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Originally published In Press as doi:10.1074/jbc.M309971200 on December 4, 2003

J. Biol. Chem., Vol. 279, Issue 9, 7395-7404, February 27, 2004
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Comparative Vibrational Spectroscopy of Intracellular Tau and Extracellular Collagen I Reveals Parallels of Gelation and Fibrillar Structure*

Laura J. Juszczak{ddagger}

From the Department of Physiology and Biophysics, Albert Einstein College of Medicine, Yeshiva University, Bronx, New York 10461

Received for publication, April 8, 2003 , and in revised form, December 3, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The N-terminal tau 2–19 peptide undergoes gelation, syneresis, and aggregation over a period of years. These changes may be approximated on a shorter time scale by agitation and partial dehydration. The anomalously enhanced (229 nm) ultraviolet resonance Raman (UVRR) imide II band reveals a common structural feature for gels of nondehydrated tau 2–19 and collagen I and insoluble paired helical filaments (PHFs) and collagen I of weak hydrogen bonding at proline carbonyls. Anomalous UVRR enhancement of amide bands at 229 nm results from gel structure, as demonstrated by increased amide absorption at the red edge for tau 2–19 gel and implies the involvement of water in gel structure. In aged, dehydrated tau 2–19 gel, proline carbonyls lose their bonds to water and tyrosine becomes deprotonated, as demonstrated by UVRR spectroscopy. The Fourier transform infrared (FTIR) amide I band shows that antiparallel {beta}-sheet structure increases with syneresis in the tau 2–19 hydrogel. The comparison of FTIR results for PHFs with collagen I gel and polyproline demonstrates that the secondary structure of PHFs is polyproline II. One implication of this assignment is that the fibrillation of hydrophilic tau is thermodynamically driven by the entropy gained as hydrogen-bonded water is freed, as for collagen I. The FTIR results also show that peptide domains culled from a longer protein do not necessarily fold into identical secondary structures. A pathological, sequential mechanism of gelation, syneresis, and fibrillation for tau in AD is suggested and is supported by the observation of amorphous neurofibrillary tangle development and fibrillation in vivo.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The cytomatrix is a living, insoluble, thixotropic gel (1) that is dynamic because its structure may change from fluid (sol) to elastic (gel) and back again; the filamentous network is constantly changing (2). Its constituent, organized structures are polymeric microtubules, actin microfilaments, and intermediate filaments (2). The assembly of this gel network and the composite fibrils is assisted through interactions with a number of other proteins. Within the set of microtubule assembling proteins, tau is of particular interest.

In healthy neurons, tau, through its control of the reversible polymerization of microtubules, participates in the extension of both axons and neuritic processes, also known as neural growth cone development, and in the development of neural polarity (3). In a post-translationally modified state, however, tau is responsible for several neurodegenerative diseases or tauopathies, one of which is Alzheimer's disease (AD),1 characterized by intraneuronal inclusions of phosphorylated tau that are initially amorphous. Over time, the intraneuronal inclusions of tau isoforms (4–6) self-assemble into paired helical filaments (PHFs) (7, 8); these are responsible for the neurofibrillary tangles found in AD.

Braak and Braak (9–11) have demonstrated the correlation between cognitive decline and the development of the tau neurofribrillary tangles. The progression of neural destruction in AD begins with loss or retraction of distal dendrites, accompanied by the formation of neuropil threads and an incipient neurofibrillary tangle in the soma, followed by loss of proximal dendrites, increased tau aggregation, and destruction of the soma (11). One of the central questions regarding the development of AD and other tauopathies is the significance of the development of the neurofibrillary tangles. Are they merely markers of the progress of the disease, or are they, in fact, the causative agent of neuronal death (12)?

Within the cytomatrix, actin microfilaments, keratin intermediate filaments, and microtubules cooperate in control of several cell activities including shape and cell membrane adhesion (13). Cellular growth and differentiation result in the build-up of tensile and compressive forces within the cytomatrix (14, 15). This cytoskeletal tension is transmitted to binding sites in the inert extracellular matrix via transmembrane integrin receptors (16) and other focal adhesion proteins (17). In fact, cell shape is controlled through a delicate balance of signal (18) and adhesive factors (19) between the cytomatrix and the extracellular matrix.

Cell adhesion substrates include extracellular matrix proteins such as collagens (19, 20). The most abundant protein of the extracellular matrix is collagen I, a fibrillar triple helical protein that forms gel networks in irregular connective tissue (21–23). Tau, like collagen I, is very hydrophilic (24) and shares other characteristics; both are proline-rich and nonglobular, self-assemble into fibrils, and are found in vivo as several isoforms.

The spectroscopic and electron microscopic results presented here demonstrate other common features of structure and behavior between collagen I and tau 2–19 peptide and full-length protein: precedence of a gel phase leading to fibril formation, the involvement of proline residues in weak hydrogen bonding of both the gel matrix and fibril, and a polyproline II secondary structure (PHFs). These results are discussed with respect to the cytoskeletal changes that might ensue from the generation of pathological intraneuronal compressive forces by a collagen-like gel and fibril (12).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Lithium thiocyanate, glycerol, bovine skin gelatin type B, and insoluble bovine Achilles tendon collagen type I were purchased from Sigma and used without further purification.

Peptide Synthesis—N-terminal tau 2–19 peptide (primary sequence AEPRQEFEVMEDHAGTYG) and tau 309–326 (primary sequence VYKPVDLSKVTSKCGSLG), a segment from the tau protein pseudorepeat region, were synthesized using solid phase peptide synthesis methodology on an ABI 433A peptide synthesizer at a 0.1 mM scale by the standard protocol for Fmoc chemistry (25). The reagents piperidine, diisopropylethylamine, N-methylpyrrolidine, dichloric methane, Fmoc amino acids, and preloaded resin and a HOBT/HBTU* activation kit were purchased from Applied Biosystems (Foster City, CA). Peptide resin was cleared and deprotected by a mixture of tetrafluoroacetic acid, water, phenol, and triisopropylsilane in a ratio of 88:5:5:2 for 2 h. The peptide was precipitated by ether, washed, and lyophilized. The peptide was characterized by matrix-assisted laser desorption ionization time-of-flight mass spectrometry high pressure liquid chromatography (Voyager DE STR, PerSeptive Biosystems, Foster City, CA) using a C8 reverse phase column (HP1090; Hewlett Packard Co.) and amino acid analysis (Hewlett Packard Amino Quant).

For the C-terminally biotinylated tau 2–19, Fmoc {epsilon}-aminocaproic acid (Bachem California, Inc., Torrance, CA) was inserted as a biotin linker during automatic peptide synthesis. After deprotection, the Fmoc group resin was washed with Me2SO; 0.244 g (1 mmol) of (+)-biotin (Sigma) was dissolved in 5 ml of a 1:1 solution of dimethylformamide and Me2SO. To this was added 2.1 ml of a 0.45 M HOBT/HBTU activation solution and 3 ml of diethyl ether. The resulting activated biotin solution was added to 0.1 mmol of the Fmoc group resin and stirred overnight. The resulting resin was washed successively with a 1:1 solution of dimethylformamide and Me2SO plus dimethylformamide and dichloromethane to remove the excess biotin. The biotinylated resin was cleared as above.

Preparation of Tau 2–19 Samples—Freeze-dried tau 2–19 was dissolved in deionized water to a concentration of 6 mM and stored undisturbed at 4 °C for 5 years. The tau 2–19 was found to have gelled after this time; UVRR results were recorded on the undiluted gel. Typically, polyanions such as perchlorate (0.2 M) (26, 27) are added as internal peak calibration standards in the acquisition of UVRR data. However, polyanions are known catalysts of in vitro PHF formation (28, 29), so an internal standard was not added to the aged tau 2–19 gel. The gel was cloudy after curing, so an absorption spectrum could not be recorded. When the gel was pipetted into a quartz tube, the gel became less viscous, exhibiting a gel phenomenon known as shear thinning; upon standing, the gel recovers its original viscosity (30). The gel was subsequently dissolved and diluted to 4 mM by the addition of a 100-µl aliquot of 9.1 M lithium thiocyanate. The UV laser beam (1.8 milliwatts) that was incident on the quartz NMR tube holding the dissolved tau 2–19 sample during the UVRR experiment caused the formation of a translucent tau film on the tube. The remaining tau solution was removed from the quartz tube, and UVRR results were recorded for the film alone. The tau film was found to be soluble in 3 M lithium thiocyanate solution. The tau 2–19 solution decanted from the quartz tube was spun at high speed (estimated as 1 kHz), and the resulting aggregated tau 2–19 was pelleted. The aggregate was resuspended in deionized water, and UVRR results were likewise recorded for it.

A 10 mM solution of lyophilized tau 2–19 with biotin on the C terminus was freshly prepared in deionized water for UVRR spectroscopy.

In trials to reproduce the UVRR results of the cured tau 2–19 gel on a shorter time scale, a fresh aliquot of lyophilized tau 2–19 was dissolved in deionized water at the gelation concentration of 10 mM and stored undisturbed at 4 °C. This aliquot gelled within 2 months. A UVRR spectrum of this gel was acquired periodically up to 9 months of age. Additional 10 mM tau 2–19 aliquots were subject to different treatments: 1) heating in a sealed ampule to 95 °C for 5 days; 2) vigorous stirring at room temperature for 19 h; 3) vigorous stirring followed by partial drying under a nitrogen stream; 4) adjusting the pH to 10.5; and 5) dissolution in 10 mM sodium chloride (31).

Preparation of PHF Samples—Immunoaffinity-purified PHFs (32, 33) at 2 mg/ml in Tris-buffered saline (10 mM Tris, 0.15 M sodium chloride, pH 7.4) were a gift of Dr. Peter Davies. PHFs dissolved in 9 M LiSCN were dialyzed exhaustively against distilled water and recovered as a solid by drying under a nitrogen gas stream at room temperature prior to incorporation in a KBr pellet. Additionally, it was found that PHFs dissolved in 9 M LiSCN, as for tau 2–19 in LiSCN, formed a translucent film when exposed to the UV laser beam.

Preparation of Collagen Samples—Insoluble bovine collagen type I was suspended in Tris-buffered saline at a concentration of 25 mg/ml. A bovine skin gelatin gel was prepared by dissolution in deionized water and in D2O at a 50 mg/ml concentration at ~60 °C with stirring.

Negative Stain Electron Microscopy—Solid tau 2–19 samples, gel and aggregate, were transferred to grids and negatively stained with 1% uranyl acetate. The grids were then blotted dry before immediate observation in a JEOL 100CX II transmission electron microscope (JEOL USA, Inc.) at 80 kV.

UVRR Spectroscopy—An argon ion laser system, described elsewhere (34), was used to generate continuous wave, 228.9-nm light at an incident power of 1.8 milliwatts. The samples were chilled to 10 ± 4 °C, rastered, and spun in order to minimize UV-induced photodamage. The UVRR spectra are an average of a set of 3-min acquisitions, collected over an 805–1680-cm–1 frequency window; see Figs. 1, 6, and 7 for numbers of acquisitions. The frequency scale of each spectrum is calibrated against the peaks of two reference solvents, indene and toluene, and is accurate to ±1cm–1. The software program Grams/32 AI, version 6.00 (Galactic Industries Corp., Salem, NH) was used to determine peak heights (full width at half-maximum) and for curve fits to complex bands. Fitting parameters were not restricted.



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  		FIG. 1.
UVRR spectra (229 nm) for tau 2–19 in various phases. a, 5-year-old cured gel originally prepared at 6 mM concentration (three 3-min acquisitions). b, difference spectrum where the minuend is the gelatin spectrum (a) and the subtrahend is the solution spectrum described in c. c, the spectrum for the C-terminally biotinylated tau in aqueous solution (three 3-min acquisitions). d, gelatin dissolved in 3 M lithium thiocyanate (final concentration) (four 3-min acquisitions). e, aggregate; film spectrum is identical (data not shown; five 3-min acquisitions for each).

 



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  		FIG. 6.
UVRR spectra (229 nm). a, bovine skin gelatin prepared at 50 mg/ml in deionized water (three 3-min acquisitions); b, 2-month-old tau 2–19 gel prepared at 10 mM concentration in deionized water (one 3-min acquisition).

 



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  		FIG. 7.
UVRR spectra (229 nm). a, bovine Achilles tendon collagen type I (25 mg/ml) suspended in Tris-buffered saline (three 3-min acquisitions); b, tau PHFs (2 mg/ml) suspended in Tris-buffered saline (10 3-min acquisitions).

 
FTIR Spectroscopy—FTIR spectra were acquired on a Nicolet (Madison, WI) Magna 600 FTIR spectrometer. Samples were prepared either as potassium bromide pellets or as smears on zinc selenide windows. For the tau 2–19 samples, the peptide concentration was 10 mM in D2O. Protein samples embedded in KBr were not subjected to potentially denaturing high pressure, since the pellets were formed using a screw-type press, which allows only for pressures applicable by hand-held wrenches. The number of accumulated scans varied; instrumental resolution was set to 4 cm–1.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
UVRR Spectroscopy of Tau 2–19 Peptide Phases—UVRR results (229-nm excitation) for the 5-year-old tau 2–19 peptide in four different phases are given in Fig. 1. The spectral window has been truncated to the 1100–1680-cm–1 region, since the bands of interest are concentrated here. Fig. 1, spectrum a shows the UVRR results for 5-year-old, cured tau 2–19 gel, whereas results for solution phase tau 2–19 with the C-terminally linked biotin, which prevents aggregation, are given in Fig. 1, spectrum c. The UVRR difference spectrum between the cured gel (Fig. 1, spectrum a) and solution phase (Fig. 1, spectrum c) of tau 2–19 is given in Fig. 1, spectrum b, where the 1616-cm–1 peaks were normalized before spectral subtraction. The addition of lithium thiocyanate (final thiocyanate concentration = 3 M), a known disrupter of water structure (35), to the cured tau 2–19 gel yields a noncloudy liquid, implying tau dissolution. The UVRR spectrum for the thiocyanate solution of the cured tau 2–19 gel is given in Fig. 1, spectrum d. The UVRR result for the tau 2–19 aggregate, described under "Materials and Methods," is given in Fig. 1, spectrum e. The result for the tau 2–19 film, also described above, is superimposable on those for the tau 2–19 aggregate (Fig. 1, spectrum e) and therefore is not shown.

The peak frequency of the vibrational bands for the UVRR results of Fig. 1 are summarized in Table I. As expected for excitation at 229 nm, the solution phase UVRR spectrum for biotin-linked tau 2–19 in Fig. 1, spectrum c, exhibits strong vibrational bands for the single tyrosine residue: Y8a at 1616 cm–1 (ring C-C stretch), a Y8b shoulder at 1600 cm–1, Y9a at 1178 cm–1 (ring C- H bend), and Y7a at 1208 cm–1 (C{beta}-C{gamma} stretch) (36–39). Less intense bands are also seen at 1238, 1551, and 1579 cm–1.


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  		TABLE I
Vibrational band frequencies (cm–1) with attribution for tau 2–19 phases

The results correspond to UVRR results for tau 2–19 phases given in Fig. 1, a, c, d, and e, respectively.

 
The UVRR results for the cured, tau 2–19 gel in Fig. 1, spectrum a, display not only the aforementioned tyrosine peaks but also strongly (and unexpectedly) enhanced amide bands at 1238 cm–1 (C{alpha}H mode), 1298 cm–1 (amide III (Am III)), 1384 cm–1 (amide S), 1550 cm–1 (amide II (Am II)), and 1579 cm–1 (amide II) (36, 40). A very weakly enhanced imide II band at 1445 cm–1, due to a carbonyl stretching mode of the single proline, is also seen (41). The presence of this band is also not expected, since proline is resonantly enhanced at a shorter wavelength, ~218 nm. The shoulder at 1218 cm–1 is an unassigned amide mode that has also been found in the UVRR spectrum (229 nm) of undiluted N-methylacetamide at –30 °C.2

The difference spectrum shown in Fig. 1, spectrum b, clarifies the amide band positions for the 5-year-old, cured tau gel results in Fig. 1, spectrum a. It also reveals a residual peak at 1600 cm–1, which is attributed to deprotonation of some unspecified population of tyrosines within the gel rather than to the single phenylalanine, because of the predominance of tyrosinate peaks in Fig. 1, spectra d and e, and the presence of a weak Y9a' band at 1168 cm–1 (42, 43). The presence of tyrosinate also suggests that the 1579-cm–1 band contains a tyrosinate Y8b' component (42, 43).3 The 1218-cm–1 shoulder of Fig. 1, spectrum a, is revealed in the difference spectrum, spectrum b of Fig. 1. Amide I bands, which appear at or above a frequency of ~1630 cm–1, are not seen (37, 44).

The addition of lithium thiocyanate to the cloudy, cured tau 2–19 gel resulted in a clear solution and yielded the UVRR results shown in Fig. 1, spectrum d. The spectrum is now less complex; tyrosine bands are now shifted to tyrosinate positions. The Y8a' band lies at 1602 cm–1, and the Y8b' band lies at 1583 cm–1; low frequency Y9a' and Y7a' bands are poorly resolved. The 1555-cm–1 band is attributed to amide II. The 1583-cm–1 band may also contain a weak amide II component, although a three-component curve fit (data not shown) to this high frequency cluster of bands (1555–1602 cm–1) yielded a reduced {chi}2 of <1, suggesting that both amide II and Y8b' bands do not contribute.

The UVRR results for the tau 2–19 aggregate are shown in Fig. 1, spectrum e; the tau 2–19 film results are similar (data not shown). The configuration of high frequency bands (1555–1602 cm–1) for the tau peptide aggregate is the same as for the dissolved tau gel (Fig. 1, spectrum d). At the low frequency end of the spectrum, however, bands are better resolved. Peaks appear at 1168 and 1182 cm–1; these are assigned as Y9a' and F9a, respectively. A weak amide III band appears at 1302 cm–1, and a weak imide II band is found at 1445 cm–1.

Electron Microscopy of Tau 2–19 Phases—Electron micrographs at x 20,000 magnification of the tau 2–19 aggregate and 5-year-old gel in the presence of uranyl acetate stain are shown in Fig. 2. The aggregate, Fig. 2A, does not stain, indicating that water is excluded from the structure, whereas the cured gel, Fig. 2B, is stained; water permeates the gel structure.



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  		FIG. 2.
Electron micrographs of tau 2–19 in two different phases, x 20,000 magnification, 1% uranyl acetate stain. A, aggregate (negative staining); B, aged or cured gel (positive staining).

 
UVRR Spectroscopy of Tau 2–19 Gels Alternately Processed— Clearly, tracking the structural change evident in the 5-year-old tau 2–19 gel at earlier time points is of interest. UVRR results for 10 mM samples of tau 2–19 under varying solution conditions and processing methods are given in Fig. 3. The high frequency quartet of peaks from the UVRR results of the 5-year-old, cured tau gel are reproduced in Fig. 3, spectrum a, for ease of comparison. The UVRR results for tau 2–19 vigorously stirred for 19 h at room temperature are shown in Fig. 3, spectrum b. This treatment yielded a cloudy gel. The tyrosine Y8a peak at 1615 cm–1 and Y8b peak at 1597 cm–1 are clearly present; a very weak amide II band at 1555.5 cm–1 is also evident. Curve-fitting results to the broad Y8 band envelope showed that additional bands, evident as shoulders in Fig. 3, spectrum b, are located at 1603 and 1584 cm–1 (data not shown). These latter bands have counterparts in the results for the 5-year-old tau gel and became more explicit after gel drying, discussed below. Heating tau 2–10 to 95 °C for 5 days yielded a Y8 band with low frequency shoulders of greater intensity and a Y8a band slightly up-shifted to 1613 cm–1. These spectroscopic results approach those of the 5-year-old tau gel, but the physical appearance of the gel was yellow, suggesting oxidation. The UVRR results for a 10 mM tau 2–19 solution at pH 10.5 are given in Fig. 3, spectrum d. Here, the tyrosinate Y8a' peak appears at 1604 cm–1, close to the 1600-cm–1 tyrosinate peak of the 5-year-old tau gel (Fig. 3, spectrum a), and a Y8b' shoulder is located at 1585 cm–1. The amide II band is stronger than those found in the UVRR results for the heated or stirred tau gels (Fig. 3, spectra c and b, respectively).



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  		FIG. 3.
Detail of UVRR results (1520–1650 cm–1) for tau 2–19 under different conditions. a, 5-year-old cured gel. b, cloudy gel at 10 mM vigorously stirred for 19 h at room temperature. c, gel (10 mM) heated to 95 °C for 5 days. d, 10 mM solution at pH 10.5.

 
In order to determine the age at which a tau 2–19 gel stored at 4 °C attains the spectroscopic markers of the 5-year-old tau gel (Fig. 1, spectrum a), a solution at the gelation onset concentration of 10 mM was prepared. UVRR spectra of this tau gel were taken at the 2, 5, 7, 8, and 9 months of age time points; the results are superimposable, and the result at the 9 months of age time point is shown in Fig. 4, spectrum a. The 9-month-old tau gel is clear in physical appearance. The tyrosine Y8 band (Fig. 4, spectrum a) has a shape and frequency characteristic of solution phase tau 2–19 (Fig. 1, spectrum c). However, a broad, fairly intense imide II peak (1465 cm–1), indicating weak hydrogen bonding at the proline carbonyl (41), also appears (Fig. 4, spectrum a). It is relatively more intense than the imide II band appearing in the results for the 5-year-old gel (Fig. 1, spectrum a). Thus, the UVRR results for the 9-month-old gel indicate structural differences relative to the tau-(2–19) gel similarly processed (quiescent, stored at 4 °C) but allowed to age for 5 years. The UVRR results in Fig. 4, spectrum b, are for the tau 2–19 gel processed by vigorous stirring for 19 h at room temperature (Fig. 3, spectrum b); note the lack of the imide II band at ~1465 cm–1. It is reproduced for comparison with the UVRR results for the tau 2–19 gel, which was similarly stirred and subsequently dehydrated, shown in Fig. 4, spectrum c. (This gel was also very cloudy, the significance of which will be discussed below.) The low frequency peaks of the tyrosine Y8 band at 1597 and 1584 cm–1 and the amide II band at 1555 cm–1 have gained intensity vis-à-vis the UVRR results for the tau gel, which has only been stirred (Fig. 4, spectrum b). Here, too, the imide II band is absent. Curve fit results to the Y8 band reveal an additional peak at 1603 cm–1. Thus, the cloudy physical appearance of the stirred, dehydrated tau gel, together with the incipient quartet of UVRR peaks (Fig. 4, spectrum c), indicates that vigorous, prolonged stirring followed by dehydration puts the tau 2–19 peptide gel on the path to spectroscopic equivalence with the 5-year-old tau gel (Fig. 1, spectrum a).



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  		FIG. 4.
UVRR (229 nm) for tau 2–19 gels formulated under different conditions. a, nonstirred, clear, 9-month-old gel at 10 mM stored at 4 °C. b, cloudy gel at 10 mM stirred vigorously for 19 h at room temperature. c, sample in b partially dehydrated under a nitrogen stream for 40 min at room temperature. Curve-fitting results for the high frequency peaks are shown as dashed line curves.

 
FTIR Spectroscopy of Tau 2–19 Gels Alternately Processed— In order to ascertain the presence of secondary structure in the tau 2–19 gels, FTIR results were acquired for a 1-week-old tau 2–19 gel, nonstirred process, stored at 4 °C (Fig. 5, spectrum a) and for a tau gel vigorously stirred for 18 h, followed by dehydration (Fig. 5, spectrum b). Both gels were prepared in D2O under a nitrogen-purged, water-free atmosphere. The amide I region, which reports on secondary structure, is shown. The nonambiguous features of the amide I band for the tau 2–19 gel produced by the nonstirred process (Fig. 5, spectrum a) are a fairly sharp peak at 1628 cm–1, a strong shoulder or buried peak at 1673 cm–1, and a low frequency shoulder at 1609 cm–1; these peak values were obtained by curve fitting (data not shown). The amide I region for the stirred, dehydrated tau gel (Fig. 5, spectrum b) has a similar contour, but the obvious peaks (1601, 1620, and 1672 cm–1; curve fitting results not shown) are more intense, better defined, and down-shifted relative to the nonstirred gel results. Inverted second derivatives of these FTIR amide I bands were also calculated and are shown below each FTIR result. The second derivative peaks are in good agreement with those obtained by curve fitting, albeit frequency up-shifted a few wave numbers relative to peaks predicted by curve fitting.



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  		FIG. 5.
FTIR spectra of tau 2–19 gels in D2O; 100 scans, 4-cm–1 resolution with inverted second derivative spectra, below. a, 1-week-old gel, nonstirred process; b, gel vigorously stirred for 18 h followed by dehydrating under a nitrogen gas stream for 1 h.

 
UVRR Spectroscopy of Tau 2–19 and Collagen I Gels—Since comparison between a well characterized protein and one that is not can often provide insight into the nature of the latter, a spectroscopic comparison of UVRR results for the nonstirred tau 2–19 gel at 2 months of age (Fig. 6, spectrum b) was made with those for a nonaged bovine skin collagen I gel (Fig. 6, spectrum a). As for the tau 2–19 peptide gels, peaks appear in the bovine collagen gel UVRR results that were not resonantly enhanced with 229-nm excitation for proteins in aqueous solution. Most noticeable is the very strong imide II peak at 1467 cm–1. Since collagen isoforms are ~25–30% proline, a large absorption cross-section predicts a strong imide II peak. The Y8 tyrosine band shape, with a peak at 1615 cm–1, is typical of solution phase results. The bovine collagen gel spectrum has an amide II band at 1555 cm–1, which is conspicuously absent in the nonstirred, 2-month-old tau gel. This difference will be important to the discussion of the PHF secondary structure below.

UVRR Spectroscopy of PHFs and Insoluble Collagen I—PHFs represent the end point in a transition from molecular disorder to order. Like the N-terminal tau 2–19 gel (Fig. 2B), they are stained by uranyl acetate; water is incorporated in their structure (7). The UVRR result for PHFs, composed of full-length tau isoforms, is given in Fig. 7, spectrum b, along with the result for insoluble bovine Achilles tendon collagen type I (Fig. 7, spectrum a). Both spectra show a tyrosine Y8a band at 1616 cm–1 and a strong broad, imide band at 1466–1468 cm–1. For full-length tau isoforms, the proline content is ~10–12%, whereas the proline content of collagen type I isoforms is ~25%; thus, the area of the imide II band scales accordingly. An amide II band at 1555 cm–1 is strongly enhanced in the PHF spectrum and weakly enhanced in the collagen type I results. A weak and broad amide III band at 1305 cm–1 is found in the PHF results (Fig. 7, spectrum b). Last, a broad, weak amide S band appears at ~1380 cm–1 in both spectra.

FTIR Spectroscopy of PHFs, Collagen I, and Polyproline— FTIR results were acquired for two forms of PHFs; these are presented, along with results for other illustrative proteins, in Fig. 8. Results for freshly isolated PHFs in the KBr pellet are given in Fig. 8, spectrum a. The peaks, located at 1294, 1404, 1460, 1552, and 1631 cm–1, are remarkably sharp; this is a consequence of the limited conformational motion possible in the solid state. Curve fits to the amide I (1631 cm–1) and amide II (1552 cm–1) bands are included, but the amide I results will be focused on; it is composed of a Lorentzian band at 1631 cm–1 and a Gaussian at 1665 cm–1. A small peak at 1746 cm–1 has been magnified for comparative purposes and will be discussed below. The amide I region for three proteins is given in Fig. 8, spectrum b. These include amide I results for PHFs recovered from dissolution in LiSCN, with a broad Lorentzian peak (1631 cm–1) corresponding to that for freshly isolated PHFs (Fig. 8, spectrum a), a single broad amide I band at 1643 cm–1 for polyproline, and a similar Lorentzian-shaped amide I band at 1653 cm–1 for bovine skin gelatin. The 1746-cm–1 peaks for the recovered PHF and polyproline results have been magnified. The high frequency region of the FTIR spectrum for insoluble bovine Achilles tendon collagen is given in Fig. 8, spectrum c. Although this sample was similarly embedded in KBr pellet, the amide I band at ~1650 cm–1 is not as sharp as that for the PHFs (Fig. 8, spectrum a); the amide I band for collagen is clearly composed of two Lorentzian peaks, determined from curve fitting and centered at 1645 and 1686 cm–1. Another feature of the bovine collagen FTIR spectrum is the strong, sharp band at 1746 cm–1, which may be attributed to type I {beta}-turns (45) but has also been found in the FTIR amide I band for other proteins with PII secondary structure (46).



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  		FIG. 8.
FTIR spectra of nonglobular proteins, 4-cm–1 resolution. a, PHFs, 1000 scans, potassium bromide pellet. Curves fitted to the 1552- and 1631-cm–1 peaks are shown as dashed lines. b, polyproline, 100 scans, potassium bromide pellet (solid line); paired helical filaments dialyzed from LiSCN solution, 100 scans, potassium bromide pellet (heavy solid line); bovine skin gelatin gel in D2O, 100 scans (dashed line). c, insoluble bovine Achilles tendon collagen, type I, 100 scans, potassium bromide pellet. Curves fitted to high frequency peaks are shown as dotted lines.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Water and Hydrogen Bonding in the Tau 2–19 Gel Network
By definition, a gel is a disordered network of molecules linked through some type of intermolecular bonding (47, 48). For a gel in a water-rich environment, which is called a hydrogel, water participates in hydrogen bonding through bridging of charged groups (30, 49, 50). Self-assembled hydrogels, such as the tau 2–19 peptide gel, are characterized by the property of shear thinning, as observed here, wherein the gel viscosity decreases upon application of a force, such as that imposed by pipetting, and increases once the force is removed (30). This property (the ease of breaking and reforming of the intermolecular gel structure) also supports the idea that water participates in intermolecular hydrogen bonding. The positive staining of the 5-year-old tau 2–19 gel with aqueous uranyl acetate (Fig. 2B) also indicates that it is a hydrogel and supports the role of water as a bridging element in maintaining gel structure. Last, the disappearance of the amide bands with dissolution of the 5-year-old tau gel by the addition of a high concentration of lithium thiocyanate (Fig. 1d), which is known to disrupt water structure (35), further supports this interpretation.

The disappearance of the amide bands from the UVRR results for the tau 2–19 film and aggregate grown from the lithium thiocyanate solution (Fig. 1, spectrum e) suggests a water-excluded environment for the amides, as found in globular proteins, under these processing conditions; the lack of uranyl acetate staining seen in the transmission electron microscopy image of the tau 2–19 aggregate (Fig. 2A) affirms this interpretation.

The Evidence for Water-bridged Amide Hydrogen Bonding
The anomalous appearance of the amide and imide II vibrational bands in the UVRR results for the tau 2–19 gels (Fig. 1, spectrum a, and Fig. 4, spectra a and c) implies heterogeneous broadening at the red edge of the corresponding absorption bands and/or a red shift in the absorption maxima such that they are on resonance with the 229-nm exciting laser beam. A comparison of far UV absorption spectra for a 6-month-old gel phase tau 2–19 (nonstirred process), a solution phase tau 309–326, and solution phase tau 2–19 with C-terminally linked biotin, shown in Fig. 9, supports this idea; the gel phase tau 2–19 is greatly red edge-broadened relative to tau 309–326 and noticeably broadened relative to the biotin-linked tau 2–19. Furthermore, the UVRR results presented in Fig. 1 demonstrate that water plays a role in the tau gel structure and that it is responsible for the anomalous appearance of imide and amide bands with 229-nm excitation. The relatively simple UVRR result for the biotinylated tau 2–19 peptide in aqueous solution (Fig. 1, spectrum c) is similar to those for other peptides reported in the literature where a longer wavelength has been used for excitation (51, 52). If the criterion for the red edge broadening and/or red shift of the maximum of the amide absorption band (Fig. 9) was only the ordering of water via hydrogen bonding about exposed amides and carbonyls, then the UVRR results of these solution phase peptides, with 229–240-nm excitation, would similarly show strongly enhanced amide vibrational bands. Nor can intermolecular amide-carbonyl hydrogen bonding account for the amide absorption red shift, since resonantly enhanced amide bands for the secondary structural elements, {alpha}-helices and {beta}-sheets, of globular proteins are optimally enhanced at ~210 nm (53), and 229-nm excited UVRR results for globular proteins are dominated by vibrational bands of the aromatic residues, tyrosine and tryptophan (54–57).



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  		FIG. 9.
Absorption spectra (10 mM, 0.5-mm path; normalized at 190 nm) for tau 309–326 (solid line), tau 2–19 with C-terminally linked biotin (dashed line), and 6-month-old tau 2–19 gel, nonstirred process (heavy solid line).

 
Interpretation of the Tau 2–19 UVRR Results
Presence of Amide S Bands Signals Lack of {alpha}-Helices—The appearance of an amide S band (~1384 cm–1) in an UVRR spectrum signals the absence of {alpha}-helical structure (40, 58). The amide III mode frequency has recently been shown to depend on not only the amide backbone dihedral angle, {psi} (59), but also on {phi} (60). Additionally, the amide III mode is coupled to the CH bending motions of the amide S band through the NH bending mode (40, 58). These vibrations mix when the dihedral angle, {psi}, takes a value characteristic for {beta}-sheet structures, polyproline II helices, and random coils (i.e. {psi} ~ 120°). At a {psi} value characteristic of {alpha}-helices, {psi} ~ 60°, the vibrations do not mix, and the amide S band does not appear (58). Since an amide S band is evident in the UVRR results for the 5-year-old (Fig. 1, spectrum a) and, to a lesser extent, the 2-month-old (Fig. 6, spectrum b) and the stirred, dehydrated (1404 cm–1; data not shown) tau 2–19 gels, {alpha}-helical structure is not incipient in the tau 2–19 gel curing process.

Weak Hydrogen Bonding at the Tau 2–19 Proline Residue— Takeuchi and Harada (41) found that the frequency of the imide II carbonyl-nitrogen stretch mode is sensitive to the strength of hydrogen bonding at the carbonyl. For a nonbonded carbonyl, the mode frequency is ~1445 cm–1, whereas a strongly hydrogen-bonded carbonyl mode appears at ~1485 cm–1. Since the imide II band position is 1445 cm–1 for both the cured tau 2–19 gel (Fig. 1, spectrum a) and the tau 2–19 aggregate (Fig. 1, spectrum e), the prolyl carbonyl in these phases is predicted to be nonbonded. For the tetrapeptide, tuftsin, the imide II band appears at 1466 cm–1, indicating a weaker hydrogen bond strength than that found for X-proline dipeptide hydrogen bound to water (i.e. 1475 cm–1) (41). The imide II band for the 2-month-old tau 2–19 hydrogel (Fig. 6, spectrum b) also appears at 1466 cm–1; its broadness suggests a spread in the range of C=O hydrogen bond strengths. A hydrogen bond strength somewhat less than that for C=O bound to water would be expected for a C=O bound to a bridging water (C=O ... H-O-H ...). There is no clear imide II band in the UVRR results for the stirred, dehydrated tau 2–19 gel (Fig. 4, spectrum c).

Interpretation of the Tau 2–19 FTIR Results
The amide I band results for the 1-week-old tau 2–19 gel, nonstirred process (Fig. 5, spectrum a) and for the stirred, dehydrated tau 2–19 gel (Fig. 5, spectrum b) demonstrate that within the disordered gel network, reorganization of peptide chains to a recognizable secondary structure is occurring. Gel networks are known to be dynamic structures where reorganization processes may occur over time (61). The more sharply defined peaks for the stirred, dehydrated tau gel (Fig. 5, spectrum b) relative to those for the nonstirred tau gel (Fig. 5, spectrum a) suggest that the former gel possesses greater secondary structural organization. The cloudy physical appearance of the stirred, dehydrated tau gel results from a refractive index difference created by an incomplete phase separation of the peptide into discrete islets in a gelation process known as syneresis (61); these islets of concentrated peptide may facilitate the development of secondary structure.

The two most prominent peaks in the amide I band for the stirred, dehydrated tau gel (Fig. 5, spectrum b) are located at 1620 and 1672 cm–1. The frequency position of these bands and their splitting (52 cm–1), resulting from transition dipole coupling of nearest neighbor peptide groups, strongly suggests the inception of an antiparallel {beta}-sheet structure in the gel (45, 62). Since FTIR peaks below 1620 cm–1 have been noted for {beta}-sheet (63), the strong shoulder in the amide I results for the stirred, dehydrated gel is also ascribed to {beta}-sheet structure. The remaining disordered fraction of peptide in the gel undoubtedly yields an FTIR signal; the broad, unresolved absorbance region between the 1620 and 1672 cm–1 peaks is assigned to this fraction.

A parallel assignment of amide I peaks can be made for the nonstirred tau gel (Fig. 5, spectrum a): antiparallel {beta}-sheet structure for the 1628 and 1673-cm–1 peaks, {beta}-sheet structure for the 1609-cm–1 shoulder, and disordered gel matrix to the region between the 1628 and 1673-cm–1 peaks. Significantly, the nonstirred tau gel remained transparent so that syneresis has not taken place. In the absence of localized concentrations of peptide, the development of secondary structure is not as readily facilitated, and thus amide I peaks are weaker for the nonstirred tau gel.

Changes in the Tau 2–19 Gel Structure with Time
The UVRR and FTIR results for the tau 2–19 gels show that the involvement of proline in gel structure decreases, whereas the development of {beta}-sheet secondary structure increases, with the onset of syneresis, mechanical agitation followed by dehydration, or gel aging. In the UVRR results for the optically clear, nonstirred, 9-month-old (Fig. 4, spectrum a) and 2-month-old (Fig. 6, spectrum b) tau 2–19 gels, a proline band at 1465 cm–1 unexpectedly appears, indicating a carbonyl hydrogen bond strength somewhat less than what is expected for hydrogen bonding to water (41), whereas tyrosine peaks are like those expected for solution phase. In the FTIR results for the clear, nonstirred, 1-week-old tau gel (Fig. 5, spectrum a), the inception of {beta}-structure is indicated, but peaks are not as well defined as for the optically cloudy gel (Fig. 5, spectrum b).

In the UVRR results for cloudy, stirred, and dehydrated gels of low age (Fig. 4, spectra b and c), the proline peak is absent, whereas the intensity of the low frequency bands in the tyrosine/amide II quartet increases. At the extreme of 5 years of age, a nonstirred yet cloudy tau 2–19 gel (Fig. 1, spectrum a) has a fully developed quartet of bands at high frequency, with both tyrosine and tyrosinate bands present, an imide II band at 1445 cm–1 for the nonbonded prolyl carbonyl, and an amide S band that precludes {alpha}-helical structure. In the limit of aggregate formation (Fig. 1, spectrum e), UVRR data for tau 2–19 aggregates shows that the prolyl carbonyl remains nonbonded and that the single tyrosine is deprotonated. The UVRR results imply that only the structure of the optically transparent tau 2–19 gels parallels that of PHFs in that the imide II band frequency for both proteins shows proline carbonyl hydrogen bonding.

Comparative Spectroscopic Analysis of PHFs
Collagens are a complex, multi-isoform set of generally nonglobular proteins containing stretches of proline residues which may (e.g. collagen I) or may not (e.g. collagen VI) be arranged in a periodic repeating motif, with N- and C-terminal propeptide sequences having no periodic repeat sequence of amino acids, and containing water-bridged interamide hydrogen bonds (64–66). Collagen I in the fibril state is a coiled-coil of three collagen I isoforms with a polyproline II (PII) secondary structure (64, 66). Subsequent to proteolysis by procollagen peptidases, which removes frayed sequences at the termini (67, 68), the primary structure of the collagen chains consists overwhelmingly of a 3-amino acid repeat of the type X-Y-Gly, where X is often proline and Y is often hydroxyproline (64, 66). The PII structure of collagen isoforms is stabilized by stereochemical interactions and does not require intrachain hydrogen bonds (69). X-ray diffraction results show that the interchain interactions, however, are stabilized by both interamide hydrogen bonds and water-bridged hydrogen bonds (64, 66); main chain carbonyls are also hydrogen-bonded to water molecules (66). Numerous collagen isoforms exist (65), and although domains of PII structure are often proline-rich, the prolines are not necessarily arranged in a regular sequence pattern as in collagen I (see, for example, the human collagen VI {alpha}1 chain).4 This, of course, is the case for the proline-rich domain of the tau isoforms. Indeed, Ramachandran has shown that all amino acids can be accommodated in the PII conformation (64). Thus, the comparison between the proline- and water-rich structure of collagen I and the tau PHF is valid and is supported both by the similarity of the UVRR results given in Fig. 7 and the correspondence of FTIR peaks in Fig. 8.

The one-to-one correspondence in UVRR peaks within the 1350–1650 cm–1 frequency window (Fig. 7) suggests a similarity in secondary structure for bovine Achilles tendon collagen type I and PHFs. Solid state PHFs, reconstituted from dissolution in LiSCN (Fig. 8, spectrum b), yield a single Lorentzian amide I band when probed by FTIR spectroscopy, as do solid state polyproline and bovine skin gelatin gel. Clearly, a PII structure is indicated for the reconstituted PHFs. Freshly isolated PHFs in the solid state result in an unambiguous amide I peak at 1631 cm–1 as well (Fig. 8, spectrum a) and a second, broader amide I band at 1665 cm–1. Again, the 1631-cm–1 peak suggests a PII structure for the virgin PHFs; the second amide band at 1665 cm–1 does not necessarily indicate that a second structural element is present, as discussed below, although the FTIR band frequency is consistent with either random coil or {alpha}-helical structure (63). The latter assignment seems unlikely, since the amide S band in the UVRR results precludes {alpha}-helical structure (Fig. 7, spectrum b).

The amide I band for insoluble bovine Achilles tendon collagen, type I (Fig. 8, spectrum c) is also resolvable into two components, yet x-ray diffraction data shows the molecular structure of collagen to be solely PII (64). In their infrared spectroscopic study of undenatured bovine Achilles tendon collagen, Susi et al. (70) similarly recorded an amide I band resolvable into two peaks, which they attributed to two populations of carbonyl groups within the structural repeat unit. The 1631-cm–1 peak of the PHFs may similarly arise from heterogeneity in carbonyl binding.

PII Structure Predicted for PHFs
The vibrational spectroscopic evidence presented here strongly supports the assignment of a PII structure to PHFs. Phosphotungstate staining results for the PHFs also support the assignment of PII structure to PHFs, since the fibrils have been shown to consist of subunit domains of greater and lesser stain inclusion, corresponding to fibril domains of greater and lesser hydrophilicity (7). The assignment of polyproline II secondary structure to PHFs is not excluded by Congo red staining, since dense collagen fibers, known to possess polyproline II secondary structure, are also stained by Congo red (71).

Factors Affecting Secondary Structure
The FTIR spectroscopic results presented here show that peptide domains culled from a longer protein will not necessarily fold into the same secondary structure and that the physical process applied to gelling peptides will affect the structure (31). The apparent discrepancy between reported CD results for various tau constructs (72–75) suggests that solution phase secondary structure can also depend on one or more factors such as protein concentration (76), time, the construct used, and ionic strength (73). Also, CD and other spectroscopic techniques have demonstrated that many proteins classified as "random coil" in fact possess PII structure (46, 69, 77–80). Indeed, the CD spectrum for the tau construct K12, which is reported as similar to results for full-length tau, exhibits a strong, negative peak at ~198 nm and is negative over the 190–250-nm spectral window (76). A similar CD result has been reported for collagen I (80).

Pathological Tau Gelation within the Cytomatrix and the Contractility of Dendrites
Previous studies of tau fibrillation have emphasized the kinetics of the process (81, 82), yet concepts from polymer science have not been applied. As for collagen I in the extracellular matrix and actin in the cytoskeletal matrix, spontaneous fibrillogenesis or self-assembly is preceded by a gelation phase (23). Prefibrillar amorphous tau deposits and the masses of tangled tau fibrils seen in vivo, as well as the formation of a tau film from a dissolved PHF solution reported here, suggest a gel precursor for PHFs. Indeed, a reticulated structure, seen in the tau 2–19 gel electron micrograph in Fig. 2, spectrum b, is also seen in an image of PHFs incubated at 90 °C for 5 min (73) and in that of the cytomatrix (2).

The gel state is characterized by processes that are ongoing beyond the initial gelation event. These may include reprecipitation and separation into solid-liquid or solid-solid phases (61). The 5-year incubation period leading to the development of syneresis within the nonstirred tau 2–19 peptide concurs with the decades time frame for development of tau neurofibrillary tangles in vivo, supporting the idea that tau fibrillation occurs via a gel curing process. Comparison of collagen I with tau and PHFs also suggests a solution to the paradox of tau fibrillation; the hydrophilicity of tau seems to countermand its fibrillation. Nevertheless, the assembly of hydrophilic collagen I into a triple helix is spontaneous and endothermic, thermodynamically driven by the increase in entropy accompanying the loss of bound water (23).

Cellular differentiation entails a delicate balance of transmembrane signaling and adhesive forces between the cytoskeletal matrix and the extracellular matrix (83). On the outside of the cell membrane, the extracellular matrix controls the structure of the cytomatrix and therefore controls cell shape (19), yet within the cell, the development of cellular protrusions involves rapid and reversible sol-gel transitions of the actin and cytoskeleton aided by an array of auxiliary proteins (23, 84). Microtubules appear to stabilize specialized cell extensions by counterbalancing the mass force produced in the cytoskeleton (85) and are involved in actin cytoskeletal organization (13).

The microtubules and actin microfilaments act as a single integrated lattice with built-in redundancy such that the load-bearing function of the cytomatrix may be maintained when the functioning of either element is compromised (12, 19, 86). Indeed, microtubule disassembly alone does not prevent dendrite growth, since the use of extracellular matrix as a cell adhesion substrate, in the absence of microtubules, is sufficient to induce dendrite elongation in PC12 cells (86). Apparently, dendrite growth on extracellular matrix results in less internal cellular tension, diminishing the need for the compressive support of microtubules (86). However, cell spreading is totally halted when microtubule and actin microfilament function is simultaneously compromised (19). Since distal, followed by proximal, dendritic processes are found to retract as neurofibrillary tangles appear (11), both microtubule and actin microfilament function must be compromised in AD. This suggests a nonspecific method of action for the neurofibrillary tangles.

From a purely biophysical perspective, PC12 dendrites and chick dorsal root ganglion neurons have been found to behave as viscoelastic solids under applied forces (87). Generation of low levels of cellular tension and the ensuing retraction response is the reverse of growth, which requires higher levels of cellular tension (87, 88). This cellular tension model has been used to explain the complete retraction of severed dendrites in PC12 and chick sensory cells (87).

The gelation-associated processes of syneresis and fibrillation potentially generate cytoplasmic forces that are responsible for the dendritic retraction seen in AD. It is foreseen that the process is set in motion by the permeation of the cytomatrix gel by phosphorylated tau, which subsequently gels to form an enmeshed network of tau gel and cytomatrix gel. The initial effect of an increase in cytoskeletal viscosity may be to retard the sol-gel transitioning of the cytoskeletal network and therefore to interfere with signaling pathways. Indeed, cognitive impairment in AD has been shown to precede tau fibril formation (9, 10).

Subsequent tau gel syneresis, followed by fibrillation and the accompanying loss of bound water, leads to contraction of the tau mass. This contraction constitutes the application of an unmitigated, pathological force on both the actin microfilaments and microtubules of the cytomatrix. Studies of the effect of compressive forces in dendrites have shown that the force applied regulates microtubule polymerization (14). An unmitigated force is expected to lead to massive microtubule depolymerization, denying specialized cell extensions the necessary mechanical support (89). The stress force generated by tau gel contraction and fibrillation may also destabilize the balance of tensile and adhesion forces across the actin-transmembrane integrin-extracellular matrix bonds. The expected outcome is dendritic retraction and cell rounding, since cytoskeletal supports of cell shape will have been compromised. The delicate balance of tensile forces between the cytomatrix and the extracellular matrix is thus destroyed by the matching of the collagenic extracellular matrix with an intracellular collagen-like network of tau. This scenario, where cellular differentiation is reversed and cytomatrix dynamics are fouled, is predicted to lead to neuronal death.

Conclusion—This vibrational spectroscopic comparison of tau PHFs with collagen I and polyproline has demonstrated that the secondary structure of PHFs is polyproline II. This structural comparison also leads to an explanation for the paradox of how a hydrophilic protein such as tau could spontaneously form fibrils; as for collagen I, fibrillation may be thermodynamically driven by the entropy gained as hydrogen-bonded water is freed. The N-terminal tau 2–19 peptide has been shown to form a hydrogel with distinct spectroscopic markers when aged for years. The spectroscopic characteristics of this tau peptide gel may be approximated on a shorter time scale by vigorous agitation and partial dehydration. A common structural feature for nondehydrated tau 2–19 gel as well as PHFs, collagen I gel, and insoluble collagen I is weak hydrogen bonding at the carbonyl of proline residues. Several additional threads of evidence point to a pathological, sequential mechanism of gelation, syneresis, and fibrillation for tau in AD. It is hypothesized that phosphorylated tau intercalates into the cytomatrix and enmeshes it by formation of its own gel structure. The resulting tau matrix, beyond the control of the cell, is thought to interfere with the dynamic rearrangement of the actin microfilaments and related signaling pathways. Subsequent syneresis and fibrillation of the tau gel produces intracellular stress forces, overwhelming the supportive cell scaffolding elements of microtubules and actin microfilaments along with their integrin-mediated bonds to extracellular collagen I. The retraction of neuronal dendrites, as found in preaptotic neurons, is predicted.


   FOOTNOTES
 
* The transmission electron microscopy work was supported by the Department of Physiology and Biophysics of the Albert Einstein College of Medicine. This work was also supported by a Resnick Gerontology Grant. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence may be addressed: Dept. of Physiology and Biophysics, Albert Einstein College of Medicine, Yeshiva University, 1300 Morris Park Ave., Bronx, NY 10461. Tel.: 718-430-3214; Fax: 718-430-8819; E-mail: juzak{at}aecom.yu.edu.

1 The abbreviations used are: AD, Alzheimer's disease; FTIR, Fourier transform infrared; PII, polyproline II; PHF, paired helical filaments; UVRR, ultraviolet resonance Raman; Fmoc, N-(9-fluorenyl)methoxycarbonyl; HBTU, O-(benzotriazol-1-yl)-N,N,N',N'-tetramethyluronium hexafluorophosphate; HOBT, 1-hydroxybenzo-triazole. Back

2 L. J. Juszczak, unpublished data. Back

3 The tyrosinate Y8b' mode is reported (43) as occurring at 1558 cm-1, whereas the data therein show that it is actually at ~1585 cm-1. The phenylalanine F8b mode is reported at 1593 cm-1, and the F8a is reported at 1604 cm-1 (42), whereas tyrosinate Y8a' is reported at 1601 cm-1. Since the tyrosinate Y8b' mode is very similar to the F8b mode, Y8b' would therefore be expected at ~1590 cm-1 and not at 1556 cm-1 as reported. The data therein show, in fact, a shoulder at ~1590 cm-1 on the Y8a' band. Back

4 Swiss-Prot number P12109 [GenBank] . Back


   ACKNOWLEDGMENTS
 
I thank Dr. Joel M. Friedman for use of the W. M. Keck Foundation Biomolecular Laser Research Center facilities. I thank Dr. Peter Davies for providing both tau peptides and PHF samples and Dr. Lisa Mints for peptide synthesis. I also thank Frank Mancuso for technical assistance in the transmission electron microscopy imaging. Last, I thank Dr. David Hamerman, Director of the Resnick Gerontolgy Center at the Montefiore Medical Center (Bronx, NY), for a Resnick Gerontology Grant initiating my work in this field.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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