- Research article
- Open Access
The proline-rich domain of tau plays a role in interactions with actin
- Hai Jin He†1, 2,
- Xing Sheng Wang†2,
- Rong Pan2, 2,
- Dong Liang Wang2,
- Ming Nan Liu1 and
- Rong Qiao He2, 2Email author
© He et al; licensee BioMed Central Ltd. 2009
Received: 10 May 2009
Accepted: 08 November 2009
Published: 08 November 2009
The microtubule-associated protein tau is able to interact with actin and serves as a cross-linker between the microtubule and actin networks. The microtubule-binding domain of tau is known to be involved in its interaction with actin. Here, we address the question of whether the other domains of tau also interact with actin.
Several tau truncation and deletion mutants were constructed, namely N-terminal region (tauN), proline-rich domain (tauPRD), microtubule binding domain (tauMTBD) and C-terminal region (tauC) truncation mutants, and microtubule binding domain (tauΔMTBD) and proline-rich domain/microtubule binding domain (tauΔPRD&MTBD) deletion mutants. The proline-rich domain truncation mutant (tauPRD) and the microtubule binding domain deletion mutant (tauΔMTBD) promoted the formation of actin filaments. However, actin assembly was not observed in the presence of the N-terminal and C-terminal truncation mutants. These results indicate that the proline-rich domain is involved in the association of tau with G-actin. Furthermore, results from co-sedimentation, solid phase assays and electron microscopy showed that the proline-rich domain is also capable of binding to F-actin and inducing F-actin bundles. Using solid phase assays to analyze apparent dissociation constants for the binding of tau and its mutants to F-actin resulted in a sequence of affinity for F-actin: tau >> microtubule binding domain > proline-rich domain. Moreover, we observed that the proline-rich domain was able to associate with and bundle F-actin at physiological ionic strength.
The proline-rich domain is a functional structure playing a role in the association of tau with actin. This suggests that the proline-rich domain and the microtubule-binding domain of tau are both involved in binding to and bundling F-actin.
Tau is an important microtubule-associated protein, promoting microtubule assembly and stabilizing microtubules [1–3]. The protein is recognized as a multifunctional molecule that interacts with actin in addition to microtubules [4–12], and is involved in the organization of the cytoskeletal network [4, 5]. Actin monomers (G-actin) were found to form gels in the presence of tau . According to Farias and colleagues , the association of tau with tubulin immobilized on a solid phase support system is inhibited by actin monomers, and a higher inhibition can be attained with preassembled actin filaments. Interestingly, tau can interact with F-actin, resulting in bundles of F-actin. MacLean-Fletcher and Pollard  have observed that tau dramatically induces an increase in the viscosity of actin filaments. Using electron microscopy tau has been shown to be capable of bundling microfilaments. Examination of morphological aspects of microtubules and actin filaments which coexist in vitro revealed associations between both cytoskeletal filaments, and in some cases, the presence of fine filamentous structures bridging these polymers . Several reports have demonstrated that tau interacts with actin in vivo. Sub-portions of tau co-immunoprecipitated with actin filaments have been found in various cell types . As described by Yu and colleagues , under NGF stimulation, tau is distributed at the ends of cellular extensions, where it associates with actin in a microtubule-independent manner in PC12 cells. Moreover, Fluga and co-workers  have provided evidence that tau induces changes in the organization and stability of neuronal actin filaments, which in turn contributes to Alzheimer's-like neurodegeneration in Drosophila and mouse model systems. This further demonstrates the physiological importance of interactions between tau and actin.
According to Buee and colleagues , tau consists of four parts: the N-terminal region, the proline-rich domain (PRD), the microtubule-binding domain (MTBD) and the C-terminal region. The microtubule binding domain has been reported to bind to actin [7, 9], but no data is available for the other regions bound to actin. It has been proposed that the proline-rich domain of tau participates in interactions with microtubules [16–18]. Interactions between tau and DNA have been studied in our laboratory , and PRD and MTBD were found to associate cooperatively with the minor groove in DNA double strands. These results intrigued and led us to investigate whether the proline-rich domain of tau also participates in interactions with actin.
Tau binds to G-actin and F-actin from skeletal muscle and platelets
Human actin has three subtypes, alpha actin being found primarily in muscle, and beta and gamma actin in other tissues. The interaction of tau with alpha actin has been well studied, however, little attention has been given to beta and gamma actin. Since tau mainly exists in neurons, beta and gamma actin are the subtypes of actin that tau can encounter. In this work we mainly used skeletal muscle actin. Platelet actin (a mixture of beta and gamma actin) was also employed to test whether subtypes of actin differ in their interactions with tau.
Biochemical Binding Parameters from Solid Phase Assays
2.10 ± 0.07
0.031 ± 0.007
2.31 ± 0.05
0.030 ± 0.004
1.90 ± 0.06
0.029 ± 0.005
2.01 ± 0.04
0.027 ± 0.003
The proline-rich domain is involved in binding to G-actin
Results indicated that the deletion of MTBD did not eliminate the ability of tau to bind to actin since both ΔMTBD and G-actin were present in protein pellets as shown by SDS-PAGE (panel a, Figure 2). Under our experimental conditions, the tauN, tauC and tauΔPRD&MTBD (panel b, Figure 3) mutants exhibited no association with actin in co-sedimentation assays, indicating that the proline-rich domain may be an essential domain involved in the association of tau with actin. Incubation of the tauPRD mutant with G-actin gave rise to a positive result in co-sedimentation assays, and tauPRD, tauMTBD, tau and BSA alone used as negative controls did not show any deposits in co-sedimentation assays (Additional file 2). To judge the efficiency of bundling during the reaction, both the pellet and supernatant fraction were loaded in gels (panel a, Figure 3). The result showed that most of the G-actin was in the pellet fraction under the experimental conditions, presenting the formation of F-actin bundles. Taken together, these results suggest that the proline-rich domain is involved in the association of tau with G-actin.
Platelet actin was also used in low speed co-sedimentation assays. Results were similar to those obtained for skeletal muscle actin (panel b, Figure 2). Under the same conditions, tauPRD bound to skeletal muscle and platelet actin without discrimination, i.e. tauPRD was capable of binding to alpha-, beta- and gamma-actin. These results again indicate that in addition to the microtubule-binding domain, the proline-rich domain played an important role in the association of tau with actin.
The proline-rich domain binds to F-actin and promotes F-actin bundling
Biochemical Binding Parameters from Solid Phase Assays
2.31 ± 0.05
0.03 ± 0.004
0.81 ± 0.05
0.57 ± 0.09
1.09 ± 0.01
0.29 ± 0.01
TauPRD induces F-actin bundles at physiological ionic strength
F-actin alone as a control did not assemble in the presence or absence of NaCl but existed in filaments of similar diameter (8.40 ± 1.65 nm ~9.95 ± 2.09 nm) (Additional file 3). On the other hand, some actin globules were observed when the NaCl concentration was increased to 500 mM, suggesting that actin filaments started to collapse at these high salt concentrations. Tau, tauPRD or tauMTBD appeared as particles without any filaments or bundles in the absence of actin under these experimental conditions (Additional file 4).
Studies on the interaction of tau with actin have generally focussed on the relationship between MTBD and alpha-actin [7, 9]. Here we show that the proline-rich domain of tau is also capable of binding to actin, promoting G-actin assembly and F-actin bundling. Our results indicate that: (1) tau is still able to interact with actin after the microtubule binding domain has been deleted (mutant tauΔMTBD); (2) the isolated proline-rich domain alone was able to associate with actin; (3) the tauN, tauC truncation mutants, and the ΔPBD&MTBD deletion mutant showed no interaction with actin; and (4) the isolated proline-rich domain was still able to induce F-actin bundling.
Compared to the microtubule binding domain, the proline-rich domain of tau is a relatively less-well characterized domain. However, evidence has shown that this domain is functionally important and participates in multiple biological processes. PRD has been demonstrated to be an indispensable domain for tau's stabilization of microtubules [20, 21]. Further work has shown that PRD may interact with the N-terminal region from another tau molecule to form dimers . Tau was reported to associate with the SH3 domain of fyn and src via its proline-rich domain . Here we show another function of proline-rich domain. As shown in the results above, PRD binds to actin with a lower affinity than native tau (Table 2). The value of Kapp for the complete protein is much lower than that for either tauPRD or tauMTBD. This suggests again that both PRD and MTBD are involved in interactions with actin.
To investigate whether tau or its mutants binds with actin under physiological ionic strength, as suggested by Roger and colleagues , we observed the interaction of tau with actin at different concentrations of NaCl. Our results showed that the interaction between tau and actin decreased significantly with increasing ionic strength. However, high ionic strength could not eliminate the interaction completely. At physiological ionic strength, a relatively weak interaction was still observed using high speed co-sedimentation and electron microscopy. We consider high-speed co-sedimentation and electron microscopy are more sensitive and accurate methods than low-speed co-sedimentation. Our results indicate that interactions between F-actin and tau, tauPRD and tauMTBD persist at physiological ionic strength.
Tau and MAP2c are two major microtubule binding proteins that are considered to be potential cross-linkers between microtubules and actin microfilaments. However, the interaction of MAP2c with actin may be different in nature. Konati and colleagues found that tau and MAP2c have different behaviour when studying their effect on actin filament viscosity . Compared with MAP2c, the binding of tau to F-actin was relatively weak at physiological ionic strength . According to Yamauchi and co-workers , phosphatidyl-inositol completely disrupts MAP2c-induced bundles. However, tau-induced actin bundles are unaffected by phosphatidyl-inositol. These reports indicate that tau and MAP2c behave differently in their interactions with cytoskeletal components.
Tang and coworkers have proposed the polyelectrolyte theory for F-actin bundling similar to DNA condensation . The general behaviour is dictated by the polyelectrolyte nature of F-actin, which causes a class of nonspecific binding by ligands that carry several net positive charges including divalent metal ions and basic polypeptides. Such bundling is induced by electrostatic force, and usually does not require a specific binding site. Tau is a positively charged protein, with a PI of 9.39 (predicted with Lasergene). Moraga and co-workers proposed that electrostatic forces are involved in the interaction between actin and a tau fragment containing a repetitive sequence from the MTBD domain, because selective carbamoylation resulted in a complete loss of the peptide induction of actin bundles . We have shown that the interaction of tau with F-actin is influenced strongly by ionic strength.
These reports provide evidence that tau may act like polycations to induce F-actin to form bundles. The charge distribution in tau is depicted in Additional file 5 which shows that MTBD and PRD are both highly positively charged, while the N-terminal and C-terminal regions are neutral or negatively charged according to Wang and coworkers . We hypothesize that electrostatic force is the basis of the interaction between tauPRD and actin. The relatively weak and nonspecific nature of the electrostatic forces between tau and F-actin does not necessarily mean that the interaction is of no relevance in vivo. Filament concentration could be higher than we can achieve in vitro. Besides, several reports have confirmed the physiological importance of this interaction [4, 13, 14].
In this work, using co-sedimentation assays and solid phase assays we have shown that the proline-rich domain (PRD) of tau binds with G-actin and F-actin. The PRD domain induced G-actin to form filamentous actin and promoted F-actin to form bundles as observed under both atomic force microscopy and electron microscopy. The promotion of actin bundles in the presence of PRD was also observed in the presence of NaCl under physiological conditions. According to the results presented here and other reports in the literature on MTBD associations with actin, it is suggested that both PRD and MTBD are involved in the association of tau with actin.
Construction of mutants
Constructs of tau mutants (tauN, tauPRD, tauMTBD, tauC, tauΔMTBD and tauΔPRD&MTBD) were prepared by PCR or megaprimer PCR amplification  using human tau23 clones as templates. Primers used are listed in Additional file 1. They were then subcloned into the prokaryotic expression vector pET-28a(+) (Novagen, Germany) as NcoI and XhoI fragments. These five mutants each contained a His-tag at the C-terminus. The clones were transformed into E. coli BL21 (DE3) cells after their nucleotide sequences had been confirmed by sequencing.
Expression and purification of tau and its mutants
The tau mutants with His-tags were purified with Ni-NTA resin columns (QIAGEN, Holland) according to the manufacturer's instructions except that cell lysates were boiled for 5 min and centrifuged before loading on the resin. Protein samples were concentrated and then purified further with HiTrap desalting columns (Amersham Pharmacia Biotech, Switzerland). Each purified mutant exhibited a single protein band on Tris-Tricine gels (Additional file 6-a, b). Low molecular weight protein markers (SIBAS, Shanghai, China), mixed with aprotinin (MW 6,500), were used as molecular markers.
The shortest isoform of human tau (tau23) was purified with Q-Sepharose and SP-Sepharose chromatography (Amersham Pharmacia Biotech, Switzerland) as described by Goedert and Jakes . Protein concentrations were measured with BCA protein assay kits (Pierce, USA).
Western blotting of tau or its mutants
Tau mutants were run on Tris-Tricine gels. The bands on the gels were verified by staining with monoclonal anti-His antibodies (Novagen, Germany) (Additional file 1-b). Tau was run on a 12% SDS-PAGE (Additional file 6-a) and verified by staining with tau-13 antibodies (Santa Cruz, USA). All proteins and mutant peptides employed in this work showed single bands on SDS-PAGE gels or Western blots (Additional file 1).
Rabbit skeletal muscle global alpha actin was purified as described by Spudich and Watt . To improve purity, actin was assembled in a buffer containing a low ATP concentration (0.2 mM) during purification. Human platelet G-actin (a mixture of 85% beta and 15% gamma isoforms) was obtained from Cytoskeleton Inc. and was resuspended in buffer A (2 mM Tris-HCl pH 8.0, 0.2 mM ATP, 0.5 mM beta-mercaptoethanol, 0.2 mM CaCl2, and 0.005% NaN3). G-actin proteins were stored at -70°C after freezing in small volumes in liquid nitrogen. These samples were thawed rapidly with gentle agitation under running water at room temperature . Both skeletal muscle and platelet G-actin showed single bands on SDS-PAGE gels (Figure 2 and Additional file 6). To obtain F-actin, G-actin was polymerized in a buffer containing 100 mM KCl, 2 mM MgCl2 and 0.2 mM ATP at 25°C for about 90 min followed by centrifugation (80,000 g, 4°C, and 3 h). The pellet was resuspended in buffer F (100 mM KCl, 1 mM MgCl2, 0.1 mM CaCl2, 0.2 mM ATP, 1 mM Tris-HCl, pH 8.0, 1 mM NaN3).
Solid phase assays
Experimental conditions were as described by Farias and co-workers  with some modifications. The protein (skeletal muscle G-actin, platelet G-actin, skeletal muscle F-actin or platelet F-actin, 5 μg/ml, 50 μl, in 20 mM Tris pH 8.0 buffer) was coated on 96-well microtiter plates (Costar, USA), and incubated at 37°C for 2 h to allow adhesion to the polystyrene surface. After washing three times with PBST (PBS containing 0.2% Tween 20), the sites were saturated by incubation with 200 μl of blocking agent (PBS containing 5% non-fat milk) at 37°C for 30 min. Wells were then washed with binding buffer (50 mM Hepes containing 0.5 mM EGTA and 0.5 mM MgCl2, pH 7.5) and 50 μl of different concentrations of tau or its mutants were added. After incubation at 37°C for 45 min, wells were washed with PBST. Primary antibodies (tau-13, Santa Cruz, USA) for tau, and an anti-His-Tag monoclonal antibody (Novagen) for tau mutants were added and incubated at 37°C for 40 min. Plates were washed with PBST before the addition of the second antibody (1:1,000 dilution, 100 μl) labelled with horseradish peroxidase (HRP), and incubated at 37°C for 30 min. After the wells were washed with PBST, binding of actin with tau was detected using 100 μl of TMB buffer (6 μg/ml TMB, 0.045% H2O2, 0.1 M PB, pH 6.0) for 10 min, and the reaction was terminated with 50 μl of 2 M H2SO4. The binding of tau with actin was recorded by measuring the net change in absorbance at 450 nm by using an automatic solid phase assay plate reader (Thermo, USA). Correas and co-workers did not add excess ATP to the reaction in their study of the interaction of G-actin with tau. Similarly, Roger and coworkers  did not use excess ATP either in studying the bundling of tau with F-actin. Furthermore, Sattilaro and colleagues  have reported that formation of MAP-2-actin bundles is inhibited by millimolar concentrations of ATP. Thus, in this work, excess ATP was not used in the reaction of tau with G-actin or F-actin.
Tau or its mutants was incubated with G-actin or F-actin (from skeletal muscle or platelet) in binding buffer in the presence or absence of NaCl (50 - 500 mM) at 37°C for 40 min, and then centrifuged at either 25,000 g for 30 min or 100,000 g for 1 h at 4°C. The pellet was resuspended and then boiled for 5 min, followed by electrophoresis on SDS-PAGE gels.
Atomic force microscopy (AFM) analysis
G-actin, tau and its mutants were incubated in binding buffer at 37°C for 40 min, and G-actin alone and tau alone were used as controls. Samples were diluted 2 - 20 times with 20 mM Tris-HCl (pH 8.0). A 10 μl drop of the protein sample was deposited on freshly cleaved mica, allowed to stand for 5 min in air, and then washed with three 200 μl aliquots of buffer solution before drying for 4 min in a stream of nitrogen. Tapping mode AFM was performed using a Nanoscope IIIa Multimode-AFM (Veeco Instruments, USA) under ambient conditions. Silicon tips (TESP, Switzerland) with a resonance frequency of about 250 kHz were used at a scan rate of 1-2 Hz. Once the tip was engaged, the set point value was adjusted to minimize the force exerted on the sample while maintaining the sharpness of the image.
Tau (or its mutants) and F-actin were incubated in binding buffer in the presence or absence of NaCl (50 - 500 mM) at 37°C for 40 min. To observe the interaction of F-actin with tau or its mutants, F-actin alone was used as a control under the same conditions. Samples were placed on 300-mesh carbon-coated copper grids for 1 min, washed with H2O and negatively stained with 1% uranyl acetate for 1 min. The specimens were examined with a Tecnai 20 electron microscope (Philips, Holland).
We are grateful to Dr. Goedert (University of Cambridge, Cambridge, U.K.) for kindly providing the human tau clone. We thank Mei Hua Qu and Ying Liu for many valuable suggestions. Ruigang Su and Wei Xu helped with the EM experiments. The Natural Sciences Foundation of China (NSFC 30621004), Project 973 (No. 2006CB500703 and No. 2010CB912303) and the Knowledge Innovation Process Foundation of the Chinese Academy of Sciences (KSCX2-YW-R-119) jointly supported this research.
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