Microtubules are dynamic, labile structures that undergo tight regulation by numerous proteins. Microtubules provide important functions for most cell types, including structural support, intracellular transport, and chromosome separation during cell division. Perhaps the most well-studied microtubule associated protein to date is tau protein, since changes in tau splicing and tau mutations lead to a neurodegenerative disease called frontotemporal dementia (FTD) with parkinsonism linked to chromosome 17 (FTDP-17), a clinical syndrome that leads to neuronal loss and dementia. Tau is also a key component of one of the pathological hallmarks of Alzheimer’s disease (AD), the neurofibrillary tangle (NFT). Tau is uniquely localized in neurons and is associated with microtubules only in axons where the microtubules adopt an asymmetric conformation. Understanding the way in which tau and microtubules interact in living cells is crucial to understanding how disruption of this process might lead to cell dysfunction or death.
Approximately 50 tau gene mutations are associated with FTDP-17. Many of the mutations are within the introns adjacent to exon 10 and have been shown to alter tau splicing to favor inclusion of exon 10, which encodes an additional microtubule binding domain. Tau isoforms containing exon 10 are referred to as 4 repeat (4R) isoforms, whereas those lacking exon 10 are referred to as 3 repeat (3R) isoforms. Alterations of exon 10 splicing are also seen in nonfamilial cases of FTD, 1, 2, 3, 4, 5, 6, 7, 8 supranuclear palsy,8, 9, 10 and in some instances of AD.3, 8, 11, 12 We have also recently shown that alternative splicing of an amino terminal domain (exon 2) as well as exon 10 occur differentially in AD and control brain tissue.12 Other FTDP-17 tau mutations lead instead to amino acid changes, often centered near the microtubule binding domains, and these are also believed to alter tau-microtubule dynamics. One of the most common of these mutations is the P301L mutation.4, 13, 14, 15
These studies have provided valuable information about the marked effect tau splicing has in terms of association with neurodegeneration, but they have lacked a functional readout for the effect tau alterations have on intact microtubules within neurons. In isolated tau/tubulin preparations, the presence of the fourth microtubule binding repeat in tau appears to stabilize microtubules, unlike the isoforms that contain only three repeats.16, 17 By contrast, and somewhat contrary to expectations based on these in vitro data, recent fluorescence recovery after photobleaching (FRAP) data suggest that 3R tau and 4R tau have very similar properties with regard to tau/tubulin association in intact retinal ganglion cells.18
We have explored this issue using an alternative approach to examine the effect of tau splice forms on microctubules in intact neurons. Second harmonic generation (SHG) is a nonlinear optical microscopy method with inherent properties that make it suitable for visualizing microtubule structure in living cells. In SHG, light is generated by a structure at exactly one-half the wavelength of incident light; unlike fluorescence, the SHG signal can be elicited by a broad range of incident light, and the emitted photons “track” at exactly one-half the wavelength of incident light. Moreover, the process is nonabsorptive, so there is no photobleaching or free radical generation (for review, see Ref. 19). SHG arises only from inherently asymmetric chemical structures, including some biological molecules. A number of proteins have been shown to generate second harmonic signals, including collagen,20, 21 myosin,22, 23, 24 and axonal microtubules,19, 25, 26 with axonal microtubules being the weakest source of SHG.
In this study, we took advantage of the observation that axonal microtubules give rise to an SHG signal due to their asymmetric structure.26 A disruption of axonal microtubules causes the loss of SHG signal.26 We utilized SHG as a functional readout for microtubule structure in living cultured neurons and examined the effect of the presence of exogenous tau in transfected neurons. Our results clearly demonstrate that the presence of exogenous tau significantly enhanced SHG signal. Surprisingly, and in contrast to extrapolation from biophysical measures of isolated tau/microtubule interaction assays, our results suggest that the 3R and 4R isoforms of tau, and even a P310L FTDP-17-associated mutant form of tau, all seem to alter microtubules/SHG to a similar extent.
Cell Culture and Western Blot Analysis
Primary cortical neurons were prepared from cerebral cortices of CD-1 strain mouse embryos (day 15 to 17 of gestation). Cortices were dissociated by trypsinization for at room temperature, and cells were resuspended in Neurobasal (NB) medium supplemented with 10% fetal bovineserum (FBS), glutamine (Gln), penicillin, and streptomycin, then centrifuged at for and resuspended in NB/B27 [NB medium containing 2% (v/v) B-27 supplement], penicillin, streptomycin, and Gln. The cells were then plated at on round coverslips (Hemogenix, Colorado Springs, CO), and coated with Poly-D-Lysine in cell culture dishes (Corning, Lowell, MA). Neurons were grown for in vitro and transiently transfected with the indicated constructs using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol.
H4 human neuroglioma cells were maintained in Opti-MEM (Gibco, Carlsbad, CA) supplemented with 10% (v/v) FBS, penicillin, and streptomycin (Gibco, Carlsbad, CA). Where indicated, cells were transiently transfected with the identified constructs using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol.
Tau constructs were subcloned into either mRFP-N1 or eCFP-N1 vectors (Clontech, Mountainview, CA). Using , , , or in pcDNA3.1(+) as templates, we performed polymerase chain reaction using the following primers: -CTCGAGATGGCTGAGCCCCGCCAGGAGTTCGAAG- (forward) and -GGATCCAAACCCTGCTTGGCCAGGGAGGCAGAC- (reverse), digested with Xho1 and BamH1 and ligated into appropriate color vectors. The integrity of each construct was verified using DNA sequencing analysis.
H4 human neuroglioma cells were transiently transfected with Lipofectamine 2000 according to the manufacturer’s protocol and collected post-transfection in 2X sodium dodecyl sulfate (SDS) lysis buffer ( Tris-Cl, pH 7.5, 2% SDS, ethylenediaminetetraacetic acid, ethylene glycol tetraacetic acid, and 10% glycerol supplemented with a protease inhibitor pellet obtained from Gibco, Carlsbad, CA). Cell lysates were then sonicated for and boiled for . Samples were centrifuged at room temperature for at . Sample concentrations were determined using the bicinchoninic acid assay (Pierce, Rockford, IL). Samples were diluted to in 2X SDS running buffer and loaded onto precast 4 to 12% Tris-Glycine gradient polyacrylamide gels (Invitrogen, Carlsbad, CA), electrophoresed, and transferred to polyvinylidene fluoride membranes. Membranes were blocked for at room temp in Tris Buffered Saline Tween-20 (TBST) ( Tris-Cl, pH 7.6, NaCl, 0.05% Tween 20) and 5% milk and incubated in TBST and milk at overnight in the indicated primary antibody. Membranes were then washed 3 times for in TBST at room temperature and incubated for with appropriate horseradish peroxidase-conjugated secondary antibody in TBST and milk. Membranes were washed 3 times for in TBST at room temperature and visualized with enhanced chemiluminescence (Amersham, Piscataway, NJ).
Live Cell Imaging and SHG
Primary neurons on coverslips were fitted in a perfusion or culture (Hemogenix, Colorado Springs, CO) closed chamber with a silicone gasket between glass coverslips. The chamber contained Hank’s balanced salt solution (HBSS) prewarmed to , and it was affixed in the heated stage of a Zeiss Axiovert 200 inverted microscope. All images were acquired on a Zeiss LSM-510 META confocal microscope fitted with a laser for red fluorescence protein (RFP) fluorescence and a Coherent (Santa Clara, CA) chameleon mode-locked Ti/Saph tunable laser with pulse width and average power with linear polarization at the sample, for cyan fluorescent protein (CFP) and SHG imaging. All images were acquired using a Zeiss numerical aperture (NA) Plan-NEOFLUAR water/oil immersion lens for epifluorescence and an infinity-corrected Zeiss -NA PlanAPOCHROMAT dry lens affixed to a custom condenser tube (fabricated by Zeiss, Jana, Germany) affixed to the condenser housing, which projected the forward-propagating signal onto a photomultiplier tube (PMT, Hamamatsu). A bandpass filter was fitted between the condenser housing and the PMT. This aided in both reducing background fluorescence at the SHG PMT and in verifying the SHG nature of the signal observed, since light was used to induce the SHG signal. The resulting signal should pass through the filter unobstructed, while producing an SHG signal using should result in a loss of the SHG signal, because the resulting signal would be blocked by the bandpass filter. The bandpass filter also ensured that no fluorescent signal from the fluorophores would pass through to the PMT, since CFP emits around and RFP emits around , and both are blocked by the bandpass filter. To further ensure the signal obtained was from SHG and not autofluorescence, a polarizer was placed in front of the SHG PMT, and the polarizer was rotated to demonstrate the polarized nature of the SHG signal. Since the SHG signal is a polarized signal, rotating the polarizer perpendicular with respect to the laser polarization should result in a loss of SHG signal, in contrast to fluorescence emissions.
Images were collected using Zeiss LSM 3.5 software and exported as JPEG images for analysis in Image J. The Neuron J macro plugin for Image J was used to trace and quantify the SHG signal in all sample images, and the resulting intensity data were analyzed using Graphpad Prism 4 (San Diego, CA) to carry out t-tests or one-way analyses of variance (ANOVAs). Data were presented as for each condition, and results were considered significant if .
Second Harmonic Generation Signal is Detectable in Single Neuronal Processes
Recent work has shown that an SHG signal arises mainly from axonal processes in neurons, not from the cell body or dendrites,26 in acute hippocampal slices. Therefore, we sought to determine whether it was possible to visualize an SHG signal from a single neuronal process in a cultured neuron that would allow us to visualize intact axonal microtubules in a living neuron. Figure 1 shows the morphology of a single neuron transfected with RFP and visualized with the red fluorescence signal. The RFP was excited with a laser, and the emitted light at was directed to the PMT through a bandpass filter. Similarly, the SHG signal was obtained with excitation and collected in the forward direction after passing the signal through a bandpass filter. A weak SHG signal was observed in single processes extending from each neuron. Several characteristics suggest that this was SHG: the signal was seen only in the axon (as expected given the asymmetric microtubule structure unique to axons), whereas tau and autofluorescent moieties are most strongly seen in the cell body. This signal was lost when treated with colchicine, indicating that it was microtubule dependent. We inserted a polarizer in the light path between the sample and the PMT. Rotating the polarizer between 0 and should either allow a polarized signal to pass or block a polarized signal, depending on the orientation, while fluorescent signals should be unaffected by the orientation of the polarizer. As expected, when the polarizer was in the position, the SHG signal was strong. However, when the polarizer was shifted to the position, the SHG signal was completely lost. As another test, we changed excitation wavelength to , and the SHG signal observed in a 375 to 425 emission filter was lost, as would be expected for SHG (Fig. 2 ). Finally, we demonstrated that if the optics were reconfigured to detect the back-propagating signal, the signal was undetectable, again as expected for SHG—which is predominantly forward-propagating, unlike a fluorescent signal.
Overexpression of Tau Increases the SHG Signal
Next, we examined whether altering tau levels would enhance the SHG signal. As a first step, we characterized tau expression in H4 cells. Figure 3 shows a western blot analysis of various tau constructs expressed in H4 cells to verify the biochemical properties of each construct. The different isoforms’ molecular weights corresponded to the presence or absence of the fourth microbutubule binding repeat (4R, representing exon 10) or the 2 amino-terminal inserts . All constructs migrated to the expected molecular weight, indicating that the proteins were expressed and processed appropriately in cells.
We examined whether microtubules in H4 neuroglioma cells, which are morphologically flat with several long processes, would show SHG signal. No SHG signal could be detected from H4 cells transfected with CFP [Fig. 3b]. By contrast, imaging of H4 cells transfected with tau showed robust tau expression in the cell body and processes; interestingly, the SHG signal arose primarily from perinuclear compartments where tau levels were highest, and where the microtubule network appears to reorganize within the perikeria [Fig. 3c].
Next we examined whether transfection of tau would enhance SHG in neurons, and if so, in which cellular compartment. Given that tau is a microtubule binding protein that promotes the assembly and stability of microtubules in axons (reviewed in Ref. 27), it might follow that expression of tau would increase axonal SHG signal. To further test this hypothesis, and also to determine if the presence of a fluorescent tag on tau would change the SHG, we transfected neurons with different isoforms of either TauRFP, untagged tau cotransfected with RFP, or RFP alone. SHG signal was measured by tracing SHG positive neurites using the Neuron J macro for Image J and analyzing the intensity across the entire length of the process.
Since transfection efficiency was very low in cultured neurons with our transfection method, it was possible to visualize the processes of a single neuron and follow them for many hundreds of microns without interference from the processes of neighboring, untransfected neurons. When TauRFP-transfected neurons were examined, the RFP fluorescence and SHG signals were distinct [Fig. 4a ]: SHG signal was present in only one process, whereas the TauRFP fluorescence signal was present in multiple processes and the cell body. To verify that the signal obtained in the SHG channel was indeed SHG and not contaminating fluorescence, the wavelength of the excitation laser was changed from . As expected, the SHG signal was abolished, since the bandpass filter in front of the SHG detector blocked emission [Fig. 4b]. As a further control, we added a bandpass filter to the turret, and we were able to switch between the and filters. Reconfiguring the detectors demonstrated that the signal was substantially forward-propagating and could not be detected in the back-propagating configuration (data not shown). As expected, when we used light and the filter, there was a detectable SHG signal. To test the hypothesis that the SHG signal observed was due to microtubule structures, sister cultures of TauRFP-transfected neurons were treated with colchicine for and imaged as above. Since colchicine completely disrupts microtubule stability, any SHG signal arising from microtubules should be attenuated. As expected, treatment of cells with colchicine led to the complete loss of SHG signal [Fig. 4c]. These results clearly demonstrate that SHG signal is obtainable from a single neuronal process that is presumably the axon, and that the source of SHG signal within that process is the microtubule network.
As an additional control, we inserted a Berek compensator (New Focus, San Jose, CA) into the excitation light path (to circularly polarize the laser source) and examined the SHG signal from the processes. Figure 5 shows that, as expected, even when the excitation beam was circularly polarized, only one process showed a strong SHG signal.
Expression of either tagged or untagged tau led to a significant and equal increase in SHG signal (Fig. 6 ) over that of RFP alone ( , , and ; for both RFP versus TauRFP and RFP versus ). The presence of either RFP or CFP fusion constructs on the C-terminus of tau did not further enhance or decrease the SHG signal from microtubules when compared to tranfection with untagged tau .
One possibility that could account for increased SHG signal from tau-transfected axons is that expression of tau significantly increases the overall diameter of the axon, and therefore increases the signal by giving rise to a signal from a larger axonal cross-section. To rule this out, images of axons of both RFP- or TauRFP-transfected neurons were examined in Image J, and the axonal diameter was measured at least distal to the axon hillock and before the first major bifurcation of the axon. Three separate measurements per axon were averaged, and a two-tailed t-test revealed that there was no significant difference between RFP- and TauRFP-transfected axons. Therefore, tau expression does not increase SHG simply by increasing the size of the axon (data not shown).
Different Tau Isoforms Do Not Significantly Differ in Enhancing SHG
In in vitro assays, 4R tau binds microtubules with a greater affinity and polymerizes the formation of microtubules to a greater extent than 3R tau16, 28 (reviewed in Ref. 29). Having shown that tau enhances SHG signal in neurons, we next sought to determine whether different tau isoforms would differentially affect SHG signal. Tau-CFP constructs were generated for ease of planned co-transfection experiments, and in all instances they were equivalent to the transfection protocol utilized for the Tau-mRFP constructs used above. Neuronal cultures were transfected with one of the following constructs: CFP, , , , or . SHG signal was determined for each condition. As expected, the expression of any isoform of tau enhanced the observed SHG signal compared to empty-vector or CFP-transfected neurons (Fig. 7 ). Surprisingly, however, there was no significant difference between exon 10-containing or exon 10-skipping constructs, indicating that the presence of 3 or 4 microtubule binding repeats did not significantly affect tau’s ability to enhance microtubule SHG signal in living neurons. No morphological changes in axon shape were noted qualitatively, and measuring the axon diameter revealed no difference between tau-transfected and control neurons.
Mutations in the tau gene found in FTDP-17 have been shown to decrease the in vitro association of tau with microtubules and the rate of microtubule polymerization. 13, 17, 30, 31, 32, 33, 34 To determine if tau mutations alter axon-derived SHG in living neurons, we transfected neurons with and examined the effect on SHG signal. Surprisingly, the mutant tau enhanced the SHG signal observed in the transfected neurons (Fig. 8 ) to essentially the same extent as other tau isoforms.
Our earlier studies suggested that axonal microtubule structures gave rise to SHG, as detected in bundles of axons in slice preparations,26 and we proposed that the signal was due to a unique structure of axonal microtubules. Our current experiments demonstrate that the unique SHG signal can be detected in single axons. The signal is lost with the addition of the pharmacological disruption of microtubules by colchicine, and enhanced (only in axons) by transfection with the axonal microtubule binding protein tau, suggesting that tau-microtubule interactions gave rise to the enhanced signal. These data are consistent with the idea that the signal seen in axon bundles originates from the axonal microtubule structure itself rather than from extracellular compartments such as collagen or basement membrane. This observation further supports the idea that the source of SHG signal in axons is the asymmetric polarity of axonal microtubules.
Interestingly, H4 neuroglioma cells have very little or no SHG signal present, even in the many small neurites that encircle the cells. However, when tau is transfected into H4 cells, the microtubule network, along with tau, rearranges into circular filamentous structures that give rise to significant SHG signals. There are also multiple processes that can be observed with SHG signal-transfected H4 cells. These data suggest that H4 cells differ significantly from neurons in their processing of tau and the organization of microtubules within processes, and highlight the importance of studying tau-microtubule interactions in axonal compartments.
Our results, following tau transfections, were unexpected from several perspectives: overexpressed tau appeared in the soma and all processes of a neuron, and localized with tubulin in all cellular compartments. However, SHG signal was observed only in single processes. These data suggest that tau-tubulin interactions in the soma or dendritic processes do not stabilize microtubules into the same conformation in all cell compartments, and instead, that a unique conformation occurs in axons. The enhancement of SHG signal in these axonal compartments is consistent with the idea that excess tau (in the axon) can stabilize microtubules or enhance microtubule bundling; if so, these data using an in vivo biophysical measure in neocortical neurons complement and extend observations in retinal ganglion cells (RGCs) in which tau overexpression led to functional impairments in organelle transport.33, 35, 36, 37, 38, 39 Consistent with this idea, it was demonstrated by Mandelkow that overexpressed tau leads to an increased density of microtubule bundles in axons,40 and we postulate that this bundling underlies the enhanced SHG signal. Although it is not clear how this concept of being an improved SHG source compares to microtubule stability measured in vitro, our data demonstrate that overexpressed tau changes the biophysical properties of the axon in a striking fashion. One possibility that could account for increased SHG signal from tau-transfected axons is that expression of tau significantly increases the overall diameter of the axon, and therefore increases the signal from a larger axonal cross-section. To rule this out, images of axons of both RFP- or TauRFP-transfected neurons were examined in Image J, and the axonal diameter was measured at least distal to the axon hillock and before the first major bifurcation of the axon. Three separate measurements per axon were averaged, and a two-tailed T-test revealed that there was no significant difference between RFP- and TauRFP-axons. Therefore, tau expression does not increase SHG by increasing the size of the axon (data not shown).
We hypothesized that 4R tau would enhance SHG to a greater extent than 3R tau. Our results suggest, however, that the presence of the fourth microtubule binding repeat had very little effect on microtubule SHG signal, implying that within the microenvironment of an intact axon, the presence of the fourth microtubule binding repeat does not significantly change the tau-induced microtubule conformation. This is consistent with the findings of Konzack, who demonstrated by using both FRAP and fluorescence correlation spectroscopy that the dwell time of a single molecule of tau on microtubules is very short, and that there was no significant difference between 3R and 4R tau with respect to how rapidly it diffused through the RGC axon or moved off and on the microtubules.18 These data suggest that within the physiological environment of an axon, the tau isoform has little effect on microtubule-tau interactions.
The picture that emerges from our data is that tau interacts with microtubules in a specific asymmetric conformation in axons to generate a unique conformation that supports SHG. Using this assay of tau-microtubule interactions, it appears that the presence of the fourth microtubule binding repeat may exert less effect on the tau-microtubule interaction in axons than previously thought. Thus, although changes in tau splicing are clearly important in FTDP-17 and related conditions, our current data, as well as that of Konzack, 18 suggest that the elevation of 4R tau may have additional pathogenic mechanisms independent of differential 3R versus 4R effects on axonal tau-microtubule interactions.
The authors would like to thank Dr. Jennifer M. Gass and Dr. Michael L. Hutton of the Mayo Clinic for their generous gifts of the , , , and isoforms. We would also like to thank Brad Reynolds of Zeiss Microimaging for assistance with modifications of the LSM 510 instrumentation. This study was supported by National Institutes of Health AG 026249, EB000768, and P50 AG 05134.