Human tissue was obtained from the University of Miami Brain Endowment Bank (NIH, Bethesda, MD, United States). Brain sections were paraffin embedded and supplied at 5-μm thickness. For immunohistochemistry, we studied samples from five PD patients and five age-matched controls (Table 1). The average age and postmortem intervals for PD patients and controls were not significantly different (two-tailed Student’s t test. Age: p = 0.3713. Postmortem intervals: p = 0.2012).

Eight-to 12-week-old male C57BL/6 mice weighing 22–28 g were purchased from the Beijing Vital River Laboratory Animal Technological Company (Beijing, China). All mice were housed in specific pathogen-free facilities, maintained on a 12-h light/dark cycle with controlled temperature and air humidity, and allowed free access to food and water. All experimental procedures were approved by the Institutional Animal Care and Use Committee (IACUC), Sun Yat-Sen University, Guangzhou, China. Mice were assigned randomly for treatment.

rAAV2/9-TH-Tfe3-3×Flag-WPRE (AAV-TFE3) contained the murine Tfe3 cDNA fused to that of the 3×Flag epitope, and rAAV2/9-TH-3×Flag-WPRE (AAV-Flag) containing only the cDNA of 3×Flag epitope was used as a control. rAAV2/9-hSyn-EGFP-miR30a-shTfe3-WPRE (AAV-shTFE3) was used to target murine Tfe3 gene and rAAV2/9-hSyn-EGFP-miR30a-shScramble-WPREs (AAV-shScr) was used as a control. The Tfe3 shRNA sequence was 5′-GCG​ACA​GAA​GAA​AGA​CAA​TCA-3’. The scrambled shRNA sequence was 5′-CCT​AAG​GTT​AAG​TCG​CCC​TCG-3′, non-target in mice. All the viruses were generated and packaged by BrainVTA (Wuhan, China) and PackGene Biotech (Guangzhou, China). The titer of all the viruses in this study was 1x10¹² gc/mL. The AAV viruses were delivered at 100 nl/min unilaterally into the right SN. The coordinates indicating distance (mm) from the bregma were as follows: anteroposterior −2.9, mediolateral +1.3, and dorsoventral −4.35. After the injection, the needle remained in place for at least 5 min to prevent retrograde flow along the needle track. After surgery, the mice were warmed under a heating pad until they awakened.

MPTP (Sigma, Shanghai, China) treatments were performed as previously described (Li et al., 2020). Briefly, ten-to 12-week-old male mice were intraperitoneally injected (i.p.) with MPTP (dissolved in saline, 30 mg/kg body weight) or saline for 5 days at 24 h intervals. The mice were intraperitoneally injected with MPTP 1 week after the stereotaxic administration of the AAV virus. For immunofluorescence and immunohistochemistry, the mice were sacrificed 1 day and 21 days after the last MPTP injection, respectively.

For real-time PCR, mice were sacrificed by cervical dislocation and the brains were rapidly removed and washed with ice-cold PBS. The SN tissue was rapidly dissected on ice and stored at −80°C prior to further experiments. For immunofluorescence and immunohistochemistry, mice were anesthetized with chloral hydrate (400 mg/kg, i. p.) and then intracardially perfused with ice-cold PBS for 3 min, followed by cold 4% PFA at a flow rate of 10 ml/min for 8 min. Mouse brains were removed and post-fixed overnight in 4% PFA at 4°C and then immersed in 20% and 30% sucrose. Tissues were embedded in embedding media O.C.T (#4583, SAKURA) and cut into 20 μm-thick cryosections for immunofluorescence and 40 μm-thick cryosections for immunohistochemistry.

Total RNA was extracted from the SN tissue using TRIzol reagent (Thermo Fisher, United States), and the RNA concentration was determined spectrophotometrically (Beckman DU 730). NovoScript^® Plus All-in-one 1st Strand cDNA Synthesis SuperMix (gDNA Purge) (E047-01B, novoprotein, China) was used to synthesize the cDNA, which was then amplified using a NovoStart^® SYBR qPCR SuperMix Plus (E096-01A, novoprotein, China) with specific primers for real-time PCR analysis. All reactions were carried out using the Light Cycler 480 System (Roche, Switzerland). The following primer sequences were used: Tfe3, forward: 5′- ATC​TCT​GTG​ATT​GGC​GTG​TCT-3′, reverse: 5′-GAA​CCT​TGA​GTA​CCT​CCC​TGG- 3'; Map1lc3b, forward: 5′- CGC​TTG​CAG​CTC​AAT​GCT​AAC-3′, reverse: 5′- CTC​GTA​CAC​TTC​GGA​GAT​GGG-3′; Lamp1, forward: 5′-TGC​TCC​GGG​ATG​CCA​CTA​T-3′, reverse: 5′- TGT​TGT​CCT​TTT​TCA​GGT​AGG​TG-3′; Ctsd, forward: 5′- CGA​TTA​TCA​GAA​TCC​CTC​TGC​G-3′, reverse: 5′- GGT​CTT​AGG​CGA​TGA​CTG​CAT-3′; Sqstm1/p62, forward: 5′- GAA​CTC​GCT​ATA​AGT​GCA​GTG​T-3′, reverse: 5′- AGA​GAA​GCT​ATC​AGA​GAG​GTG​G-3'; Actin, forward: 5′-GGC​TGT​ATT​CCC​CTC​CA TCG-3′, reverse: 5′-CCA​GTT​GGT​AAC​AAT​GCC​ATG​T-3'.

Immunofluorescence analysis was performed as previously described (Li et al., 2020). Briefly, free-floating sections were pre-incubated in blocking solution containing 5% normal donkey serum at room temperature for 1 h. Then primary antibodies were dissolved in diluent and incubated with sections overnight at 4°C. The following primary antibodies were used: TH (1:1,000, #AB9702, Millipore), TFE3 (1:500, #ab93808, Abcam), LC3 (1:100, #2775, Cell signaling Technology), Lamp1 (1:1,000, #1D4B-C, DSHB), CatD (1:200, #SC-6486, Santa Cruz Biotechnology), and p62 (1:1,000, #GP62-C, Progen Biotechnik). After three washes, the sections were then incubated with the secondary antibodies (Thermo Fisher), which were conjugated with Alexa 488, Alexa 555, or Alexa 647, at room temperature for 1 h. Finally, the sections were visualized under a confocal laser scanning microscope (LSM 780, Carl Zeiss) and the immunofluorescent images were quantified using ImageJ software.

For human samples, paraffin-embedded human brain tissue slices were deparaffinized in xylene and rehydrated in graded alcohols. Antigens were retrieved by microwave heating with citrate buffer (pH 6) for a total of 30 min. Then, tissue sections were incubated in 3% H₂O₂ at room temperature for 30 min to block nonspecific peroxidase-activity and then blocked in 2.5% normal horse blocking serum for 1 h at room temperature. Tissue sections were first incubated with Iba1 (1:1,000, #ab5076, Abcam) overnight at 4°C, washed, incubated with the secondary antibody (#MP-5405, Vector Labs) for 1 h at room temperature, and then visualized using an ImmPACT Vector Red substrate kit (#SK-5105, Vector Labs). Subsequently, the tissue sections were similarly incubated with TFE3 (1:500, #HPA023881, Sigma) and the secondary antibody (#MP-7401, Vector Labs) and visualized by DAB (#SK-4100, Vector Labs). Next, tissue sections were dehydrated in graded alcohols and xylene before mounting with permanent mounting medium (#H-5000, Vector Labs). Finally, the sections were examined under a light microscope (Eplice Ni-E, Nikon).

The optical density of TFE3 staining on human brain samples was analyzed with the aid of Image Pro Plus 6 software. The regions of the nuclei and cytoplasm of neuromelanin-positive cells were defined by experienced researchers. The area with neuromelanin was excluded to avoid the interference of neuromelanin. The density of DAB staining in the nuclear and cytoplasmic regions was measured separately, and then the ratio of nuclear/cytoplasmic TFE3 expression was calculated in each cell.

For mouse samples, the immunohistochemical method was performed as previously described (Li et al., 2020). Briefly, cryo-coronal sections (40 μm) encompassing the entire midbrain and striatum were collected serially. Sections were incubated with anti-TH antibody (1:10,000, #AB152, Merck Millipore) overnight at 4°C and then visualized using the Vectastain Elite ABC Kit (#PK-6101, Vector Labs) and the DAB Peroxidase Substrate Kit (#SK-4100, Vector Labs), following the manufacturer’s protocol.

For Nissl staining, adjacent midbrain sections were mounted on the slides, stained with 0.5% cresyl violet after de-fatting, dehydrated and cover-slipped.

To estimate the number of TH-positive and Nissl-positive neurons in the SN, stereological counting was performed as previously described (Li et al., 2020). Briefly, stereological cell counting analyses were performed using a stereological method of optical fractionator with the aid of Stereo Investigator (MicroBrightField Inc.). Every fourth 40 μm-thick sections from the midbrain (AP −2.7 to −3.8 mm) were collected. The SN was delineated under a ×4 objective, and the actual counting was performed under a ×100 objective. Stereological counting was performed in a double-blind fashion. The optical density of striatal TH-positive fibers was estimated using ImageJ software. The region of the dorsal striatum was determined as previously described (Li et al., 2020). All analyses were performed blinded to the treatments.

GraphPad Prism version 8.0 (GraphPad Software) was used for the statistical analysis. All data are presented as mean ± standard error of the mean (SEM). When comparing data from two groups, a two-tailed Student’s t test was used. When there were two variables, ANOVA followed by Tukey’s multiple comparisons test was used. For all analyses, statistical significance was considered when probability value of p < 0.05.

To investigate whether TFE3 is involved in the pathogenesis of PD, we first analyzed TFE3 protein expression levels in postmortem human brains. Immunohistochemistry for TFE3 was performed on the midbrain sections of postmortem brains from both PD and control subjects. The brownish pigment neuromelanin served as a convenient marker for dopaminergic neurons, and TFE3 immunostaining was visualized with gray-black substrate (DAB with Nickel) to best distinguish it from neuromelanin. TFE3 staining was evident in the SN dopaminergic neurons of control brains and it was localized to the nucleus and cytoplasm (Figure 1A). In PD patients, we found that TFE3 staining in the nuclei was reduced in the SN dopaminergic neurons (Figure 1A).

The shuttling of MiTF/TFE family members between the nucleus and cytoplasm usually occurred in a variety of stress condition (Raben and Puertollano, 2016). Therefore, we performed quantitative analysis of nuclear-to-cytoplasmic TFE3 immunostaining in individual neuromelanin-positive neurons in five PD and five control subjects. The quantitative results showed that the nuclear/cytoplasmic ratios of TFE3 were decreased in PD patients, demonstrating a reduction of nuclear TFE3 expression in the dopaminergic neurons of PD patients (Figure 1B). These results suggest that the transcriptional activity of TFE3 is repressed in SN dopaminergic neurons of PD patients.

Although TFE3 is reported to be a crucial gene that regulates autophagy and lysosomal biogenesis, the function of TFE3 in dopaminergic neurons is undefined. Therefore, we assessed whether TFE3 is involved in autophagy of dopaminergic neurons by TFE3 knockdown. We delivered AAV-shScr or AAV-shTFE3 to the unilateral SN of mice by stereotaxic injections. Five weeks following AAV injection, the mice were sacrificed.

AAV-shTFE3 injection significantly decreased Tfe3 mRNA levels in the SN of mice (Figure 2A). Then, we measured the transcriptional levels of autophagy-lysosomal genes targeted by TFE3. The mRNA levels of Map1lc3b, Lamp1 and Cstd were diminished in the AAV-shTFE3 group, although only the decrease in Lamp1 was statistically significant (Figure 2A). No significant change was observed in the mRNA levels of Sqstm1/p62 (Figure 2A). To precisely assess the effect of TFE3 knockdown on autophagy of dopaminergic neurons, we observed the autophagy-lysosomal proteins in situ by immunofluorescence. Our results showed that TFE3 was distributed in both the cytoplasm and the nuclei of dopaminergic neurons (Figure 2B), as observed in human (Figure 1A). TFE3 of dopaminergic neurons was dramatically reduced in the AAV-shTFE3 group (Figures 2B,C). LC3, an autophagosome marker, was significantly decreased in the dopaminergic neurons of mice injected with AAV-shTFE3 compared with AAV-shScr (Figures 2D,E). The magnified image showed that the number of LC3-positive dots of dopaminergic neurons was also markedly reduced in the AAV-shTFE3 group (Figure 2D), implicating a blockade of autophagosome formation by TFE3 knockdown . Lamp1, a lysosome marker, and CatD, a lysosomal protease, showed decreased expression levels in dopaminergic neurons of mice injected with AAV-shTFE3 (Figure 2F–I). Similarly, magnified images showed that the numbers of Lamp1-and CatD-positive dots of dopaminergic neurons were reduced in the AAV-shTFE3 group (Figures 2F,H), indicating that the number and activity of lysosomes were reduced by TFE3 knockdown. Conversely, p62, an autophagy receptor, was significantly up-regulated in the dopaminergic neurons of mice injected with AAV-shTFE3 (Figures 2J,K). Furthermore, magnified image showed that TFE3 knockdown promoted p62 aggregates formation (Figure 2J), suggesting compromised autophagic degradative capacity. These results demonstrate that TFE3 inhibition of dopaminergic neurons leads to autophagy flux disruption, supporting the notion that reduced nuclear TFE3 in dopaminergic neurons may result in the downregulation of autophagy in PD patients.

Previous studies show that defective autophagy often causes dopaminergic neurons loss (Friedman et al., 2012; Tang et al., 2015a; Niu et al., 2016; Ren et al., 2018). Since autophagy was severely impaired in dopaminergic neurons by TFE3 knockdown, we examined the impact of TFE3 knockdown on dopaminergic neuronal survival. We performed an unbiased stereological analysis of dopaminergic neurons in the SN of mice 3 months after the stereotaxic administration of AAV-shScr or AAV-shTFE3. Unbiased stereological estimation revealed a 56.7% loss of TH-positive neurons in the AAV-shTFE3 group relative to the AAV-shScr group (Figures 3A,B). Consistently, there was a 43.8% reduction of Nissl-positive neurons in the SN in the AAV-shTFE3 group compared with the AAV-shScr group (Figures 3C,D), suggesting an actual dopaminergic neuronal loss rather than a loss of TH expression. In parallel with the results in the SN, densitometric analysis showed that TFE3 knockdown resulted in a 64.3% decrease in TH-positive fibers in the striatum (Figures 3E,F). Taken together, these results indicate that TFE3-mediated autophagy is required for the survival of dopaminergic neurons.

Because TFE3 knockdown contributes to autophagy dysfunction of dopaminergic neurons, we investigated whether TFE3 overexpression can enhance autophagy in dopaminergic neurons. To activate TFE3, we delivered Flag-tagged murine Tfe3 under the TH promoter by AAV (AAV-TFE3). AAV-Flag was used as a control. Both viruses were injected unilaterally in the SN of mice. Two weeks after AAV injection, the mice were sacrificed.

Contrary to TFE3 knockdown, AAV-TFE3 injection significantly increased Tfe3 mRNA levels in the SN of mice (Figure 4A). Autophagy-lysosomal genes reported to be targeted by TFE3 were up-regulated, except the increase in Sqstm1/p62 was not statistically significant (Figure 4A). Likewise, to precisely examine the effect of TFE3 overexpression on autophagy of dopaminergic neurons, we observed the autophagy-lysosomal proteins in situ by immunofluorescence. We found that TFE3 protein expression was remarkably upregulated in the dopaminergic neurons of mice injected with AAV-TFE3 compared with AAV-Flag (Figures 4B,C). In contrast to TFE3 knockdown, the protein expression levels of LC3, Lamp1 and CatD were significantly increased in the dopaminergic neurons of mice injected with AAV-TFE3 (Figure 4D–I). Moreover, magnified images showed that the numbers of LC3-, Lamp1-and CatD-positive dots were increased, demonstrating that TFE3 overexpression promoted autophagosome formation and lysosomal biogenesis (Figures 4D,F,H). Although a slight increase of p62 was also observed in the AAV-TFE3 group, magnified images showed that there were no p62 aggregates (Figures 4J,K). These findings indicate that TFE3 overexpression enhances autophagy flux in dopaminergic neurons.

Previous studies showed lysosomal depletion and reduced autophagosomes in the MPTP mouse model of PD (Dehay et al., 2010; Bove et al., 2014; Zhu et al., 2020). Our results showed that TFE3 overexpression can activate the biogenesis of autophagosomes and lysosomes. Hence, an MPTP mouse model was used to determine whether TFE3 overexpression could counteract autophagy defects in dopaminergic neurons. One week after AAV injection, the mice were challenged with MPTP. For immunofluorescence analysis, mice were sacrificed 1 day after the last MPTP injection.

In mice injected with AAV-Flag, MPTP resulted in decreased protein expression levels of LC3 compared with those in Saline-treated mice (Figures 5A,B). TFE3 overexpression completely reversed MPTP-induced downregulation of LC3 in dopaminergic neurons (Figures 5A,B). Similarly, TFE3 overexpression significantly rescued the decrease of Lamp1 and CatD in the dopaminergic neurons of MPTP-treated mice, compared to the AAV-Flag group (Figures 5C–F). These results demonstrate that TFE3 overexpression can reverse MPTP-induced autophagy dysfunction in dopaminergic neurons.

Our results showed that TFE3 overexpression restored autophagic activity of dopaminergic neurons in the MPTP model. Activation of autophagy in dopaminergic neurons often exerts a neuroprotective effect. Therefore, we evaluated whether TFE3 overexpression has a neuroprotective effect. For immunohistochemical analysis, mice were sacrificed 21 days after the last MPTP injection.

We systematically analyzed dopaminergic neuron numbers in the SN of mice after unilateral injection of AAV viruses. Unbiased stereological estimation revealed a 50.5% loss of TH-positive neurons induced by MPTP injection in the AAV-Flag group, while only 7.4% of the TH-positive neurons degenerated in the AAV-TFE3 group (Figures 6A,C). Nissl staining revealed similar trends and statistical results (47.8 and 7.1% loss in the AAV-Flag and AAV-TFE3 groups, respectively; Figures 6B,D), suggesting an actual dopaminergic neuronal loss rather than a loss of TH expression. The loss of dopaminergic neurons in the SN was accompanied by fewer TH-positive terminals in the striatum. Next, we quantified TH-positive terminals in the striatum using densitometry. In parallel with the results in the SN, TFE3 overexpression mitigated MPTP-induced loss of TH-positive fibers in the striatum (Figures 6E,F). These findings demonstrate that TFE3 overexpression prevents MPTP-induced dopaminergic neurodegeneration.