2021 Volume 3 Issue 1 Pages 9-16
Inositol polyphosphate 5-phosphatase K (INPP5K) is an endoplasmic reticulum (ER)-residing phosphoinositide 5-phosphatase that de-phosphorylates PI(4,5)P2, thereby regulating ER morphology. Here, we show that INPP5K interacts with β-tubulin through its 5-phosphatase domain and localizes to microtubules. A cluster of basic amino acids within the 5-phosphatase domain of INPP5K is responsible for this interaction. Alteration of these amino acids to alanine (INPP5K 4A mutant) abolished INPP5K localization to the ER. Expression of the INPP5K 4A mutant induced ER swelling and the withdrawal of ER tubules. Taken together, these findings suggest that INPP5K connects the ER to microtubules and regulates ER morphology.
● INPP5K binds to tubulin through basic amino acid clusters within the 5-phosphatase domain.
● INPP5K connects the endoplasmic reticulum to microtubules.
● INPP5K-Tubulin interaction regulates ER morphology.
The endoplasmic reticulum (ER) is the largest intracellular continuous membrane organelle and enables protein synthesis, ion homeostasis, quality control of newly synthesized proteins, and delivery of proteins to other organelles. Once the threshold of misfolded protein accumulation has been overwhelmed, a signal activates the unfolded protein response (UPR) to synthesize ER chaperones, such as GRP78 and GRP94, which refold the misfolded proteins in the ER lumen and translocate them to the cytoplasm for degradation by the ubiquitin-proteasome system or autophagic degradation. Activation of the UPR increases the ER capacity to fold proteins and maintain ER homeostasis by increasing the size of the ER [1].
The ER network is found in all eukaryotic cellsand is very dynamic in mammalian cells [2]. It undergoes continuous remodeling in shape and volume through remodeling of different subdomains, e.g., reshaping of tubules into sheets and vice versa, to adapt to cellular conditions [3,4,5,6]. The ER is composed of tubules and sheets, and the ratio of tubules to sheets varies between cell lines. ER morphology, dynamics, and subdomain composition are dependent on the actin cytoskeleton and microtubules [7,8,9,10]. Depolymerization of actin filaments by latrunculin A treatment decreased the number of ER sheets [11], demonstrating the connection of the ER network to the actin cytoskeleton. In contrast, manipulation of microtubules shifts ER morphology toward tubules. Depolymerization of microtubules with nocodazole treatment in Huh-7 cells shifts ER morphology toward sheets, decreasing the number of tubules and generating stacked membrane sheets [11]. Microtubule stabilization by taxol results in longer, less fenestrated ER sheets [12]. Distinct ER microdomains have been reported to interact with microtubules, and a number of ER proteins that facilitate ER-microtubule interaction have been found to affect ER morphology [13, 14]. The sheet-localizing proteins Climp-63 and p180 bind directly to microtubules, and their overexpression induces ER sheet proliferation [15,16,17]. Stromal interaction molecule 1 (STIM1), an ER transmembrane protein that mediates store-operated calcium entry, binds directly to EB1 and localizes at growing microtubule ends, which is necessary for ER tubule formation and extension [18].
Inositol polyphosphate 5-phosphatase K (INPP5K), also referred to as skeletal muscle- and kidney-enriched inositol polyphosphate 5-phosphatase (SKIP) or mouse putative phosphatase (Pps), is a phosphoinositide phosphatase that hydrolyzes PI(4,5)P2 and PIP3 [19,20,21]. Human bi-allelic mutations that decrease the phosphatase activity of INPP5K give rise to congenital muscular dystrophy, characterized by cataracts, intellectual impairments, and short stature [22, 23]. Arl6IP1, a reticulon-like protein with the ability to shape the ER membrane to generate and stabilize ER tubules, binds to the SKIP C-terminal homology (SKICH) domain of INPP5K and mediates the anchoring of INPP5K to the ER, while INPP5K is enriched in newly formed ER tubules along microtubules [24]. The knockdown of INPP5K results in the ER remodeling: a decrease in ER tubules and an increase in ER sheets. Although emerging evidence suggests the importance of ER remodeling in the development of neuronal axons and dendrites as well as in the pathogenesis of hereditary spastic paraplegia, an inherited neuronal disorder [25, 26], the molecular basis underlying the impact of INPP5K on ER morphological changes remains unclear. In this study, we aimed to demonstrate that INPP5K interacts with tubulin and connects the ER to microtubules and that the expression of the defective tubulin-binding mutant of INPP5K results in ER swelling.
Rabbit polyclonal and mouse monoclonal antibodies against INPP5K (catalog no. C135932 and C308501) were purchased from Lifespan Biosciences (Seattle, WA, USA). Anti-β-tubulin rabbit polyclonal antibody (catalog no. 2146) was purchased from Cell Signaling Technology (Beverly, MA, USA). Anti-FLAG M2 monoclonal antibody (catalog no. F3165), anti-FLAG M2 agarose affinity gels (catalog no. A2220), anti-α-tubulin mouse monoclonal antibody (catalog no. T6074), and insulin were purchased from Sigma Aldrich (St. Louis, MO, USA).
ConstructsINPP5K wild type (INPP5K WT) expression vectors were generated by introducing cDNAs encoding human INPP5K into pEGFP (enhanced green fluoresence protein)-C1 (Clontech Laboratories, Mountain View, CA, USA), and p3×FLAG-CMV8 (Sigma Aldrich) vectors. A phosphatase-negative D310G mutant (D310G), a SKICH domain alone mutant (INPP5K 318–448), and a mutant lacking SKICH domain position 1 to 317 (INPP5K 1–317), a mutant of the basic amino acid cluster within the phosphatase domain (INPP5K 4A; INPP5K p.K261A/K262A/R263A/K264A), were generated by PCR and cloned into expression vectors. The pDsRed2-ER expression vector was purchased from Takara Bio (Kusatsu, Japan).
Cell culture and transfection of plasmids, and RNA interference (RNAi)HeLa and C2C12 cell lines were purchased from the American Type Culture Collection (Manassas, VA, USA). The 293F cells were purchased from Thermo Fisher Scientific (Waltham, MA, USA). HeLa and C2C12 cells were grown in Dulbecco’s modified Eagle’s medium (DMEM; Wako Pure Chemical Industries) containing 4.5 g/l glucose, 10% fetal bovine serum (FBS), and antibiotics (100 µg/ml streptomycin and 100 units/ml penicillin) at 37°C in an atmosphere containing 5% CO2. The 293F cells were cultured in FreeStyleTM 293 serum-free medium at 37°C in an atmosphere containing 5% CO2. For transfection, cells were cultured to 50% confluence and transfected with 1 µg of plasmid DNA. The cells were then cultured in DMEM containing 10% FBS for 48 hr. Small interfering RNA (siRNA) duplexes were purchased from Thermo Fisher Scientific. The following oligonucleotides were used in this study: control, 5′-GAGCAACTGCGTGTCGAATCTCTTA-3′; mouse Pps, 5′-GAGTCAACGTCTGCCTGAAGCTTTA-3′. Twenty nanomoles of control and INPP5K siRNA duplexes were transfected into C2C12 myoblast cells. The cells were cultured in DMEM containing 10% FBS for 24 hr. Subsequently, 1 µg of plasmid DNA was transfected into the cells for re-expression, and the cells were cultured for another 48 hr.
ImmunoblottingForty-eight hours after transfection, the cells were serum-starved in serum-free medium for 24 hr. Cells were then washed with phosphate-buffered saline and lysed with cell lysis buffer containing 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 2 mM EDTA, 5 mM NaF, 1 mM Na3VO4, 1 mM dithiothreitol, 1 mM PMSF, and 1% Triton X-100. Cell lysates were centrifuged at 14,000 × g for 10 min, and the supernatants were solubilized in sodium dodecyl sulfate (SDS) sample buffer, resolved by SDS-polyacrylamide gel electrophoresis (SDS-PAGE), and transferred onto a nitrocellulose membrane. For western blot analysis, the membrane was incubated with the primary antibody at 25°C for 2 hr, followed by washing with Tris-buffered saline (25 mM Tris-HCl [pH 7.5], 150 mM NaCl) containing 0.05% Tween-20. The membrane was then incubated with the secondary antibody at room temperature for 2 hr. Densitometry was used to quantify protein levels.
ImmunoprecipitationCell lysates were subjected to immunoprecipitation in cell lysis buffer with anti-FLAG antibody at 4°C for 4 hr. Twenty-five micrograms of protein A-agarose beads (Thermo Fisher Scientific) was added 2 hr before the end of the incubation period. The beads were washed five times with wash buffer (20 mM Tris-HCl [pH 7.5], 150 mM NaCl, 2 mM EDTA, 5 mM NaF, 1 mM Na3VO4, 1 mM DTT, 1 mM PMSF, and 1% Nonidet P-40) before the immunoprecipitates were subjected to SDS-PAGE. For immunoprecipitation using the FLAG antibody, lysates were incubated with anti-FLAG M2 agarose affinity gel beads for 4 hr at 4°C in cell lysis buffer and then washed five times with wash buffer prior to immunoblotting.
ImmunofluorescenceForty-eight hours after transfection, cells were serum-starved for 24 hr and then stimulated with insulin (100 nM) for 10–30 min. The cells were fixed with 3.7% formaldehyde, permeabilized with PBS containing 0.2% Triton X-100 for 10 min, and incubated with 1% FBS in PBS for 30 min to block non-specific antibody binding. The cells were then immunostained with the first antibody for 1 hr at room temperature and then incubated with the appropriate fluorescein-labeled secondary antibody. F-actin was visualized using AlexaFluor-647-labeled phalloidin (Life Technologies). After a brief wash with PBS, coverslips were mounted onto slides using PermaFluor Aqueous Mounting Medium (Thermo Fisher Scientific), and cells were observed under a FluoView 1000-D confocal microscope (IX81; Olympus, Tokyo, Japan) equipped with 473-, 568-, and 633-nm diode lasers (Olympus) through a 60 Å ~ oil immersion objective, NA 1.35 objective lens (Olympus) and using the FluoView software (Olympus). The acquired images were processed using Photoshop (Adobe Systems, San Jose, CA, USA).
Measurement of ER areaThe ER area of each cell was quantified by measuring the extent of DsRed2-Sec61β staining. For each cell, the percentage area occupied by the ER in the cytoplasm was determined using the Image J 1.47 software. Thirty cells were measured for each experiment.
Statistical analysisPrior to analysis, all data were visually inspected to ensure that they were normally distributed. Differences between treatments were determined using Student’s t-test. All values are presented as mean ± standard deviation.
To identify the INPP5K-interacting proteins, 3xFLAG-tagged INPP5K was expressed in adherent 293F cells, and lysates were immunoprecipitated using an anti-FLAG antibody. A band with a molecular weight of approximately 80 kDa was previously identified as GRP78, also referred to as BiP, by mass spectrometric analysis [27]. Another protein with a molecular weight of approximately 50 kDa was identified as β-tubulin (Fig. 1A). INPP5K consists of an N-terminal 5-phosphatase catalytic domain (INPP5K 1-317) and an INPP5K C-terminal homology (SKICH) domain (INPP5K 318–448) (Fig. 1B). Immunoprecipitation analysis using 293F cell lysates showed that the phosphatase domain (3×FLAG INPP5K 1-317), but not the SKICH domain alone (3×FLAG INPP5K 318–448), interacted with α-tubulin (Fig. 1C). To identify direct binding between INPP5K and β-tubulin, a glutathione S-transferase (GST) pulldown assay using purified GST-fused INPP5K protein and α-tubulin was performed. Neither the GST-INPP5K WT nor the GST-INPP5K mutants directly bound α-tubulin (data not shown), indicating that α-tubulin indirectly binds to the 5-phosphatase domain of INPP5K. INPP5K possesses a cluster of basic amino acids within the 5-phosphatase domain, containing K261, K262, R263, and K264. To identify the role of these amino acids in the interaction with tubulin, these amino acids were substituted with alanine (INPP5K K261A/K262A/R263A/K264A, Fig. 1B) to form the mutant INPP5K 4A. An immunoprecipitation assay using HeLa cells showed that, unlike the 3×FLAG INPP5K WT, the 3×FLAG INPP5K 4A mutant abolished its interaction with α- and β-tubulin (Fig. 1D). An ER-delocalizing INPP5K D361A mutant was immunoprecipitated with α- and β-tubulin (Fig. 1D).
Inositol polyphosphate 5-phosphatase K (INPP5K) interacts with tubulin. (A) Identification of tubulin as an INPP5K-binding protein by mass spectrometry: The 293F cells were transfected with 3×FLAG-tagged INPP5K WT, and the lysates were immunoprecipitated with an anti-FLAG antibody. Non-transfected cells were used as a control. Arrows on the right side indicate the position of tubulin, GRP78, and the 3×FLAG INPP5K WT. (B) Domain structures of INPP5K and INPP5K mutants used in this study. (C) The 293F cells were made to express 3×FLAG-tagged wild-type INPP5K (INPP5KWT), INPP5K 1–317, and INPP5K 318–448. Cell lysates were immunoprecipitated with an anti-FLAG antibody. Arrows on the left side indicate the positions of 3×FLAG INPP5K WT, 3×FLAG INPP5K 1–317, and 3×FLAG INPP5K 318–448, and the position of tubulin is indicated on the right side. (D) HeLa cells were made to express 3×FLAG-tagged INPP5K WT, INPP5K D361A, and INPP5K 4A mutants, and the lysates were immunoprecipitated with an anti-FLAG antibody. The results of western blot analysis using anti-α-tubulin and β-tubulin antibodies are shown.
The SKICH domain is necessary for INPP5K localization to the ER and plasma membrane [28]. This suggests that INPP5K interacts with microtubules. The immunofluorescence of α-tubulin in HeLa cells showed that many of microtubule filaments co-localized with EGFP-INPP5K WT, especially at the cell periphery (Fig. 2A and 2B). The INPP5K phosphatase domain (EGFP-INPP5K 1–317) exhibited partial co-localization with α-tubulin, whereas the SKICH domain (EGFP-INPP5K 318–448) did not (Fig. 2C and 2D). The EGFP-INPP5K 4A mutant showed cytosolic localization and did not co-localize with α-tubulin (Fig. 2B). These results indicate that the basic amino acids within its phosphatase domain mediate the interaction of INPP5K with microtubules.
Distribution of Inositol polyphosphate 5-phosphatase K (INPP5K) and microtubules in C2C12 cells. C2C12 cells were made to express enhanced green fluoresence protein (EGFP) (A), EGFP-INPP5K WT (B), EGFP- INPP5K 1–317 (C), INPP5K 318–448 (D), and EGFP-INPP5K 4A (E). α-Tubulin was visualized using an α-tubulin antibody. Enlarged images of the boxed areas are shown in the lower panels. Scale bar, 20 µm.
It has been reported that depletion of INPP5K or inhibition of its phosphatase activity induces ER swelling [24]. Knockdown of INPP5K in DsRed2-ER-expressing C2C12 cells led to an increase in the ER area, with the retraction of ER tubules (Fig. 3A). To determine the significance of the INPP5K-microtubule interaction for ER morphology, we expressed EGFP-human INPP5K WT or mutants along with DsRed2-ER in INPP5K-depleted C2C12 cells containing the ER-targeting sequence of calreticulin and the ER retention sequence KDEL fused to the DsRed2 vector (Fig. 3B). Although the re-expression of the control INPP5K WT induced ER retraction, expression of the INPP5K 4A mutant as well as the phosphatase-dead INPP5K D310G mutant did not change the amount of ER area compared to that in the control GFP-expressing cells (Fig. 3B and 3C). These results suggest that the interaction between INPP5K and microtubules is likely to mediate the control of ER morphology.
Inositol polyphosphate 5-phosphatase K (INPP5K) changes endoplasmic reticulum (ER) morphology. (A) C2C12 cells were transfected with control or INPP5K- siRNA together with DsRed2-ER. F-actin was visualized using Alexa Fluor 647-conjugated phalloidin. Scale bar, 20 µm. (B) C2C12 cells were transfected with enhanced green fluoresence protein (EGFP), EGFP-INPP5K WT, EGFP- INPP5K D310G, and EGFP- INPP5K 4A together with INPP5K siRNA and DsRed2-ER. Enlarged images of the boxed areas are shown in the lower panels. Scale bar, 20 µm. (C) The compositions of the ER area were determined using the intensities of DsRed2-ER in A and B. Mean ± standard error was calculated for each condition, and the number of cells analyzed is indicated.
INPP5K represents a molecular link between ER stress and insulin-dependent glucose uptake in skeletal muscle [29]. Here, we demonstrated, by immunoprecipitation analysis, that INPP5K interacts with tubulin through its basic amino acid cluster within the 5-phosphatase domain. These basic amino acid clusters are INPP5K-specific and are not conserved among other 5-phosphatases. However, the in vitro binding assay of the purified proteins did not show direct binding between them and INPP5K. Although this does not rule out the possibility that posttranslational modification may be necessary for this direct binding, it may also be possible that INPP5K and tubulin bind indirectly. Desmoplakin, periplakin, and envoplakin are members of the plakin family of intermediate filament-binding proteins. Basic amino acid clusters of these proteins recognize acidic side chains of vimentin intermediate filaments [30, 31]. Although further binding assays between INPP5K and intermediate filaments are required, it may be possible that INPP5K interacts with microtubules via intermediate filaments. Intracellular localization analysis revealed that INPP5K localizes to microtubules via the 5-phosphatase domain. Thus, INPP5K might be one of the proteins that connects microtubules and the ER, such as STIM1, because INPP5K anchors onto the ER membrane depending on its interaction with the ER membrane protein Arl6IP1 via its C-terminal SKICH domain [32]. However, based on the results of the study reporting these findings, the SKICH domain alone exhibited cytosolic localization and was no longer anchored to the Arl6IP1-positive ER tubules [24]. The INPP5K 4A mutant showed cytosolic localization and did not localize to the ER, suggesting that not only the SKICH domain, but also the 5-phosphatase domain, may be necessary for INPP5K anchoring at the ER membrane. In contrast, the ER-delocalizing INPP5K D361A mutant within the SKICH domain was immunoprecipitated with α- and β-tubulin, suggesting that ER localization is not necessary for INPP5K to interact with microtubules.
Our results also suggest that INPP5K may be a molecular link between the ER and microtubules and may thus be necessary for maintaining ER architecture, since the number of ER-residing proteins that mediate ER-microtubule interplay have been found to affect ER morphology. Depletion of INPP5K in C2C12 cells leads to a decrease in ER tubules and ER sheet expansion. These effects were rescued by the expression of INPP5K WT but not by the INPP5K 4A mutant or the phosphatase-dead D310G mutant, which failed to induce ER retraction (Fig. 3). The congenital muscular dystrophy-related INPP5K mutant, p.I363T, is defective for interaction with Arl6IP1 and ER localization and thus causes ER swelling. Based on our results, in addition to the localization of INPP5K at the ER via Arl6IP1, its association with microtubules may play a role in the regulation of ER morphology [24]. Since ER/sarcoplasmic reticulum (SR) is necessary for store-operated calcium entry-induced muscle excitation-contraction [3] and ER membrane expansion is induced by an excess UPR or by a defect in the degradation of unfolded proteins [33], INPP5K may function in ER homeostasis functions, including Ca2+ homeostasis, lipid transfer, and protein quality control. Although further validation of the molecular mechanisms of the INPP5K-mediated control of ER architecture needs to be conducted, our findings provide new insights into the regulation of ER functions.
A number of proteins involved in shaping the ER network are mutated in neurological disorders, particularly hereditary spastic paraplegia (HSP), an inherited neurological disorder characterized by spastic weakness and loss of muscle control, emphasizing the importance of proper ER morphology in the maintenance of highly polarized neurons [34]. More than half of HSP cases result from autosomal dominant mutations in atlastin-1 (SPG3A) or receptor expression enhancing protein 1 (REEP1; SPG31). REEP1 forms protein complexes with atlastin-1 and spastin, and in the tubular ER of COS7 cells; REEP1 proteins are required for the formation of an ER tubular network in vitro, while REEP1 binds microtubules and promotes the alignment of ER tubules along the microtubules in COS7 cells. One study showed that the SPG31 mutant REEP1 did not interact with microtubules and disrupted the ER network, which raises the possibility that the proper regulation of ER morphology may play an essential role in the development and function of neurons [35,36,37]. In addition, REEP1 has been shown to promote ER stress resistance and to prevent the formation of Tau aggregates and Tau-mediated cytotoxicity [37]. Pathological aggregation of Tau is a common characteristic observed in various neurological diseases, suggesting the importance of ER morphology in the pathogenesis of neurological disorders. The INPP5K gene has been reported to be upregulated in regenerating mammalian spinal cord axons. Overexpression of INPP5K in acutely dissociated mouse cortical neurons enhanced neurite outgrowth and increased the number of branches per neuron after spinal cord injury [38]. The finding that depletion of Arl6IP1, an ER membrane INPP5K-binding protein, results in degenerative axonal diseases [39, 40] and fragmentation of the smooth ER in axons [41] raises the possibility that INPP5K may link the ER to the microtubules and help coordinate the progression of neuronal growth through the extension of ER tubules. In the skeletal muscle, ER/SR is well developed and controls various functions, including the regulation of Ca2+ homeostasis during muscle excitation-contraction. Although little is known about the importance of ER morphology in the induction of muscle disorders, our results may help understand the role of INPP5K in the pathogenesis of skeletal muscle disorders.
In conclusion, our results provide evidence that INPP5K connects the ER and microtubules and modulates ER morphology. Expression of the INPP5K 4A mutant caused defective assembly of microtubules and thereby induced ER swelling. These actions of INPP5K appear to be independent of INPP5K phosphoinositide phosphatase activity. Our results provide new evidence to better understand the roles of INPP5K in the control of ER shape and distribution in normal cellular function.
The authors declare no conflicts of interest.
This work was supported by a grant to T.I. from the Japan Society for the Promotion of Science (JSPS; Kakenhi Grant Number, 19K07368). These funding agencies had no role in the study design, data collection, data analysis, decision to publish, or manuscript preparation.
