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Division of Regulatory Glycobiology, Institute of Molecular Biomembrane and Glycobiology, Tohoku Medical and Pharmaceutical University, 4-4-1 Komatsushima, Aoba-ku, Sendai Miyagi 981-8558, Japan,
Division of Regulatory Glycobiology, Institute of Molecular Biomembrane and Glycobiology, Tohoku Medical and Pharmaceutical University, 4-4-1 Komatsushima, Aoba-ku, Sendai Miyagi 981-8558, Japan,
Department of Life Science and Biotechnology, National Institute of Advanced Industrial Science and Technology, 1-1-1 Umezono, Tsukuba, Ibaraki 305-8568, Japan, and
Division of Regulatory Glycobiology, Institute of Molecular Biomembrane and Glycobiology, Tohoku Medical and Pharmaceutical University, 4-4-1 Komatsushima, Aoba-ku, Sendai Miyagi 981-8558, Japan,
Division of Regulatory Glycobiology, Institute of Molecular Biomembrane and Glycobiology, Tohoku Medical and Pharmaceutical University, 4-4-1 Komatsushima, Aoba-ku, Sendai Miyagi 981-8558, Japan,
Aberrant N-glycan sialylation of glycoproteins is closely associated with malignant phenotypes of cancer cells and metastatic potential, which includes cell adhesion, migration, and growth. Recently, phosphatidylinositol 4-kinase IIα (PI4KIIα), which is localized to the trans-Golgi network, was identified as a regulator of Golgi phosphoprotein 3 (GOLPH3) and of vesicle transport in the Golgi apparatus. GOLPH3 is a target of PI4KIIα and helps anchor sialyltransferases and thereby regulates sialylation of cell surface receptors. However, how PI4KIIα-mediated sialyation of cell surface proteins is regulated remains unclear. In this study, using several cell lines, CRISPR/Cas9-based gene knockout and short hairpin RNA–mediated silencing, RT-PCR, lentivirus-mediated overexpression, and immunoblotting methods, we confirmed that PI4KIIα knockdown suppresses the sialylation of N-glycans on the cell surface, in Akt phosphorylation and activation, and integrin α3-mediated cell migration of MDA-MB-231 breast cancer cells. Interestingly, both integrin α3β1 and PI4KIIα co-localized to the trans-Golgi network, where they physically interacted with each other, and PI4KIIα specifically associated with integrin α3 but not α5. Furthermore, overexpression of both integrin α3β1 and PI4KIIα induced hypersialylation. Conversely, integrin α3 knockout significantly inhibited the sialylation of membrane proteins, such as the epidermal growth factor receptor, as well as in total cell lysates. These findings suggest that the malignant phenotype of cancer cells is affected by a sialylation mechanism that is regulated by a complex between PI4KIIα and integrin α3β1.
Alteration of the sialylation of glycoproteins has often been observed in several types of malignant tumors, such as those found in breast, ovary, and colorectal cancers (
). Sialylation is linked via either an α2,3 or an α2,6 bond to Gal/GalNAc and via an α2,8 bond to sialic acid in glycoproteins through a group of sialyltransferases. The up-regulation of cell surface sialic acid is thought to control cell phenotypes such as cell adhesion, migration, immune response, apoptosis, and cell epithelial–mesenchymal transition (EMT).
). Some transcription factors are critical for transcriptional activation of sialytransferases in cancer cells. For example, the expression level of α2,6-galactoside sialyltransferase 1 (ST6GAL1) is up-regulated by the RAS oncogene and increases the α2,6 sialylation of β1 integrins, which promotes integrin-mediated cell migration, adhesion, and cell proliferation (
As described above, sialylation levels are mainly dependent on their gene expression levels, whereas other mechanisms for regulation are usually neglected. However, Popoff and co-workers (
) independently reported that Golgi phosphoprotein 3 (GOLPH3), which has been identified as an oncogenic protein and is increased in several human solid tumors (
), could anchor sialyltransferases to regulate sialylation on cell surface receptors without regulating the gene expression levels of sialyltransferases. In particular, suppression of GOLPH3 attenuated the levels of cellular sialylation and integrin-dependent cell migration. Furthermore, tumor formation was significantly reduced in mice implanted with GOLPH3 shRNA-expressing cells (
). GOLPH3 has multiple cellular functions in vesicle trafficking and in support of Golgi apparatus structure, which has specific affinity for phosphatidylinositol 4 (PI4P) (
). Given the importance of GOLPH3 in sialylation and cell functions, it is plausible that PI4P expression at the TGN may influence sialylation.
Integrins are heterodimeric cell surface adhesion receptors and major carriers of sialyation. The interaction between integrin and the extracellular matrix (ECM) is essential for cell adhesion, migration, viability, and proliferation (
β1 integrin inhibitory antibody induces apoptosis of breast cancer cells, inhibits growth, and distinguishes malignant from normal phenotype in three dimensional cultures and in vivo.
). In fact, glycosylation is a key regulator that plays important roles in modulating integrin functions. For example, integrin α5β1 binding to fibronectin and integrin-mediated cell spreading and migration are modulated by overexpression of glycosyltransferase genes such as N-acetylglucosaminyltransferase III and V or ST6GAL1 (
Expression of N-acetylglucosaminyltransferase III suppresses α2,3-sialylation, and its distinctive functions in cell migration are attributed to α2,6-sialylation levels.
). In addition, sialylated N-glycans on the membrane-proximal domain of integrin β1 play crucial roles in integrin activation and in complex formation between integrin and EGF receptors and syndecan-4 to regulate cell migration and proliferation (
In contrast to the regulation of integrin functions from the extracellular domain, it is well known that integrin function is regulated by its association with cytoplasmic molecules such as focal adhesion kinase and phosphatidylinositol 3-kinase (
). Other studies have detected PI4K activity in the immune complex with integrin β1, suggesting that integrin might also regulate the biosynthesis of PI4P (
Highly stoichiometric, stable, and specific association of integrin α3β1 with CD151 provides a major link to phosphatidylinositol 4 kinase, and may regulate cell migration.
In this study, to further understand the underlying mechanism for GOLPH3 expression on sialylation and cell functions, we investigated the effects of PI4KIIα, which is one of the regulators of GOLPH3 in breast cancer MDA-MB-231 cells. We found that the sialylation on integrins, Akt phosphorylation, and integrin α3-mediated cell migration all were significantly inhibited in PI4KIIα knockdown cells. It was interesting that overexpression of both PI4KIIα and integrin α3 greatly increased sialylation. Conversely, knockout of integrin α3 significantly inhibited sialylation in membrane proteins. These findings suggest a novel mechanism for sialylation, which suggests a new concept for the regulation of glycosylation in cell biology.
Results
We recently reported that the expression of GOLPH3 up-regulated cell surface sialylation and cell migration (
). However, the molecular mechanism of posttranslational modification of sialylation on the cell surface remains unclear. Considering that GOLPH3 exhibits a highly selective affinity for PI4P that is similar to the canonical pleckstrin homology (PH) domain in the TGN (
), we hypothesized that the expression of PI4P in the TGN could affect sialylation and cell properties. PI4P is produced mainly by PI4Ks, which are classified as type II and III according to their sensitivity to inhibitors (
). Therefore, we chose PI4KIIα for functional analysis, and established a line of doxycycline (DOX)–controlled PI4KIIα silencing in MDA-MB-231 breast cancer cells.
Knockdown of PI4KIIα affected the sialylation of N-glycans
First, we verified the expression levels of PI4KIIα and sialyltransferases (ST3GAL3, ST3GAL4, and ST6GAL1) that involve major sialylated N-glycans (
). RT-PCR analysis showed that the expression levels of PI4KIIα mRNA were significantly decreased in knockdown cells, whereas the expression levels of ST6GAL1 for α2,6 sialylation and those of ST3GAL3 and ST3GAL4 for α2,3 sialylation were all similar in both cells (Fig. 1A). Second, we verified the expression levels and the distributions of PI4P in KD cells. The PH domain of four-phosphate adaptor protein 1 (FAPP1), which preferentially recognizes PI4P, is known for its use as a monitor for PI4P distribution (
A plasma membrane pool of phosphatidylinositol 4-phosphate is generated by phosphatidylinositol 4-kinase type-IIIα: studies with the PH domains of the oxysterol binding protein and FAPP1.
). In control cells, PI4P was localized near the medial Golgi marker GM130, as expected (Fig. 1B). In contrast, PI4P staining was significantly decreased in knockdown (KD) cells (Fig. 1B). These data confirmed that PI4KIIα is mainly involved in the synthesis and localization of PI4P in the TGN, which agrees with previous reports (
Figure 1Effects of PI4KIIα gene knockdown on the sialylation of N-glycans. A, RT-PCR using total RNA extracted from the control (Ctrl) and PI4KIIα KD of MDA-MB-231 cells were carried out to examine the expression levels of PI4KIIα and sialyltransferases that involve sialylated N-glycans. The expression level of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as a loading control. ST3GAL, β-galactoside α2,3-sialyltransferase. B, the amounts and distribution of PI4P in KD and Ctrl cells were examined by expressing mRFP-FAPP, which specifically binds to PI4P. These cells were also immunostained with anti-GM130 antibody, which is a medial Golgi marker. The immunostaining intensities for PI4P were quantified using ImageJ software, averaged by cell numbers, and normalized to the control values (percent). Values represent the means ± S.E. (n = 10). **, p < 0.01 (Welch’s t test). Scale bar = 10 μm. C, Cells were incubated with biotin-conjugated SNA or WGA and then fixed and visualized by Alexa Fluor–streptavidin conjugates. Scale bar = 20 μm. D, preparation of PA oligosaccharides and calculation of ratios of sialylated N-glycans versus total N-glycans as described under “Experimental procedures.” Data were normalized to the control. E, equal amounts of cell lysates from Ctrl and KD cells were precipitated by SSA-agarose (preferentially recognizes the α2,6 sialylation) and probed with anti-β1 antibody (top panel). The expression levels of integrin β1 in total cell lysates were verified by Western blotting with anti-β1 antibody (center panel). α-Tubulin was used as a loading control. KD and Ctrl cells refer to transfectants with DOX-inducible knockdown of PI4KIIα cultured with or without DOX, respectively. IB, immunoblot.
Next, we tested whether decreased PI4P affects sialylation expression on the cell surface. As shown in Fig. 1C, top panels, the intensities of the staining in KD cells with Sambucus nigra agglutinin (SNA) lectin, which preferentially recognizes α2,6 sialylation of glycoproteins, was weaker than that in control cells. However, the intensities of staining with WGA lectin, which preferentially recognizes GlcNAc residues and hybrid-type and lactohexose N-glycans (
), were almost equal in both cells (Fig. 1C, bottom panels). These data suggested that the expression levels of sialylated glycans were decreased in KD cells. Furthermore, the decrease was confirmed by quantitative analysis using HPLC. Total pyridylaminated (PA) N-glycans of cells were examined via anion exchange chromatography, and the ratios of sialylated N-glycans versus total N-glycans were calculated. The expression levels of sialylated N-glycans were significantly suppressed in KD cells (Fig. 1D). A similar change in α2,6 sialylation on specific membrane proteins such as integrin β1 was also consistently observed (Fig. 1E) using Sambucus sieboldiana agglutinin (SSA)–agarose. These data suggest that the expression levels of PI4P in the TGN are important for sialylation.
Knockdown of PI4KIIα significantly inhibited cell migration and Akt phosphorylation
Alterations in N-glycosylation affect many cellular events involved in cellular signaling and cell migration. Here we examined the effects of PI4KIIα KD on cell migration and Akt activation. As shown in Fig. 2A, cell migration on laminin-332 in a Transwell assay was significantly reduced in KD cells compared with control cells. The expression levels of integrin α3β1, a major receptor for laminin-332 and α5β1 on the cell surface were similar in both cells (Fig. 2B). Activation of Akt is required for integrin-mediated cell migration (
). As shown in Fig. 2C, the level of phosphorylated Akt was apparently decreased in KD cells compared with control cells. These data suggest that PI4P in the TGN plays an important role in Akt activation and in integrin α3–mediated cell migration.
Figure 2Comparison of cell migration and cellular signaling between Ctrl and KD cells. A, cell migration toward laminin-332 was determined by a Transwell assay as described under “Experimental procedures.” The quantitative data were obtained by counting the cell number of five random fields. The p values were calculated using Welch’s t test. Values represent the means ± S.E. **, p < 0.01. B, comparison of the expression levels of α3β1 and α5β1 integrins on the cell surface between Ctrl and KD cells. Both Ctrl and PI4KIIα KD cells were incubated with (bold line) or without (gray shadow) antibodies against integrin α3, α5, or β1 and subjected to FACS analysis. C, comparison of Akt activation upon FBS stimulation between Ctrl and KD cells. After starvation, cells were supplied with fresh medium with (+) or without (−) 3% FBS. The cell lysates were then analyzed by Western blotting with anti-p-Akt or total Akt antibody.
Complex formation between integrin α3 and PI4KIIα is important for sialylation
As described above, PI4KIIα regulated sialylation and the integrin α3–mediated phenotype. Therefore, we examined the interaction between PI4KIIα and integrin α3 in the Golgi apparatus. First, we used immunostaining to assess the intracellular distributions of PI4KIIα and α3. Immunostaining showed that PI4KIIα was localized in the vicinity of GM130 and that it was extensively co-localized with integrin α3 (Fig. 3A). Notably, co-immunoprecipitation with anti-integrin β1 showed that it was α3, rather than α5, that interacted with PI4KIIα in 293T cell stable expression of PI4KIIα and/or either integrin α3 or α5 (Fig. 3B). The interaction between PI4KIIα and α3 was further reciprocally confirmed by immunoprecipitation with either anti-α3 antibody (Fig. 3C) or anti-GFP antibody (Fig. 3D).
Figure 3Interaction between integrin α3β1 and PI4KIIα and the effect on sialylation. A, MDA-MB-231 cells overexpressed with GFP-PI4KIIα were fixed and then stained with either anti-GM130 (top panel) or anti-integrin α3 (bottom panel) antibody, followed by detection using anti-mouse Alexa 647 conjugates. Scale bar = 20 μm. B, three overexpressed 293T cells were established by lentivirus infection: PI4KIIα alone; PI4KIIα, GFP-tagged α3 and β1; and PI4KIIα, GFP-tagged α5 and β1 integrin. Equal amounts of cell lysates were immunoprecipitated (IP) with anti-integrin β1 antibody, followed by detection with the indicated antibodies. The cell lysates were used as input to show similar expression levels of PI4KIIα, integrin α3β1, or α5β1 among these cells. C and D, reciprocal immunoprecipitation was performed using either anti-α3 antibody (C) or anti-GFP for tagged PI4KIIα (D) using cell lysates from 293T cells stably overexpressed with or without GFP tagged PI4KIIα, which were further transiently transfected with or without α3 and β1 integrin. The cell lysates were used as input to show similar expression levels of integrin α3 and GFP-PI4KIIα. E, equal amounts of cell lysates were precipitated with either SSA-, MAM-, or ConA-conjugated agarose, followed by detection with antibody against integrin β1. F, 293T cells expressing the indicated genes were incubated with (bold line) or without (gray shading) biotin-conjugated MAM or SNA and subjected to FACS analysis.
Next we wondered whether this interaction was required for efficient sialylation. To address this question, we precipitated those cell lysates with SSA or Maackia amurensis mitogen (MAM) agarose, which preferentially recognize α2,6-sialylated and α2,3-sialylated N-glycans, respectively, and then performed Western blotting against integrin β1. Interestingly, both sialylated N-glycans were up-regulated on integrin β1 in cells that expressed both PI4KIIα and integrin α3 but not in those that expressed PI4KIIα and α5 (Fig. 3E, top and center panels). In contrast to SSA or MAM lectin blotting, ConA lectin blots showed a similar level in both integrin α3- and α5-expressing cells (Fig. 3E, bottom panel). ConA lectin equally recognized both mature and immature forms of integrin β1 in α3-expressing cells but only the most immature form of β1 in α5-expressing cells. These data further suggest the importance of α3 for sialylation. Data obtained from flow cytometry analysis via MAM lectin consistently supported this observation, although SSA lectin showed no significant changes in these cells (Fig. 3F). Therefore, these results clearly suggest that intracellular complex formation between PI4KIIα and α3 regulates the biosynthesis of sialylated N-glycans.
Expression of integrin α3 is important for efficient sialylation
To further verify the importance of complex formation between PI4KIIα and integrin α3 for cellular sialylation, we used the CRISPR/Cas9 system to construct an integrin α3 knockout (KO) MDA-MB-231 cell line. Flow cytometric analysis revealed a KO efficiency of more than 95% (Fig. 4A), which also was confirmed by Western blotting with anti-integrin α3 (Fig. 4B). Notably, the band mobility of integrin β1 on SDS-PAGE in KO cells was faster than in the control cells, suggesting that integrin α3 deficiency leads to an accumulation of the immature β1 form without sialylation. It is well known that integrin α3 is important for laminin-mediated cell adhesion, migration, and several forms of cellular signaling (
Suppression of Integrin α3β1 in breast cancer cells reduces cyclooxygenase-2 gene expression and inhibits tumorigenesis, invasion, and cross-talk to endothelial cells.
), expression of α3β1 regulated cell morphology and that of the actin cytoskeleton by promoting lamellipodium and filopodium formation on laminin-332, a specific ligand for integrin α3, whereas lack of α3 blocked their formation (Fig. 4C). There were no significant differences between two cells spread on collagen, which is a specific ligand for integrin α1 and α2. Furthermore, cell spreading and migration on laminin-332 were inhibited in KO cells compared with control cells (Fig. 4, D and E).
Figure 4Effects of integrin α3 deficiency on cell spreading and migration. Establishment of integrin α3 KO MDA-MB-231 cells using the CRISPR/Cas9 system is described under “Experimental procedures.” The efficiency of α3 KO was assessed via flow cytometric analysis (A) and Western blotting (B). A, Both α3 KO and Ctrl cells were stained with (bold line) or without (gray shading) anti-α3 or β1 integrin antibody and subjected to FACS analysis. B, the same amount of cell lysate was subjected to Western blotting to detect the indicated antibodies. C, cells were replated on coverslips coated with the indicated ECM, followed by incubation for 30 min, and were then fixed and stained with Alexa Fluor–conjugated phalloidin. Representative images are shown. Scale bar = 10 μm. D, cells were detached and then replated on laminin 332-coated dishes. After incubation for 15 min, the cells were fixed. The ratios of cell spreading were calculated. Cell spreading was defined as a cell spread of more than 0.025 μm2. Values represent the means ± S.E. (n = 5). *, p < 0.05 (Welch’s t test). E, cell migration toward laminin-332 was determined using a Transwell assay. Scale bar = 100 μm. The quantitative data were obtained from three independent experiments. Values represent the means ± S.E. (n = 6). *, p < 0.05 (Welch’s t test).
Next, we compared total sialylated N-glycan levels between KO and control cells, which were quantified using HPLC. Consistent with the data from PI4KIIα KD cells, the ratios of sialylated N-glycans versus total N-glycans were greatly suppressed in KO cells (Fig. 5A). Because N-glycans of integrin β1 could be directly influenced by α3 knockout (Fig. 4B, center panel), here we also analyzed sialylation on the EGF receptor, which has been reported to affect receptor activation (
). As expected, sialylation of EGF receptors obtained from KO cells was also decreased compared with control cells (Fig. 5B). To investigate whether the phenomenon also occurred in other cell lines, we examined HeLa cells. The knockout efficiency of α3 was assessed via flow cytometry analysis (Fig. 5C). Interestingly, a similar tendency of sialylation decline was observed in α3 KO HeLa cells compared with control cells (Fig. 5D). To thoroughly examine the glycosylation states caused by knockout of integrin α3 or knockdown of PI4KIIα, we utilized MS-based approaches to identify the N-glycan and O-glycan structures (Fig. 6, A and B). Consistently, hyposialylation in N-glycans was observed by attenuation of either integrin α3 or PI4KIIα (Fig. 6A), which further suggests the importance of this pathway in the biosynthesis of sialylated N-glycans. In contrast to N-glycans, there were no significant changes in the sialylation of O-glycans among these cells (Fig. 6B). It was also notable that a decrease in α2,3 sialylation in N-glycans could slightly increase α2,6 sialylation in total cell lysates of α3 KO and PI4KIIα KD cells (Fig. 6A), which seems contrary to the observation for integrin β1 (Fig. 3E). Further study is needed to fully elucidate the mechanism. Nevertheless, these data suggest that complex formation of α3 and PI4KIIα could act as a novel regulator for the sialylation of N-glycans.
Figure 5Deficiency of integrin α3 decreased sialylation of N-glycans in different cell lines. A, N-glycans released from both Ctrl and α3 KO MDA-MB-231 cells were pyridylaminated and verified as populations of sialylated N-glycans. The relative ratios of sialylated N-glycans versus total N-glycans were calculated by subtracting the neutral portions from the total and then dividing by the total. The control data were set as 1. B, equal amounts of cell lysates for control and α3 knockout cells were precipitated via SSA-agarose and probed with anti-EGFR antibody (top panel). The same cell lysates were also probed with anti-EGFR antibody as a loading control (bottom panel). IP, immunoprecipitation. C, α3 KO of HeLa cells was established using the CRISPR/Cas9 system and verified via flow cytometric analysis. D, the ratios of sialylated N-glycans were analyzed as described in A.
Figure 6MS spectra of glycans obtained from cultured cells. A, MS spectra of N-linked glycans. The reducing ends of the N-linked glycans were derivatized with aoWR (
). The sialic acids of the N-linked glycans were differentially amidated with methylamine (+13.0 Da) for α2,3-sialic acids and isopropylamine (+41.1 Da) for α2,6-sialic acids via the SALSA method (
). MDA-MB-231 and α3 KO cells refer to parent and α3 knockout cells, respectively. KD and control cells of MDA-MB-231 cell line refer to transfectants with DOX-inducible knockdown of PI4KIIα cultured with or without DOX, respectively. B, MS spectra of O-linked glycans. The O-linked glycans were obtained as the corresponding alditols and then permethylated.
In this study, we found that PI4KIIα expression played an important role in α3β1 integrin-mediated cell migration, cellular signaling, and the expression of sialylation; PI4KIIα could specifically associate with α3 but not α5, and complex formation between PI4KIIα and α3 enhanced sialylation; and deletion of integrin α3 significantly inhibited not only cell adhesion and migration but also sialylation. These observations are the first to directly demonstrate a novel regulatory mechanism for sialylation, which may also partially explain the previous observation that GOLPH3 is a special regulator in the sialylation of N-glycans and a part of the signaling events that could influence mTOR signaling and tumor progression (
Many cancers are associated with sialylated structures such as sialyl Tn, sialyl Lewis antigen (sLe), α2,6-sialylated lactosamine, polysialic acid, and gangliosides (
Varki A. Cummings R.D. Esko J.D. Stanley P. Hart G.W. Aebi M. Darvill A.G. Kinoshita T. Packer N.H. Prestegard J.H. Schnaar R.L. Seeberger P.H. Essentials of Glycobiology. Cold Spring Harbor Laboratory Press,
Cold Spring Harbor, NY2015: 597-609
). The altered expression of these structures in cancer cells could be the result of multiple mechanisms. Loss of expression or excessive expression of certain sialyltransferases is frequently observed. For example, up-regulated expression of α2,3 and α2,6 sialyltransferases has been observed in many cancers, such as colon cancer, breast cancer, liver cancer, cervical cancer, choriocarcinomas, acute myeloid leukemias, and some malignancies of the brain (
). Interestingly, high expression of ST6GAL1 has been correlated with human-induced pluripotent stem cells and cancer stem cells, indicating that sialylation may be involved in maintaining some aspects of stem cell behavior (
ST6Gal-I protein expression is upregulated in human epithelial tumors and correlates with stem cell markers in normal tissues and colon cancer cell lines.
). In fact, ST6GAL1 expression is required for transforming growth factor β–induced EMT. Knockdown of ST6GAL1 prevented a transforming growth factor β–induced increase in cell migration (
Given the accumulating evidence of the importance of sialylation in cancer progression, much attention has been paid to elucidating the regulatory mechanisms of its expression. The expression of sialylation on a tumor cell surface can be modulated at different levels (
). The most frequently observed mechanism is modulation of transcription. For example, ST6GAL1 expression is induced by the ras oncogene in NIH3T3 cells via its promoter (
). Beyond its promoter activity, in this study, we clearly showed that sialylation could also be regulated by complex formation of PI4KIIα and integrin α3β1 on a posttranscriptional level. In fact, the PI4P-binding Golgi protein GOLPH3, which functions in secretory trafficking in the Golgi, is also important for the localization of several glycosyltransferases (
). Our previous study showed that knockdown of GOLPH3 leads to down-regulation of both α2,3 and α2,6 sialylation but has no effect on the transcription of sialyltransferases. Therefore, regulation of biosynthetic glycan is not only dependent on the expression levels of glycosyltransferases but also on those of substrates, chaperones, and on the environment of the Golgi apparatus.
Considering that PI4P is a relatively abundant phosphoinositide that is required for the maintenance and function of the Golgi apparatus, which includes intracellular trafficking, it is plausible that PI4K could be involved in membrane transport from the trans-Golgi network to the plasma membrane. In fact, PI4KIIα is known to associate with several cellular receptors such as EGFR, E-cadherin, LDL receptor-related protein, and the Fas receptor (
). In addition, alteration of the N-glycosylation of such receptors also regulates the functions of these receptors. For example, N-glycosylation is required for EGFR trafficking and has an effect on its endocytosis (
Overexpression of N-acetylglucosaminyltransferase III enhances the epidermal growth factor-induced phosphorylation of ERK in HeLaS3 cells by up-regulation of the internalization rate of the receptors.
). Given the importance of sialylation as described here, it is reasonable to expect up-regulation of both α3β1 and PI4KIIα in cancer cells.
These facts raise the question of why the interaction of PI4KIIα with integrin α3β1 but not of α5β1 regulates sialylation. So far, the association underlying the mechanism between α3β1 and PI4KIIα remains unclear. However, the specificity of α3β1 could be due to its interaction with the tetraspanin family, such as CD151, CD63, and CD9. In fact, other studies have reported that integrin α3β1 and the tetraspanin family can interact with type II of PI4K (
Highly stoichiometric, stable, and specific association of integrin α3β1 with CD151 provides a major link to phosphatidylinositol 4 kinase, and may regulate cell migration.
). In addition, palmitoylation could also be a plausible factor because PI4Ks are proteins with membrane association and activity that are highly dependent on such a modification (
). We assume that the association between PI4KIIα and α3, but α5, might also relate to tetraspanin or palmitoylation.
It is noteworthy that both cell spreading and migration on laminin 332 of integrin α3 KO cells were significantly suppressed (Fig. 4), which further supports the notion that α3β1 is a major receptor of lamin-332 compared with integrin α6β1 and α6β4 in epithelial cells (
Suppression of Integrin α3β1 in breast cancer cells reduces cyclooxygenase-2 gene expression and inhibits tumorigenesis, invasion, and cross-talk to endothelial cells.
). In this study, α3 deficiency significantly inhibited sialylation, indicating a novel link between sialylation and this integrin, which demonstrated the importance for both forms of functional expression. Although the MS analysis clearly showed that the complex between PI4KIIα and integrin α3 regulates the biosynthesis of sialylation on N-glycans but not O-glycans, we could not completely exclude other possibilities for the regulation of N-glycan structures other than sialylation or gangliosides. In addition, it has been reported that GOLPH3 containing a PI4P binding domain could control Golgi localization of core 2 N-acetylglucosaminyltransferase 1 for the biosynthesis of O-glycans (
). Thus, further studies are required to explain the underlying mechanism for different influences, such as localization of glycosyltransferase, pH environment (
) through this system. Taken together, this study may provide a new concept for the regulation of glycosylation and could suggest insights for the development of cancer treatment.
Experimental procedures
Cell lines and cell culture
The HeLa and 293T cells were obtained from the RIKEN Bioresource Research Center (Japan). The MDA-MB-231 cells were purchased from the ATCC. All cell lines were maintained at 37 °C in DMEM containing 10% FBS in a humidified atmosphere of 5% CO2.
PCR for mRNA expression analysis
Total RNA was prepared with TRIzol (Invitrogen), and 1.0 μg of RNA was reverse-transcribed using the PrimeScript RT reagent kit (Takara Bio Inc.) according to the manufacturer’s instructions. PCR primers against ST3GAL3, ST3GAL4, ST6GAL1, and GAPDH have been described previously (
). The following oligonucleotides were inserted into pENTR/H1/TO (Invitrogen): 5′-CACCAGAAGCAGAACCTCTTCCTGATGATATGTGCATCAGGAAGAGGTTCTGCTTCT-3′ and 5′-AAAAAGAAGCAGAACCTCTTCCTGATGCACATATCATCAGGAAGAGGTTCTGCTTCT-3′ (
). The use of LR Clonase allowed the inserted oligo to be transferred to CS-RfA-ETBsd, which is an encoding DOX-dependent transactivator for shRNA expression. The resultant vector was then transfected into 293T cells with packaging plasmids of calcium phosphate for the preparation of viruses. MDA-MB-231 cells were then infected with the obtained viruses and selected for stable integration with 1 μg/ml blasticidin. shRNA-mediated silencing of PI4KIIα was induced by addition of 1 μg/ml DOX for 72 h in the established cell line, and cells cultured in DOX-free medium were used as a control.
Gene introduction using the lentivirus system
The cDNA sequences for the PH domain of human FAPP1, PI4KIIα, and integrin α3 were cloned from HeLa cells and inserted into pENTR vectors (pENTR/d-TOPO cloning kit, Invitrogen). To obtain the N-terminal GFP-tagged PI4KIIα, C-terminal mRFP-tagged FAPP1 and a C-terminal GFP-tagged integrin α3, in-fusion enzyme (Clontech) was used with standard PCR protocols. The linkers for GFP-PI4KIIα, mRFP-FAPP1, and α3-GFP were 5′-GGGGS-3′, 5′-KNPPVAT-3′ (
A plasma membrane pool of phosphatidylinositol 4-phosphate is generated by phosphatidylinositol 4-kinase type-IIIα: studies with the PH domains of the oxysterol binding protein and FAPP1.
), and 5′-LELKLRILQSTVPRARDPPVAT-3′, respectively. The resultant cDNAs were confirmed by DNA sequencing using an ABI Prism 3130 sequencer (Applied Biosystems Japan Ltd., Tokyo, Japan). The subcloned cDNAs were transferred into CSII-EF-Rfa (
) via LR Clonase (Invitrogen) for lentivirus production. The resultant vectors (CSII-EF-GFP-PI4KIIα/mRFP-FAPP1/α3-GFP) and the previously constructed integrins of either α5-GFP- or β1-overexpressed lentiviral vectors (CSII-EF-α5GFP (
)) were then transfected into 293T cells with packaging plasmids via calcium phosphate during the preparation of viruses. Either MDA-MB-231 or 293T cells were then infected with the obtained viruses for further experiments.
Cell migration (Boyden chamber assay)
Cell Migration was performed as described previously with minor modifications (
N-glycosylation of the beta-propeller domain of the integrin α5 subunit is essential for α5β1 heterodimerization, expression on the cell surface, and its biological function.
). Each Transwell (BD BioCoat control inserts, 8.0-μm inserts; BD Biosciences) was coated only on the bottom side with 1 μg/ml laminin-332 (Oriental Yeast Co., Ltd.) in PBS containing 0.1% BSA at 4 °C for 24 h and then blocked with 5% BSA in DMEM at 37 °C for 1 h. Cells were trypsinized and suspended in DMEM containing 1% FBS. The suspended cells were centrifuged, and the cell pellets were resuspended in an assay medium (0.1% BSA in DMEM containing 1% FBS) and diluted to 3 × 105 cells/ml; cell viability was confirmed by trypan blue staining. Cell suspensions were then added to the Transwell. Following incubation, the membranes were fixed with 4% paraformaldehyde and stained with 0.5% crystal violet solution (Merck) for 24 h. After washing the Transwells with PBS, cells that migrated to the lower side were counted using a phase-contrast microscope.
Immunostaining
Cells (1 × 105) were plated on glass coverslips (MatTek), precoated with or without ECM for 24 h, fixed with 4% paraformaldehyde for 15 min, permeabilized with 0.1% Triton X-100 for 10 min, and blocked with PBS containing 3% BSA and 0.01% Tween 20 (PBSBT) for 1 h. Cells were then incubated with the indicated primary antibodies (GM130, BD Biosciences, 610822; α3, Millipore, P1B5) at 4 °C overnight, followed by incubation with Alexa Fluor 647–conjugated secondary antibodies (Molecular Probes) for 1 h. For lectin staining, cells on coverslips were incubated with either biotinylated SNA lectin (Vector Laboratories, B-1305) or wheat germ agglutinin (WGA) lectin (J-Oil Mills, J220) at 4 °C for 30 min in DMEM, fixed and blocked with PBSBT for 1 h, followed by incubation with streptavidin-conjugated Alexa Fluor 647 at 4 °C for 1 h in the dark. All samples were mounted using Prolong Diamond Antifade mounting medium (Molecular Probes). Images were acquired by sequential excitation using an Olympus FV1000 laser-scanning confocal microscope with an UPlanSApo ×60/1.35 oil objective operated with F10-ASW version 4.02 software.
Cell spreading and adhesion experiment
Coverslips were coated with 1 μg/ml laminin-332 or 2 mg/ml gelatin in PBS containing 0.1% BSA at 4 °C for 24 h and then blocked with 5% BSA in DMEM at 37% for 1 h. Cells were replated on the coverslips in DMEM with 0.1% BSA. After incubation for 10 min, the areas of adherent cells were measured using ImageJ. We defined cells spread at more than 0.025 μm2 as adherent cells. After incubation for 30 min, the cells were fixed with 4% paraformaldehyde and stained with phalloidin–Alexa Fluor 647 (Molecular Probes).
Immunoprecipitation and Western blotting
Immunoprecipitation was performed as described previously with minor modifications (
An N-glycosylation site on the β-propeller domain of the integrin α5 subunit plays key roles in both its function and site-specific modification by β1,4-N-acetylglucosaminyltransferase III.
). Briefly, cells were gently rinsed three times with PBS at room temperature and solubilized in cold lysis buffer A (20 mm Tris-HCl (pH 7.4), 150 mm NaCl, and 1% Brij98), which included protease inhibitors. Protein concentrations of lysates were determined via BCA assay (Pierce). The lysates were immunoprecipitated using either anti-GFP-agarose (MBL), anti-integrin α3 antibody (Millipore, P1B5), or anti-integrin β1 (P5D2, DSHB) with Ab-Capcher Protein A-R28 (Protenova, Tokushima, Japan) for 1 h at 4 °C and gentle rotation. After washing with lysis buffer, the immunoprecipitates were subjected to SDS-PAGE. The proteins were probed with anti-integrin α3 (Santa Cruz Biotechnology, sc-6592) or anti-GFP antibody (Rockland, 600-101-215) and then detected using anti-goat IgG-conjugated horseradish peroxidase (Santa Cruz Biotechnology) with an Immobilon Western chemiluminescent horseradish peroxidase substrate (Millipore). For analysis of the lectin precipitates, we used lysis buffer B (20 mm Tris-HCl (pH 7.4), 150 mm NaCl, and 1% Triton X-100). Cell lysates were precipitated with either SSA-agarose (J-Oil Mills, J318), MAM (also called MAL-1)–agarose (J-Oil Mills, J310) or concanavalin A (ConA)–agarose (J-Oil Mills, J303), which specifically recognize either α2,6- and α2,3-sialylation or total N-glycans, respectively. The precipitants were detected using either anti-integrin β1 (BD Biosciences, 610468) or anti-EGFR (Cell Signaling Technology, 4267). Antibody agents against Akt (9272) and p-Akt (4060) were purchased from Cell Signaling Technology. α-Tubulin (Sigma, T6199) was used as a loading control.
Flow cytometric analysis
Flow cytometric analysis was performed as described previously with minor modifications (
N-glycosylation of the beta-propeller domain of the integrin α5 subunit is essential for α5β1 heterodimerization, expression on the cell surface, and its biological function.
An N-glycosylation site on the β-propeller domain of the integrin α5 subunit plays key roles in both its function and site-specific modification by β1,4-N-acetylglucosaminyltransferase III.
). Briefly, semiconfluent cells were detached from the culture dishes using trypsin containing 1 mm EDTA. The cells were subsequently stained with or without primary mouse anti-α3 (P1B5), anti-β1 (P5D2), and anti-α5 integrin (Millipore, HA5) in PBS, followed by incubation with Alexa Fluor 647–conjugated secondary antibodies. For staining cells with biotin-conjugated lectins (MAM and SNA), we used PBS buffer containing 0.5 mm CaCl2 and MgCl2. Flow cytometric analysis was performed using a FACSCalibur flow cytometer and Cell Quest Pro software (BD Biosciences).
PA oligosaccharide preparation of N-glycosylation and quantitative analysis of sialylated N-glycans by anion exchange HPLC
To release N-glycans from glycoproteins, the lyophilized cell pellets (1–2 mg) were heated with 200 μl of anhydrous hydrazine at 100 °C for 10 h using an oil bath. Removal of hydrazine and acetylation of N-glycans were carried out with a graphite carbon column (
). The released N-glycans were then 2-pyridylaminated using the Pyridylamination Manual Kit (Takara Bio Inc.), and then the excess 2-aminopyridine was removed via phenol–chloroform extraction (
). The prepared PA oligosaccharides treated with or without neuraminidases were subjected to a HPLC system (JASCO) equipped with a TSKgel DEAE-5PW column (7.5 × 75 mm, Tosoh) and analyzed as described previously (
Expression of N-acetylglucosaminyltransferase III suppresses α2,3-sialylation, and its distinctive functions in cell migration are attributed to α2,6-sialylation levels.
). HPLC chromatogram data were analyzed using chromNAV software (JASCO). The amounts of total and neutral N-glycans were calculated based on the peak area of neutral fraction with or without neuraminidase treatment. Relative ratios of sialylated N-glycans versus total N-glycans were calculated by subtracting the neutral portions from the total and then dividing by the total. The data were normalized to the control as 1.
Generation of CRISPR/Cas9-based integrin α3 KO cells
CRISPR/Cas9-based integrin α3 cells were established as described previously (
). Briefly, the single guide RNA-specifying oligo sequences spanning human integrin α3 (forward, 5′-CACCGCATCGGGCACAGCGAGCTCC-3′; reverse, 5′-AAACGGAGCTCGCTGTGCCCGATGC-3) were chosen from the human KO library (
). Then the oligos were cloned into pSpCas9 (BB)-2A-GFP, which was a kind gift from Dr. Feng Zhang (Addgene plasmid ID 48138). The plasmid was transfected into cells according to the manufacturer’s instructions (Amaxa Cell Line Nucleofector kitV). After 4 days of transfection, GFP-positive cells were sorted using a FACSAria II (BD Bioscience). Following 10-day culture, α3-negative and GFP-negative cells were sorted, and the procedure was repeated twice. The KO cells were defined by flow cytometry and Western blot analysis as described above.
Mass spectrometry of glycans
Total plasma membrane proteins of cells were prepared using a plasma membrane protein extraction kit (101Bio). For N-glycan analysis, a previous report (
) suggests that the pellets are dissolved and denatured and then digested by glycopeptidase F (Takara Bio Inc.). The released N-glycans were captured on hydrazide beads (BlotGlyco, 5 mg, Sumitomo Bakelite Co., Ltd.) using a process recommended by the manufacturer. Sialic acids of the N-glycans on the beads were then differentially amidated with methylamine for α2,3-sialic acids and isopropylamine for α2,6-sialic acids using the sialic acid linkage–specific alkylamidation (SALSA) method (
), and purified according to the manufacturer’s instructions. The obtained glycan solutions were mixed with a 2,5-dihydroxybenzoic acid solution (10 mg/ml in 30% acetonitrile) at a ratio of 1:10. Aliquots (0.5 μl) of the mixed solutions were deposited onto a MALDI target plate and dried. MS spectra were acquired with a MALDI quadrupole ion trap TOF mass spectrometer (AXIMA-QIT, Shimadzu Corp.). For O-glycan analysis, pellets of the membrane fractions were dissolved in a 1% SDS solution. Aliquots (10 μl) of the solution were transferred into 90 μl of 0.6 m sodium borohydride containing 60 mm NaOH and incubated at 45 °C for 16 h. Then the mixtures were neutralized, diluted to 1 ml, and applied onto a solid-phase extraction cartridge (Sep-Pak C18 Vac 1cc, 50 mg, Waters Corp.), and then the cartridge was washed with distilled water (1 ml). The eluents and washings were combined and lyophilized. The obtained residues were dissolved in 1% acetic acid in methanol (100 μl) and evaporated using a centrifuge evaporator. The procedure of dissolution and evaporation was repeated. The obtained residues were permethylated according to previous reports (
). The permethylated glycans were dissolved in a 2,5-dihydroxybenzoic acid solution (20 μl, 10 mg/ml in 30% acetonitrile), and aliquots (0.5 μl) of the mixed solutions were deposited onto a MALDI target plate and dried. MS spectra were acquired with a MALDI-TOF mass spectrometer (Ultraflex, Bruker Daltonik).
Author contributions
T. I., T. F., and J. G. conceptualization; T. I., S. I., A. K., Y. W., and J. G. data curation; T. I., S. I., A. K., Y. W., and J. G. formal analysis; T. I. and J. G. supervision; T. I., S. I., A. K., Y. W., T. F., and J. G. validation; T. I., S. I., A. K., and Y. W. investigation; T. I., S. I., A. K., Y. W., T. F., and J. G. methodology; T. I. and S. I. writing-original draft; T. I., S. I., A. K., Y. W., T. F., and J. G. writing-review and editing; S. I. visualization; A. K. and J. G. funding acquisition.
Acknowledgments
We thank Dr. Takashi Nishikaze for informative suggestions regarding the SALSA method and Dr. Wai Wai Thet Tin for technical assistance with glycan analysis.
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N-glycosylation of the beta-propeller domain of the integrin α5 subunit is essential for α5β1 heterodimerization, expression on the cell surface, and its biological function.
An N-glycosylation site on the β-propeller domain of the integrin α5 subunit plays key roles in both its function and site-specific modification by β1,4-N-acetylglucosaminyltransferase III.
This work was supported in part by Grants-in-Aid for Scientific Research 15H04354 (to J. G.) and JP17H03808 (to A. K.) from the Japan Society for the Promotion of Science, National Natural Science Foundation of China Grant 31670807, and Grant-in-Aid for Scientific Research on Innovative Areas 18H04868 (to J. G.) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan. The authors declare that they have no conflicts of interest with the contents of this article.
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