Rewiring of neuronal metabolism promotes neurodegenerative recovery caused by mitochondrial dysfunction

Present *Current address: Cologne 50931, Germany, Cologne Excellence Cluster Research on Cellular Stress Response in Aging-related Diseases (CECAD).
The neurodegeneration of mitochondrial diseases is considered irreversible because the metabolic plasticity of neurons is limited, but the effect of mitochondrial dysfunction on the cell autonomy of neuronal metabolism in the body is poorly understood. Here, we introduce the cell-specific proteome of Purkinje neurons with progressive OXPHOS deficiency caused by disrupted mitochondrial fusion dynamics. We found that mitochondrial dysfunction triggered a profound change in the field of proteomics, ultimately leading to the sequential activation of precise metabolic programs before cell death. Unexpectedly, we determined the obvious induction of pyruvate carboxylase (PCx) and other peroxidases related to supplementing the TCA cycle intermediates. The inhibition of PCx exacerbated oxidative stress and neurodegeneration, indicating that atherosclerosis has a protective effect in neurons lacking OXPHOS. The restoration of mitochondrial fusion in terminally degenerated neurons completely reverses these metabolic characteristics, thereby preventing cell death. Our findings identify previously unknown pathways that confer resilience to mitochondrial dysfunction and show that neurodegeneration can be reversed even in the late stages of the disease.
The central role of mitochondria in maintaining neuronal energy metabolism is emphasized by the extensive neurological symptoms associated with human mitochondrial diseases. Most of these diseases are caused by gene mutations that regulate mitochondrial gene expression (1, 2) or gene destruction related to mitochondrial dynamics, which indirectly affect the stability of mitochondrial DNA (mtDNA) (3, 4). Work in animal models has shown that in response to mitochondrial dysfunction in surrounding tissues, conservative metabolic pathways (5-7) can be activated, thus providing important insights into the pathogenesis of these complex diseases. In stark contrast, our understanding of the metabolic changes of specific cell types caused by the general failure of brain mitochondrial adenosine triphosphate (ATP) production is fundamental (8), emphasizing the need to identify therapeutic targets that can be used to prevent or prevent disease. Prevent neurodegeneration (9). The lack of information is the fact that nerve cells are widely considered to have very limited metabolic flexibility compared to the cell types of surrounding tissues (10). Given that these cells play a central role in coordinating the supply of metabolites to neurons to promote synaptic transmission and respond to injury and disease conditions, the ability to adapt cell metabolism to the challenging conditions of brain tissue is almost limited to glial cells (11-14). In addition, the inherent cellular heterogeneity of brain tissue largely hinders the study of metabolic changes that occur in specific neuronal subgroups. As a result, little is known about the exact cellular and metabolic consequences of mitochondrial dysfunction in neurons.
In order to understand the metabolic consequences of mitochondrial dysfunction, we isolated Purkinje neurons (PNs) in different stages of neurodegeneration caused by the destruction of mitochondrial outer membrane fusion (Mfn2). Although Mfn2 mutations in humans are associated with a form of hereditary motor sensory neuropathy known as Charcot-Marie-Tooth type 2A (15), the conditional destruction of Mfn2 in mice is a well-recognized induction of oxidation​​​ Phosphorylation (OXPHOS) dysfunction method. The various neuronal subtypes (16-19) and the resulting neurodegenerative phenotype are accompanied by progressive neurological symptoms, such as movement disorders (18, 19) or cerebellar ataxia (16). By using a combination of label-free quantitative (LFQ) proteomics, metabolomics, imaging, and virological methods, we show that progressive neurodegeneration strongly induces pyruvate carboxylase (PCx) and other factors involved in arteriosclerosis of PNs in vivo The expression of enzymes. To verify the relevance of this finding, we specifically down-regulated the expression of PCx in Mfn2-deficient PNs, and found that this operation aggravated oxidative stress and accelerated neurodegeneration, thus proving that azoospermia confers cell death Metabolic adaptability. Severe expression of MFN2 can completely rescue the terminal degeneration PN with severe OXPHOS deficiency, massive consumption of mitochondrial DNA, and apparently broken mitochondrial network, which further emphasizes that this form of neurodegeneration can even recover in the advanced stage of disease before cell death.
In order to visualize the mitochondria in Mfn2 knockout PNs, we used a mouse strain that allows Cre-dependent mitochondria to target yellow fluorescent protein (YFP) (mtYFP) (20) Cre expression and checked the mitochondrial morphology in vivo. We found that the destruction of the Mfn2 gene in PNs would lead to the gradual division of the mitochondrial network (Figure S1A), and the earliest change was found at 3 weeks of age. In contrast, the substantial degeneration of the PN cell layer, as evidenced by the loss of Calbindin immunostaining, did not begin until 12 weeks of age (Figure 1, A and B). The time mismatch between the earliest changes in mitochondrial morphology and the visible onset of neuronal death prompted us to investigate the metabolic changes triggered by mitochondrial dysfunction before cell death. We developed a fluorescence-activated cell sorting (FACS)-based strategy to isolate YFP (YFP+)-expressing PN (Figure 1C), and in control mice (Mfn2 + / loxP :: mtYFP loxP- stop-loxP: : L7-cre), hereinafter referred to as CTRL (Figure S1B). The optimization of the gating strategy based on the relative intensity of the YFP signal allows us to purify the YFP+ body (YFPhigh) of PNs from non-PNs (YFPneg) (Figure S1B) or putative fluorescent axon/dendritic fragments (YFPlow; Figure S1D, left), confirmed by confocal microscope (Figure S1D, right). In order to verify the identity of the classified population, we conducted LFQ proteomics and then principal component analysis, and found that there is a clear separation between YFPhigh and YFPneg cells (Figure S1C). YFPhigh cells showed a net enrichment of known PNs markers (ie Calb1, Pcp2, Grid2 and Itpr3) (21, 22), but no enrichment of proteins commonly expressed in neurons or other cell types (Figure 1D) ). A comparison between samples in classified YFPhigh cells collected in independent experiments showed a correlation coefficient> 0.9, demonstrating good reproducibility between biological replicates (Figure S1E). In summary, these data validated our plan for acute and specific isolation of feasible PN. Because the L7-cre driver system used induces mosaic recombination in the first week after delivery (23), we started to culling mice from CTRL and conditional (Mfn2 loxP / loxP :: mtYFP loxP-stop-loxP :: L7-cre) Collect neurons. After recombination is completed, it is called Mfn2cKO at 4 weeks of age. As the end point, we selected 8 weeks of age when the PN layer was still intact despite the obvious mitochondrial fragments (Figure 1B and Figure S1A). In total, we quantified a total of 3013 proteins, of which about 22% were based on MitoCarta 2.0 annotations based on the mitochondrial proteome as mitochondria (Figure 1E) (Figure 1E) (24). The differential gene expression analysis performed at week 8 showed that only 10.5% of all proteins had significant changes (Figure 1F and Figure S1F), of which 195 proteins were down-regulated and 120 proteins were up-regulated (Figure 1F). It is worth noting that the “innovative pathway analysis” of this data set shows that the differentially expressed genes mainly belong to a restricted set of specific metabolic pathways (Figure 1G). Interestingly, although the down-regulation of pathways related to OXPHOS and calcium signaling confirms the induction of mitochondrial dysfunction in fusion-deficient PNs, other categories that mainly involve amino acid metabolism are significantly up-regulated, which is in line with the metabolism that occurs in mitochondrial PNs. Rewiring is consistent. disfunction.
(A) Representative confocal photographs of cerebellar sections of CTRL and Mfn2cKO mice showing progressive loss of PNs (calbindin, gray); nuclei were counterstained with DAPI. (B) Quantification of (A) (one-way analysis of variance, ***P<0.001; n = 4 to 6 circles from three mice). (C) Experimental workflow. (D) Heat map distribution of markers specific to Purkinje (top) and other cell types (middle). (E) Venn diagram showing the number of mitochondrial proteins identified in the classified PN. (F) Volcano plot of differentially expressed proteins in Mfn2cKO neurons at 8 weeks (significance cut-off value of 1.3). (G) The creativity pathway analysis shows the five most important up-regulation (red) and down-regulation (blue) pathways in the Mfn2cKO PN classified as 8 weeks. The average expression level of each detected protein is shown. Grayscale heat map: adjusted P value. ns, not important.
Proteomics data showed that the protein expression of complexes I, III, and IV gradually decreased. Complexes I, III, and IV all contained essential mtDNA-encoded subunits, while complex II, which was only nuclear-coded, was basically unaffected (Figure 2A and Figure S2A). . Consistent with the proteomics results, immunohistochemistry of cerebellar tissue sections showed that the MTCO1 (mitochondrial cytochrome C oxidase subunit 1) subunit level of complex IV in PN gradually decreased (Figure 2B). The mtDNA-encoded subunit Mtatp8 was significantly reduced (Figure S2A), while the steady-state level of the nuclear-encoded ATP synthase subunit remained unchanged, which is consistent with the known stable ATP synthase subassembly F1 complex when mtDNA expression is stable. The formation is consistent. Interrupt (7). Evaluation of the mtDNA level in the sorted Mfn2cKO PNs by real-time polymerase chain reaction (qPCR) confirmed the gradual decrease in mtDNA copy number. Compared with the control group, at 8 weeks of age, only about 20% of the mtDNA level Was retained (Figure 2C). Consistent with these results, the confocal microscopy staining of Mfn2cKO PNs to detect DNA shows the time-dependent consumption of mitochondrial nucleotides (Figure 2D). We found that only some candidates involved in mitochondrial protein degradation and stress response were up-regulated, including Lonp1, Afg3l2 and Clpx, and OXPHOS complex assembly factors. No significant changes in the levels of proteins involved in apoptosis were detected (Figure S2B). Similarly, we found that the mitochondria and endoplasmic reticulum channels involved in calcium transport have only minor changes (Figure S2C). In addition, the evaluation of autophagy-related proteins found no significant changes, which is consistent with the visible induction of autophagosomes observed in vivo by immunohistochemistry and electron microscopy (Figure S3). However, the progressive OXPHOS dysfunction in PNs is accompanied by obvious ultrastructural mitochondrial changes. Mitochondrial clusters can be seen in the cell bodies and dendritic trees of Mfn2cKO PNs aged 5 and 8 weeks, and the inner membrane structure has undergone profound changes (Figure S4, A and B). Consistent with these ultrastructural changes and a significant decrease in mtDNA, analysis of acute cerebral cerebellar slices with tetramethylrhodamine methyl ester (TMRM) showed that the mitochondrial membrane potential in Mfn2cKO PNs was significantly decreased (Figure S4C).
(A) Time course analysis of the expression level of OXPHOS complex. Only consider proteins with P<0.05 at 8 weeks (two-way ANOVA). Dotted line: No adjustment compared to CTRL. (B) Left: An example of a cerebellar section labeled with anti-MTCO1 antibody (scale bar, 20 μm). The area occupied by Purkinje cell bodies is covered by yellow. Right: Quantification of MTCO1 levels (one-way analysis of variance; n = 7 to 20 cells analyzed from three mice). (C) qPCR analysis of mtDNA copy number in the sorted PN (one-way analysis of variance; n = 3 to 7 mice). (D) Left: An example of a cerebellar slice labeled with an anti-DNA antibody (scale bar, 20 μm). The area occupied by Purkinje cell bodies is covered by yellow. Right: Quantification of mtDNA lesions (one-way analysis of variance; n = 5 to 9 cells from three mice). (E) An example of an acute cerebellar section showing mitoYFP + Purkinje cells (arrow) in a whole-cell patch clamp recording. (F) Quantification of IV curve. (G) Representative recordings of depolarizing current injection in CTRL and Mfn2cKO Purkinje cells. Top trace: The first pulse that triggered AP. Bottom trace: Maximum AP frequency. (H) Quantification of postsynaptic spontaneous inputs (sPSPs). The representative recording trace and its zoom ratio are shown in (I). One-way analysis of variance analyzed n = 5 to 20 cells from three mice. Data are expressed as mean±SEM; *P<0.05; **P<0.01; ***P<0.001. (J) Representative traces of spontaneous AP recorded using the perforated patch clamp mode. Top trace: Maximum AP frequency. Bottom trace: zoom of a single AP. (K) Quantify the average and maximum AP frequency according to (J). Mann-Whitney test; n = 5 cells were analyzed from four mice. Data are expressed as mean±SEM; not important.
Obvious OXPHOS damage was detected in the 8-week-old Mfn2cKO PN, indicating that the physiological function of neurons is severely abnormal. Therefore, we analyzed the passive electrical characteristics of OXPHOS-deficient neurons at 4 to 5 weeks and 7 to 8 weeks by performing whole-cell patch clamp recordings in acute cerebellar slices (Figure 2E). Unexpectedly, the average resting membrane potential and input resistance of Mfn2cKO neurons were similar to the control, although there were subtle differences between cells (Table 1). Similarly, at 4 to 5 weeks of age, no significant changes in the current-voltage relationship (IV curve) were found (Figure 2F). However, no Mfn2cKO neurons 7 to 8 weeks old survived the IV regimen (hyperpolarization step), indicating that there is a clear sensitivity to hyperpolarization potential at this late stage. In contrast, in Mfn2cKO neurons, the depolarizing currents that cause repetitive action potential (AP) discharges are well tolerated, indicating that their overall discharge patterns are not significantly different from those of 8-week-old control neurons (Table 1 and Figure 2G). Similarly, the frequency and amplitude of spontaneous postsynaptic currents (sPSCs) were comparable to those of the control group, and the frequency of events increased from 4 weeks to 5 weeks to 7 weeks to 8 weeks with a similar increase (Figure 2, H and I). The period of synaptic maturation in PNs (25). Similar results were obtained after perforated PNs patches. This configuration prevents the possible compensation of cellular ATP defects, as might happen in whole-cell patch clamp recording. In particular, the resting membrane potential and spontaneous firing frequency of Mfn2cKO neurons were not affected (Figure 2, J and K). In summary, these results indicate that PNs with obvious OXPHOS dysfunction can cope with high-frequency discharge patterns well, indicating that there is a compensation mechanism that allows them to maintain near-normal electrophysiological responses.
Data are expressed as mean ± SEM (one-way analysis of variance, Holm-Sidak’s multiple comparison test; *P<0.05). The unit number is indicated by brackets.
We set out to investigate whether any category in the proteomics dataset (Figure 1G) includes pathways that can counteract severe OXPHOS deficiency, thereby explaining why affected PN can maintain near-normal electrophysiology (Figure 2, E to K). . Proteomics analysis showed that the enzymes involved in the catabolism of branched chain amino acids (BCAA) were significantly up-regulated (Figure 3A and Figure S5A), and the final product acetyl-CoA (CoA) or succinyl CoA can supplement the tricarboxylates in arteriosclerosis Acid (TCA) cycle. We found that the content of BCAA transaminase 1 (BCAT1) and BCAT2 both increased. They catalyzed the first step of BCAA catabolism by generating glutamate from α-ketoglutarate (26). All subunits that make up the branched chain keto acid dehydrogenase (BCKD) complex are upregulated (the complex catalyzes the subsequent and irreversible decarboxylation of the resulting BCAA carbon skeleton) (Figure 3A and Figure S5A). However, no obvious changes in BCAA itself were found in the sorted PN, which may be due to the increased cellular uptake of these essential amino acids or the use of other sources (glucose or lactic acid) to supplement the TCA cycle (Figure S5B). PNs lacking OXPHOS also showed increased glutamine decomposition and transamination activities at 8 weeks of age, which can be reflected by the up-regulation of the mitochondrial enzymes glutaminase (GLS) and glutamine pyruvate transaminase 2 (GPT2) (Figure 3, A and C). It is worth noting that the up-regulation of GLS is limited to the spliced ​​isoform glutaminase C (GLS-GAC) (the change of Mfn2cKO/CTRL is approximately 4.5-fold, P = 0.05), and its specific up-regulation in cancer tissues Can support mitochondrial bioenergy. (27).
(A) The heat map shows the fold change in protein level for the specified route at 8 weeks. (B) Example of a cerebellar slice labeled with anti-PCx antibody (scale bar, 20 μm). The yellow arrow points to the Purkinje cell body. (C) Time course protein expression analysis identified as an important candidate for atherosclerosis (multiple t-test, *FDR <5%; n = 3-5 mice). (D) Above: A schematic diagram showing the different ways of entering the labeled carbon contained in the [1-13C]pyruvate tracer (ie, via PDH or trans-arterial route). Bottom: The violin chart shows the percentage of single-labeled carbon (M1) converted to aspartic acid, citric acid and malic acid after labeling acute cerebellar slices with [1-13C]pyruvate (paired t-test; ** P <0.01). (E) Comprehensive time history analysis of the indicated path. Only consider proteins with P<0.05 at 8 weeks​​. Dashed line: no adjustment value (two-way analysis of variance; * P <0.05; *** P <0.001). Data are expressed as mean±SEM.
In our analysis, BCAA catabolism has become one of the key up-regulation pathways. This fact strongly suggests that the ventilation volume entering the TCA cycle may be changed in PN lacking OXPHOS. This may represent a major form of neuronal metabolic rewiring, which may have a direct impact on neuronal physiology and survival during the maintenance of severe OXPHOS dysfunction. Consistent with this hypothesis, we found that the main anti-atherosclerotic enzyme PCx is up-regulated (Mfn2cKO/CTRL changes approximately 1.5 times; Figure 3A), which catalyzes the conversion of pyruvate to oxaloacetate (28), which is believed to be in brain tissue The expression in is restricted to astrocytes (29, 30). Consistent with the proteomics results, confocal microscopy showed that PCx expression was specifically and significantly increased in OXPHOS-deficient PNs, while PCx reactivity was mainly restricted to the adjacent Bergmann glial cells of the control (Figure 3B). To functionally test the observed upregulation of PCx, we treated acute cerebellar slices with [1-13C]pyruvate tracer. When pyruvate was oxidized by pyruvate dehydrogenase (PDH), its isotope label disappeared , But is incorporated into the TCA cycle intermediates when pyruvate is metabolized by vascular reactions (Figure 3D). In support of our proteomics data, we observed a large number of markers from this tracer in the aspartic acid of Mfn2cKO slices, while citric acid and malic acid also had a moderate trend, although not significant (Figure 3D).
In the dopamine neurons of MitoPark mice with mitochondrial dysfunction caused by dopamine neurons specifically destroying the mitochondrial transcription factor A gene (Tfam) (Figure S6B), PCx expression was also significantly up-regulated (31), indicating that acetone acid arteriosclerosis The occurrence of the disease is regulated during the dysfunction of neuronal OXPHOS in the body. It is worth noting that it has been found that unique enzymes (32-34) that may be expressed in neurons that may be associated with arteriosclerosis are significantly up-regulated in PNs lacking in OXPHOS, such as propionyl-CoA carboxylase (PCC-A), Malonyl-CoA converts propionyl-CoA to succinyl-CoA and mitochondrial malic enzyme 3 (ME3), whose main role is to recover pyruvate from malate (Figure 3, A and C) (33, 35). In addition, we found a significant increase in the Pdk3 enzyme, which phosphorylates and thus inactivates PDH (36), while no changes were detected in the Pdp1 enzyme that activates PDH or the PDH enzyme complex itself (Figure 3A). Consistently, in Mern2cKO PNs, the phosphorylation of the α1 subunit α (PDHE1α) subunit of the pyruvate dehydrogenase E1 component of the PDH complex in Ser293 (known to inhibit the enzyme activity of PDH) was enhanced ( Figure S6C) (Figure S6C). Pyruvate has no vascular access.
Finally, we found that the super pathway of serine and glycine biosynthesis, the related mitochondrial folate (1C) cycle and proline biosynthesis (Figure 1G and Figure S5C) are all significantly up-regulated, according to reports, during the activation process. The surrounding tissues are activated with mitochondrial dysfunction (5-7). Confocal analysis supporting these proteomics data showed that in PN with OXPHOS missing, cerebellar slices of 8-week-old mice were subjected to serine hydroxymethyltransferase 2 (SHMT2), a key enzyme of the mitochondrial folate cycle. Significant immune response (Figure S5D). In 13 CU-glucose-incubated acute cerebellar slices, metabolic tracing experiments further confirmed the up-regulation of serine and proline biosynthesis, indicating that the flux of carbon isoforms into serine and proline increased (Figure S5E). Since the reactions promoted by GLS and GPT2 are responsible for the synthesis of glutamate from glutamine and the transamination between glutamate and α-ketoglutarate, their upregulation indicates that OXPHOS-deficient neurons have an increased demand for glutamate , This may be aimed at maintaining the increased biosynthesis of proline (Figure S5C). In contrast to these changes, a proteomic analysis of cerebellar astrocytes from PN-specific Mfn2cKO mice showed that these pathways (including all antiperoxidases) did not change significantly in expression, thus demonstrating This metabolic redirection is selective to degraded PN (Fig. S6, D to G).
In summary, these analyses revealed significantly different patterns of temporal activation of specific metabolic pathways in PNs. Although abnormal neuronal mitochondrial function can lead to early atherosclerosis and 1C remodeling (Figure 3E and Figure S5C), and even predictable changes in the expression of I and IV complexes, the changes in serine de novo synthesis are only It only became apparent in the late stages. OXPHOS dysfunction (Figure 3E and Figure S5C). These findings define a sequential process in which the stress-induced mitochondrial (1C cycle) and cytoplasmic (serine biosynthesis) response synergistically with the increase in atherosclerosis in the TCA cycle to reshape neuronal metabolism.
8-week-old OXPHOS-deficient PNs can maintain high-frequency excitation activity and undergo significant metabolic reconnection to compensate for mitochondrial dysfunction. This discovery raises an interesting possibility that even at this moment, these cells may also Receive therapeutic intervention to delay or prevent neurodegeneration. Late. We resolved this possibility through two independent interventions. In the first method, we designed a Cre-dependent adeno-associated virus (AAV) vector so that MFN2 can be selectively expressed in OXPHOS-deficient PNs in vivo (Figure S7A). The AAV encoding MFN2 and the fluorescent reporter gene mCherry (Mfn2-AAV) were verified in primary neuron cultures in vitro, which caused MFN2 to be expressed in a Cre-dependent manner and rescued the mitochondrial morphology, thereby preventing neuromutation in Mfn2cKO neurons ( Figure S7, B, D and E). Next, we conducted in vivo experiments to stereotactically deliver 8-week-old Mfn2-AAV to the cerebellar cortex of Mfn2cKO and control mice, and analyzed 12-week-old mice (Figure 4A). The treated Mfn2cKO mice died (Figure 1, A and B) (16). Viral transduction in vivo resulted in selective expression of PN in some cerebellar circles (Figure S7, G and H). The injection of the control AAV expressing only mCherry (Ctrl-AAV) had no significant effect on the degree of neurodegeneration in Mfn2cKO animals. In contrast, the analysis of Mfn2cKOs transduced with Mfn2-AAV showed a significant protective effect of the PN cell layer (Figure 4, B and C). In particular, the neuron density seems to be almost indistinguishable from the control animals (Figure 4, B and C, and Figure S7, H and I). The expression of MFN1 but not MFN2 is equally effective in saving neuronal death (Figure 4C and Figure S7, C and F), which indicates that the expression of ectopic MFN1 can effectively supplement the lack of MFN2. Further analysis at the single PN level showed that Mfn2-AAV largely rescued the ultrastructure of mitochondria, normalized mtDNA levels, and reversed the high expression of the anti-angiogenesis marker PCx ​​(Figure 4, C to E ). Visual inspection of the rescued Mfn2cKO mice in a resting state showed that their posture and motor symptoms (movement S1 to S3) were improved. In conclusion, these experiments show that delayed reintroduction of MFN2 into PNs severely deficient in OXPHOS is sufficient to reverse mtDNA consumption and induce atherosclerosis, thereby preventing axon degeneration and neuronal death in vivo.
(A) A scheme showing the experimental schedule for injecting AAV encoding MFN2 when the indicated metabolic pathway is activated. (B) Representative confocal photographs of 12-week-old cerebellar sections transduced with Mfn2cKO mice at 8 weeks and labeled with anti-Calbindin antibody. Right: Scaling of axon fibers. The scale of the axon zoom is 450 and 75 μm. (C) Left: Quantification of Purkinje cell density in the AAV transduction loop (AAV+) (one-way analysis of variance; n = 3 mice). Right: mtDNA focus analysis in transduced PN at week 12 (unpaired t-test; n = 6 cells from three mice). * P <0.05; ** P <0.01. (D) Representative transmission electron micrographs of PNs of Mfn2cKO cerebellar sections transduced with the indicated viral vectors. The pink mask illustrates the area occupied by dendrites, and the yellow dotted square illustrates the zoom provided on the right; n represents the nucleus. Scale bar, 1μm. (E) shows an example of PCx staining in PN transduced at 12 weeks. Scale bar, 20μm. OE, overexpression; FC, fold change.
Finally, we investigated the importance of peroxidase-induced cell survival in PNs that have experienced OXPHOS dysfunction. We generated mCherry encoding AAV-shRNA (short hairpin RNA) specifically targeting mouse PCx mRNA (AAV-shPCx), and injected the virus or its scrambled control (AAV-scr) into the cerebellum of Mfn2cKO mice. The injection was performed in the fourth week of age (Figure 5A) to achieve effective PCx knockdown during the period when PCx expression increased (Figure 3C) and the PN cell layer was still intact (Figure 1A). It is worth noting that knocking down PCx (Figure S8A) leads to a significant acceleration of PN death, which is the only restricted ring of infection (Figure 5, B and C). In order to understand the mechanism of the metabolic effects induced by PCx up-regulation, we studied the redox status of PNs after PCx ​​knockdown and AAV-mediated optical biosensor Grx1-roGFP2 were simultaneously expressed (Figure S8, B to D) to evaluate glutathione The relative change of peptide redox potential (38). Then, we performed two-photon fluorescence lifetime imaging microscopy (FLIM) in acute brain slices of 7-week-old Mfn2cKO or control littermates to detect potential changes in cytoplasmic redox status after verifying FLIM conditions (Figure S8, E to G). The analysis showed a significant increase in the oxidation state of a single Mfn2cKO PNs lacking PCx expression, which is different from control neurons or Mfn2cKO PNs expressing only scrambled shRNA (Figure 5, D and E). When PCx expression was down-regulated, the percentage of Mfn2cKO PNs showing a highly oxidized state increased by more than three times (Figure 5E), indicating that PCx up-regulation maintained the redox capacity of degenerated neurons.
(A) A scheme showing the experimental schedule for injecting AAV encoding shPCx when the indicated metabolic pathway is activated. (B) Representative confocal photographs of 8-week-old cerebellar sections in Mfn2cKO mice transduced and labeled with anti-calcineurin antibody at 4 weeks. Scale bar, 450μm. (C) Quantification of Purkinje cell density in AAV-transduced loops (one-way analysis of variance; n = 3 to 4 mice). Data are expressed as mean±SEM; ***P<0.001. (D) Representative FLIM picture shows the average life span of 7-week-old PN expressing glutathione redox sensor Grx1-roGFP2 under the specified experimental conditions. LUT (look-up table) ratio: survival time interval (in picoseconds). Scale bar, 25μm. (E) The histogram shows the distribution of Grx1-roGFP2 lifetime values ​​from (D) (n=158 to 368 cells in two mice under each condition). The pie chart above each histogram: shows the number of cells with significantly longer (red, oxidized) or shorter (blue, reduced) lifespan values, which exceed 1 SD of the average lifespan value in CTRL-AAV-scr. (F) The proposed model shows the protective effect of upregulation of neuronal PCx.
All in all, the data we provide here show that the re-expression of MFN2 can completely rescue advanced PN with severe OXPHOS deficiency, severe mtDNA depletion, and extremely abnormal ista-like morphology, thereby providing continuous progress even in advanced diseases. Neurodegeneration provides reversible evidence of the stage before cell death. This degree of metabolic flexibility is further emphasized by the ability of neurons to induce atherosclerosis (a rewiring of the TCA cycle), which inhibits PCx expression in PNs lacking OXPHOS and enhances cell death, thereby playing a protective role (Figure 5F).
In this study, we provided evidence that the response of PNs to OXPHOS dysfunction is to gradually converge to TCA cycle atherosclerosis through the differential activation pathway activated by metabolic programs. We confirmed the proteomic analysis with many complementary methods and revealed that when challenged by severe mitochondrial dysfunction, neurons have a previously unknown form of metabolic elasticity. To our surprise, the entire rewiring process does not necessarily mark the terminal metabolic state that accompanies neurodegeneration gradually and irreversibly, but our data suggests that it may constitute a maintenance neuron even in the stage before cell death Functional compensation mechanism. This finding indicates that neurons have a considerable degree of metabolic plasticity in the body. This fact proves that the later reintroduction of MFN2 can reverse the expression of key metabolic markers and prevent PN degeneration. On the contrary, it inhibits atherosclerosis and accelerates nerves. transsexual.
One of the most fascinating findings in our research is that PNs lacking OXPHOS can modify the TCA cycle metabolism by up-regulating enzymes that specifically stimulate arteriosclerosis. Metabolic rearrangement is a common feature of cancer cells, some of which rely on glutamine to supplement TCA cycle intermediates to produce reducing equivalents, which drive the respiratory chain and maintain the production of lipid and nucleotide biosynthesis precursors (39 , 40). A recent study showed that in peripheral tissues experiencing OXPHOS dysfunction, the reconnection of glutamine/glutamate metabolism is also a prominent feature (5, 41), where the direction of glutamine entry into the TCA cycle depends on Because of the severity of OXPHOS injury (41). ). However, there is lack of clear evidence on any similarity of neuronal metabolic plasticity in the body and its possible relevance in the disease context. In a recent in vitro study, primary cortical neurons were shown to mobilize glutamate pools for neurotransmission, thereby promoting oxidative metabolism and atherosclerosis under metabolic stress conditions (42). It is worth noting that under the pharmacological inhibition of the TCA cycle enzyme succinate dehydrogenase, pyruvate carboxylation is believed to maintain the synthesis of oxaloacetate in cultured cerebellar granule neurons (34). However, the physiological relevance of these mechanisms to brain tissue (where atherosclerosis is believed to be mainly confined to astrocytes) still has important physiological significance (43). In this case, our data shows that PNs damaged by OXPHOS in the body can be switched to BCAA degradation and pyruvate carboxylation, which are the two main sources of supplementation of TCA pool intermediates. Although the putative contribution of BCAA catabolism to neuronal energy metabolism has been proposed, in addition to the role of glutamate and GABA for neurotransmission (44), there is still no evidence for these mechanisms in vivo. Therefore, it is easy to speculate that dysfunctional PNs can automatically compensate for the consumption of TCA intermediates driven by the assimilation process by increasing atherosclerosis. In particular, upregulation of PCx may be required to maintain an increased demand for aspartic acid, which has been shown in proliferating cells with mitochondrial dysfunction (45). However, our metabolomics analysis did not reveal any significant changes in the steady-state level of aspartic acid in Mfn2cKO PNs (Figure S6A), which presumably reflects the different metabolic utilization of aspartic acid between proliferating cells and post-mitotic neurons . Although the exact mechanism of PCx upregulation in dysfunctional neurons in vivo remains to be characterized, we demonstrated that this premature response plays an important role in maintaining the redox state of neurons, which was demonstrated in FLIM experiments on cerebellar slices. In particular, preventing PNs from up-regulating PCx can lead to a more oxidized state and accelerate cell death. The activation of BCAA degradation and the carboxylation of pyruvate are not ways to characterize the peripheral tissues of mitochondrial dysfunction (7). Therefore, they seem to be a priority feature of OXPHOS-deficient neurons, even if not the only feature, which is important for neurodegeneration. .
Cerebellar disease is a heterogeneous type of neurodegenerative disease that usually manifests as ataxia and often damages PNs (46). This neuron population is particularly vulnerable to mitochondrial dysfunction because their selective degeneration in mice is sufficient to reproduce many of the motor symptoms that characterize human spinocerebellar ataxia (16, 47, 48). According to reports, a transgenic mouse model with a mutant gene is associated with human spinocerebellar ataxia and has mitochondrial dysfunction (49, 50), emphasizing the importance of studying the consequences of OXPHOS deficiency in PNPH. Therefore, it is particularly suitable to effectively isolate and study this unique neuron population. However, given that PNs are very sensitive to pressure and account for a low proportion of the entire cerebellar cell population, for many omics-based studies, selective separation of them as whole cells is still a challenging aspect. Although it is almost impossible to achieve absolute lack of contamination of other cell types (especially adult tissues), we combined an effective dissociation step with FACS to obtain a sufficient number of viable neurons for downstream proteomics analysis, and have Quite high protein coverage (about 3000 proteins) compared with the existing data set of the whole cerebellum (51). By preserving the viability of whole cells, the method we provide here allows us not only to check the changes in the metabolic pathways in the mitochondria, but also to check the changes in its cytoplasmic counterparts, which complements the use of mitochondrial membrane label enrichment specific to cell types The new method for the number of mitochondria in complex tissues (52, 53). The method we describe is not only related to the study of Purkinje cells, but can be easily applied to any type of cell to address metabolic changes in diseased brains, including other models of mitochondrial dysfunction.
Finally, we have identified a therapeutic window during this metabolic rearrangement process that can completely reverse the key signs of cellular stress and prevent neuronal degeneration. Therefore, understanding the functional implications of the rewiring described here may provide fundamental insights into possible treatments for maintaining neuronal viability during mitochondrial dysfunction. Future research aimed at dissecting changes in energy metabolism in other brain cell types is needed to fully reveal the applicability of this principle to other neurological diseases.
MitoPark mice have been described previously (31). C57BL/6N mice with loxP flanking Mfn2 genes have been described previously (18) and crossed with L7-Cre mice (23). The resulting double heterozygous progeny were then crossed with homozygous Mfn2loxP/Mfn2loxP mice to generate Purkinje-specific gene knockouts for Mfn2 (Mfn2loxP/Mfn2loxP; L7-cre). In a subset of mating, the Gt (ROSA26) SorStop-mito-YFP allele (stop-mtYFP) was introduced through additional crosses (20). All animal procedures were performed in accordance with European, national and institutional guidelines and approved by LandesamtfürNatur of Umwelt and Verbraucherschutz, North Rhine-Westphalia, Germany. Animal work also follows the guidance of the European Federation of Laboratory Animal Sciences Associations.
After anesthetizing the pregnant woman’s cervical dislocation, the mouse embryo is isolated (E13). The cortex was dissected in Hanks’ Balanced Salt Solution (HBSS) supplemented with 10 mM Hepes and passed on Dulbecco’s Modified Eagle’s Medium containing papain (20 U/ml) and cysteine ​​(1μg/ml). Incubate the tissue in DMEM) and dissociate it by enzymatic digestion. Ml) at 37°C for 20 minutes, and then mechanically milled in DMEM supplemented with 10% fetal bovine serum. Cells were seeded on glass coverslips coated with polylysine at a density of 2×106 per 6 cm culture dish or at a density of 0.5×105 cells/cm2 for imaging analysis. After 4 hours, the medium was replaced with Neurobasal serum-free medium containing 1% B27 supplement and 0.5 mM GlutaMax. The neurons were then maintained at 37°C and 5% CO2 throughout the experiment, and fed once a week. In order to induce recombination in vitro, 3μl (24-well culture dish) or 0.5μl (24-well plate) of the following AAV9 virus vector was used to treat neurons on the second day in vitro: AAV9.CMV.PI.eGFP. WPRE.bGH (Addgene, catalog number 105530-AAV9) and AAV9.CMV.HI.eGFP-Cre.WPRE.SV40 (Addgene, catalog number 105545-AAV9).
Mouse Mfn1 and Mfn2 complementary DNA (obtained from Addgene plasmid #23212 and #23213, respectively) are marked with the V5 sequence (GKPIPNPLLGLDST) at the C-terminus, and are fused with mCherry in frame through the T2A sequence. Grx1-roGFP2 is a gift from Heidelberg TP Dick DFKZ (Deutsches Krebsforschungszentrum). By replacing the tdTomato cassette using conventional cloning methods, the cassette was subcloned into the pAAV-CAG-FLEX-tdTomato backbone (Addgene reference number 28306) to generate pAAV-CAG-FLEX-mCherry-T2A-MFN2-V5, pAAV-CAG- FLEX-mCherry-T2A-MFN1-V5 and pAAV-CAG-FLEX-Grx-roGFP2 vectors. A similar strategy was used to generate the control vector pAAV-CAG-FLEX-mCherry. In order to generate the AAV-shPCx construct, a plasmid AAV vector (VectorBuilder, pAAV [shRNA] -CMV-mCherry-U6-mPcx- [shRNA#1]) is required, which contains the DNA sequence encoding the shRNA targeting mouse PCx (5′CTTTCGCTCTAAGGTGCTAAACTCGAGTTTAGCACCTTAGAGCGAAAG 3′) Under the control of the U6 promoter, mCherry is used under the control of the CMV promoter. The production of auxiliary AAV vectors was carried out according to the manufacturer’s instructions (Cell Biolabs). In short, use a transfer plasmid carrying mCherry-T2A-MFN2-V5 (pAAV-CAG-FLEX-mCherry-T2A-MFN2-V5), mCherry-T2A-MFN1-V5 (pAAV-CAG-FLEX-mCherry) transiently Transfection of 293AAV cells-T2A-MFN1-V5), mCherry (pAAV-CAG-FLEX-mCherry) or Grx-roGFP2 (pAAV-CAG-FLEX-Grx-roGFP2) coding gene, as well as coding AAV1 capsid protein and accessory protein Packaging plasmid plasmid, using calcium phosphate method. The crude virus supernatant was obtained by freeze-thaw cycles in a dry ice/ethanol bath and lysed cells in phosphate buffered saline (PBS). The AAV vector was purified by discontinuous iodixanol gradient ultracentrifugation (24 hours at 32,000 rpm and 4°C) and concentrated using an Amicon ultra-15 centrifugal filter. Genome titer of AAV1-CAG-FLEX-mCherry-T2A-MFN2-V5 [2.9×1013 genome copy (GC)/ml], AAV1-CAG-FLEX-mCherry (6.1×1012 GC/ml), AAV1-CAG- FLEX was as previously described (54), measured by real-time quantitative PCR (qPCR) -MFN1-V5 (1.9×1013 GC/ml) and AAV1-CAG-FLEX-Grx-roGFP2 (8.9×1012 GC/ml).
Primary neurons were scraped off in ice-cold 1x PBS, pelleted, and then homogenized in 0.5% Triton X-100 / 0.5% sodium deoxycholate/PBS lysis buffer containing phosphatase and protease inhibitor (Roche). Protein quantification was performed by using the bicinchoninic acid assay (Thermo Fisher Scientific). The proteins were then separated by SDS-polyacrylamide gel electrophoresis, and then blotted onto a polyvinylidene fluoride membrane (GE Healthcare). Block non-specific sites and incubate with the primary antibody (see Table S1 for details) in 5% milk in TBST (Tris-buffered saline with Tween), washing steps and secondary antibody in TBST Incubate. Incubate with primary antibody overnight at +4°C. After washing, apply the secondary antibody for 2 hours at room temperature. Subsequently, by incubating the same blot with an anti-β-actin antibody, the same loading was confirmed. Detection by converting to chemiluminescence and enhancing chemiluminescence (GE Healthcare).
The neurons previously seeded on glass coverslips were fixed with 4% paraformaldehyde (PFA)/PBS at the specified time point at room temperature for 10 minutes. The coverslips are first permeated with 0.1% Triton X-100/PBS for 5 minutes at room temperature, and then in blocking buffer [3% bovine serum albumin (BSA)/PBS]. On the second day, the coverslips were washed with blocking buffer and incubated with the appropriate fluorophore-conjugated secondary antibody for 2 hours at room temperature; finally, the samples were washed thoroughly in PBS with 4′,6-diamidino-2 -Phenylindole (DAPI) is counterstained and then fixed on the microscope slide with Aqua-Poly/Mount.
Mice (male and female) were anesthetized by intraperitoneal injection of ketamine (130 mg/kg) and xylazine (10 mg/kg) and administered subcutaneously with carprofen analgesic (5 mg/kg) , And placed in a stereotactic instrument (Kopf) equipped with a warm pad. Expose the skull and use a dental drill to thin the part of the cerebellar cortex corresponding to the mis bone (from lambda: tail 1.8, lateral 1, corresponding to lobules IV and V). Use a curved syringe needle to carefully create a small hole in the skull to avoid disrupting the vasculature below. Then the thin drawn glass capillary is slowly inserted into the micro-hole (from -1.3 to -1 on the ventral side of the dura mater), and 200 to 300 nl AAV is injected into the micro-injector (Narishige) with manual syringes (Narishige) several times at low pressure over a period of time 10 to 20 minutes window. After the infusion, place the capillary for another 10 minutes to allow the virus to spread completely. After the capillaries are withdrawn, the skin is sutured carefully to minimize wound inflammation and allow the animal to recover. The animals were treated with analgesics (caspofen) for several days after the operation, during which time their physical condition was carefully monitored and then they were euthanized at the stated time point. All procedures were carried out in accordance with European, national and institutional guidelines and were approved by LandesamtfürNatur of Umwelt and Verbraucherschutz, North Rhine-Westphalia, Germany.
The animals were anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg), and the heart was perfused with 0.1 M PBS first, and then with 4% PFA in PBS. The tissue was dissected and fixed in 4% PFA/PBS overnight at 4°C. A vibrating knife (Leica Microsystems GmbH, Vienna, Austria) was used to prepare sagittal sections (50 μm thick) from the fixed brain in PBS. Unless otherwise specified, staining of free-floating sections was performed as described above (13) at room temperature and stirring. In short, first, the obtained slices were permeabilized with 0.5% Triton X-100/PBS for 15 minutes at room temperature; for some epitopes (Pcx and Shmt2), by in tris-EDTA buffer at 80°C (PH 9) heat the slices for 25 minutes instead of this step. Next, the sections were incubated with primary antibody (see Table S1) in blocking buffer (3% BSA/PBS) at 4°C overnight with stirring. The next day, the sections were washed with blocking buffer and incubated with the appropriate fluorophore-conjugated secondary antibody for 2 hours at room temperature; finally, the sections were washed thoroughly in PBS, counter-stained with DAPI, and then fixed with AquaPolymount On a microscope slide.
A laser scanning confocal microscope (TCS SP8-X or TCS Digital Light Sheet, Leica Microsystems) equipped with a white light laser and a 405 diode ultraviolet laser was used to image the sample. By exciting the fluorophore and collecting the signal with Hybrid Detector (HyDs), LAS-X software was used to collect stacked images conforming to Nyquist sampling in sequential mode: for non-quantitative panels, it is highly dynamic signals (for example, in somatic cells and dendrites) mtYFP) Use HyD to detect the number of PNs in BrightR mode). Gating of 0.3 to 6 ns is applied to reduce background.
Real-time imaging of sorted cells. After sorting in Neurobasal-A medium containing 1% B27 supplement and 0.5 mM GlutaMax, cells were immediately seeded on poly-l-lysine-coated glass slides (μ-Slide8 Well, Ibidi, catalog number 80826) , And then keep it at 37°C and 5% CO2 for 1 hour to allow the cells to settle. Real-time imaging was performed on a Leica SP8 laser scanning confocal microscope equipped with a white laser, HyD, 63×[1.4 numerical aperture (NA)] oil objective lens and a heating stage.
The mouse was quickly anesthetized with carbon dioxide and decapitated, the brain was quickly removed from the skull, and cut into 200μm thick (for 13C labeling experiment) or 275μm thick (for two photon experiments) sagittal section filled with the following materials The ice cream (HM-650 V, Thermo Fisher Scientific, Walldorf, Germany) is filled with the following substances: 125 mM ice-cold, carbon-saturated (95% O2 and 5% CO2) low Ca2 + artificial cerebrospinal fluid (ACSF) NaCl, 2.5 mM KCl, 1.25 mM sodium phosphate buffer, 25 mM NaHCO3, 25 mM glucose, 0.5 mM CaCl2 and 3.5 mM MgCl2 (osmotic pressure of 310 to 330 mmol). Transfer the obtained brain slices to a pre-incubation chamber containing higher Ca2 + ACSF (125.0 mM NaCl, 2.5 mM KCl, 1.25 mM sodium phosphate buffer, 25.0 mM NaHCO3, 25.0 mM d-glucose, 1.0 mM CaCl2 and 2.0 mM MgCl2) Medium) pH 7.4 and 310 to 320 mmol).
During the imaging process, the slices were moved into a dedicated imaging room, and the experiment was performed under continuous ACSF perfusion at a constant temperature of 32° to 33°C. A multiphoton laser scanning microscope (TCS SP8 MP-OPO, Leica Microsystems) equipped with a Leica 25x objective lens (NA 0.95, water), Ti: Sapphire laser (Chameleon Vision II, Coherent) was used for slice imaging. FLIM module (PicoHarp300, PicoQuant).
FLIM of Grx1-roGFP2. The changes in the cytoplasmic redox state of PNs were measured by two-photon FLIM in sagittal brain slices, where the Grx1-roGFP2 biosensor targeted PNs. In the PN layer, the acquisition field is selected about 50 to 80 μm below the slice surface to ensure that there is a viable PN (that is, the lack of beaded structure or neuronal morphological changes along the dendrites) and the double positive roGFP2 sensor and AAV encoding shRNA PCx or its control sequence (each co-expressing mCherry). Collect single-stack images with 2x digital zoom [excitation wavelength: 890 nm; 512 nm 512 pixels]. Detection: internal HyD, fluorescein isothiocyanate (FITC) filter group] and image averaging within 2 to 3 minutes are used to ensure that enough photons are collected (1000 photons in total) for curve fitting. The sensitivity of the Grx1-roGFP2 probe and the verification of FLIM conditions were performed by monitoring the lifespan value of roGFP2 when adding exogenous 10 mM H2O2 to the perfusion ACSF (to maximize oxidation, resulting in increased lifespan), and then adding 2 mM dithiothreitol (minimizes the degree of reduction, resulting in a decrease in lifespan) (Figure S8, D to G). Use FLIMfit 5.1.1 software to analyze the acquired results, fit the single exponential decay curve of the entire image to the measured IRF (instrument response function), and χ2 is approximately 1. To calculate the lifetime of a single PN, the mask around the nerve body was manually drawn, and the average lifetime in each mask was used for quantification.
Mitochondrial potential analysis. After the acute section was incubated with 100 nM TMRM directly added to the perfused ACSF for 30 minutes, the mitochondrial potential changes of PNs were measured by a two-photon microscope. TMRM imaging was performed by exciting the probe at 920 nm and using internal HyD (tetramethylrhodamine isothiocyanate: 585/40 nm) to collect signals; by using the same excitation wavelength but using a different internal HyD (FITC :525/50) to image mtYFP. Use ImageJ’s Image Calculator plug-in to evaluate mitochondrial potential at the single cell level. In short, the plug-in equation: signal = min (mtYFP, TMRM) is used to identify the mitochondrial region that shows the TMRM signal in Purkinje Somali in the single-stack confocal image of the corresponding channel. Then the pixel area in the resulting mask is quantified, and then normalized on the corresponding threshold single-stack image of the mtYFP channel to obtain the mitochondrial fraction showing the mitochondrial potential.
The image was deconvoluted with Huygens Pro (Scientific Volume Imaging) software. For the scanned pictures of tiles, the montage of a single tile is made using the automatic stitching algorithm provided by LAS-X software. After image calibration, use ImageJ and Adobe Photoshop to further process the image and uniformly adjust the brightness and contrast. Use Adobe Illustrator for graphic preparation.
mtDNA focus analysis. The number of mtDNA lesions was quantified on cerebellar sections labeled with antibodies against DNA by confocal microscope. Each target area was created for the cell body and the nucleus of each cell, and the respective area was calculated using the Multi Measure plug-in (ImageJ software). Subtract the nuclear area from the cell body area to obtain the cytoplasmic area. Finally, the Analyze Particles plug-in (ImageJ software) was used to automatically quantify the cytoplasmic DNA points indicating mtDNA on the threshold image, and the obtained results were normalized to the PN average of CTRL mice. The results are expressed as the average number of nucleosides per cell.
Protein expression analysis. Use ImageJ’s Image Calculator plug-in to evaluate protein expression in PN at the single cell level. In short, in the single-layer confocal image of the corresponding channel, through the equation: signal = min (mtYFP, antibody), the mitochondrial region that shows immunoreactivity to a certain antibody in Purkina is identified. Then the pixel area in the resulting mask is quantified, and then normalized on the corresponding threshold single-stack image of the mtYFP channel to obtain the mitochondrial fraction of the displayed protein.
Purkinje cell density analysis. The Cell Counter plug-in of ImageJ was used to evaluate Purkinje density by dividing the number of Purkinje cells counted by the length of the cerebellar ring occupied by the counted cells.
Sample preparation and collection. The brains from the control group and Mfn2cKO mice were fixed in 2% PFA/2.5% glutaraldehyde in 0.1 M phosphate buffer (PB), and then coronal sections were prepared using ciliates (Leica Mikrosysteme GmbH, Vienna, Austria) (Thickness 50 to 60 μm) Then fix in PB buffer in 1% os tetraoxide and 1.5% potassium ferrocyanide at room temperature for 1 hour. The sections were washed three times with distilled water, and then stained with 70% ethanol containing 1% uranyl acetate for 20 minutes. The sections were then dehydrated in graded alcohol and embedded in Durcupan ACM (Araldite casting resin M) epoxy resin (Electron Microscopy Sciences, catalog number 14040) between silicon-coated glass slides, and finally at 60°C Polymerize in the oven for 48 hours. The cerebellar cortex area was selected and 50 nm ultrathin sections were cut on Leica Ultracut (Leica Mikrosysteme GmbH, Vienna, Austria) and picked on a 2×1 mm copper slit grid coated with polystyrene film. The sections were stained with a solution of 4% uranyl acetate in H2O for 10 minutes, washed with H2O several times, then with Reynolds lead citrate in H2O for 10 minutes, and then washed with H2O several times. Micrographs were taken with a transmission electron microscope Philips CM100 (Thermo Fisher Scientific, Waltham, MA, USA) using a TVIPS (Tietz Video and Image Processing System) TemCam-F416 digital camera (TVIPS GmbH, Gauting, USA). Germany).
For mice infected with AAV, the brain was separated and sliced ​​into a 1 mm thick sagittal section, and the cerebellum was examined using a fluorescence microscope to identify the AAV-infected ring (that is, mCherry expressing). Only experiments in which AAV injection results in a very high transduction efficiency of the Purkinje cell layer (ie almost the entire layer) in at least two consecutive cerebellar rings are used. The AAV-transduced loop was microdissected for overnight post-fixation (4% PFA and 2.5% glutaraldehyde in 0.1 M cocoate buffer) and further processed. For EPON embedding, the fixed tissue was washed with 0.1 M sodium cocoate buffer (Applichem), and incubated with 2% OsO4 (os, Science Services; Caco) in 0.1 M sodium cocoate buffer (Applichem) 4 Hours, then wash for 2 hours. Repeat 3 times with 0.1 M cocamide buffer. Subsequently, the ascending series of ethanol was used to incubate each ethanol solution at 4°C for 15 minutes to dehydrate the tissue. The tissue was transferred to propylene oxide and incubated overnight in EPON (Sigma-Aldrich) at 4°C. Place the tissue in fresh EPON at room temperature for 2 hours, and then embed it at 62°C for 72 hours. Use an ultramicrotome (Leica Microsystems, UC6) and a diamond knife (Diatome, Biel, Switzerland) to cut 70 nm ultrathin sections, and stain with 1.5% uranyl acetate for 15 minutes at 37°C, and stain with lead citrate solution 4 minutes. The electron micrographs were taken using a JEM-2100 Plus transmission electron microscope (JEOL) equipped with Camera OneView 4K 16-bit (Gatan) and DigitalMicrograph software (Gatan). For analysis, electron micrographs were acquired with 5000× or 10,000× digital zoom.
Morphological analysis of mitochondria. For all analyses, the contours of individual mitochondria were manually outlined in digital images using ImageJ software. Different morphological parameters are analyzed. Mitochondrial density is expressed as a percentage obtained by dividing the total mitochondrial area of ​​each cell by the cytoplasm area (cytoplasm area = cell area-cell nucleus area) × 100. The roundness of mitochondria is calculated with the formula [4π∙(area/perimeter 2)]. The ista morphology of mitochondria was analyzed and divided into two categories (“tubular” and “blister”) according to their main shapes.
Autophagosome/lysosome number and density analysis. Use ImageJ software to manually outline the contours of each autophagosome/lysosome in the digital image. Autophagosome/lysosome area is expressed as a percentage calculated by dividing the total autophagosome/lysosome structure area of ​​each cell by the cytoplasm area (cytoplasm area=cell area-nucleus area)×100. The density of autophagosomes/lysosomes is calculated by dividing the total number by the number of autophagosome/lysosome structures per cell (in terms of cytoplasmic area) (cytoplasmic area = cell area-nuclear area).
Labeling for acute sectioning and sample preparation. For experiments that require glucose labeling, transfer the acute brain slices to a pre-incubation chamber, which contains saturated carbon (95% O2 and 5% CO2), high Ca2 + ACSF (125.0 mM NaCl, 2.5 mM KCl, 1.25 mM sodium phosphate buffer, 25.0 mM NaHCO 3, 25.0 mM d-glucose, 1.0 mM CaCl 2 and 2.0 mM MgCl 2, adjusted to pH 7.4 and 310 to 320 mOsm), in which glucose is 13 C 6- Glucose substitution (Eurisotop, catalog number CLM-1396). For experiments that require pyruvate labeling, transfer the acute brain slices to higher Ca2 + ACSF (125.0 mM NaCl, 2.5 mM KCl, 1.25 mM sodium phosphate buffer, 25.0 mM NaHCO3, 25.0 mM d-glucose, 1.0 mM CaCl2 and Add 2.0mM MgCl2, adjust to pH 7.4 and 310 to 320mOsm), and add 1mM 1-[1-13C]pyruvate (Eurisotop, catalog number CLM-1082). Incubate the sections for 90 minutes at 37°C. At the end of the experiment, the sections were quickly washed with an aqueous solution (pH 7.4) containing 75 mM ammonium carbonate, and then homogenized in 40:40:20 (v:v:v) acetonitrile (ACN): methanol: water. After the sections were incubated on ice for 30 minutes, the samples were centrifuged at 21,000 g for 10 minutes at 4°C, and the clear supernatant was dried in a SpeedVac concentrator. The resulting dried metabolite pellet was stored at -80°C until analysis.
Liquid chromatography-mass spectrometry analysis of 13 C-labeled amino acids. For liquid chromatography-mass spectrometry (LC-MS) analysis, the metabolite pellet was resuspended in 75μl of LC-MS grade water (Honeywell). After centrifugation at 21,000 g for 5 minutes at 4°C, 20 μl of the clarified supernatant was used for amino acid flux analysis, while the rest of the extract was immediately used for anion analysis (see below). Amino acid analysis was performed using the previously described benzoyl chloride derivatization protocol (55, 56). In the first step, 10μl of 100 mM sodium carbonate (Sigma-Aldrich) was added to 20μl of metabolite extract, and then 10μl of 2% benzoyl chloride (Sigma-Aldrich) was added to the LC grade ACN. The sample was vortexed briefly and then centrifuged at 21,000 g for 5 minutes at 20°C. Transfer the cleared supernatant to a 2 ml autosampler vial with a conical glass insert (200 μl volume). The samples were analyzed using the Acquity iClass ultra-high performance LC system (Waters) connected to the Q-Exactive (QE)-HF (Ultra High Field Orbitrap) high-resolution precision mass spectrometer (Thermo Fisher Scientific). For analysis, 2μl of the derivatized sample was injected into a 100×1.0 mm high-strength silica T3 column (Waters) containing 1.8μm particles. The flow rate is 100μl/min, and the buffer system consists of buffer A (10 mM ammonium formate and 0.15% formic acid in water) and buffer B (ACN). The gradient is as follows: 0%B at 0 minutes; 0%B. 0 to 15% B at 0 to 0.1 minutes; 15 to 17% B at 0.1 to 0.5 minutes; B at 17 to 55% at 0.5 to 14 minutes; B at 55 to 70% at 14 to 14.5 minutes; at 14.5 to 70 to 100% B at 18 minutes; 100% B at 18 to 19 minutes; 100 to 0% B at 19 to 19.1 minutes; 0% B at 19.1 to 28 minutes (55, 56). The QE-HF mass spectrometer operates in positive ionization mode with a mass range of m/z (mass/charge ratio) of 50 to 750. The applied resolution is 60,000, and the gain control (AGC) ion target obtained is 3×106, and the maximum ion time is 100 milliseconds. The heated electrospray ionization (ESI) source operates at a spray voltage of 3.5 kV, a capillary temperature of 250°C, a sheath airflow of 60 AU (arbitrary units), and an auxiliary airflow of 20 AU. 250°C. The S lens is set to 60 AU.
Anion chromatography-MS analysis of 13C labeled organic acids. The remaining metabolite precipitate (55μl) was analyzed using a Dionex ion chromatography system (ICS 5000+, Thermo Fisher Scientific) connected to a QE-HF mass spectrometer (Thermo Fisher Scientific). In short, 5μl of metabolite extract was injected into a Dionex IonPac AS11-HC column equipped with HPLC (2 mm×250 mm, particle size 4μm, Thermo Fisher Scientific) in push-in partial loop mode with a filling ratio of 1. )on. Dionex IonPac AG11-HC guard column (2 mm x 50 mm, 4μm, Thermo Fisher Scientific). The column temperature is maintained at 30°C, and the autosampler is set to 6°C. Use a potassium hydroxide cartridge provided with deionized water to generate a potassium hydroxide gradient through the eluent generator. Separation of metabolites at a flow rate of 380μl/min, applying the following gradient: 0 to 3 minutes, 10 mM KOH; 3 to 12 minutes, 10 to 50 mM KOH; 12 to 19 minutes, 50 to 100 mM KOH; 19 to 21 minutes , 100 mM KOH; 21 to 21.5 minutes, 100 to 10 mM KOH. The column was re-equilibrated under 10 mM KOH for 8.5 minutes.
The eluted metabolites are combined with a 150μl/min isopropanol supplement stream after the column and then directed to a high-resolution mass spectrometer operating in negative ionization mode. MS is monitoring the mass range from m/z 50 to 750 with a resolution of 60,000. The AGC is set to 1×106, and the maximum ion time is kept at 100 ms. The heated ESI source was operated at a spray voltage of 3.5 kV. The other settings of the ion source are as follows: capillary temperature 275°C; sheath gas flow, 60 AU; auxiliary gas flow, 20 AU at 300°C, and S lens setting to 60 AU.
Data analysis of 13C labeled metabolites. Use TraceFinder software (version 4.2, Thermo Fisher Scientific) for data analysis of isotope ratio. The identity of each compound was verified by a reliable reference compound and independently analyzed. In order to perform isotope enrichment analysis, the area of ​​the extracted ion chromatogram (XIC) of each 13C isotope (Mn) was extracted from [M + H] +, where n is the carbon number of the target compound, used to analyze amino acids or [ MH] + is used to analyze anions. The mass accuracy of XIC is less than five parts per million, and the accuracy of RT is 0.05 minutes. The enrichment analysis is performed by calculating the ratio of each detected isotope to the sum of all isotopes of the corresponding compound. These ratios are given as percentage values ​​for each isotope, and the results are expressed as molar percent enrichment (MPE), as described previously (42).
The frozen neuron pellet was homogenized in ice-cold 80% methanol (v/v), vortexed, and incubated at -20°C for 30 minutes. Vortex the sample again and stir at +4°C for 30 minutes. The sample was centrifuged at 21,000 g for 5 minutes at 4°C, and then the resulting supernatant was collected and dried using a SpeedVac concentrator at 25°C for subsequent analysis. As described above, LC-MS analysis was performed on the amino acids of the sorted cells. Using TraceFinder (version 4.2, Thermo Fisher Scientific), data analysis was performed using the monoisotopic mass of each compound. Quantile normalization of metabolite data was performed using the preprocessCore software package (57).
Slice preparation. The mouse was quickly anesthetized with carbon dioxide and decapitated, the brain was quickly removed from the skull, and the ice-filled vibrating knife (HM-650 V, Thermo Fisher Scientific, Walldorf, Germany) was used to cut it into 300 to 375 μm sagittal sections Cold carbon gasification (95% O2 and 5% CO2) Low Ca2 + ACSF (125.0 mM NaCl, 2.5 mM KCl, 1.25 mM sodium phosphate buffer, 25.0 mM NaHCO3, 25.0 mM d-glucose, 1.0 mM CaCl2 and 6.0 mM MgCl2 Adjust to pH 7.4 and 310 to 330 mOsm). Transfer the obtained brain slices to a chamber containing higher Ca2 + ACSF (125.0 mM NaCl, 2.5 mM KCl, 1.25 mM sodium phosphate buffer, 25.0 mM NaHCO3, 25.0 mM d-glucose, 4.0 mM CaCl2 and mM 3.5 MgCl2) pH 7.4 and 310 to 320 mOsm). Store the slices for 20 to 30 minutes so that they can be restored before recording.
recording. A microscope stage equipped with a fixed recording chamber and a 20x water immersion objective lens (Scientifica) was used for all recordings. The putative Purkinje cells were identified by (i) body size, (ii) anatomical location of the cerebellum, and (iii) expression of the fluorescent mtYFP reporter gene. The patch pipette with a tip resistance of 5 to 11 megohms is pulled out by a borosilicate glass capillary (GB150-10, 0.86 mm×1.5 mm×100 mm, Science Products, Hofheim, Germany) and a horizontal pipette Instruments (P-1000, Sutter), Novato, CA). All recordings were performed by ELC-03XS npi patch clamp amplifier (npi electronic GmbH, Tam, Germany), which was controlled by the software Signal (version 6.0, Cambridge Electronic, Cambridge, UK). The experiment was recorded at a sampling rate of 12.5 kHz. The signal is filtered with two short-pass Bessel filters with cutoff frequencies of 1.3 and 10 kHz respectively. The capacitance of the membrane and the pipette is compensated by the compensation circuit using the amplifier. All experiments were performed under the control of an Orca-Flash 4.0 camera (Hamamatsu, Gerden, Germany), which was controlled by the Hokawo software (version 2.8, Hamamatsu, Gerden, Germany).
Routine whole-cell configuration and analysis. Immediately before recording, fill the pipette with the internal solution containing the following substances: 4.0 mM KCl, 2.0 mM NaCl, 0.2 mM EGTA, 135.0 mM potassium gluconate, 10.0 mM Hepes, 4.0 mM ATP (Mg), 0.5 mM Guanosine triphosphate (GTP) (Na) and 10.0 mM creatinine phosphate were adjusted to pH 7.25, and the osmotic pressure was 290 mOsm (sucrose). Immediately after applying a force of 0 pA to rupture the membrane, the resting membrane potential was measured. The input resistance is measured by applying hyperpolarized currents of -40, -30, -20, and -10 pA. Measure the magnitude of the voltage response and use Ohm’s law to calculate the input resistance. Spontaneous activity was recorded in a voltage clamp for 5 minutes, and sPSC was identified and measured in Igor Pro (version 32 7.01, WaveMetrics, Lake Oswego, Oregon, USA) using a semi-automatic recognition script. The IV curve and steady-state current are measured by clamping the battery at different potentials (starting from -110 mV) and increasing the voltage in 5 mV steps. The production of AP was tested by applying a depolarizing current. Clamp the cell at -70 mV while applying a depolarizing current pulse. Adjust the step size of each recording unit separately (10 to 60 pA). Calculate the maximum AP frequency by manually counting the pulse spikes that cause the highest AP frequency. The AP threshold is analyzed by using the second derivative of the depolarization pulse that first triggers one or more APs.
Perforated patch configuration and analysis. Perform perforated patch recording using standard protocols. Use an ATP- and GTP-free pipette that does not contain the following ingredients: 128 mM gluconate K, 10 mM KCl, 10 mM Hepes, 0.1 mM EGTA and 2 mM MgCl2, and adjust to pH 7.2 (using KOH). ATP and GTP are omitted from the intracellular solution to prevent uncontrolled permeability of the cell membrane. The patch pipette is filled with an amphotericin-containing internal solution (approximately 200 to 250 μg/ml; G4888, Sigma-Aldrich) to obtain a record of the perforated patch. Amphotericin was dissolved in dimethyl sulfoxide (final concentration: 0.1 to 0.3%; DMSO; D8418, Sigma-Aldrich). The concentration of DMSO used had no significant effect on the neurons studied. During the punching process, the channel resistance (Ra) was continuously monitored, and the experiment was started after the amplitude of Ra and AP stabilized (20-40 minutes). Spontaneous activity is measured in a voltage and/or current clamp for 2 to 5 minutes. Data analysis was performed using Igor Pro (version 7.05.2, WaveMetrics, USA), Excel (version 2010, Microsoft Corporation, Redmond, USA) and GraphPad Prism (version 8.1.2, GraphPad Software Inc., La Jolla, CA). United States). In order to identify spontaneous APs, IgorPro’s NeuroMatic v3.0c plug-in is used. Automatically identify APs using a given threshold, which is adjusted individually for each record. Using the spike interval, determine the spike frequency with the maximum instantaneous spike frequency and the average spike frequency.
PN isolation. By adapting to the previously published protocol, PNs was purified from the mouse cerebellum at a specified stage (58). In short, the cerebellum was dissected and minced in ice-cold dissociation medium [without HBSS Ca2+ and Mg2+, supplemented with 20 mM glucose, penicillin (50 U/ml) and streptomycin (0.05 mg/ ml)], and then digest the medium in papain [HBSS, supplemented with 1-cysteine·HCl (1 mg / ml), papain (16 U / ml) and deoxyribonuclease I (DNase I; 0.1 mg/ml)] Treat for 30 minutes at 30°C. First wash the tissues in HBSS medium containing egg mucus (10 mg/ml), BSA (10 mg/ml) and DNase (0.1 mg/ml) at room temperature to prevent enzymatic digestion, and then in the HBSS medium containing 20 mM glucose Gently grinding in HBSS, penicillin (50 U/ml), streptomycin (0.05 mg/ml) and DNase (0.1 mg/ml) release single cells. The resulting cell suspension was filtered through a 70μm cell strainer, then the cells were pelleted by centrifugation (1110 rpm, 5 minutes, 4°C) and resuspended in sorting medium [HBSS, supplemented with 20 mM glucose, 20% fetal bovine ) Serum, penicillin (50 U/ml) and streptomycin (0.05 mg/ml)]; evaluate cell viability with propidium iodide and adjust the cell density to 1×106 to 2×106 cells/ml. Before flow cytometry, the suspension was filtered through a 50 μm cell strainer.
Flow cytometer. Cell sorting was performed at 4°C using FACSAria III machine (BD Biosciences) and FACSDiva software (BD Biosciences, version 8.0.1). The cell suspension was sorted using a 100 μm nozzle under a pressure of 20 psi at a rate of ~2800 events/sec. Since traditional gating criteria (cell size, bimodal discrimination, and scattering characteristics) cannot ensure the correct isolation of PN from other cell types, the gating strategy is set based on the direct comparison of the YFP intensity and autofluorescence in mitoYFP+ ​​and the control mitoYFP − Mice. YFP is excited by irradiating the sample with a 488 nm laser line, and the signal is detected using a 530/30 nm band pass filter. In mitoYFP+ ​​mice, the relative strength of the Rosa26-mitoYFP reporter gene is also used to distinguish neuronal body and axon fragments. 7-AAD is excited with a 561 nm yellow laser and detected with a 675/20 nm bandpass filter to exclude dead cells. In order to separate astrocytes at the same time, the cell suspension was stained with ACSA-2-APC, then the sample was irradiated with a 640 nm laser line, and a 660/20 nm bandpass filter was used to detect the signal.
The collected cells were pelleted by centrifugation (1110 rpm, 5 minutes, 4°C) and stored at -80°C until use. Mfn2cKO mice and their litter pups are classified on the same day to minimize procedural variability. FACS data presentation and analysis were performed using FlowJo software (FlowJo LLC, Ashland, Oregon, USA).
As mentioned above (59), real-time PCR is used to isolate DNA from the sorted neurons for subsequent mtDNA quantification. The linearity and threshold sensitivity were initially tested by running qPCR on different numbers of cells. In short, collect 300 PN in a lysis buffer consisting of 50 mM tris-HCl (pH 8.5), 1 mM EDTA, 0.5% Tween 20 and proteinase K (200 ng/ml) and incubate at 55°C 120 minutes. The cells were further incubated at 95°C for 10 minutes to ensure complete inactivation of proteinase K. Using a TaqMan probe (Thermo Fisher) specific to mt-Nd1, mtDNA was measured by semi-quantitative PCR in the 7900HT Real-Time PCR system (Thermo Fisher Scientific). Science, catalog number Mm04225274_s1), mt-Nd6 (Thermo Fisher Scientific, catalog number AIVI3E8) and 18S (Thermo Fisher Scientific, catalog number Hs99999901_s1) genes.
Proteome sample preparation. By heating the solution at 95°C for 10 minutes and sonicating, in the lysis buffer [6 M guanidine chloride, 10 mM tris(2-carboxyethyl) phosphine hydrochloride, 10 mM chloroacetamide and 100 mM tris- Lyse frozen neuron pellets in HCl]. On Bioruptor (Diagenode) for 10 minutes (30 seconds pulse / 30 seconds pause period). The sample was diluted 1:10 in 20 mM tris-HCl (pH 8.0), mixed with 300 ng trypsin gold (Promega), and incubated overnight at 37°C to achieve complete digestion. On the second day, the sample was centrifuged at 20,000 g for 20 minutes. The supernatant was diluted with 0.1% formic acid, and the solution was desalted with self-made StageTips. The sample was dried in a SpeedVac instrument (Eppendorf concentrator plus 5305) at 45°C, and then the peptide was suspended in 0.1% formic acid. All samples were prepared simultaneously by the same person. In order to analyze astrocyte samples, 4 μg desalted peptides were labeled with a tandem mass tag (TMT10plex, catalog number 90110, Thermo Fisher Scientific) with a peptide to TMT reagent ratio of 1:20. For TMT labeling, 0.8 mg of TMT reagent was resuspended in 70 μl of anhydrous ACN, and the dried peptide was reconstituted into 9 μl of 0.1 M TEAB (triethylammonium bicarbonate), to which 7 μl of TMT reagent in ACN was added. The concentration was 43.75%. After 60 minutes of incubation, the reaction was quenched with 2 μl of 5% hydroxylamine. The labeled peptides were collected, dried, resuspended in 200μl of 0.1% formic acid (FA), divided into two, and then desalted using self-made StageTips. Using UltiMate 3000 ultra high performance liquid chromatograph (UltiMate 3000 ultra high performance liquid chromatograph), one of the two halves was fractionated on a 1mm x 150mm Acquity chromatographic column filled with 130Å1.7μm C18 particles (Waters, catalog No. SKU: 186006935). Thermo Fisher Scientific). Separate peptides at a flow rate of 30μl/min, separate from 1% to 50% buffer B for 85 minutes with a stepwise gradient of 96 minutes, from 50% to 95% buffer B for 3 minutes, then 8 minutes for 95 % Buffer B; Buffer A is 5% ACN and 10 mM ammonium bicarbonate (ABC), and buffer B is 80% ACN and 10 mM ABC. Collect fractions every 3 minutes and combine them into two groups (1 + 17, 2 + 18, etc.) and dry them in a vacuum centrifuge.
LC-MS/MS analysis. For mass spectrometry, the peptides (number r119.aq) were separated on a 25 cm, 75 μm inner diameter PicoFrit analytical column (new objective lens, part number PF7508250) equipped with 1.9 μm ReproSil-Pur 120 C18-AQ medium (Dr. Maisch, mat), Use EASY-nLC 1200 (Thermo Fisher Scientific, Germany). The column was maintained at 50°C. Buffers A and B are 0.1% formic acid in water and 0.1% formic acid in 80% ACN, respectively. Peptides were separated with a stepwise gradient of 200 nl/min from 6% to 31% buffer B for 65 minutes, and from 31% to 50% buffer B for 5 minutes. The eluted peptides were analyzed on an Orbitrap Fusion mass spectrometer (Thermo Fisher Scientific). Peptide precursor m/z measurement is performed with a resolution of 120,000 in the range of 350 to 1500 m/z. Using 27% normalized collision energy, the strongest precursor with a charge state of 2 to 6 is selected for high energy C trap dissociation (HCD) cleavage. The cycle time is set to 1 s. The m/z value of the peptide fragment was measured in the ion trap using the smallest AGC target of 5×104 and the maximum injection time of 86 ms. After fragmentation, the precursor was placed on the dynamic exclusion list for 45 s. TMT-labeled peptides were separated on a 50 cm, 75 μm Acclaim PepMap column (Thermo Fisher Scientific, catalog number 164942), and the migration spectra were analyzed on an Orbitrap Lumos Tribrid mass spectrometer (Thermo Fisher Scientific) equipped with high-field asymmetric waveform ions (FAIMS) equipment (Thermo Fisher Scientific) operates at two compensation voltages of −50 and −70 V. MS3 selected based on the synchronization precursor is used for TMT report ion signal measurement. The peptide separation was carried out on EASY-nLC 1200, using a 90% linear gradient elution, with a buffer concentration of 6% to 31%; buffer A was 0.1% FA, and buffer B was 0.1% FA and 80% ACN. The analytical column is operated at 50°C. Use FreeStyle (version 1.6, Thermo Fisher Scientific) to split the original file according to the FAIMS compensation voltage.
Protein identification and quantification. Using the integrated Andromeda search engine, the original data was analyzed using MaxQuant version 1.5.2.8 (https://maxquant.org/). In addition to the Cre recombinase and YFP sequences obtained from Aequorea victoria, peptide fragment spectra were searched for the canonical sequence and isoform sequence of the mouse reference proteome (Proteome ID UP000000589, downloaded from UniProt in May 2017) . Methionine oxidation and protein N-terminal acetylation were set as variable modifications; cysteine ​​carbamoyl methylation was set as fixed modifications. The digestion parameters are set to “specificity” and “trypsin/P”. The minimum number of peptides and razor peptides used for protein identification is 1; the minimum number of unique peptides is 0. Under the conditions of peptide map matching, the protein identification rate was 0.01. The “Second Peptide” option is enabled. Use the “match between runs” option to transfer successful identifications between different original files. Use LFQ minimum ratio count 1 for label-free quantification (LFQ) (60). The LFQ intensity is filtered for at least two valid values ​​in at least one genotype group at each time point, and is extrapolated from a normal distribution with a width of. 0.3 and move down 1.8. Use Perseus computing platform (https://maxquant.net/perseus/) and R (https://r-project.org/) to analyze the LFQ results. A two-way moderate t test from the limma software package was used for differential expression analysis (61). Exploratory data analysis is performed using ggplot, FactoMineR, factoextra, GGally and pheatmap. The TMT-based proteomics data was analyzed using MaxQuant version 1.6.10.43. Search for raw proteomics data from UniProt’s human proteomics database, which was downloaded in September 2018. The analysis includes the isotope purity correction factor provided by the manufacturer. Use limma in R for differential expression analysis. The original data, database search results, and data analysis workflow and results are all stored in the ProteomeXchange alliance through the PRIDE partner repository with the data set identifier PXD019690.
Functional annotations enrich the analysis. The Ingenuity Pathway Analysis (QIAGEN) tool was used to determine the richness of the functional annotation terms of the data set at 8 weeks (Figure 1). In short, the quantitative protein list obtained from LC-MS/MS (tandem mass spectrometry) data analysis is used with the following filter criteria: Mus musculus is selected as the species and background, and the category shows the P value adjusted by Benjamini for enrichment 0.05 or lower is considered significant. For this graph, the top five excess categories in each cluster based on the adjusted P value are shown. Using multiple t-test, using the two-stage linear boost program of Benjamini, Krieger, and Yekutieli (Q = 5%), time-course protein expression analysis is performed on the important candidates identified in each category, and each row is analyzed separately. There is no need to adopt a consistent SD.
In order to compare the results of this study with published databases and generate a Venn diagram in Figure 1, we combined the quantitative protein list with MitoCarta 2.0 annotations (24). Use the online tool Draw Venn Diagram (http://bioinformatics.psb.ugent.be/webtools/Venn/) to generate the diagram.
For detailed information on the statistical procedures used for proteomics analysis, please refer to the corresponding section of Materials and Methods. For all other experiments, detailed information can be found in the corresponding legend. Unless otherwise specified, all data are expressed as mean ± SEM, and all statistical analyses were performed using GraphPad Prism 8.1.2 software.
For supplementary materials for this article, please see http://advances.sciencemag.org/cgi/content/full/6/35/eaba8271/DC1
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