IntroductionThe trace element (TE) zinc (Zn) is essential for the catalytic, structural, and regulatory function of proteins and therefore is crucial for numerous physiological processes, such as the immune system and metabolism. As there is no Zn reservoir in the human body, it must be ingested daily to compensate for endogenous losses1. Zn deficiency is one of the most prevalent micronutrient deficiencies, affecting approximately 17% of the global population, thereby representing a significant health and economic burden worldwide2. The primary cause of Zn deficiency is either insufficient oral Zn intake or diets with low Zn bioavailability in the intestine, where Zn absorption via specific Zn/iron-regulated transporter-like proteins (ZIPs) and Zn transporters (ZnTs) occurs3. Symptoms of Zn deficiency encompass immune dysfunction, increased incidence of infection, slow wound healing, and metabolic disturbances. In addition, Zn deficiency significantly impacts gastrointestinal health, including altered intestinal homeostasis, increasing the likelihood of infection and, in severe cases, diarrhea2. The underlying (cellular) processes in the intestine leading to these symptoms, however, remain to be fully understood.A crucial protective barrier for the intestinal epithelium is provided by mucus, a viscoelastic hydrogel-like substance that shields the underlying epithelium against chemical, mechanical, and pathogenic damage4. Mucus plays an essential role in the absorption of (micro)nutrients, such as Zn. In fact, mucus can bind TEs (reviewed in5), prolonging their residence time at the epithelial surface and thereby increasing their bioavailability6,7. Intestinal mucus consists of 90–95% water and up to 5% mucins8, highly glycosylated proteins classified as either secreted or membrane-bound. To date, 21 mucins belonging to the MUC gene family have been identified8, of which MUC2 is the main secreted mucin produced by goblet cells in the small and large intestine4. Mucin biosynthesis is a complex, multistep process involving dimerization and extensive N- and O-glycosylation9. N-glycosylation initiates in the endoplasmic reticulum (ER), where mannose (Man)-rich glycans are assembled and processed before being transported to the Golgi apparatus for conversion into complex and hybrid type N-glycans (Fig. 1A). Additionally, the Golgi apparatus serves as the primary site for O-N-acetyl-galactosamine (GalNAc) glycosylation, which is initiated by the attachment of GalNAc to Ser/Thr-residues10. Driven by the activity and presence of specific glycosyltransferases (GTs), this so-called Tn antigen is extended to up to 8 different core O-glycan structures, which can be further elongated and capped by the addition of galactose (Gal), fucose (Fuc) and/or sialic acid (NeuAc) (summarized in Fig. 1B)8. In particular, O-glycosylation is crucial for mucins’ high water-binding capacity, leading to the hydrogel-like viscoelastic properties of mucus and protecting the protein backbone against bacterial degradation8. The produced O-glycome plays a vital role in host-microbiome cross-talk, serving as a primary determinant of gut microbial composition and diversity11. Accordingly, goblet cells are essential regulators of intestinal homeostasis, playing a crucial role in epithelial protection, immune defense, and nutrient absorption via mucus production. Disruption of mucin production or glycosylation can impair the mucus barrier12 and has been linked to the pathogenesis of intestinal disorders, including inflammatory bowel disease (IBD)12, and increased susceptibility to infection13.Fig. 1Full size imageSchematic illustration of the N- and O-glycan biosynthesis in the secretory pathway and the impact of Zn deficiency. (A) N-glycans are formed through a multi-step process. High Man structures linked to dolichoyl-pyrophosphate (DOL-PP) are transferred via the oligosaccharyltransferase (OST) protein complex to Asn residues. These high Man structures are subsequently processed by cleaving, the protein undergoes folding and is transported to the Golgi, where N-glycans are further modified to generate high-Man complex type and hybrid type structures10. (B) Tn antigen (Ser/Thr residues linked with GalNAc) are elongated with Gal or GlcNAc, catalyzed by C1GALT1 or B3GNT6, respectively, which can lead to the formation of the core 1 or 3, respectively. These core O-glycans can be further processed to core 2 or 4 by glucosaminyl (N-acetyl) transferase (GCNT)1 or GCNT3, respectively. Core structures can then be further elongated8 or terminated by Neu5Ac (sialic acid), Fuc, Man, or sulfate4,8. Glycan structures were drawn with GlycoGlyph14. Red arrows show where Zn deficiency impacts gene/protein expression.In the 1970 s, changes in mucin composition were reported in Zn-deficient (ZD) rats and sheep, with an increase in mucin sialylation observed under ZD conditions15. Previous work from our group showed that, in the HT-29-MTX goblet cell model, the O-glycosylation pattern of secreted mucins was profoundly altered during Zn deficiency, with a reduction in more complex O-glycan structures16. In addition, the mRNA expression of several GTs was deregulated, and MUC2 was upregulated in ZD cells16. Despite these observations, the molecular mechanisms linking Zn deficiency to altered mucin glycosylation and production are still not understood. Furthermore, no information exists on how the N-glycan pattern changes during Zn deficiency. This would be highly relevant as N-glycosylation is not only important for mucin maturation17 but also essential for maintaining cellular homeostasis and health, as it is crucial for determining protein structure and stability, modulating protein expression, and various cellular processes, including cell-cell communication, immune response, and protein folding10. The importance of (sub)cellular Zn status for glycosylation was indicated as knocking-out (KO) the Zn transporter solute carrier (SLC)39A9 (ZIP9), which resides in the secretory pathway18, led to diminished formation of core 1 O-glycans and complex type N-glycans18,19. Moreover, reduction of Golgi-stored Zn by KO of ZnT7, which shuttles Zn back into the Golgi20,21, altered N-glycosylation in vitro22. However, it remains unclear how a (sub)cellular Zn deficit alters the expression or activity of GTs and how these changes impact mucin structure and composition. Therefore, the aim of this study was to examine the role of Zn in mucin formation and glycosylation at the cellular level using the HT-29-MTX goblet cell model. We investigated how Zn deprivation affects human goblet cells’ Zn homeostasis, mucin production, and global glycosylation patterns by applying a transcriptomic approach to analyze changes in mucins, GTs, and Zn transporters. Additionally, we employed mass spectrometry (MS)-based techniques to screen the changes in mucinome and glycan composition of cellular and secreted mucins.ResultsThe Zn level in the secretory pathway of goblet cells is diminished in ZD cellsTo assess Zn levels within the secretory pathway, we utilized the low-molecular-weight fluorescent probe Zinpyr-1 (ZP1), which demonstrated significant co-localization with BODIPY™ TR Ceramide and ER tracker in living HT-29-MTX cells (Fig. 2A-B). Co-localization analysis shows proportional co-distribution of both markers with ZP1, as at least 98.5% or 97.6%, respectively, of ZP1 correlates with the stained secretory pathway confirming that the ZP1 signal accurately reports the Zn status of the secretory pathway (Fig. 2A). The specificity of the ZP1 probe for Zn was validated using the cell-permeant chelator Tetrakis-(2-pyridylmethyl)ethylendiamin (TPEN), which successfully abolished the fluorescent signal. Of note, to induce Zn deficiency, cells were treated with Zn-depleted medium using Chelex®. To control for changes caused by treating medium with Chelex® resin that were not attributable to Zn-depletion, cells were cultured with Chelex®-treated medium with replenished Zn content (Zn-adequate (ZA)). Cellular distribution of ZP1 did not change with cellular Zn-content, as ZP1 correlates with both TR-Ceramide and ER-marker in control (CTR), ZA and ZD cells (TR-Ceramide: 99.4%, ZA 98.3, ZD 99.8%; ER-tracker: CTR 98.2%, ZA 97.7%, ZD 98%, Fig. 2D). In accordance with the significantly reduced total cellular Zn content previously observed in ZD goblet cells16, the concentration of free Zn was found to be markedly decreased in ZD HT-29-MTX cells when measured with ZP1 (Fig. 2E). With the spatial analysis of the sensor, this demonstrates that Zn in the secretory pathway was decreased.Fig. 2Full size imageChanges of subcellular free Zn after Zn deprivation and localization of ZP1 in the secretory pathway of the HT-29-MTX goblet cell model. (A, B) LSM images show the subcellular distribution of free Zn (green) detected with the fluorescent Zn sensor ZP1 and either the Golgi apparatus labelled with the BODIPY™ TR coupled sphingolipid ceramide (red) or the ER stained with ER-Tracker™ Blue-White DPX in CTR HT-29-MTX cells, cultured for 2 d and treated with ZnSO₄ or the Zn chelator TPEN (A). Organelle distribution was additionally confirmed in CTR, ZA, and ZD cells upon 14 d cultivation (B). Images are representative of n = 3–4 independent experiments. Co-localization of the two organelle probes with ZP1 was supported by a Mander’s overlap coefficient (MOC), comparing data of 6–20 images, analyzed in n = 3–4 replicates (C, D). Presented is the quantification of intracellular free Zn in CTR, ZA, and ZD HT-29-MTX cells, measured using the fluorescent Zn probe ZP1 in a 96-well approach (E). Data are presented as mean + SD of at least n = 3 replicates. Statistically significant differences between treatments were tested with one-way ANOVA followed by Tukey post hoc test (*p