA
   loss by reducing oxidative stress.

Figure 9.

   Figure 9
   [186]Open in a new tab

   RVG‐nDMC attenuates oxidative stress in in vivo and in vitro PD models.
   A) Ultrastructural analysis of mitochondria in the SN of Control, MPTP,
   and MPTP + RVG‐nDMC groups. Red arrows indicate damaged mitochondria in
   the MPTP group. Scale bars, left, 2 µm; right, 500 nm. B)
   Quantification of the number of mitochondria in the three groups. n =
   10 per group. C,D) Representative blots and quantification showing the
   Bcl‐2 and Bax expression levels in the SN for the three groups. n = 6
   per group. E) Mito‐Sox was used to stain live cells upon RVG‐nDMC and
   NAC treatment in control or MPP^+‐treated cells. Scale bars, upper, 50
   µm; lower, 10 µm. F) Representative graphs of ROS generation upon
   RVG‐nDMC or NAC treatment in control or MPP^+‐treated cells. Scale
   bars, 50 µm. G,H) Quantification of the relative Mito‐Sox and ROS
   fluorescence intensity. n = 6 per group. I,J) The content of SOD and
   GSH upon RVG‐nDMC or NAC treatment in control or MPP^+‐treated cells. n
   = 3 per group. K,L) Relative GSH/GSSG and NADPH/NADP^+ ratio after
   RVG‐nDMC or NAC treatment in control or MPP^+‐treated cells. n = 3 per
   group. Results are expressed as the mean ± SEM. ^** p < 0.01 vs
   Control. ^## p < 0.01, ^# p < 0.05 vs MPTP or MPP^+ group. Statistical
   significance was determined by one‐way ANOVA and Tukey tests for
   post‐hoc comparisons.

   Previously, DMC was revealed to exert neuroprotection by inducing
   autophagy in yeast and mice.^[ [187]^16 ^] Consequently, we evaluated
   the effects of RVG‐nDMC on autophagic indicators in PD mice. RVG‐nDMC
   slightly increased the number of autophagosomes in the SN of PD mice,
   though the differences were not statistically significant (Figure
   S12A,B, Supporting Information). Furthermore, the RVG‐nDMC group had
   increased LC3 II/I expression as compared with the MPTP group (Figure
   S12C,D, Supporting Information), suggesting that RVG‐nDMC might also
   induce autophagy in the subacute PD model mice, though the link between
   the neuroprotection of RVG‐nDMC in PD model mice and its regulation of
   autophagy is unclear.

2.7. RVG‐nDMC Administration Downregulates the Inflammatory Reaction

   Because RVG‐nDMC can also be internalized by microglia (Figure [188]3I;
   Figures S4B and 5B, Supporting Information); and microglial‐mediated
   neuroinflammation underlies the pathogenesis of PD,^[ [189]^44 ^] we
   further examined the effects of RVG‐nDMC on the inflammatory cytokine
   expression profile. nDMC and RVG‐nDMC decreased mRNA levels of
   pro‐inflammatory cytokines, including IL‐1β and TNF‐α in the SN, and
   TNF‐α in the striatum of PD mice (Figure S13A,B,E, Supporting
   Information), suggesting general antiinflammatory properties of DMC.
   However, RVG‐nDMC showed no obvious effects on IL‐1β mRNA levels in the
   striatum or IL‐6 mRNA expression in the SN and striatum of PD mice
   (Figure S13C,D,F, Supporting Information). Because these cytokines are
   secreted proteins, we also examine their protein levels by ELISA.
   RVG‐nDMC significantly decreased the protein levels of IL‐1β, TNF‐α,
   and IL‐6 in the SN, as well as IL‐1β and TNF‐α in the striatum of PD
   mice (Figure S14A–E, Supporting Information), which is consistent with
   the mRNA results. Moreover, in multianalyte suspension arrays, RVG‐nDMC
   increased the expression of antiinflammatory cytokines, including IL‐4
   and IL‐10, in the serum of PD mice (Figure S15C,G, Supporting
   Information). These results are consistent with an additional role for
   RVG‐nDMC in protecting against PD by downregulating the inflammatory
   reaction.

3. Conclusion

   In this study, an RVG‐nDMC “Trojan horse” delivery system was
   constructed for brain‐targeted delivery of the antiaging drug DMC for
   PD intervention. RVG‐nDMC penetrated the BBB for enhanced delivery of
   DMC to the DA neurons and microglia in the SNpc in PD mice and
   ameliorated motor deficits and nigral DA neuron death without
   exhibiting adverse effects in the major organs after systemic
   administration. Moreover, RVG‐nDMC reversed TH ubiquitination and
   degradation, as well as oxidative stress, and exerted antiinflammatory
   effects in PD mice. Taken together, these findings identify RVG‐nDMC as
   a potentially powerful and clinically feasible approach for
   parkinsonian neuroprotection.

4. Experimental Section

Materials

   DMC was purchased from Sigma‐Aldrich (St. Louis, MO, USA).
   Poly(ethylene glycol) methyl ether‐block‐poly(l‐lactide‐co‐glycolide)
   (PLGA20K‐b‐mPEG2K) and PLGA20K‐PEG5K‐MAL were purchased from Xi'an
   ruixi Biological Technology Co. Ltd. (Xi'an, China). Cy5.5 was
   purchased from Beijing Solarbio science & technology co. Ltd. (Beijing,
   China). MPP^+ and MPTP were purchased from Sigma‐Aldrich (St. Louis,
   MO, USA). Anti‐TH (F‐11, sc‐25269), dopamine transporter (DAT,
   sc‐32258), and PDH‐E1α (sc‐377092) antibodies were purchased from Santa
   Cruz Biotechnology (Dallas, TX, USA). Anti‐synapsin І (#5297),
   Iba1(#17198), synaptotagmin (#14558), syntaxin (#18572), PSD‐95
   (#3450), and Bax (#14796) antibodies were purchased from Cell Signaling
   Technology (Danvers, MA, USA). Bcl‐2 (16396‐1‐AP), α‐Tubulin
   (11224‐1‐AP), and GAPDH (60004‐1) antibodies were purchased from
   Proteintech Group (Rosemont, IL, USA). Antimicrotubule‐associated
   protein‐1 light chain 3 (LC3) II/І (ab48394) and p62 (ab5416)
   antibodies were purchased from Abcam (Cambridge, MA, USA). TH
   (Phospho‐Ser19) (E1A0037A), TH (Phospho‐Ser31) (E1A0038A), and TH
   (Phospho‐Ser40) (E1A0039A) antibodies were purchased from Enogene
   (Nanjing, China). DyLight 488 goat antimouse IgG (H+L) (70‐GAM4882) and
   DyLight 594 goat antirabbit IgG (H+L) (70‐GAR5942) antibodies were
   purchased from Multi Sciences (Hangzhou, China). Horseradish peroxidase
   (HRP)‐labeled goat antirabbit IgG, HRP‐labeled goat antimouse IgG and
   Mito‐tracker were purchased from Beyotime Biotechnology (Shanghai,
   China). MitoSOX Red Mitochondrial Superoxide Indicator was purchased
   from Thermo Fisher Scientific (Waltham, MA, USA).

Preparation of RVG‐nDMC

   A mixture of DMC (4 mg), PLGA20K‐b‐mPEG2K (18 mg), and
   PLGA20K‐PEG5K‐MAL (2 mg) in CHCl[3] (1 mL) was added to ethanol
   solution (4%) under sonication for 5 min. The ratio of PLGA20K‐b‐mPEG2K
   and PLGA20K‐b‐PEG5K‐MAL was optimized to balance the stability with the
   targeting ability of RVG‐nDMC, which was also done in a previous
   report.^[ [190]^17 ^] nDMC was obtained by evaporating the solution at
   30 °C to remove the CHCl[3]. To prepare RVG‐nDMC, YC‐30 peptide was
   conjugated to nDMC via a maleimide–thiol reaction. Briefly, nDMC was
   mixed with an excess amount of YC‐30 peptide and incubated in PBS for
   12 h at room temperature (RT). Subsequently, the solution was dialyzed
   against distilled water to remove the unconjugated YC‐30.

Characterizations of nDMC and RVG‐nDMC

   The size distribution and ζ‐potential of nDMC and RVG‐nDMC were
   measured on a Zetasizer Nano ZS (Malvern, U.K.). The morphology of nDMC
   and RVG‐nDMC was observed by transmission electron microscopy
   (JEM‐1400PLUS, Japan). The absorption spectra were obtained by UV/vis
   spectrometry (Lambda 35, Perkin‐Elmer, USA). The fluorescence spectra
   were analyzed by fluorescence spectroscopy (LS55 luminescence
   spectrometer (Perkin‐Elmer).

Drug Release Analysis

   The drug release kinetics of nDMC and RVG‐nDMC were measured using a
   dialysis method under physiological conditions. In brief, 1 mL nDMC and
   RVG‐nDMC (containing 2 mg mL^−1 DMC) were suspended in PBS (1 mL, pH
   7.4) in a dialysis bag (molecular weight cutoff, 100 kDa). Then, the
   solutions were dialyzed against PBS (30 mL, pH 7.4) at 37 °C. At
   different time points, the absorbance of the DMC was measured by UV–vis
   spectrometry, and the drug release kinetics were calculated by using a
   standard curve of DMC in PBS with the absorption at 350 nm.

Drug Loading and Encapsulation Efficiency

   Freeze‐dried nDMC and RVG‐nDMC were dissolved in CHCl[3]. Subsequently,
   the solution was rotary evaporated to remove CHCl[3]. The DMC was
   dissolved in distilled water and measured by UV/vis spectrometry. The
   loading efficiency and encapsulation efficiency were calculated as
   follows:

   LE = weight of DMC in nDMC or RVG‐nDMC/weight of nDMC or RVG‐nDMC ×
   100%

   EE = weight of DMC in nDMC or RVG‐nDMC/weight of total DMC × 100%

Animals

   Eight‐week‐old C57BL/6 mice, weight about (24 ± 2) g, were purchased
   from SPF Biotechnology Co., Ltd. (Beijing, China). The mice were housed
   in the experimental animal center of Guangzhou Medical University with
   a 12 h dark/light cycle at ambient temperature (22 ± 1) °C and relative
   humidity (60 ± 5)%. The experimental methods used were in compliance
   with the Institutional Animal Care and Use Committee of Guangzhou
   Medical University (Approval number: GY2020‐041) and National Institute
   of Health guidelines on the care and use of animals (NIH Publications
   No. 8023, revised 1978).

PD Mouse Model Induction and Drug Treatment

   The MPTP‐induced PD mouse model was established according to the
   previous reports.^[ [191]^45 ^] MPTP was administrated
   intraperitoneally continuously over 5 days to induce the subacute PD
   model. Simultaneously, nDMC, RVG‐nDMC (equivalent dose of DMC), and
   unloaded RVG vehicle were administered intravenously starting on the
   day of the first MPTP injection once every other day for 12 days. After
   3 days, behavioral tests were carried out.

Assessment of Brain Targeting Delivery

   Mice were intravenously injected with nDMC or RVG‐nDMC (200 µL,
   containing 3 mg mL^−1 DMC and 20 µg mL^−1 Cy5.5). Afterward, the real
   time fluorescence was quantitatively analyzed by using the IVIS imaging
   system (Excitation wavelength: 674 nm, Emission wavelength: 692 nm).
   After 48 h, the major organs (heart, liver, spleen, lung, kidneys,
   brain) were sacrificed and analyzed by using the IVIS imaging system.
   Simultaneously, photoacoustic (PA) images of Cy5.5 were obtained and
   quantitatively analyzed on a preclinical photoacoustic computerized
   tomography scanner (Vevo LAZR‐X, Canada) at specified time intervals.
   Cy5.5 was used to label RVG‐nDMC for in vitro and in vivo
   pharmacokinetics analysis.

Open Field Test

   Mice were placed in the bottom center of a box and filmed by
   EthoVisione XT software (Beijing, China) for 15 min. The movement
   speed, total travelled distance, and time spent in the central area
   were recorded.

Rotarod Test

   Mice were trained on the Rotarod (Ugo basile SRL, Gemonio, VA, Italy)
   at a speed of 10 rpm for 3 days. During the test, the speed of the
   Rotarod accelerated from 4 rpm uniformly to 40 rpm within 5 min, and
   the time to falling was recorded.

Pole‐Climbing Test

   Mice were placed head up on the top of a pole with a length of 75 cm
   and diameter of 9 mm. The total time that it took the mice to climb to
   the ground was recorded.

Grasping Test

   Two front paws of mice were suspended on a horizontal wire (diameter l
   mm, 30 cm from the ground) for 10 s, and their abilities to grasp a
   wire with their hind legs was measured. The grasping score was as
   follows: The mouse grasped the wire with both hind legs, 3; the mouse
   grasped the wire with only one hind limb, 2; the mouse could not grasp
   the wires with either hind limb, 1; the mouse dropped, 0. Results were
   averaged from three measurements.

MN9D Cell Culture

   MN9D cells were purchased from American Type Culture Collection (ATCC,
   Manassas, VA, USA) and were cultured in Dulbecco's Modified Eagle
   Medium (GIBCO, Carlsbad, CA, USA) containing 8% fetal bovine serum
   (GIBCO, Carlsbad, CA, USA), 2 U mL^−1 penicillin (Beyotime
   Biotechnology), and 2 mg mL^−1 streptomycin (Beyotime Biotechnology) in
   an incubator at 37 °C and 5% CO[2].

Cell Viability Assay

   A MN9D cell suspension (≈1 × 10^4 cells) was added to 96‐well plates.
   After treatment with DMC, nDMC, or RVG‐nDMC with or without 1000 ×
   10^−6 m MPP^+ for 48 h, CCK‐8 solution was added to each well, and the
   plates were incubated for 1.5 h. The absorbance at 450 nm was measured
   with a microplate reader (PerkinElmer). Data were obtained from three
   separate experiments, each of which were performed in triplicate.

Mitochondrial Superoxide Imaging of Live Cells

   MitoSOX (Thermo Fisher Scientific) and Hoechst 33342 Staining (Beyotime
   Biotechnology) working solutions were added to MN9D cells. After
   incubation for 10 min at 37 °C, the cells were washed three times, and
   the images were scanned under a confocal laser‐scanning microscope
   (SP8; Leica).

ROS Detection Assay

   2,7‐dichlorofluorescein diacetate (DCFH‐DA) (Beyotime Biotechnology)
   and Hoechst 33342 Staining (Beyotime Biotechnology) working solutions
   were added to MN9D cells. After incubation for 20 min at 37 °C, the
   cells were washed three times, and images were scanned under a confocal
   laser‐scanning microscope (SP8; Leica).

SOD Detection Assay

   A Cu/Zn‐SOD Assay Kit with WST‐8 (Beyotime Institute of Biotechnology,
   Shanghai, China) was used to determine intracellular SOD activity.
   Briefly, MN9D cells were homogenized and incubated with WST‐8/enzyme
   working solution at 37 °C for 30 min. The absorbance was measured at a
   wavelength of 450 nm with a microplate reader (PerkinElmer). The
   protein concentrations were determined using BCA assay. The results
   were expressed as units per mg protein (U mg^−1 protein).

GSH and GSSG Detection Assay

   GSH and GSSG detection was performed using the GSH/GSSG kit (S0053,
   Beyotime Institute of Biotechnology, Shanghai, China) according to the
   manufacturer's instructions. Cells were collected and Protein Removal
   solution was added. The samples were then freeze‐thawed twice rapidly
   in liquid nitrogen and 37 °C water alternatively. After centrifugation
   at 10 000 × g for 10 min, the supernatant was used for the
   determination of total glutathione. GSH scavenging reagent working
   solution was added to degrade GSH. Then total glutathione working
   solution and NADPH solution were added to the supernatant to detect the
   concentration of total glutathione and GSSG. The GSH concentration was
   obtained from the total glutathione concentration minus the double GSSG
   concentration, and the GSH/GSSG ratio was determined. The absorbance at
   412 nm was measured with a microplate reader (PerkinElmer). Data were
   obtained from three separate experiments, each of which were performed
   in triplicate.

NADPH/NADP^+ Ratio Determination

   The MN9D cellular NADPH/NADP^+ ratio was measured using the
   NADPH/NADP^+ assay kit (S0179, Beyotime Institute of Biotechnology,
   Shanghai, China) according to the manufacturer's instructions. Briefly,
   the samples were extracted from MN9D cells with extraction buffer and
   deproteinized with a spin column. Next, the extracted samples were
   resuspended in the NADP^+/NADPH extraction buffer included in the
   NADP^+/NADPH quantification kit. A series of standards were also
   prepared and measured colorimetrically, and the resulting standard
   curve was then used to calculate the ratio of NADPH/NADP^+ in the
   samples. The absorbance at 450 nm was measured with a microplate reader
   (PerkinElmer). Data were obtained from three separate experiments, each
   of which were performed in triplicate.

RNA Sequencing Analysis

   RNA sequencing was performed according to the previous report.^[
   [192]^46 ^] Briefly, total RNA was extracted from nigral samples using
   Trizol (Life Technologies, Carlsbad, CA, USA). Next, RNA libraries were
   prepared using the NEBNext UltraTM RNA Library Prep Kit for Illumina
   (NEB, USA) according to the manufacturer's instructions. The qualities
   of purified libraries were assessed on the Agilent Bioanalyzer 2100
   system. Clustering of the index‐coded samples was performed on a cBot
   Cluster Generation System using the TruSeq PE Cluster Kit v3‐cBot‐HS
   (Illumia), according to the manufacturer's instructions. After cluster
   generation, the samples were sequenced on an Illumina HiSeq 2500
   platform (San Diego, CA). HTSeq v0.6.0 was used to count the read
   numbers mapped to each gene, and fragments per kilobase of
   transcript‐per‐million mapped reads (FPKM) of each gene were calculated
   based on the length of the gene and read counts mapped to the analyzed
   gene. DEG analysis was performed using the DESeq2 R package (1.10.1).
   The resulting p‐values were adjusted using the Benjamini‐Hochberg
   approach for controlling false discovery rates. The functional
   enrichment of DEGs was analyzed by the KEGG database.

Metabolomics Analysis of Nigral Samples

   Nigral tissues were overlaid with 80% methanol solution containing 0.1%
   formic acid (FA) and vortexed. After centrifugation at 15 000 rpm, 4 °C
   for 10 min, the supernatant was collected for liquid
   chromatography‐tandem mass spectrometry (LC‐MS/MS) analysis. LC‐MS/MS
   analysis was performed using a Vanquish UHPLC system (Thermo Fisher)
   coupled with an Orbitrap Q Exactive series mass spectrometer (Thermo
   Fisher) as previously described.^[ [193]^47 ^] Samples were injected
   into a Hyperil Gold column (100 × 2.1 mm, 1.9 µm) using a 16 min linear
   gradient at a flow rate of 0.2 mL min^−1. The Q Exactive series mass
   spectrometer was operated in positive/negative polarity mode with a
   spray voltage of 3.2 kV, capillary temperature of 320 °C, sheath gas
   flow rate of 35 arb, and aux gas flowrate of 10 arb. The metabolites
   data analysis were performed using the KEGG
   ([194]http://www.genome.jp/kegg/), HMDB ([195]http://www.hmdb.ca/) and
   Lipidmaps ([196]http://www.lipidmaps.org/) databases, as previously
   described.^[ [197]^47 ^]

TEM

   The ultrastructural morphology of mitochondria was analyzed by TEM as
   described previously.^[ [198]^48 ^] Briefly, fixed tissues were
   dehydrated with an increasing gradient of alcohol and acetone, and then
   the tissues were embedded with 812 embedding agent (SPI‐Pon 812 Epoxy
   Resin Monomer; SPI, Shanxi, China). The sliced sections were
   double‐stained with uranium lead and dried overnight at RT, and the
   photographs were imaged and analyzed by transmission electron
   microscopy (HT7700; Hitachi, Tokyo, Japan).

Quantitative Analysis of Neurotransmitters

   The nigral levels of DA, DOPAC, 5‐HT, and 5‐HIAA were determined using
   HPLC‐MS/MS. Nigral tissues were homogenized with methanol containing
   0.1% FA, and supernatants were obtained after centrifugation at 14 000
   rpm for 10 min. Then the samples were subjected to HPLC‐MS/MS analysis.
   HPLC‐MS/MS analysis was performed using UltiMate 3000 RS (Thermo
   Fisher) coupled with Q Exactive (Thermo Fisher). The samples were
   injected into a Waters T3 column (150 × 2.1 mm, 3 µm) at a flow rate of
   0.3 mL min^−1. The eluent for the aqueous phase mode was 0.1% FA in
   water; and the eluent for the organic phase was 0.1% ethyl formate. The
   Q Exactive series mass spectrometer was operated in positive/negative
   polarity mode with a spray voltage of 3.2 kV, a capillary temperature
   of 300 °C, and a sheath gas flow rate of 40 arb. The chromatogram
   acquisition and integration of each analyte were processed by Xcalibur
   4.0 software (Thermo Fisher). The peak area of each analyte was taken
   as the ordinate and the concentration as the abscisic coordinate, and a
   standard curve was obtained by regression with the weighted
   coefficient. The sample concentration was calculated from the standard
   curve.

Western Blot Assay

   Substantia nigra tissue was collected, and total protein was extracted
   with RIPA Lysis Buffer (Beyotime Biotechnology) and quantified using
   the BCA Protein Assay Kit (Beyotime Biotechnology). The samples were
   separated by SDS‐PAGE and transferred to PVDF membranes. Then the
   membranes were blocked with 5% BSA and incubated with primary antibody.
   After incubation with secondary antibody, chemiluminescence was
   visualized on the GeneGnome XRQ Chemiluminescence imaging system (Gene
   Company, Hong Kong, China). Image J software was used to analyze the
   optical density of bands.

Co‐Immunoprecipitation

   Nigral tissues were lysed by IP lysis buffer, and then the lysates were
   centrifuged at 12 000 rpm at 4 °C for 10 min, and supernatant was
   collected. TH or Ub antibody was added to the supernatant, with IgG
   antibody as a negative control. After incubation at 4 °C overnight,
   HRP‐conjugated Protein A Sepharose beads were added to the supernatant,
   and the samples were incubated at 4 °C for 4 h and then boiled and
   subjected to western blot assay.

Immunohistochemistry and Immunofluorescence Assays

   Embedded mouse brains were cut into 15 µm sections with a freezing
   microtome (Leica), and the slices were incubated with corresponding
   primary antibodies overnight at 4 °C. For the immunohistochemistry
   assay, brain slices were incubated with a secondary antibody labeled
   with biotin and stained with diaminobenzidine (DAB). Images were
   scanned under a microscope (Leica CS2, Hamburg, Germany). For the
   immunofluorescent assay, brain slices were incubated with
   fluorescent‐labeled secondary antibody, and DAPI was used to stain
   nuclei. Images were scanned under a confocal laser‐scanning microscope
   (SP8; Leica). Quantitative analysis was performed using the Image‐Pro
   Plus 6.0 photogram analysis system (IPP 6.0, Media Cybernetics,
   Bethesda, MD, USA).

Quantitative RT‐PCR

   Total RNA from tissues was extracted in Trizol (Invitrogen, San Diego,
   CA, USA). The genomic DNA was removed by using DNA Eraser Buffer
   (Takara, Otsu, Japan). RNA was reverse transcribed into cDNA, and qPCR
   was performed with TB Green Premix Ex Taq (Takara). The thermal
   parameters were as follows: 95 °C for 5 s, 55 °C for 30 s, and 72 °C
   for 30 s by 40 cycles. The primer sequences were as follows: IL‐1β
   forward 5’‐ AATGCCACCTTTTGACAGTGAT‐3’ and reverse 5’‐
   TGCTGCGAGATTTGAAGCTG‐3’; TNF‐α forward 5’‐ CACGTCGTAGCAAACCACC‐3’ and
   reverse 5’‐ TGAGATCCATGCCGTTGGC‐3’; IL‐6 forward
   5’‐AGGATACCACTCCCAACAGACC‐3’ and reverse 5’‐AAGTGCATCATCGTTCATACA‐3’;
   GAPDH forward 5’‐ACGGGAAGCTCACTGGCATGGCCTT‐3’ and reverse
   5’‐CATGAGGTCCACCACCCTGTTGCTG‐3’.

Enzyme‐Linked Immunosorbent Assay (ELISA)

   Mouse nigral and striatal IL‐1β, TNF‐α, and IL‐6 concentrations were
   measured using ELISA kits (Shanghai Enzyme‐linked Biotechnology Co.,
   Ltd., Shanghai, China) as described in a previous study.^[ [199]^47 ^]
   Briefly, nigral and striatal samples were homogenized in ice‐cold PBS
   containing protease inhibitors, and the homogenates were then
   centrifuged to obtain the supernatants. The resulting samples were
   incubated with the indicated antibody‐coated plates and horseradish
   peroxidase (HRP)‐labeled secondary antibody at 37 °C for 1 h. After
   incubation with chromogenic agents A and B at 37 °C for 15 min, the OD
   values were measured by Multiscan Spectrum (PerkinElmer, MA, USA) at
   450 nm. The protein concentrations were determined using BCA assay. The
   results were expressed as pg per mg protein (pg mg^−1 protein).

Multianalyte Suspension Arrays Application

   Each sample in 96‐well microtiter plates was incubated with bead
   mixture (100 microspheres with type‐specific probes in 25 µL) for 2 h
   at RT, and subsequently with denatured biotin‐labeled detective
   antibody complex (50 µL) for 1 h at RT. After three washes, the samples
   with microspheres were incubated with streptavidin‐labelled
   phycoerythrin (50 µL) for 30 min at RT in the dark. Finally, the
   reaction product was detected by a Luminex 100 analyzer instrument
   (Luminex Corporation, Austin, USA). Each reaction was repeated in
   triplicate.

Statistical Analysis

   Data are presented as mean ± standard error of the mean (SEM). For
   behavioral experiments, 9–10 per group were used for measurement. For
   in vitro immunofluorescence experiments, 6 per group were used for
   measurement. For in vivo immunofluorescence, immunohistochemistry,
   metabolomic analysis, and western blot experiments, 6 per group were
   used for measurement. For ultrastructural analysis using TEM, 10 per
   group were used for measurement. Data were analyzed using the unpaired
   Student's t‐test or one‐way ANOVA followed by a Tukey's post‐hoc test,
   where appropriate. The Hotelling T2 test was used to evaluate outliers
   in the metabolomic analysis. Differences with a p‐value < 0.05 were
   considered statistically significant. All statistical analyses were
   performed using GraphPad Prism 8.0 (GraphPad Software, La Jolla, CA).
   Representation of the p‐value was ^* p < 0.05, ^** p < 0.01.

Conflict of Interest

   The authors declare no conflict of interest.

Supporting information

   Supporting Information
   [200]Click here for additional data file.^ (3.6MB, pdf)

Acknowledgements