Background
The primary etiologies of neurodegenerative disorders, including Alzheimer’s disease (AD), frontotemporal dementia (FTD) and Parkinson’s disease (PD), remain largely unknown, but common pathological features suggest a role for altered protein degradation. For instance, proteinaceous intracellular inclusions composed in part of aggregated α-synuclein protein, termed Lewy bodies, typify PD brain pathology, whereas neurofibrillary tangles (NFT) and Pick bodies containing phosphorylated tau protein are commonly found in the context of taupathies such as AD and FTD. Rare, inherited familial forms of neurodegenerative diseases [
1] are caused by mutations in genes encoding these accumulated proteins, such as α
-synuclein [
2,
3] in PD and tau in FTD, but the vast majority of patients do not harbor known mutations. Thus, it has been hypothesized that in these ‘sporadic’ cases, pathological inclusions may reflect broadly defective protein degradation through mechanisms such as the ubiquitin-proteasome system (UPS) [
4] and macroautophagy [
5,
6]. The latter is of particular interest because of its apparent role in the degradation of protein aggregates and inclusions [
7].
Macroautophagy is a pathway of bulk cytoplasmic protein and organelle degradation characterized by double-membrane vesicles that engulf cargo and target it to lysosomes for degradation [
8]. The pathway is typically induced in the context of starvation or other stressors. Defects in the macroautophagy process may theoretically occur at a variety of steps, from the initial formation of a pre-autophagosome limiting membrane, to the ultimate fusion of mature autophagosomes with the lysosomal compartment [
9]. Macroautophagy defects have been well described on pathological analyses of brain sections from patients with a variety of neurodegenerative disorders, including AD, PD and FTD [
5,
10]. Furthermore, inherited genetic forms of neurodegeneration are associated with mutations in the macroautophagy-lysosomal pathway [
11,
12]. Finally, as macroautophagy dysfunction is a well-documented feature of aging, it has been implicated in the age-dependent nature of the major neurodegenerative disorders [
5,
9,
10].
Genetically altered mice that are deficient in essential macroautophagy pathway components, Atg5 or Atg7, throughout neural development, display reduced neuronal survival and harbor ubiquitin-positive inclusions in the cell soma [
13‐
16]. But surprisingly, prevention of inclusion formation in the context of Atg7-deficiency by a second genetic ablation of p62, which encodes an ubiquitin-binding protein associated with autophagosomes, does not suppress neurodegeneration, arguing against a toxic role for inclusions [
17]. Thus, the mechanism of neuronal loss with macroautophagy deficiency, and how this relates to neurodegeneration, remains unclear.
Here we generated conditional Atg7-deficient mice specifically within mature CNS neurons. Atg7-deficient neurons were defective in the initiation of macroautophagy, and displayed a progressive degeneration with prominent inclusions that harbor ubiquitin, p62, phosphorylated tau and GSK3β. The mutant mice exhibited behavioral deficits consistent with the pathological changes. Furthermore, pharmacological or genetic suppression of tau phosphorylation effectively inhibited neurodegeneration in the context of Atg7-deficiency in vivo.
Discussion
Here we investigated mechanisms of neurodegeneration downstream of Atg7-deficiency, and describe the pathological accumulation of GSKβ and phospho-tau proteins. A striking feature of neuropathology in the context of Atg7-deficiency is the redistribution of GSK3β to inclusions. We note that both GSK3β and phospho-tau are reported to be found in inclusions in tauopathy patient brain [
39‐
43]. However, it is important to emphasize that Atg7-deficiency does not appear to induce a full tauopathy pathology, as not all phospho-tau epitopes are observed (e.g., PHF1 antibody is negative, Figure
4e), and amyloid staining with Thioflavin S, as well as electron microscopic analysis, do not support the presence of mature NFTs. A similar phospho-tau pattern has previously been suggested to represent an early ‘pre-tangle’ pathological state [
32], thought to reflect non-fibrillar tau aggregation prior to assembly into PHFs. Such non-fibrillar hyperphosphorylated tau, rather than mature NFTs, may be the relevant toxic form
in vivo in the context of neurodegeneration and behavioral impairment [
44]. Hoozemans
et al. reported phospho-tau-positive pre-tangles with accumulation of GSK3β, ubiquitin and p62 in postmortem specimens of AD patients [
45], reminiscent of pathology in
Atg7-deficient neurons
in vivo. Phospho-tau pathology as seen in Atg7-deficient animals may broadly relate to neuronal dysfunction in neurodegeneration, as macroautophagy deficiency and phospho-tau are commonly observed in a broad array of neurodegenerative disorders including AD, PD, tauopathy, huntington disease, amyotrophic lateral sclerosis, and Gaucher disease [
6,
46‐
49]. Although genetic mutations in
ATG7 have not been described in human disease, mutations within other components of the macroautophagy-lysosomal pathway underlie tauopathies [
50], consistent with our observations in the mouse model.
The
in vivo pharmacological and genetic ‘rescue’ studies herein suggest a role for phospho-tau accumulation in neurodegeneration downstream of Atg7-deficiency. In contrast, prior attempts to rescue macroautophagy-deficiency associated neurodegeneration by preventing the formation of aggregates, by generation of double-knockout mice deficient in Atg7 as well as p62, were unsuccessful [
17], suggesting that inclusion formation
per se is insufficient for degeneration. It is interesting to note that nonetheless, p62 deletion does rescue the Atg7 deficiency-associated cell loss in hepatocytes [
17], and thus degenerative pathways downstream of macroautophagy loss appear cell type-specific. Furthermore, within the CNS, various neuronal subtypes appear to be differentially affected by macroautophagy deficiency. Purkinje neurons deficient in Atg7 display axonal swellings and are rapidly lost [
51]. TH-positive midbrain DA neurons display axonal dystrophy and degeneration, ubiquitin/p62-positive inclusions, and delayed cell loss and locomotor dysfunction [
52]. Although tau pathology was not investigated in these other models, staining for the Parkinson’s disease associated proteins α-synuclein and leucine rich repeat kinase-2 (LRRK2) was reported in Atg7-deficient DA neurons [
52]. We failed to detect evidence of α-synuclein accumulation in our analysis of either midbrain DA neuron-selective or forebrain neuron-selective Atg7-deficient mice detailed above (data not shown). Such discrepancies may reflect differences in the selectivity or timing of the CRE-mediated deletion strains used in the different studies, or selective sensitivity to macroautophagy loss across distinct neuron types. We note that phospho-tau pathology was apparent in the context of either midbrain DA neuron-selective or forebrain neuron-selective Atg7-deficiency.
The molecular basis of GSK3β and phospho-tau accumulation in Atg7-deficient neurons remains to be elucidated. We cannot exclude the possibility that GSK3β accumulation is a secondary effect of phospho-tau accumulation. A recent study described intracellular redistribution of GSK3β to multivesicular bodies, albeit in the context of Wnt pathway modulation [
53]. As multivesicular bodies directly associate with the macroautophagy machinery, it is possible that GSK3β degradation is selectively modified with macroautophagy loss [
54]. Although GSK3β is a strong candidate for the relevant upstream kinase, we hypothesize the involvement of other kinase pathways, particularly given the multiple targets of the pharmacological kinase inhibitor used, Alsterpaullone. Furthermore, Alsterpaullone-mediated protection may be mediated through targets in addition to tau, which would be of further interest.
We propose a role for basal macroautophagy in regulating the metabolism of phospho-tau proteins at physiological or pre-pathological state (Figure
5e). In the context of macroautophagy loss, GSK3β and phospho-tau are accumulated, reminiscent of early pathology that precedes human tauopathy. It is interesting to note that both GSK3β and tau are believed to be potent upstream regulators of macroautophagy [
55‐
58]. We hypothesize that this may reflect a feedback loop, where defective macroautophagy leads progressively to more accumulation of phospho-tau and GSK3β, and in turn the accumulated phospho-tau and GSK3β both induce macroautophagy activity. Initially such feedback may be effective, although the accumulated proteins form inclusions. But once macroautophagy deficiency is complete, as in late-stage disease or in knockout mice, this feedback would be ineffective. Thus, such a feedback circuit may be an important pathway to rejuvenate the macroautophagy pathway, which is known to wane with aging [
59].
Methods
Animal
CamK-Cre transgenic mice,
Dat
Cre/+
mice,
Atg7
flox/flox
mice, hAPP-Tg and
tau KO mice, used in this study were generated previously [
19,
20,
37,
60‐
62].
CamK-Cre Tg and
tau KO mice were purchased from Jackson Laboratories. All animals were maintained in the animal facility of the Columbia University Medical Center. Experimental protocols were approved by the Institutional Animal Care and Use Committee. Genomic DNA extracted from mouse tails was amplified by PCR for genotyping using standard methods. The PCR primers are the followings: 5’-AGA TGT TCG CGA TTA TC-3’, 5’-AGC TAC ACC AGA GAC GG-3’ for Cre transgene; 5’-TGC TCT GTG AAC TGC CCT GTT T-3’, 5’-TGT TCC TGT GCA CTG CCT CAT T-3’ for
Atg7 wild-type allele; 5’-CTT GGG TGG AGA GGC TAT TC-3’, 5’-AGG TGA GAT GAC AGG AGA TC-3’ for
Atg7 floxed allele.
Histology
Mice were perfused and fixed in 4% paraformaldehyde and post-fixed at 4°C overnight, 50 μm coronal brain sections were generated using a vibratome. The sections were blocked with PBS containing 5% normal donkey serum [NDS], 0.2% Triton X-100 [Tx] for 1 h, and incubated with the solution (PBS, 1% NDS, 0.2% Tx) containing primary antibody at 4°C overnight. The following antibodies were used; anti-TH (P60101, Pel-Freez), anti-TuJ1 (MMS-435P, COVANCE), anti-MAP2 (AB5622, Millipore), anti-cleaved caspase-3 [Asp175] (#9661, Millipore), anti-active caspase-3 (AB3623, Cell Signaling Technology), anti-ubiquitin (Sigma-Aldrich), anti-p62 (03-GP62-C, American Research Products), anti-Aβ [4G8] (SIG39200, COVANCE), anti-Aβ [6E10] (SIG39300, COVANCE), anti-αSynuclein (610786, BD Bioscience) (AB5038, Millipore) (ab1903, ab24715, Abcam), anti-phosph-tau TG3 and PHF1 (gifts from Dr. Peter Davies, Alberts Einstein College of Medicine), anti-phospho-tau AT8, AT100, AT180, and AT270 (Pierce), anti-total GSK3β (#9315, Cell Signaling Technology), anti-phospho-GSK3α/β [Y279/Y216] (ab52188, Abcam), anti-phospho-GSK3β [S9] (ab30619, Abcam), anti-total CRMP2 (#9393, Cell Signaling Technology), anti-phospho-CRMP2 [T514] (#9397, Cell Signaling Technology), anti-Cdk5 (MAB5410, Millipore), anti-p35/25 (#2680, Cell Signaling Technology), anti-β-catenin (#9581, 9587, Cell Signaling Technology), and anti-β-catenin (#610154, BD Biosciecnes). For secondary detection, Cy3- or FITC-conjugated antibodies were incubated for 1 h (Jackson ImmunoResearch). Photographs were taken using a Zeiss LSM 510 Meta confocal microscope.
Neuron counting
To obtain neuronal cell count, 50 μm coronal brain sections were made using a vibratome. In order to count CA1 neurons, the first 30 sections from the rostral hippocampus were stained with rabbit anti-MAP2 antibody (AB5622, Millipore) at a dilution of 1:500, as well as NeuroTraceTM Fluorescent Nissl stain (N21480, Invitrogen). MAP2-positive neurons were visualized using a Cy3-conjugated secondary antibody (Jackson ImmunoResearch). MAP2 and Nissl double-positive neurons in the CA1 regions were counted manually. In order to count TH-positive neurons, sections covering the entire substantia nigra (25-30 sections / mouse) were stained with sheep anti-TH antibody (P60101, Pel-Freez) at a dilution of 1:250. TH-positive neurons were visualized using the ABC Kit (PK6106, Vector Laboratories) and DAB (SK4100, Vector Laboratories). TH-positive neurons in the substantia nigral regions were counted manually under the light microscope.
Electron microscopy
Electron microscopic analysis was as described [
61]. Anesthetized mice were perfused and fixed in PBS containing 4% paraformaldehyde and 0.5% gultaralaldehyde. The brains were post-fixed at 4°C for 2 h, and the 80 μm vibratome sections were made. The sections were treated in 1% osmium tetroxide, then dehydrated in pure ethanol and infiltrated overnight with Epon 812. Epon was polymerized at 60°C for 24 h, cooled and embedded in a larger Epon capsule. Ultrathin sections were cut with an MT5000 ultramicrotome, stained with uranyl acetate and lead citrate. Images were taken with a JOEL 100S Electron Microscope (JOEL USA).
Tissue fractionation
Preparation of soluble and insoluble fractions was performed as described with some modifications [
14]. Cortical and hippocampal tissues from mouse brains were homogenized in 5× volume of ice-cold 0.25M sucrose buffer (50mM Tris-HCl [pH7.6]) containing protease inhibitors (P8340, Sigma) and phosphatase inhibitors (#78420, Thermo Scientific). The homogenized tissues were centrifuged at 500× g for 10 min at 4°C. The supernatants were lysed with an equal volume of cold sucrose buffer containing 1% Triton X-100. The lysates were centrifuged at 13,000× g for 15 min at 4°C. The supernatants contained the soluble fraction. The pellets were resuspended in 1% SDS in PBS (insoluble fraction). Both fractions were subjected to standard Western Blotting analysis. The antibodies used here are: anti-phospho-tau AT8, AT100, AT180, AT270, TG3 and PHF1, anti-Tau1 and anti-Actin (ab3280, Abcam). Horseradish peroxidase-conjugated secondary antibodies (Jackson ImmunoResearch) and SuperSignal West Pico or Dura (#34077, 34075, Pierce) were used for detection.
Electrophysiology
Brains from CamK-Atg7 cWT and cKO mice littermates (~12 weeks of age) were quickly removed and transverse hippocampal slices (400 μm) were isolated with a Leica VT1200 Vibratome (Leica, Bannockburn, IL), and placed in ice-cold cutting solution (CS: 110 mM Sucrose, 60 mM NaCl, 3 mM KCl, 1.25 mM NaH2PO4, 28 mM NaHCO3, 0.5 mM CaCl2, 7 mM MgCl2, 5 mM Glucose, 0.6 mM Ascorbate. Slices were placed in an interface chamber (Scientific Systems Design, Mississauga, Ontario, Canada) and maintained at 32°C in ACSF (2 ml/min) containing 125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 25 mM NaHCO3, 25 mM D-glucose, 2 mM CaCl2, and 1 mM MgCl2. All solutions were constantly caboxygenated with 95% O2 + 5% CO2. Slices were allowed to recover for 120 min on the electrophysiology rig prior to experimentation. Bipolar stimulating electrodes (92:8 Pt:Y) were placed at the border of area CA3 and area CA1 along the Schaffer-Collateral pathway. ACSF-filled glass recording electrodes (1–3 MΩ) were placed in stratum radiatum of area CA1. Basal synaptic transmission was assessed for each slice by applying gradually increasing stimuli (0.5–15V), using a stimulus isolator (A-M Systems, Carlsborg, WA) and determining the input:output relationship. All subsequent stimuli applied to slices was equivalent to the level necessary to evoke a fEPSP that was ~40% of the maximal initial slope that could be evoked. Synaptic efficacy was continuously monitored (0.05 Hz). Sweeps were averaged together every 2 min. fEPSPs were amplified (A-M Systems Model 1800) and digitized (Digidata 1440, Molecular Devices, Sunnyvale, CA) prior to analysis (pClamp, Molecular Devices, Sunnyvale, CA). Stable baseline synaptic transmission was established for 30 min. Slices were given high-frequency stimulation (HFS) to induce long-term potentiation (LTP) using one train of 100 Hz for one second. Stimulus intensity of the HFS was matched to the intensity used in the baseline recordings. fEPSP initial slopes from averaged traces were normalized to those recorded during baseline. Two-way RM-ANOVA were used for electrophysiological data analysis with p < 0.05 as significance criteria.
Fear conditioning
10-13-mon-old male CamK-Atg7 cWT or CamK-Atg7 cKO mice were used (n = 8 - 10). The mice were placed in a conditioning chamber (Med Associates) for 2 min before the onset of a tone (conditioned stimulus) (30 s, 85 dB sound at 2800 Hz) and conditioned by a single electrical foot shock (0.45 mA) in the last 2 s. The mice were left in the chamber for another 30 s and placed back into their home cage. Contextual fear learning was measured in the same chamber 24 h after the training by monitoring the freezing for 5 min without electrical shock. Cued fear learning was measured 24 h after the contextual testing. The mice were placed in a novel chamber for 2 min (pre-conditioning). After that, the mice were exposed to the conditioned stimulus for 3 min, and the freezing was monitored. Freezing behavior was scored using FreezeView software (Med Associates Inc.).
Drug injection
Five-week-old
Dat-Atg7 cWT and
Dat-Atg7 cKO mice were treated with Alsterpaullone (A1136, A.G. Scientific) [
35]. The drug was dissolved in saline containing 20% DMSO/ 25% Tween80, sonicated, and injected intraperitoneally at a dose of 5 mg/kg every day for 3 weeks. After the final injection, the mice were perfused and processed for histological analyses. We used
Dat-Atg7 cWT mice as controls for
Dat-Atg7 cKO mice, to address potential phenotypes due to Cre transgene inserted at the DAT locus [
62].
Statistical analysis
All comparisons between groups were made using the Mann-Whitney U-test (for two samples) or non-repeated measures ANOVA (for multiple samples). The values are expressed as the means ± S.E. A p value less than 0.05 is considered significant.
Acknowledgements
We would like to thank G. Di Paulo, and O. Hobert for suggestions and comments on the manuscript, R. Hen for generously providing Dat
Cre/+
mice, P. Davies for generously providing phospho-tau antibodies, E. Kominami, T. Chiba, and K. Tanaka for generously providing Atg7
flox/flox
mice, J.Q. Trojanowski and D. Dickson for electron microscopic analysis, and T. Iwasato, J. Dunning, C. Doege, H. Rhinn, D. MacLeod, W. Vanti, S. Vasishta for technical help. This work was supported by grants from Kanae Foundation for the Promotion of Medical Science, and Research Foundation ITSUU Laboratory to K.I. K.I. was a postdoctoral fellow of New York Stem Cell Foundation. This work was supported by grants from the Michael J. Fox Foundation, NINDS, and NIA to A.A.
Competing interests
The authors declare no competing interests.
Authors’ contributions
KI, JR, HK, EC, JK, and MK performed the experiments. KI, HK, EK, EC, and AA analyzed the results. KI and AA designed the study and wrote the manuscript. All authors read and approved the final manuscript.