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Address correspondence to Dr. Eva-Maria Mandelkow, Max-Planck-Unit for Structural Molecular Biology, Notkestrasse 85, 22607 Hamburg, Germany. Tel.: 49-40-8998-2810. Fax: 49-40-8971-6822. E-mail: mand{at}mpasmb.desy.de
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Abstract |
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Key Words: axonal traffic; microtubules; tau protein; amyloid precursor protein; oxidative stress
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Introduction |
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Results |
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Elevation of tau causes transport defects in primary neuronal cells
Since N2a is a neuroblastoma cell line, we wanted to test if primary neurons would show a similar behavior when tau is increased. Hippocampal neurons were prepared from the cortices of rats or mice (following Banker and Goslin, 1998) and transfected with tau by three different methods, by herpes simplex virus (HSV) or adenovirus (AV) vectors containing the longest human tau isoform (htau40) or by the calcium phosphate method (Fig. 2). The transfected tau was detected by the enhanced immunofluorescence of an antibody against human tau (Fig. 2 A, bottom right) or by an antibody against the HA tag on tau. In control rat primary neurons, mitochondria were spread throughout the cell body and the cell processes (Fig. 2 A, top left). However, when cells were transfected with tau by either method mitochondria disappeared from the neurites and became concentrated in the cell body (Fig. 2 A, bottom left). Cells transfected by the vector only did not show an effect (unpublished data). At 16 h after transfection, the average number of mitochondria found in the initial 50 µm of the cell process decreased threefold (Fig. 2 B, left), and the decrease was even more marked further toward the growth cone. Similar observations were made with peroxisomes (Fig. 2 B, right). This means that the elevation of tau has similar effects on intracellular transport in primary neurons, neuron-like cell models, or nonneuronal cells, that is, the preferential inhibition of plus-enddirected transport by kinesin motors along microtubules so that minus-enddirected transport by a dynein-like motor becomes dominant.
Tau expression makes neurites vulnerable against stress
Since the depletion of organelles from the neurites in cells with elevated tau would be expected to make them more vulnerable, we tested their susceptibility against oxidative stress. Fig. 3 illustrates the reaction of differentiated N2a cells against 250 µM H2O2. This leads to a gradual loss of cell processes and eventually to cell death (Fig. 3 A). However, in tau-transfected cells the degradation of neurites is much more rapid (Fig. 3 A, bottom), and long neurites are more vulnerable than short ones. This was quantitated separately for long (>30 µm) neurites where the decay was noticeable already at lower H2O2 and shorter times (Fig. 3 B). Starting from a cell population differentiated for 2 d where 12.3% of the cells contained a long neurite, 150 µM H2O2 caused a loss of half the long neurites (down to 5.9%) (Fig. 3 B, third bar) in the control cells within 40 min. Tau-expressing cells had somewhat fewer long neurites to begin with (9.9%), but the loss upon exposure to H2O2 was much more dramatic (33-fold, down to 0.3%) (Fig. 3 B, seventh bar). This could be explained by the depletion of peroxisomes containing catalase for the detoxification of H2O2. We added catalase (0.02 U/µl) to the medium to check that the degeneration effect was due to the extracellular added hydrogen peroxide. Catalase alone had no effect on the survival of neurites (Fig. 3 B, second and sixth bar), but when challenging the cells with H2O2 and then immediately adding the catalase it provided an efficient protection (Fig. 3 B, fourth and eighth bars).
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One concern in interpreting these results is that tau expression might have a negative effect on cell viability or the cell's capacity for detoxification, independently of tau's effect on transport after differentiation. In other words, elevated tau might somehow be toxic for the cell independently of its state of differentiation. Therefore, as a control the survival of undifferentiated N2a cells was checked by the 3-(4,5-dimethylthiazol-2yl)-2,5-diphenyltetrazolium bromide (MTT) test (which senses the reducing capacity of mitochondria [Mattson et al., 1995]) or the trypan blue test (which senses the leakiness of the plasma membrane). By both assays, survival rates were the same in tau-expressing and control cells (unpublished data). Furthermore, we compared the metabolic pathways for the detoxification of peroxides in N2a mock and tau-transfected cells. After treatment of the cells with 100 µM H2O2, the concentration of the peroxide decreased with comparable rates in both cell lines. The inhibitory effect of 3-AT was also similar (30%) and indicates that catalase is mainly responsible for H2O2 detoxification in both cell clones (unpublished data). Various concentrations (1 and 10 mM) of hydrogen peroxide were used to probe for an involvement of the glutathione system in peroxide detoxification. No immediate rise in glutathione disulfide was detectable after the addition of the peroxide (Dringen et al., 1999); thus, both cell clones make little use of glutathione peroxidase in detoxification of peroxide. Therefore, we conclude that the expression of tau is not toxic by itself, it has no negative effect on the overall biochemical pathways in the transfected cells as such, so that the decrease in neurite growth and viability must be ascribed to the inhibition of transport processes.
The trafficking of APP is inhibited by tau
APP is a membrane protein implicated in Alzheimer's disease by improper proteolysis and accumulation of its Aß peptide fragment. After synthesis in neurons, it is initially transported to the axon by Golgi-derived vesicles, and later a small fraction travels back to the dendrites by transcytosis (Simons et al., 1995). Since tau inhibits the transport of vesicles down the axon, we investigated whether tau would affect the anterograde transport of APP as well using N2a cells stably transfected with the human isoform APP695 (Thinakaran et al., 1996) and detecting it by immunofluorescence. The inhibition is indeed observed. In the absence of tau, APP-containing vesicles can be visualized in the cell body around the Golgi area and throughout the neurite (Fig. 4 A). When the cells are additionally transfected with tau, it appears throughout the neurites (Fig. 4 A, right), but APP vesicles are strongly reduced in the neurites after 12 h. Quantification shows that the level of APP-carrying vesicles in neurites is decreased about threefold (Fig. 4 B) after tau transfection comparable to the observations with Golgi-derived vesicles in general (Fig. 1 B). The results demonstrate a direct connection between the elevation of tau and the inhibition of anterograde APP trafficking.
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The same effect of overexpressed tau on the transport of APP was seen in cultured hippocampal neurons, which were simultaneously transfected with APP-YFP and CFP-htau40 by using the recombinant adenoviral vectors. A sufficient amount of doubly transfected neurons was obtained (50% of cells). Transfection of APP alone resulted in APP vesicles distributed throughout the cell body and neurites (Fig. 5 B, top left), but cells containing both transfected APP and tau showed that APP was restricted mostly to the cell body, and dendrites and axons were almost completely devoid of YFP fluorescent vesicles (Fig. 5 B, bottom left). As a consequence, only very few moving YFP fluorescent structures could be observed, moving mainly retrogradely with velocities <0.4 µm/s (in this case the axons were clearly defined by the fluorescence of tau) (Fig. 5 B, arrow).
Similar experiments were performed to observe the movement of mitochondria (stained with Mitotracker red) in RGCs (Fig. 8). Without tau, most particles were mobile with instantaneous velocities 1 µm/s, and anterograde traffic dominated (Fig. 8 A). After AV-mediated tau transfection, about half of the mitochondria became immobile over extended periods, and the rest moved mainly retrogradely (Fig. 8 B). The quantification (Fig. 8 C) shows that tau biases the particles toward the immobile and retrograde fractions without significant changes in instantaneous velocities. These results explain the gradual disappearance of mitochondria from the cell processes discussed above (Fig. 1 D).
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Discussion |
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We have tested three types of cells: differentiated neuroblastoma cells, primary hippocampal neurons from rat or mouse brain, and chick RGCs. Tau was elevated either by stable or transient transfection, transfection with viral vectors (AV or HSV), or the calcium phosphate method. We studied the distribution of cytoskeletal components (microtubules, tau, and neurofilaments), organelles (mitochondria and peroxisomes), and vesicles derived from the Golgi complex, in particular vesicles carrying APP. In addition, we monitored the growth of neurites and their response to oxidative stress. All of the observed effects support the view that tau generally inhibits transport along microtubules, preferentially in the plus-end direction, that is, toward the growth cone. This applies to organelles, vesicles, and neurofilaments (Figs. 13). The inhibition of transport is so efficient that organelles are largely excluded from cell processes, and vesicles are dramatically reduced (Fig. 1 C). In this regard, neurons are more vulnerable than the CHO cells studied earlier because diffusion is too limited. The effects of tau are observed both in neuron-like cell models (N2a), primary neurons, and RGCs, confirming that the underlying interactions between microtubules, motors, and cargo are similar.
The inhibition of organelles, vesicles, and neurofilaments is consistent with the view that these components are carried down the axon by a kinesin-dependent transport along microtubules (Hirokawa et al., 1998; Baas, 1999; Chou et al., 2001; Goldstein, 2001; Shea and Flanagan, 2001; Zhou et al., 2001). The motor-cargo connection may be direct (Liao and Gundersen, 1998; Yabe et al., 1999; Kamal et al., 2000), or it could be mediated by adaptor proteins (Morris and Hollenbeck, 1995; Ratner et al., 1998; Setou et al., 2000; Verhey et al., 2001). By contrast, the inhibition does not apply to microtubules or their associated proteins (e.g., tau itself), presumably because their transport into the cell process is mediated by a different type of mechanism. This would be consistent with the assumption that microtubules and MAPs might be carried by a dynein-mediated transport along actin microfilaments (Sheetz et al., 1998; Baas, 1999). Thus, tau highlights the difference between microtubules and neurofilaments even though both are transported as "slow components" of axonal transport. Using GFP-tagged intermediate filament subunits, several groups showed recently that they are transported by a fast motor of the kinesin family except that the average speed in axons is slowed down by frequent long pauses (Roy et al., 2000; Wang et al., 2000).
It is interesting to compare the tau-induced transport effects with the accumulation of proteins into aggresomes (Kopito, 2000). There is a superficial similarity because in both cases there is a net transport toward the microtubule minus ends around the MTOC. Aggresomes are formed by improperly folded and aggregated proteins, which are transported toward the cell center by a dynein-mediated process. They selectively contain cellular factors engaged in protein folding or degradation, such as proteasomes, heat shock proteins, and enzymes of the ubiquitin pathway, but aggresome formation does not affect the distribution of organelles such as mitochondria, peroxisomes, or the ER. Moreover, aggresomes represent the cell's response to (nearly) irreversible protein aggregation (e.g., proteins with polyglutamine stretches [Muchowski et al., 2000]); this does not occur with tau protein in transfected cells, which remains either microtubule-bound or highly soluble in the cytosol (Preuss et al., 1997; Lu and Kosik 2001). Finally, aggresomes cannot be redispersed, in contrast to the clusters of cell components described here, which dissolve again when microtubules are destroyed or when dynein-mediated transport is impaired (e.g., by dynamitin expression [Ebneth et al., 1998]).
One can expect that transport inhibition has serious consequences for the growth and survival of cell processes. We have tested this for two cases, the growth of neurites and their vulnerability to oxidative stress. Indeed, tau-transfected cells have shorter neurites (Fig. 3 A, bottom), and they become highly sensitive to oxidative conditions (H2O2). This is due to the absence of peroxisomes, since exogenous catalase can provide substantial protection (Fig. 3 B, left). The alternative pathway of detoxification by glutathione peroxidase plays little role in neurons (Dringen et al., 1999), and we have verified this for our cell lines (unpublished data). The role of catalase can be probed specifically with the catalase inhibitor 3-AT. In control cells, it amplifies the toxic effect of H2O2 but not in tau-expressing cells (Fig. 3 C). Thus, roughly speaking the loss of peroxisomes from the neurites due to elevated tau is as damaging as the direct inhibition of catalase. By the same argument, the exclusion of mitochondria from the cell processes implies a local depletion of ATP. This might be bearable for a compact cell where ATP diffuses throughout the cytoplasm but becomes a problem in extended cell processes.
Implications of tau-induced transport defects for neurodegeneration
The results described here may be important for understanding neurodegenerative disorders such as Alzheimer's disease, frontotemporal dementias, and others, which are characterized by elevated and aggregated tau protein (for reviews see Buee et al., 2000; Hutton, 2001). Alzheimer's disease is traditionally characterized by two types of protein deposits in the brain, the extracellular amyloid plaques, consisting largely of the peptide Aß, a derivative of the membrane protein APP, and the intracellular neurofibrillary tangles, consisting mostly of tau protein (Price et al., 1998). However, these visible signs of pathology must be preceded by more subtle changes. The progression of the disease correlates with the spreading of the neurofibrillary tangles, whereas the amyloid plaques have a more generalized distribution (Braak and Braak, 1991; Arriagada et al., 1992). One of the earliest detectable signs is the loss of synapses and retrograde degeneration ("dying back") of neurons, which appears to be accompanied by a decay of intracellular transport (Terry, 1998). We suggest that the data presented here speak both to the amyloid and the tau dysfunction, linking them through the impairment of intracellular traffic.
APP appears to have neurotrophic functions and is carried by kinesin-driven vesicles along axons and dendrites (Ferreira et al., 1993; Amaratunga et al., 1995; Simons et al., 1995; Kaether et al., 2000). The anterograde movement is dependent on a kinesin motor; specifically, the cytosolic COOH-terminal tail of APP interacts directly with a kinesin light chain (Kamal et al., 2000). Despite their high mobility, the APP vesicles are effectively eliminated from the axon once tau becomes elevated, and APP is found concentrated in the cell body (Fig. 5 B, bottom). This is a prominent locus for the generation of the Aß peptides; in particular, both Aß40 and the more toxic variant Aß42 are generated in the trans-Golgi network (Xu et al., 1997). Thus, if the dwell time of APP were increased by a tau-dependent retardation of traffic one would expect an increase in the production of Aß with the known pathological consequences of aggregation and toxicity. This would be analogous to the increased accumulation of Aß by other treatments, which inhibit vesicle budding or transport (Greenfield et al., 1999). We note that the inhibition of APP transport by tau is a robust phenomenon that can be achieved in different cell models, for example, by transient transfection of tau in N2a cells stably transfected with APP or transfecting primary cortical or retinal ganglion neurons with APP and tau using AV vectors. In the latter case, the transfected proteins were tagged with fluorescent markers (YFP for APP and CFP for tau), which allows one to monitor vesicle movements and tau distributions by two-color live cell imaging. Several features were notable. (a) APP vesicles can be remarkably fast: whereas the known kinesins typically move 0.60.8 µm/s in vitro, APP vesicles in cortical and RGCs can reach up to 10 µm/s. (b) Fast transport occurs mainly in the anterograde direction: retrograde movement is generally slower, corresponding to the activity of dynein (Fig. 6 C). These results are in good agreement with recent findings by Kaether et al. (2000). (c) The majority of mobile particles (80%) moves anterogradely. This is the main reason for the strong bias of transport toward the synapse, independently of other transport characteristics. (d) Transport is nearly halted in both directions when tau is increased, that is, the number of moving particles becomes minute and the few mobile ones are retrograde (Fig. 6, B and C). (e) Remarkably, the transport infrastructure is much more resistant than traffic itself: microtubule tracks survive for many hours after mitochondria, peroxisomes, neurofilaments, APP vesicles, and others have deserted the neurite and accumulated in the cell body (Fig. 3), but eventually microtubules also disappear when the cell processes degenerate. On the basis of these data, one could imagine two potential causes for neuronal damage due to transport inhibition of APP. One is the depletion of APP from the synapse where it would loose its neurotrophic function (loss of function), and the second is its retention in the cell body where increased levels of Aß would be produced (gain of toxic function). It remains to be seen which of these is more important in the context of brain tissue.
With regard to tau protein, the most visible change in Alzheimer's disease and related tauopathies is the aggregation into neurofibrillary tangles, which is accompanied by an increase in the level of tau, hyperphosphorylation, and loss of microtubule binding in the affected neurons. These observations are usually interpreted within a hypothesis where the physiological function of tau (stabilization of microtubules) is disrupted due to excess phosphorylation; the unbound tau then is thought to aggregate in a pathological manner and obstructs the cell interior, and the microtubules disassemble so that axonal transport is disrupted (for review see Buee et al., 2000). This hypothesis is based on the view that tau's role is to promote neurite outgrowth by stabilizing microtubule bundles. Indeed, tau is upregulated during neuronal differentiation, adult isoforms are generated by alternative splicing, and phosphorylation then gradually decreases, all of which favor tighter binding of tau to microtubules (Mandell and Banker, 1996). Thus, the presence of tau would appear to be beneficial for the neuron. However, several observations do not fit into this scheme: tau-deficient transgenic mice show no major phenotype, since tau can be substituted by other cofactors (MAP1b) (Takei et al., 2000), mice overexpressing tau show transport defects even though microtubules are intact and tau aggregates are absent or minor (Ishihara et al., 1999; Götz et al., 2001; Lewis et al., 2001), and tau-overexpressing flies also show defects in neuronal traffic without evidence of tau aggregation (Wittmann et al., 2001). These observations argue that even "normal" tau may be detrimental when it becomes elevated.
How can we reconcile the requirement for tau in neurite outgrowth with the damage inflicted on transport? A hint comes from the fact that the tau to tubulin ratio is normally quite low, in the range of a few percent (Cleveland et al., 1977). This means that relatively few tau molecules may suffice to initiate a growing neurite, and the tau concentration required for microtubule stabilization along the axon may be low in the presence of other stabilizing factors (consistent with the tau knockout results [Harada et al., 1994]). Thus, in a physiological environment tau's effect on transport is negligible. However, it can become noticeable if tau becomes elevated in degenerating neurons as reported by Khatoon et al. (1992). The reason for the increase in tau is unclear, but one possibility is the neuron's tendency for reactive sprouting to counteract toxic challenges in an aging brain (Savaskan and Nitsch, 2001). Once traffic jams are initiated, they are exacerbated by the loss of mitochondria and peroxisomes from the axons, eventually speeding up degeneration. This could be the situation mimicked by the cell models described here. In conclusion, tau may be beneficial for the neuron at physiologically low concentrations but becomes detrimental if the concentration is elevated. This may simply happen in response to an adverse cellular environment or a consequence of a possible transcription factor activity of the cleaved COOH-terminal stub of APP (as proposed recently [Cao and Sudhof, 2001]). In addition, the cell could make use of the "friction" of tau to actively regulate intracellular transport. This could be achieved by regulating the concentration of tau or more flexibly by the kinases and phosphatases that determine tau's affinity for microtubules. Thus, pathological "hyperphosphorylation" could be the neuron's response to a traffic jam.
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Materials and methods |
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Fusion proteins
APP-EYFP.
EYFP cDNA (CLONTECH Laboratories, Inc.) without start codon was fused to the 3' end of human APP695 cDNA omitting the stop codon. The COOH-terminal part of APP was amplified from pSG5-hAPP695 (provided by B. De Strooper, Katholieke Universiteit Leuven, Leuven, Belgium). The fragment with HA tag was subcloned into a pCR Blunt II TOPO vector using the Zero Blunt TOPO cloning kit (Invitrogen). The resulting plasmid pCR Blunt II APP-HA was used for the insertion of EYFP DNA. The obtained fragment was subcloned into pCR Blunt II TOPO vector. The EYFP fragment was purified and cloned into pCR Blunt II APP-HA vector to obtain pCR Blunt II APP-HA-EYFP. The sequences generated by PCR were checked by sequencing.
ECFP-htau40.
The plasmid pECFP-htau40 was provided by J. Kupper, Max-Plank-Institute for Biochemistry, Martinsried, Germany. To allow the cloning of ECFP-htau40 into an AV vector, the SalI restriction site was introduced at the 5' end of the ECFP-htau40 cDNA using QuikChange site-directed mutagenesis (Stratagene).
AV vectors encoding fluorescent fusion proteins
Recombinant AVs were generated following He et al. (1998). Briefly, the SalI-XbaI fragment that contained the CFP-htau40 cDNA and the HindIII-XbaI fragment that contained the APP-YFP cDNA were subcloned into the respective restriction sites of the pShuttle-CMV vector. The resulting plasmids were linearized with PmeI and cotransfected with the pAdEasy 1 vector into Escherichia coli BJ 5183 for homologous recombination. Plasmid DNA was amplified in E. coli DH10B and digested with PacI to cut out the entire recombinant adenoviral DNA, which was transfected into 911 cells, using Lipofectamine transfection (Life Technologies). The generation of the viruses in the 911 cells was monitored by fluorescence microscopy using an Axioplan fluorescence microscope (ZEISS). The cells were harvested, resuspended in PBS, and lysed by three freeze-thaw cycles. Cellular debris and nuclei were removed by centrifugation, and the virus suspension was purified by two CsCl gradient centrifugations. CsCl was removed by gel filtration and equilibrated in storage buffer (10 mM Tris/Cl, 135 mM NaCl, 3 mM KCl, 1 mM MgCl2, 10% glycerol, pH 8.0).
Transfection of primary cortex and retinal ganglion neurons
Cultures of E18 (rat) or E15 (mouse) hippocampal neurons were prepared according to Banker and Goslin (1998). Cells were plated in HBSS buffer (Biochrom) at a density of 7 x 104 cells per cm2 on a glass surface coated with poly-L-lysine (0.01% in 100 mM borate buffer, pH 8.5) and fibronectin (0.001% in HBSS; Life Technologies). To cultivate the neurons for live observation, 4.3 cm2 Lab-Tek chambers were used (Nunc) and coated as described above. Cells were transfected with tau either using a HSV vector or an AV vector. Cells were transfected between days 4 and 8 in culture. The HSV virus carrying the gene of htau40 was provided by Dr. R. Brandt, University of Heidelberg, Heidelberg, Germany. 10 µl virus suspension was added per 1.5 x 105 cells and incubated for 48 h. Then, the cells were fixed for immunofluorescence. For adenoviral transfection of APP-YFP or CFP-htau40, a 100-fold multiplicity of infection (multiplicity of infection of 100, 3 x 107 infectious particles) was applied to primary neurons. In the case of double transfections, 3 x 107 infectious particles of each recombinant AV were added. After 4 h incubation, the viral suspensions were removed. Vesicle movement was observed by confocal microscopy 2448 h after transfection.
RGCs were prepared from white leghorn chicken eyes at embryonic day 7. Glass bottom dishes were coated overnight with 4 µg/ml laminin (Sigma-Aldrich), washed with sterile H2O, and dried. Retinae were mounted on nitrocellulose filter as described previously (Walter et al., 1987) and cut with a scalpel into 12-mm-wide stripes. Retinal explant was placed into the dish, and DME-F12 media (GIBCO BRL) with 10% FCS and 0.4% methyl cellulose was added. Explants were then cultured for 24 h at 37°C, 5% CO2 and 100% relative humidity prior viral transfection. Transfection and observation was done as above.
For staining of mitochondria in RGCs, the medium was removed 2448 h after explantation of the retina and replaced with medium containing MitoTracker red 589 (Molecular Probes) at a final concentration of 12 nM or MitoTracker green FM (Molecular Probes) at a final concentration of 100 nM. Cells were incubated overnight under growth conditions. Then, the MitoTracker solution was replaced with fresh prewarmed medium, and movement of mitochondria was observed. For double labeling with CFP-tau AV, virus was added and removed after 5 h of incubation. MitoTracker red solution was added overnight. The expression of tau and the movement of mitochondria were observed by confocal microscopy 2448 h after transfection.
Antibodies and dyes
Rat monoclonal antitubulin antibody YL1/2 and mouse monoclonal antibody DM1A were purchased from Serotec and Sigma-Aldrich. Polyclonal rabbit antitau antibody K9JA was from Dako, polyclonal rabbit anti-PMP69 antibody for peroxisomes was a gift from Dr. W. Just (University of Heidelberg, Heidelberg, Germany). All fluorescently (TRITC, FITC, and AMCA) labeled secondary antibodies were from DIANOVA. Fluorescent dyes MitoTracker red and rhodamine-labeled WGA were purchased from Molecular Probes. The monoclonal mouse antibody SMI32 (Chemicon) was used for the detection of unphosphorylated neurofilaments. The monoclonal tag antibodies from mouse against HA tag (12CA5) and myc tag were obtained from Roche Diagnostics and Invitrogen. Polyclonal antibody B5 (5313) against human APP (residues 444592) was a gift from Dr. C. Haass (University of München, München, Germany), and monoclonal antibody 6E10 was from Senetek.
Immunofluorescence
Neurons and neuroblastoma cells were fixed in methanol or 2% paraformaldehyde and incubated with antibodies. Cells were examined with an Axioplan fluorescence microscope (ZEISS) equipped with an 100x oil-immersion objective and filters optimized for triple label experiments (FITC, TRITC, and AMCA fluorescence). Pictures were taken with a cooled CCD camera (Visitron) and analyzed using the MetaMorph software package.
Quantification of vesicles, organelles, and neurofilament protein in N2a cells
Peroxisomes were stained with polyclonal anti-PMP69 antibody (45 min, 1:200). Golgi-derived vesicles were stained with 10 µM rhodamine-labeled WGA (Molecular Probes) for 5 min after methanol fixation. Mitochondria were visualized by adding 400 nM MitoTracker red (Molecular Probes) for 30 min in media at 37°C before fixation. Vesicles carrying APP were stained either with monoclonal anti-myc antibody (1:200; Invitrogen) or polyclonal anti-APP antibody (1:300). After fixation and immunofluorescence labeling, the pictures were recorded and a defined area (2030 µm in length for N2a cells and 50 µm for neurons) beginning at the proximal neurite was circumscribed manually with the MetaMorph drawing tools. Usually >100 cells were recorded per experiment. Afterwards, signals of vesicles and organelles above the background threshold were visualized and counted using the threshold function of MetaMorph. Both in tau- and mock-transfected cells we quantified and compared commensurate areas.
Live cell light microscopy
For visualizing tau, primary cortex neurons (E18) were transfected at day 4 in culture with CFP-tau40 AV (or control virus) at multiplicity of infection of 30 for 24 h. For tracking APP vesicles, transfection with APP-YFP AV at multiplicity of infection of 100 was done at day 8 in culture for 4 h, and analysis was done 24 h later. For observing vesicles labeled with fluorescent lectins, cells at day 4 in culture were incubated with rhodamine-WGA (Molecular Probes) at a final concentration of 4 µg/ml for 15 min. Vesicles were tracked with an Axioplan fluorescence microscope (ZEISS). To visualize CFP fluorescence of CFP-htau40, the FITC filter set was employed; WGA-rhodaminelabeled vesicles were observed by using the TRITC filter set. The object plane was kept at 37°C by air heating. APP-YFP vesicles were tracked with a LSM 510 confocal microscope (ZEISS), equipped with an 63x oil-immersion objective, beam path, and laser settings for YFP and CFP fluorescence and a 37°C air-heated object plane. Image analysis was performed with LSM 510 software.
Response of cells to oxidative stress
N2a cells differentiated on coverslips were incubated in 150 µM H2O2 in MEM medium for 40 min or more. Cells were fixed in methanol, stained for immunofluorescence with antibodies K9JA (for tau) and YL1/2 (for tubulin), and the number and length of neurites was recorded. Protection against H2O2 was done by adding catalase (0.02 U/µl; Sigma-Aldrich) immediately after H2O2. Catalase was inhibited by 3-AT (Sigma-Aldrich) in concentrations of 0.1, 1, and 10 mM and incubating for 30 or 60 min before the addition of H2O2. The integrity of mitochondria was measured by the 3-(4,5-dimethylthiazol-2yl)-2,5-diphenyltetrazolium bromide test, and the viability of cells was measured by the trypan blue assay (Mattson et al., 1995). The capacity of the glutathione system for peroxide detoxification was determined following Dringen et al. (1999).
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Footnotes |
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Acknowledgments |
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This research was supported in part by the Deutsche Forschungsgemeinschaft.
Submitted: 13 August 2001
Revised: 21 January 2002
Accepted: 1 February 2002
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References |
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