HSPGs Mediate Tau Fibril Binding and Uptake.

Tau and TAT contain heparin-binding domains, and it is established that TAT enters cells via HSPG-mediated macropinocytosis (20–25). Given the extensive colocalization between tau fibrils and TAT after endocytosis, we hypothesized that HSPGs might also mediate cellular binding and internalization of tau aggregates. HSPGs are transmembrane and lipid-anchored cell surface receptors that interact with a variety of ligands. They are extensively sulfated, a crucial posttranslational modification. These sulfated moieties permit electrostatic interactions between the sugar polymers and short basic amino acid stretches in heparin-binding proteins.

To determine if tau fibrils and HSPGs colocalize, we treated C17.2 cells with 50 nM tau RD-488 fibrils for 60 min, removed extracellular fibrils by using trypsin, and allowed the cells to recover before immunostaining for HSPGs. We found that the HSPGs enveloped the RD-488 puncta (Fig. 2A), consistent with the tau fibrils within macropinosomes that we had observed by EM. We next investigated whether HSPGs mediate binding of tau fibrils to the cell surface. We incubated C17.2 cells with RD-488 fibrils at 4 °C in the presence or absence of HSPG inhibitors and imaged cells by using confocal microscopy. At this restrictive temperature tau fibrils are not internalized, but instead adhere to the cell membrane. We then tested two chemical inhibitors of tau/HSPG interactions. Sodium chlorate, a metabolic inhibitor, prevents proper sulfation of HSPGs. Conversely, heparin, a glycosaminoglycan, competitively inhibits tau binding to HSPGs. Both compounds dose-dependently blocked tau RD binding to the cell surface, as determined by confocal microscopy (Fig. 2B) and flow cytometry (Fig. 2 C and D). The effective concentrations were consistent with those reported previously by others (14, 26). Taken together, this work suggests that HSPGs mediate tau fibril binding to the cell surface.

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Fig. 2.

HSPGs mediate binding of tau RD fibrils to C17.2 cells. (A) Tau RD-488 fibrils colocalize with anti-HSPG antibody (10E4). (Scale bar: 10 µm.) (B) At 4 °C, tau RD-546 fibrils bind to the plasma membrane but are not internalized, a process that is inhibited by pretreatment with heparin and chlorate. HSPG inhibition abolishes the association between tau RD fibrils and C17.2 cells observed by confocal microscopy. (C and D) Flow cytometry quantification of tau fibril binding to the cell membrane in the presence of chlorate or heparin. Cells were treated with 50 nM tau RD-488 fibrils at 4 °C for 1 h. A total of 25,000 cells were analyzed for each condition, which was run in triplicate. Error bars show SEM.

We further examined the role of HSPGs for tau fibril internalization by using automated high-content microscopy (IN Cell Analyzer 1000; GE Healthcare). This method permits quantification of the percentage of cells within a population that are positive for labeled tau aggregates, and the average number of tau aggregates per cell (among positive cells). This latter quantification allows determination of the “tau aggregate burden” in tau-positive cells. We treated C17.2 cells with chlorate or heparin before exposure to 50 nM RD-488 fibrils for 3 h. After a 5-min trypsin treatment, cells were replated and imaged by confocal or automated microscopy (Fig. 3 A–D). Heparin and chlorate dose-dependently decreased the internalization of tau fibrils within the same concentration ranges that blocked cell surface binding. They also reduced the average number of tau aggregates per cell, demonstrating that inhibition of tau binding to HSPGs decreases the subsequent total tau burden in aggregate-positive cells (Fig. 3 B and C).

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Fig. 3.

HSPGs mediate tau RD fibril uptake. (A) At 37 °C, tau RD-546 fibrils are internalized by the cell, a process that is inhibited by chlorate and heparin. (Scale bar: 20 µm.) (B and C) Automated microscopy analysis of tau fibril internalization in the presence of chlorate or heparin. Cells were treated with 50 nM tau RD-488 fibrils and chlorate or heparin at 37 °C for 3 h and trypsinized before imaging. The left y-axis (blue) depicts percentage of positive cells and the right y-axis (red) depicts the average number of tau aggregates per cell. Approximately 40,000 cells were analyzed for each condition, run in quadruplicate. (D) Percent cells positive for tau fibril internalization in the presence of heparinase III or chondroitinase AC as measured by automated microscopy analysis. Approximately 40,000 cells were analyzed for each condition, run in duplicate. (E) HSPG inhibition does not affect clathrin-mediated transferrin endocytosis. Internalization of Tfn-488 (25 µg/mL) was unaltered in the presence of chlorate (63.2 mM), heparin (200 µg/mL), heparinase III, or chondroitinase AC (10² IU/mL) as measured by flow cytometry mean fluorescence intensity. A total of 25,000 cells were analyzed for each condition in duplicate. The nontreated group (NT) received no inhibitor, and data reflect uptake relative to this group. (Error bars: B, C, and E, SEM; D, range.)

Next we used enzymatic modification of HSPGs to investigate their role in fibril uptake. We pretreated C17.2 cells with increasing concentrations of heparinase III or chondroitinase AC to specifically degrade cell surface heparan or chondroitin sulfates, respectively. Heparinase III dose-dependently decreased the percentage of tau aggregate-positive cells and total aggregates per cell, whereas chondroitinase AC had no effect alone or in combination with heparinase III (Fig. 3D and Fig. S4). Thus, HSPGs, and not chondroitin sulfates, are critical for tau fibril uptake. To rule out nonspecific effects, we treated cells for 90 min with 25 µg/mL transferrin, a substrate of clathrin-mediated endocytosis, covalently labeled with Alexa Fluor-488 (i.e., Tfn-488). Heparin, chlorate, heparinase III, and chondroitinase AC had no effect on Tfn-488 uptake (Fig. 3E). Thus, tau RD fibril binding to HSPGs is critical for uptake by macropinocytosis in C17.2 cells.

Full-Length Tau Requires HSPGs to Enter Neurons.

Tau RD is commonly used because of its efficient fibril formation. However, we wanted to determine whether full-length (FL) tau fibril internalization is also mediated by HSPGs. We applied 50 nM FL tau-488 fibrils to C17.2 cells in the presence or absence of chlorate and heparin. FL tau fibril uptake was approximately two- to fourfold more sensitive to HSPG inhibition than RD fibrils (Fig. 4 A and B). Thus, FL tau aggregates also require HSPGs for cellular internalization.

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Fig. 4.

HSPGs mediate internalization of FL tau fibrils in C17.2 cells and primary hippocampal neurons. (A and B) Automated microscopy analysis of FL tau-488 fibril internalization into C17.2 cells in the presence of chlorate or heparin. Cells were treated with 50 nM FL tau-488 fibrils at 37 °C for 3 h and trypsinized before imaging. Approximately 40,000 cells were analyzed for each condition and run in quadruplicate. The left y-axis (blue) depicts percentage of positive cells and the right y axis (red) depicts the average number of tau aggregates per cell. (C) Lentivirus encoding Ext1 shRNA reduces Ext1 transcript in C17.2 cells by quantitative PCR relative to GAPDH (n = 3). (D) Knockdown of murine Ext1 by shRNA, but not luciferase shRNA, reduces the internalization of FL tau-488 fibrils into primary hippocampal neurons by mean fluorescence intensity measurements. The nontreated (NT) group received no shRNA (n = 9). Error bars show SEM. (E) Knockdown of murine Ext1 by shRNA does not reduce Tfn-488 internalization into primary hippocampal neurons (n = 4, error bars show SEM; ***P < 0.001 by one-way ANOVA).

To test the involvement of HSPGs in primary hippocampal neurons, we knocked down a key HSPG biosynthesis enzyme, Ext1, before exposure to FL tau aggregates. Although critical to synthesis of HSPGs, Ext1 is not rate-limiting in their production (27). Cells deficient in Ext1 cannot produce HSPGs, although they can produce other glycosaminoglycans (28). We first identified potent shRNAs against Ext1 by screening five different constructs in C17.2 cells by quantitative real-time (RT)-PCR and selecting a lentivirus that achieved ~90% Ext1 knockdown (Fig. 4C). Next, we transduced primary neurons and used flow cytometry to quantify aggregate uptake based on mean cell fluorescence intensity. We observed a 43% reduction of 50 nM FL tau fibril uptake relative to neurons treated with control luciferase shRNA (Fig. 4D). Ext1 knockdown did not affect Tfn-488 uptake (Fig. 4E), confirming that inhibition of tau fibril uptake in neurons is specific to the HSPG pathway, as in other cells. In summary, pharmacological, enzymatic, and genetic manipulations implicate HSPGs as key receptors for recombinant FL tau and RD fibrils in neural cell lines and primary neurons alike.

HSPG Inhibition Blocks Aggregate Propagation.

Although HSPGs mediate virtually all detectable tau uptake, it remained possible that other modes of cell entry could permit propagation of aggregation into cells. Thus, we used previously developed methods to test whether blockade of HSPGs would inhibit seeded aggregation and transcellular propagation of RD fibrils. We have previously found that fusion of RD containing a disease-associated mutation (K280) to CFP or YFP [i.e., RD(K)CFP/YFP] allows quantification of intracellular aggregation based on FRET (12). Incubation of HEK293 cells expressing tau RD-CFP/YFP with 50 nM tau RD fibrils increased intracellular aggregation as expected. However, pretreatment with heparin or heparinase III decreased induction of intracellular aggregation by recombinant fibrils (Fig. 5A) without affecting intrinsic intracellular aggregation (Fig. 5B).

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Fig. 5.

Inhibition of HSPGs blocks seeded aggregation and transcellular propagation of tau aggregation. (A) Heparin (Hep) and heparinase (h’ase) inhibit intracellular seeding by recombinant tau RD fibrils in a cell-based FRET assay. HEK293 cells cotransfected with tau RD(ΔK)-CFP/YFP were pretreated with heparinase (0.01 IU/mL) for 3 h before treatment with tau fibrils or treated with tau RD fibrils plus vehicle or heparin (6 µg/mL) for 24 h before reading FRET measurements on a plate reader. The FRET signal is shown as a percentage relative to the vehicle treated group. (B) Neither heparin nor heparinase affect cell-autonomous tau aggregation. Cells expressing RD(ΔK)-CFP/YFP were treated with vehicle or heparin (6 µg/mL) for 24 h or heparinase (0.01 IU/mL) for 27 h. A value of 100% represents baseline aggregation signal for the vehicle-treated group. (C) Heparin and heparinase block transcellular propagation. HEK293 cells expressing RD(ΔK)-CFP/YFP were cocultured with an equivalent number of cells expressing tau RD(LM)-HA for 48 h to monitor transcellular propagation of tau protein misfolding. Heparin dose dependently inhibited transcellular aggregate propagation, as did 0.01IU/mL of heparinase. The FRET signal is shown as a percentage relative to the vehicle-treated group. Error bars show SEM from four biological replicates per experiment for heparin and from six biological replicates per experiment for heparinase (***P < 0.001, **P < 0.01, Student t test).

No effective assay exists to measure transcellular propagation of tau aggregates in primary neurons. Thus, to test the role of HSPGs in this process, we used a HEK293 cell coculture assay in which donor cells express untagged tau RD that contains two disease-associated mutations, P301L and V337M [termed RD(LM)], that greatly increases its intrinsic aggregation propensity. RD(LM) aggregates are released by the donor cells into the media, taken up by aggregation-sensor cells expressing RD(K)-CFP/YFP, and nucleate further tau aggregation, as measured by FRET (12). We cultured donor cells expressing tau RD(LM) with an equal number of acceptor cells expressing RD(ΔK)-CFP/YFP. After 48 h, we observed a significant increase in FRET, indicating propagation of aggregation from donor to acceptor cells. Heparin titration and heparinase III pretreatment blocked the induced aggregation, demonstrating inhibition of transcellular propagation (Fig. 5C). Thus, HSPGs mediate propagation of aggregation from the outside to the inside of the cell as well as between cells.

HSPGs Are Required for FL Tau Fibril Entry in Vivo.

We next investigated the role of HSPGs as a receptor for tau fibrils in vivo. Heparin has strong anticoagulant properties that preclude its use within mouse brain. On the contrary, heparin mimetics that do not promote bleeding are promising therapeutic agents for wound healing and for blocking prion infectivity (29, 30). We screened a selection of synthetic heparin mimetics for activity in blocking tau aggregate uptake, and selected F6 for its potency and lack of anticoagulation activity (Fig. S5). We first tested F6 as a potential inhibitor of tau fibril internalization, seeded aggregation, and transcellular propagation in cell culture. We used polymeric dextran (PD), a nonsulfated heparin mimetic, as a negative control (Fig. S5). Based on automated microscopy analysis of C17.2 cells, F6 potently inhibited recombinant tau fibril internalization, whereas PD had no effect (Fig. 6A). In HEK293 cells, F6 also blocked seeded aggregation and transcellular propagation with an IC₅₀ similar to that of heparin (Fig. 6 B and C), and it had no effect on cell-autonomous tau aggregation (Fig. 6D).

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Fig. 6.

Heparin mimetic F6 inhibits tau fibril uptake and induction of misfolding. (A) F6 inhibits tau RD fibril internalization into C17.2 cells whereas PD has no effect. Cells were treated with 50 nM tau RD-AF488 fibrils at 37 °C for 3 h and trypsinized before replating and imaging with automated microscopy. Approximately 40,000 cells were analyzed for each condition and run in quadruplicate. (B) F6 (6 µg/mL) and heparin (6 µg/mL) equivalently inhibit seeded aggregation in a FRET-based assay. (C) F6 and heparin each block transcellular propagation between cells expressing tau RD(LM) and cells expressing RD(K)-CFP/YFP. (B and C) FRET signal is shown as a percentage relative to the vehicle treated group. (D) F6 does not inhibit cell-autonomous tau aggregation. A value of 100% represents baseline aggregation signal of the vehicle-treated group. Error bars show SEM from four biological replicates per experiment (***P < 0.001, Student t test).

To test the effect of blocking tau fibril binding to HSPGs in vivo, we coinjected 472 ng of FL tau-488 fibrils with 1 µg of F6 or PD into the cortex of 5-mo-old WT mice (Fig. 7A). After 48 h, mice were killed and cortical neurons in brain sections were identified by staining with anti-NeuN antibody. In mice coinjected with tau fibrils and PD, we observed a large number of tau fibril-positive neurons. All exhibited punctate morphology similar to our cell culture model. In mice coinjected with tau fibrils and F6, only a small fraction of neurons scored positive for tau aggregates, indicating that F6 blocked neuronal uptake (Fig. 7B). To rule out the possibility that F6 inhibited neuronal tau fibril entry as a result of cellular toxicity or other off-target effects, we injected 5 µg of Tfn-488 and PD or F6 in conjunction with FL tau fibrils labeled with Alexa Fluor-647. In mice receiving PD treatment, neurons near the injection site scored positive for tau fibrils and Tfn-488, indicating efficient uptake (Fig. 7C and Movie S1). However, in mice receiving F6 treatment, the majority of neurons had only internalized Tfn-488 (Fig. 7D). Image analysis (ImageJ; National Institutes of Health) indicated 64% of neurons were positive for tau fibrils when coinjected with PD, whereas only 30% were positive when coinjected with F6 (Fig. 7E). Similarly, F6 significantly reduced the mean fluorescence intensity for tau fibrils in neurons per animal by 50%, but not the mean fluorescence intensity of transferrin (Fig. 7 F and G). Thus, FL tau fibril uptake into neurons in vivo also requires binding to HSPGs.

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Fig. 7.

HSPGs mediate internalization of FL tau fibrils in vivo. (A) Schematic representation of stereotactic injection site. (B) Injection of FL tau fibrils with heparin mimetic F6 leads to a reduction of fibril internalization by cortical neurons. Mice (n = 3 per group) were injected with 472 ng of FL tau-488 fibrils (green) and 1 µg of PD or F6. Brain sections were immunostained with anti-NeuN antibody to label neurons (red) and counterstained with DAPI (blue). Arrowheads designate FL tau fibrils associated with neurons. (Scale bar: 10 µm.) (C and D) Heparin mimetic F6 blocks FL tau fibril uptake, but not Tfn-488. Mice (n = 4 per group) were injected with 472 ng of FL tau-647 fibrils (white), 5 µg of Tfn-488 (green), and 1 µg of PD or F6. Injection of tau fibrils and transferrin with PD resulted in entry of both proteins into neurons, whereas coinjection with F6 resulted in the selective inhibition of tau fibril entry. The grayscale images are single channel fluorescence images corresponding to the multichannel images labeled as “merged.” Arrowheads designate FL tau fibrils; long arrows designate transferrin. (E) Quantification of tau-positive neurons from experiment C and D. A total of 494 and 660 neurons were counted and averaged in the PD and F6 conditions, respectively, from four male mice per cohort. Error bars show SEM (**P < 0.01, Student t test). (F and G) Quantification of neuronal fluorescence intensity from experiment C and D. Mean fluorescence intensity of tau fibrils and transferrin were calculated per neuron and averaged from individual mice, each represented by a single data point (P = 0.028, Mann–Whitney U test, two-tailed exact significance) comparing treatment effects on tau fibril uptake.

Aggregate Uptake by Macropinocytosis.

Based on pharmacologic studies and colocalization with fluid phase markers, macropinocytosis was previously suggested as the mechanism for cell uptake of SOD1 (36). Likewise, macropinocytosis and HSPGs have been previously implicated in prion protein uptake (14, 37). In this study, we investigated the molecular basis of uptake with a multifaceted approach. We initially used EM to directly image uptake events. This indicated that the cell internalizes tau fibrils via dynamic membrane rearrangement and forms large endocytic vesicles consistent with macropinosomes. Indeed, extracellular tau fibrils stimulated fluid-phase endocytosis in a dose-dependent fashion. We subsequently observed colocalization of aggregates with labeled TAT peptide, which is known to enter cells via macropinocytosis (17–19). Other mechanisms of cell uptake have been proposed for protein aggregates, including dynamin-dependent endocytosis (31, 38) and direct permeabilization of the cell membrane (35, 39), neither of which appear to contribute to tau aggregate uptake in our studies.

Aggregate propagation requires direct contact between the macropinosome-encapsulated seed and the cytosolic monomer. However, it remains unclear how a tau aggregate traverses the membrane barrier of a macropinosome. Of note, viruses and cell-penetrating peptides such as TAT exploit macropinosomes to enter the cytosol (17, 40, 41), also by unclear mechanisms. Macropinosomes are inherently leaky in comparison with other types of endosomes (42), which may allow contents to escape, and thus permit fibrils to seed aggregation in the cytosol. We hypothesize that fibrils may actually promote their own escape from the vesicular compartment based on destabilization of the lipid bilayer. This remains to be tested, but has been proposed as a property of tau oligomers (43).

HSPGs Mediate Tau Uptake.

HSPGs participate in numerous cell surface interactions and serve as a primary receptor for macropinocytosis (19). Heparin-binding proteins interact with HSPGs on the cell surface, triggering internalization. In addition to our work with tau and α-synuclein, this mechanism has previously been demonstrated for infectious prion protein and Aβ monomer (14, 26). Binding to HSPGs requires a heparin/heparan sulfate-binding domain consisting of a stretch of positively charged lysines or arginines on the ligand. Prion protein, β-amyloid, tau, and α-synuclein all have putative heparin-binding domains (25, 44–46).

We have found a critical role for HSPGs in selectively binding and internalizing aggregated tau. We picked a tau aggregate concentration for our studies (50 nM monomer equivalent) that roughly approximates physiologic levels, based on our best estimates. Currently, it is impossible to quantify tau aggregate concentration in the brain interstitial fluid (ISF) of humans or tauopathy animal models. Our recent work used a microdialysis technique that measures only tau monomer in brain ISF. We estimated a concentration of ~250 ng/mL (~17 nM, assuming FL tau) in P301S human tau transgenic mice (47), but total ISF tau levels (including aggregates) may in fact be higher. In this study, we potently inhibited tau aggregate binding, uptake, and seeding of intracellular aggregation with multiple compounds specific to this pathway: heparinase III, an enzyme that degrades cell surface HSPGs; heparin, which blocks binding to HSPGs; and chlorate, a metabolic inhibitor of sulfation. We also tested this pathway by knockdown of Ext1, which is involved in elongation of heparan sulfate chains, and is critical to HSPG synthesis (28, 48). Genetic knockdown of Ext1 has been used extensively in cell culture to probe the involvement of HSPGs, and manipulations of this enzyme do not reduce the synthesis of other proteoglycan subtypes (14, 26, 28, 49). Ext1 knockdown inhibited the internalization of tau aggregates into primary hippocampal neurons without affecting clathrin-mediated uptake of transferrin. Thus, pharmacologic and genetic interventions demonstrate that HSPGs are critical mediators of tau fibril internalization. Tau binding to HSPGs was also required for transcellular propagation in cell culture, as introduction to the media of heparin or the heparin mimetic F6 prevented recombinant and cell-derived tau fibrils from nucleating further aggregation. Finally, we confirmed the involvement of tau/HSPG binding in vivo by stereotactic injection of tau fibrils into the cortex of WT mice. F6 blocked aggregate uptake into neurons without affecting transferrin uptake. Taken together, this work suggests that recombinant tau fibrils and tau aggregates produced within a cell use HSPG binding to seed further aggregation within recipient cells.

Tau and α-Synuclein Use Similar Modes of Cell Uptake.

Our data suggest strongly that tau and α-synuclein fibrils use the same mechanism of uptake into cells. Fibrils of these proteins each colocalize with HSPGs, TAT peptide, and with each other when analyzed by confocal microscopy. Further, both are equivalently sensitive to heparin and chlorate inhibition. These mechanisms of uptake are specific, because Htt(Q50) fibrils failed to colocalize with tau, α-synuclein, or TAT, and were not sensitive to heparin or chlorate. Htt(Q50) fibrils nonetheless seeded aggregation of intracellular Htt(Q25)CFP/YFP, indicating that at least two modes exist for propagation of seeding activity from the outside to the inside of the cell. The primary route of Htt internalization remains unclear, although undefined cell surface proteins have been implicated (34). Unlike tau and α-synuclein, Htt exon 1 does not feature a clear heparin-binding domain. Indeed, at least one study has suggested that Htt fibrils directly penetrate the cell membrane to enter the cytosol before being sequestered in aggresomes (35). These distinct uptake mechanisms have obvious implications for further studies and for consideration of therapeutic targets.

Immunofluorescence and Microscopy.

All cells were grown on µ-Slides (Ibidi) for imaging by microscopy. For propidium iodide staining, cells were coincubated with tau RD-488 fibrils, tau monomer, tau buffer, or 5% (vol/vol) ethanol and propidium iodide (5 µg/mL; Sigma) for 3 h, washed twice with PBS solution, and examined by confocal microscopy. For clathrin heavy chain (CHC) and caveolin-1 immunofluorescence, cells were treated with 150 nM tau RD-488 fibrils for 1 h, extensively washed with PBS solution, and fixed with 4% PFA. Cells were stained with α-CHC mouse monoclonal antibody (1:300; Covance) or caveolin-1 mouse monoclonal antibody (1:500; Santa Cruz), followed by secondary labeling with Alexa Fluor-546 goat anti-mouse (1:1,000; Molecular Probes) and DAPI. For phalloidin staining, cells were incubated with 150 nM tau RD-488 or Alexa Fluor 488-containing buffer for the time indicated, washed, fixed as described earlier, and stained with 33nM rhodamine-phalloidin (Invitrogen) for 20 min. For colocalization studies between tau RD and the TAT peptide, 150 nM of tau RD-488 fibrils and 5 µM TAT-TAMRA peptide (residues 47–57; AnaSpec) were coadministered to C17.2 cells for 90 min. Cells were then extensively washed with media. To quench extracellular fluorescence, trypan blue (0.05% in PBS solution; Sigma) was added to the cells for 15 s and removed. Cells were washed twice and immediately imaged live in phenol red-free DMEM (1% FBS) by using confocal microscopy with an environmental chamber (5% CO₂ at 37 °C). For colocalization studies between tau RD and HSPGs, 150 nM of tau RD-488 fibrils were administered to C17.2 cells for 90 min. Cells were trypsinized for 5 min, followed by replating on Ibidi µ-Slides. Cells were allowed to recover for 3.5 h before fixation in 4% PFA. HSPGs were immunostained with anti-10E4 antibody (1:100; US Biological), followed by secondary labeling with Alexa Fluor-546 goat anti-mouse (1:1,000; Molecular Probes), and images were captured by using confocal microscopy.

For immunohistochemistry of mouse brain sections, sections were immunostained with α-NeuN antibody (1:200; Millipore) followed by secondary labeling with Alexa Fluor-546 goat anti-mouse and DAPI. Images for quantification were collected such that only brain regions 150 to 300 µM away (medial/lateral) from the injection sites were included. This was done to avoid regions saturated or void of injected material. In the dorsal/ventral axis, images were collected along the entire length of the injection tract. Images were acquired from five sections per animal including the midpoint of the injection site, ±25 µM and ±50 µM (rostral/caudal) from the injection site. Quantification of images was conducted by using ImageJ (National Institutes of Health). For quantification of percent neurons positive for tau aggregates, regions of interest (ROIs) were drawn around cells based on NeuN staining. The fluorescence signal threshold was applied equally to all sections, and cells were counted as positive if a fluorescence puncta fell within the boundary of the ROI. For quantification of mean fluorescent intensity, cells were measured by using the same ROIs as described earlier without fluorescence signal threshold. All cells were counted that had a discernible DAPI and NeuN stain.

The 3D projection was created by acquiring a series of confocal images of a neuron at an interval of 0.2 µM sections over a span of 3 µM. VolViewer software was used to create a ray sum projection from individual images, and Adobe After Effects was used to animate the projection.

Flow Cytometry.

C17.2 cells were plated at 25,000 cells per well in a 24-well plate. The next day, cells were pretreated with the following drugs for 30 min: cytochalasin D (1 µM; Sigma), latrunculin A (3 µM; Invitrogen), amiloride hydrochloride hydrate (1 mM; Sigma), rottlerin (30 µM; Sigma), and Dynasore (80 µM; Sigma). Cells were next treated with tau RD-488 fibrils or Alexa Fluor-488–containing buffer for 3 h. Cells were harvested with 0.25% trypsin for 5 min and resuspended in HBSS plus 1% FBS and 1 mM EDTA before flow cytometry. Cells were counted in a FACScan flow cytometer (BD Biosciences) or MACSQuant VYB (Miltenyi Biotec). Each experiment was conducted three times, and 25,000 cells were counted in each individual experiment. For tau fibril-binding experiments, cells or tau fibrils were pretreated separately with the indicated concentrations of sodium chlorate or heparin (Sigma), respectively, for 12 h. Cells were then equilibrated for 4 °C for 15 min before the addition of tau fibrils for 60 min at 4 °C, suspended in cell dissociation solution (Sigma), and subjected to flow cytometry.

Automated Microscopy Analysis.

After RD-488 fibril or dextran fluorescein (70 kD, anionic, lysine fixable) treatment, cells were trypsinized for 5 min, replated on a 96-well plate, allowed to recover for 3.5 h, and fixed. To identify cell boundaries, cells were stained with 10 µg/mL of wheat germ agglutinin labeled with Alexa Fluor-647 followed by DNA staining with DAPI. Cells and intracellular puncta were visualized by automated microscopy by using an InCell Analyzer 1000 high-content microscope fitted with a 10× objective (GE Healthcare). ROIs were defined by user-assigned size and fluorescence intensity thresholds and were quantified with the Multi-Target Analysis Module of the IN Cell Analyzer 1000 Workstation 3.7 analysis software. For tau fibril internalization experiments, cells were pretreated with chlorate for 12 h or heparinase III or chondroitinase AC (Ibex) for 3 h before fibril treatment. Similarly, tau fibrils were pretreated with heparin for 12 h before addition of the heparin–fibril complexes.

Cell Culture and Transfections.

HEK293 cells were cultured in DMEM supplemented with 10% FBS, 100 µg/mL penicillin, and 100 µg/mL streptomycin. Cultures were maintained in a humidified atmosphere of 5% CO₂ at 37 °C. For transient transfections, cells plated in complete medium were transfected by using Lipofectamine 2000 (Invitrogen) and 600 ng of appropriate DNA constructs according to the manufacturer’s recommendations and harvested 24 h later for further analyses. To culture primary neurons, the cortex or hippocampus of embryonic day 18.5 mouse embryos was isolated and digested with 2 mg/mL papain and 0.1% DNase I. Neurons in Neurobasal media containing serum-free B-27 and GlutaMAX were then seeded on culture plates precoated with 10 μg/mL poly-d-lysine and 1.5 μg/cm² laminin. Medium was changed once every 4 d until neurons were ready for use.

FRET Assays.

Total tau aggregation was measured by an assay based on FRET between RD(K)-CFP/YFP, which has been described previously (12). The FRET calculations provide a reproducible measure of intracellular aggregation, and take into account the relative amounts of donor and acceptor proteins (53–55). For seeded aggregation experiments, HEK293 cells were transfected with 150 ng of tau RD(ΔK)-CFP and 450 ng of tau RD(ΔK)-YFP. Twenty-four hours later, cells were split into a 96-well plate and allowed to recover overnight. Tau RD fibrils (50 nM) were preincubated with heparin or F6 at the indicated concentrations for 12 h and then applied to cells for 24 h to induce intracellular aggregation. Heparinase III was applied directly to the cell media for 3 h before the application of tau RD fibrils for 24 h. For transcellular propagation experiments, acceptor cells were transfected as described earlier; donor cells were transfected with 600 ng of tau RD(LM)-HA. After 24 h, cells were split into 96-well plates at equal percentages of donor and acceptor populations and cocultured for 48 h in the presence or absence of heparin, F6, or heparinase III. Spectral FRET measurements (FRET/donor) were obtained by using a Tecan M1000 fluorescence plate reader according to methods described previously (12, 53). Excitation and emission optima were as follows: CFP, 435 nm excitation/485 nm emission; YFP, 485 nm excitation/527 nm emission; and FRET, 435 nm excitation/527 nm emission.

We thank Josiah Gerdts for help with Movie S1 and the Alvin J. Siteman Cancer Center at Washington University School of Medicine and Barnes–Jewish Hospital for the use of the Siteman Flow Cytometry Core, which provided flow cytometry service. This work was supported by the Tau Consortium; the Muscular Dystrophy Association; the American Health Assistance Foundation; the Ruth K. Broad Foundation; National Institutes of Health (NIH) Grants 1R01NS071835 (to M.I.D.), 1R01GM038093 (to F.M.B.), K08NS074194 (to T.M.M.), 1F31NS079039 (to B.B.H.), P50 CA94056 (Molecular Imaging Center, Mallinckrodt Institute of Radiology, Washington University School of Medicine), and P30 CA091842 (National Cancer Institute Cancer Center Support Grant to Siteman Cancer Center, Washington University School of Medicine); the Molecular Imaging Center at the Mallinckrodt Institute of Radiology; the Bridging Research with Imaging, Genomics, and High-Throughput Technologies Institute at Washington University School of Medicine; and an Anheuser-Busch/Emerson challenge gift. The Hope Center Alafi Neuroimaging Laboratory and the Bakewell Neuroimaging Core are supported in part by the Bakewell Family Foundation and NIH Neuroscience Blueprint Interdisciplinary Center Core Grant P30 NS057105 (to Washington University).