Zinc deficiency causes immune dysfunction. In T lymphocytes, hypozincemia promotes thymus atrophy, polarization imbalance, and altered cytokine production. Zinc supplementation is commonly used to boost immune function to prevent infectious diseases in at-risk populations. However, the molecular players involved in zinc homeostasis in lymphocytes are poorly understood. In this paper, we wanted to determine the identity of the transporter responsible for zinc entry into lymphocytes. First, in human Jurkat cells, we characterized the effect of zinc on proliferation and activation and found that zinc supplementation enhances activation when T lymphocytes are stimulated using anti-CD3/anti-CD28 Abs. We show that zinc entry depends on specific pathways to correctly tune the NFAT, NF-κB, and AP-1 activation cascades. Second, we used various human and murine models to characterize the zinc transporter family, Zip, during T cell activation and found that Zip6 was strongly upregulated early during activation. Therefore, we generated a Jurkat Zip6 knockout (KO) line to study how the absence of this transporter affects lymphocyte physiology. We found that although Zip6KO cells showed no altered zinc transport or proliferation under basal conditions, under activation, these KO cells showed deficient zinc transport and a drastically impaired activation program. Our work shows that zinc entry into activated lymphocytes depends on Zip6 and that this transporter is essential for the correct function of the cellular activation machinery.

Zinc is an essential element for human physiology. Zinc deficiency causes impaired body growth, neurologic disorders, and immunosuppression, leading to morbidity and increased infection risk (1). It is considered a major public health problem in developing countries, especially in groups with low zinc intake as well as those with inhibited absorption due to high levels of phytates and fiber in vegetable-based diets. Globally, the elderly, vegetarians, and people with gastrointestinal disorders are also considered to be risk populations for hypozincemia (1, 2). Due to its requirement for the proper immune function, zinc is a common component in nutritional supplements with the aim to boost the immune system. Clinical trials have proven that zinc supplementation is effective in certain types of infections (reviewed in Ref. 3). Moreover, in animal models, zinc administration has been shown to be beneficial in autoimmune diseases like autoimmune encephalomyelitis (4, 5).

At the molecular level, besides being a structural component of many proteins, zinc has also been shown to be a second messenger. Thus, zinc movements influence signaling cascades by regulating key enzymes such as AKT and GSK-3 (6, 7), as well as transcription programs by acting on transcription factors such as MTF-1 and NF-кB (8, 9). In this dynamic scenario, several transporters and zinc binding proteins work in a coordinated way to tightly regulate cytosolic zinc concentrations. There are structurally and functionally two main families of the zinc transporters. The ZnT family, with 10 members, extrudes zinc from the cytosol to organelles or outside of the cell. In contrast, the Zip family of transporters, which has 14 members, allows zinc fluxes from the extracellular medium or intracellular compartments to the cytosol (2).

Despite the importance of zinc for immune system function, relatively little is known about how this ion enters and operates in these cells. In recent years, work in transgenic mouse models have highlighted the role of Zip8 and Zip10 function in immune cells (911). Zip8 has emerged as an important regulator of innate immunity by fine-tuning the NF-кB pathway (9), whereas Zip10 plays an important role modulating the B cell Ag receptor (10, 11). In the case of T lymphocytes, zinc deficiency is known to dramatically alter T cell maturation, influence polarization of Th subsets, and impair cytokine production (1214). In this context, various lines of evidence show that lymphocytes increase cellular zinc content during activation (15, 16). EVER proteins, positive regulators of the ZnT1 transporter that imports zinc from the cytosol to the endoplasmic reticulum, are downregulated during T cell activation, favoring this way cytosolic zinc accumulation (15). Moreover, zinc influx from the extracellular compartment has been shown to fine-tune the TCR machinery, including Lck binding and phosphorylation of ZAP70 (16, 17). Nevertheless, the latter aspect remains controversial because different studies have shown conflicting results regarding the influence of zinc in IL-2 production, an important early activation marker in T cells (16, 18, 19). Based on expression analysis, several Zip transporters have been suggested to participate in T cell physiology, including Zip6, Zip8, Zip10, and Zip12 (15, 16, 19, 20). However, the existing mouse knockout (KO) models have not confirmed the importance of these transporters in T cell function. Zip6 deserves special attention because studies with Zip6 small interfering RNA have shown a reduction in zinc transport entry in T cells (16). The absence of KO model for this transporter makes it difficult to further validate this finding and to study its overall impact on T cell activation. Thus, clarifying the function of Zip6 in T cell physiology would allow a better understanding of the role of zinc in immune response, the pathology associated to zinc deficiency, and the effects obtained by zinc supplementation.

In this work, we conducted a detailed study of the impact of zinc on various aspects of T cell physiology, with a main focus on the activation process. To clarify discrepancies between previous reports, we compared results for various common activation stimuli. We characterized the expression of Zip transporters in different T cell models. Finally, we generated a Zip6KO cell line to explore how the absence of this transporter affects cellular zinc transport, cell proliferation, and activation kinetics.

We used different strains of the human leukemic Jurkat T cell line. We also acquired a stable CRISPR/Cas9–expressing Jurkat cell line (Applied StemCell) to generate a Zip6KO Jurkat line targeting two guide RNAs to the exon 3 sequence of Slc39a6 (5′-ATCTCATGGCATGGGCATCC-3′ and 5′-ACTGAACGTCACTACAGGGG-3′). We also worked with a Jurkat triple-reporter line (NF-κB-CFP, NFAT-eGFP, and AP-1-mCherry) characterized previously (21).

Cells were maintained in RPMI 1640 medium supplemented with 10% FBS and 1% penicillin/streptomycin. Where indicated, FBS was incubated according to the manufacturer’s instructions with Chelex 100 resin (Bio-Rad Laboratories) to generate Zn2+-free growth medium. Various amounts of ZnSO4 were added to the final medium to generate specific Zn2+ concentration conditions. Cells were grown at 37°C in a humidified 5% CO2 atmosphere. Lymphocytes were activated by incubating with 1 μM ionomycin and 10 nM PMA or using plate-bound 0.25 μg/ml anti-CD3 (HIT3a clone) and 1 μg/ml anti-CD28 Abs (BD Biosciences) unless a different concentration is stated. The expression of TCR complex and CD3 in our clones was validated by immunostaining and flow cytometry assays using FITC–anti-TCR and FITC–anti-CD3 Abs (eBioscience).

Expression analysis was conducted using Jurkat cells, T lymphocytes isolated from human donors, and T lymphocytes isolated from mouse spleen. Human T cells were isolated from PBMCs of healthy volunteers using enhanced human T Cell Immunocolumns (Diagnóstica Longwood). The use of human samples for this study was approved by the clinical research ethics committees at Hospital Germans Trias i Pujol and Comite Etico de Investigaciones Clinicas del Instituto Municipal de Asistencia Sanitaria. All participants provided written informed consent before donating samples.

Mouse T cells were isolated from the spleen of C57BL/6 mice using the Dynabeads Untouched Mouse T Cell kit (Invitrogen), according to the manufacturer’s instructions. Cells were activated using plate-bound 0.5 μg/ml anti-CD3 (BD biosciences) and 1 μg/ml anti-CD28 Abs (eBioscience).

Total RNA extraction (NucleoSpin RNA II kit; Macherey-Nagel) and cDNA generation were conducted using SuperScript Reverse Transcriptase system (Invitrogen) according to the manufacturer’s instructions. Quantitative PCR was performed using SYBR Green (Applied Biosystems). All PCRs used the following conditions: 95°C for 5 min; 95°C for 30 s; 60° for 30 s, 72°C for 30 s; and 72°C for 5 min, with 40 cycles of amplification. Expression levels of selected genes were normalized to those of the housekeeping genes RPL13A or MLN51 in human and β-actin in mouse. PCR primers are listed in Supplemental Table I.

Cells were seeded into 24-well plates at 1 × 105 cells per well and stimulated with ionomycin/PMA or plate-bound anti-CD3, anti-CD28 for 1 h. They were then washed twice with cold 1× PBS and lysed at 4°C with 100 μl of 1× Laemli buffer (6× stock solution: 0.018% bromophenol blue, 6% 2-ME, 0.75 M Tris-HCl, 6% SDS, 3% glycerol, pH 6.8). Lysates were sonicated for 15 s, boiled at 95°C for 5 min, placed in ice for 1 min, and then centrifuged; 40 μl of lysate was then loaded onto a 10% polyacrylamide gel. After electrophoresis, proteins were transferred to nitrocellulose membranes using the iBlot system (Invitrogen); membranes were then blocked with 5% BSA for 1 h at room temperature (RT). Primary Abs were diluted in blocking solution O/N at 4°C: p-GSK-3β (S9) (1:500, 9323S; Cell Signaling Technology), GSK-3β (1:500, 9832S; Cell Signaling Technology), p–Zap70 (Y319) (1:1000, 2701T; Cell Signaling Technology), Zap70 (1:500, 3465T; Cell Signaling Technology), and β-actin (1:2000; Sigma-Aldrich). Anti-rabbit or anti-mouse HRP secondary Abs (1:2000; GE Healthcare) were used.

Jurkat cells were seeded into 96-well plates at 5 × 104 cells per well and exposed to different concentrations of ZnSO4. At 24 h after seeding, MTT reagent was added to obtain a final concentration of 0.5 mg/ml, followed by incubation for 3 h at 37°C. The volume was then transferred into a tube, centrifuged at 13,000 rpm for 10 min, and washed once with isotonic solution. Formazan precipitates were resuspended in 200 μl of DMSO. The absorbance was read at 590 nm.

Cells were seeded at 1 × 105 cells per 10-cm plate with normal RPMI medium. After the indicated time points, cells were counted using the Neubauer chamber.

Double FITC annexin V–propidium iodide (PI) staining (BD Bioscience) was used to test for apoptosis at different zinc concentrations in resting and activated cells. After the incubation times, 24 h for resting experiments and 6 h for activated cells, cells were centrifuged at 1200 rpm, washed once with PBS, centrifuged, and resuspended in 300 μl of annexin V–binding buffer mixed with 5 μl of annexin V and 2 μl of PI. Fluorescence intensity was detected by flow cytometry using BD LSR II. Data were analyzed using Flowing Software (Perttu Terho).

Cells were incubated in normal or zinc-deficient medium with different concentrations of added zinc (0.10 or 100 μM of ZnSO4) and stimulated with ionomycin/PMA or plate-bound anti-CD3 plus anti-CD28 at indicated concentrations for 6 h.

IL-2 production analysis by flow cytometry was performed using an intracellular IL-2 staining protocol. Briefly, cells were fixed in 2% PFA in PBS for 20 min at RT. After a PBS wash, cells were permeabilized and blocked with PBS buffer containing 0.5% saponin (Sigma-Aldrich) and 1% BSA (Sigma-Aldrich). IL-2 staining was performed using anti–human-IL2 Ab (eBioscience) for 30 min at RT. Cells were then washed twice with PBS without saponin and suspended with 0.5% PFA in PBS. Flow cytometry analysis using FACSCalibur was performed of cells gated from a forward versus side scatter plot to exclude death cells. Further analysis was done using Flowing Software (Perttu Terho).

Tto quantify the secreted IL-2, after 6 h of activation, cells were centrifuged at 1200 rpm for 5 min and supernatants were collected. Then, an ELISA analysis was carried out using Human IL-2 ELISA Kit II (BD Bioscience) following the manufacturer’s instructions.

To study zinc transport capability, we determined zinc content in the various cell lines using FluoZin-3 fluorescent probe (Invitrogen). Briefly, cells were seeded at 2.5 × 105 cells per well, grown for 24 h under zero Zn2+ conditions, and then exposed to different zinc concentrations (0, 10, and 100 μM of ZnSO4) for 24 h. Cells were then loaded with 1 μM of FluoZin-3 for 30 min at RT. After washing with PBS, fluorescence was quantified using an FACSCalibur flow cytometer. Further analysis was performed using Flowing Software (Perttu Terho).

To analyze zinc transport dynamics, we used Zinquin (Sigma-Aldrich). Cells were seeded 2.5 × 105 cells per well for 24 h, incubated with Zinquin (25 μM) at 37°C (5% CO2) for 30 min, washed once, and then monitored under the fluorescence microscope using Aquacosmos software (Hamamatzu). At minute 4 we added extracellular medium with 100 μM of ZnSO4 promoting zinc transport to the cytosol. Intensity was normalized to the initial value.

The Jurkat triple-reporter cell line was seeded in different zinc concentrations (0, 10, 100 ZnSO4) and stimulated with ionomycin/PMA or plate-bound anti-CD3 plus anti-CD28. After 24 h, cells were additionally stained with TO-PRO3 (Thermo Fisher Scientific) to check cell viability. Flow cytometry analysis was performed using LSRFortessa equipment and supported by Flowing Software. First, cells were gated from a forward versus side scatter plot and then gated from a forward scatter height versus area for doublet exclusion; finally, we gated the TO-PRO3 negative cells before analyzing the reporter intensity.

Zinc is an important element for T lymphocytes, affecting maturation, cytokine production, and polarization (1, 13, 14). Zinc supplementation is a common strategy to prevent mortality in populations at risk for zinc deficiency but also is a common nutritional supplement to boost immune function in people. However, considering previous contradictory results regarding the effect of zinc supplementation on T cell physiology (16, 18, 19), we decided to study it in our system. First, to study cell viability, we incubated Jurkat cells for 24 h in different zinc content media. Our analysis showed that 100 μM zinc supplementation reduced the signal in our MTT assay (Fig. 1A). We also carried out annexin V and PI staining in cells incubated for 24 h at a zinc concentration of 10 and 100 μM (Fig. 1B). Our results confirmed that 100 μM of zinc promoted cell apoptosis in resting lymphocytes. We then studied zinc supplementation effects on activation. We detected a positive correlation between extracellular zinc content and IL-2 secretion under activation for 6 h using plate-bound anti-CD3/anti-CD28 (Fig. 1C). An annexin V and PI staining showed that apoptosis in activated cells was not induced at high zinc doses, contrary to what was observed before (Fig. 1D). We further studied the positive effect of zinc on activation in a dose-dependent manner by flow cytometry doing intracellular IL-2 staining (Fig. 1E, 1F). Using plate-bound anti-CD3/anti-CD28 with increasing concentrations of anti-CD3, we observed that 0.25 μg/ml showed the highest number of IL-2 stained cells independently of the zinc concentration (Fig. 1E). In contrast, our results also confirmed that extracellular zinc promoted the expression of IL-2. Remarkably, the presence of 100 μM of extracellular zinc boosted the strongest anti-CD3 stimulation. We performed similar studies using increasing concentrations of ionomycin plus PMA (Fig. 1F). In this case, we could not observe a correlation between extracellular zinc and IL-2 staining (Fig. 1F). Notably, although plate-bound anti-CD3/anti-CD28 triggers activation through the TCR receptor, ionomycin acts downstream, allowing divalent cations, mainly calcium, to enter from the extracellular medium. However, zinc can also enter through ionomycin pores (22) in an unspecific manner.

FIGURE 1.

Zinc in lymphocyte proliferation and activation. Jurkat cells were incubated in zinc-free medium supplemented with 0, 10, and 100 μM ZnSO4. (A) MTT assay of cells growing at different zinc doses for 24 h. (B) Apoptosis assay. Flow cytometry analyses of cells stained with PI and annexin V. Left, Representative dot blots. Right, Bar graph showing the mean of the different populations obtained with PI/annexin V staining. Live cells (LC), early apoptosis (EA), late apoptosis (LA), and dead/necrotic (D/N). (B) Cells were incubated for 24 h with a zinc concentration of 10 μM (basal) and 100 μM. (C) IL-2 secretion measurement by ELISA of cells activated with anti-CD3/anti-CD28 Abs for 6 h at different extracellular zinc concentration. **p < 0.01, Bonferroni-corrected ANOVA compared with basal condition. (D) Same apoptosis assay as in (B) but in cells activated during 6 h with anti-CD3/anti-CD28 Abs using 10 μM (basal) and 100 μM zinc concentration medium. (E and F) Bar graph representing the activation assay by flow cytometry with intracellular staining of IL-2 in cells incubated at different concentrations of zinc and activated for 6 h using 0.05, 0.25, and 0.5 μg/ml of plate-bound anti-CD3 plus anti-CD28 Abs (E) or 0.1, 1, and 2.5 μM ionomycin plus 10 nM PMA (F). *p < 0.05, **p < 0.01, ***p < 0.001, Bonferroni-corrected ANOVA compared with the weakest stimulus in each zinc concentration condition. #p < 0.05, ##p < 0.01 Bonferroni-corrected ANOVA compared between different zinc concentration conditions using the same stimulus. n is indicated in each bar.

FIGURE 1.

Zinc in lymphocyte proliferation and activation. Jurkat cells were incubated in zinc-free medium supplemented with 0, 10, and 100 μM ZnSO4. (A) MTT assay of cells growing at different zinc doses for 24 h. (B) Apoptosis assay. Flow cytometry analyses of cells stained with PI and annexin V. Left, Representative dot blots. Right, Bar graph showing the mean of the different populations obtained with PI/annexin V staining. Live cells (LC), early apoptosis (EA), late apoptosis (LA), and dead/necrotic (D/N). (B) Cells were incubated for 24 h with a zinc concentration of 10 μM (basal) and 100 μM. (C) IL-2 secretion measurement by ELISA of cells activated with anti-CD3/anti-CD28 Abs for 6 h at different extracellular zinc concentration. **p < 0.01, Bonferroni-corrected ANOVA compared with basal condition. (D) Same apoptosis assay as in (B) but in cells activated during 6 h with anti-CD3/anti-CD28 Abs using 10 μM (basal) and 100 μM zinc concentration medium. (E and F) Bar graph representing the activation assay by flow cytometry with intracellular staining of IL-2 in cells incubated at different concentrations of zinc and activated for 6 h using 0.05, 0.25, and 0.5 μg/ml of plate-bound anti-CD3 plus anti-CD28 Abs (E) or 0.1, 1, and 2.5 μM ionomycin plus 10 nM PMA (F). *p < 0.05, **p < 0.01, ***p < 0.001, Bonferroni-corrected ANOVA compared with the weakest stimulus in each zinc concentration condition. #p < 0.05, ##p < 0.01 Bonferroni-corrected ANOVA compared between different zinc concentration conditions using the same stimulus. n is indicated in each bar.

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We decided to further explore the modulation of zinc on lymphocytes by focusing on the main signaling cascades during T cell activation. To study this, we used a previously described Jurkat triple-reporter cell line for transcription factors NF-κB, NFAT, and AP-1 (21). Fig. 2A shows expression of CFP, eGFP, and mCherry reporters characterized by flow cytometry in T cells after 24 h of activation. We used different zinc concentrations and evaluated reporter expression at early (8 h) and late (24 h) time points after activation with two stimuli, PMA/ionomycin (Fig. 2B–D) or plate-bound anti-CD3/anti-CD28 Abs (Fig. 2E, 2F). Using PMA/ionomycin as activator, zinc supplementation reduced NF-κB, NFAT, and AP-1 reporter signal at 8 h while at 24 h only NF-κB remained affected, whereas NFAT and AP-1 signal tended to be higher with increasing zinc concentrations (Fig. 2B–D). In contrast, zinc improved activating signaling when using the anti-CD3/anti-CD28 stimulus, producing higher expression of NF-κB, NFAT, and AP-1 reporters under high dose of zinc at 24 h (Fig. 2E, 2F). These data further support the IL-2 staining results and suggest that zinc entry during T cell activation must occur through specific routes to modulate and promote the activation program.

FIGURE 2.

Modulation of NFAT, NF-кB, and AP-1 pathways by zinc. (A) Representative flow cytometry histograms using Jurkat triple-reporter cell line for NFAT (GFP), NF-кB (CFP), and AP-1 (mCherry) transcription factors obtained upon activation for 24 h using anti-CD3/anti-CD28 Abs. (BG). Jurkat triple-reporter line activated using PMA/ionomycin (B–D) or anti-CD3/anti-CD28 Abs (E–G). Bar graphs representing the percentage of GFP (B and E), CFP (C and F), and Cherry (D and G) positive cells at 8 and 24 h, to evaluate the activity of transcription factors NFAT, NF-кB, and AP-1, respectively (n = 3 in all conditions except in PMA/ionomycin 8 h, in which n = 6). *p < 0.05, **p < 0.01, ***p < 0.001, Bonferroni-corrected ANOVA compared with 0 μM zinc condition.

FIGURE 2.

Modulation of NFAT, NF-кB, and AP-1 pathways by zinc. (A) Representative flow cytometry histograms using Jurkat triple-reporter cell line for NFAT (GFP), NF-кB (CFP), and AP-1 (mCherry) transcription factors obtained upon activation for 24 h using anti-CD3/anti-CD28 Abs. (BG). Jurkat triple-reporter line activated using PMA/ionomycin (B–D) or anti-CD3/anti-CD28 Abs (E–G). Bar graphs representing the percentage of GFP (B and E), CFP (C and F), and Cherry (D and G) positive cells at 8 and 24 h, to evaluate the activity of transcription factors NFAT, NF-кB, and AP-1, respectively (n = 3 in all conditions except in PMA/ionomycin 8 h, in which n = 6). *p < 0.05, **p < 0.01, ***p < 0.001, Bonferroni-corrected ANOVA compared with 0 μM zinc condition.

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To determine the most probable candidate gate for zinc entry during activation, we characterized expression of the Zip family of transporters in different T lymphocyte models. In isolated human T cells, we detected the expression of most of the family members but Zip2, Zip5, and Zip12. (Fig. 3A). In addition, stimulation with anti-CD3/anti-CD28 promoted an increase in expression levels of Zip1, Zip4, Zip6, Zip7, Zip8, Zip9, Zip10, and Zip14. We then performed a similar experiment in the Jurkat cell line using the same two previous activation stimuli, anti-CD3/anti-CD28 Abs and PMA/ionomycin (Fig. 3B). Jurkat cells showed a similar expression pattern of Zip transporters as human T cells, with abundant expression of Zip1, Zip6, Zip7, Zip8, Zip9, Zip10, and Zip14, but only Zip6, Zip7, and Zip8 expression increased significantly upon anti-CD3/anti-CD28 stimulation. Intriguingly, Zip6 was not upregulated when PMA/ionomycin stimulation was performed. Finally, we used a murine model to explore the expression pattern of the Zip transporters in resting T cells isolated from the spleen, after activation with anti-CD3/anti-CD28 (Fig. 3C). We also used zinc-deficient medium and normal medium to compare early and late activation patterns. Our results showed that Zip6, Zip7, Zip9, Zip10, Zip11, and Zip14 increased their early expression and that the presence of zinc in the activation medium reduced the expression of Zip10. In summary, we observed that Zip6 was highly expressed in all models studied and was consistently upregulated during activation with anti-CD3/anti-CD28. Therefore, to study its role in lymphocyte physiology, we created knockdown of this transporter.

FIGURE 3.

ZIP transporter expression in lymphocytes. Real-time PCR was conducted in lymphocytes for the entire ZIP transporter family. Data is normalized to MCL51 (A and B) or β-actin (C) expression levels. (A) Lymphocytes extracted from three human donors activated at 24 and 48 h with anti-CD3/anti-CD28 Abs. (B) Jurkat cells treated for 24 h with PMA/ionomycin or anti-CD3/anti-CD28 Abs (n = 3). (C) T cells isolated from mouse spleen and activated at 4 and 24 h with anti-CD3/anti-CD28 Abs in medium with or without (w/o) zinc (n = 3). *p < 0.05, **p < 0.01, ***p < 0.001, Bonferroni-corrected ANOVA compared with basal condition.

FIGURE 3.

ZIP transporter expression in lymphocytes. Real-time PCR was conducted in lymphocytes for the entire ZIP transporter family. Data is normalized to MCL51 (A and B) or β-actin (C) expression levels. (A) Lymphocytes extracted from three human donors activated at 24 and 48 h with anti-CD3/anti-CD28 Abs. (B) Jurkat cells treated for 24 h with PMA/ionomycin or anti-CD3/anti-CD28 Abs (n = 3). (C) T cells isolated from mouse spleen and activated at 4 and 24 h with anti-CD3/anti-CD28 Abs in medium with or without (w/o) zinc (n = 3). *p < 0.05, **p < 0.01, ***p < 0.001, Bonferroni-corrected ANOVA compared with basal condition.

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Using a Jurkat cell line with constitutive Cas9 expression (Cas9–wild type [WT]), we generated a Zip6KO clone, with a frame shift introduced in both alleles (Fig. 4A). As far as we are aware, there is no reliable commercial Ab to check Zip6 expression, so we simply compared the RNA expression in the KO line to that in the Cas9-WT line and observed a marked change in expression of the different Zip transporters in the Zip6KO clone (Fig. 4B). The proliferation of the Zip6KO line was not markedly different from that of the control cells (Fig. 4C). We also evaluated zinc transport in the Zip6KO line compared with that in the Cas9-WT line. By monitoring zinc entry with Zinquin, we found no difference in transport kinetics between these cell lines after adding 100 μM ZnSO4 to the external solution (Fig. 4D), suggesting either that Zip6 is not essential for zinc transport under basal conditions or that the observed upregulation of other Zip transporters compensates the zinc transport activity. In line with this, incubating the cells with 0, 10, and 100 μM Zn2+ for 24 h did not significantly modify intracellular zinc content, as measured using FluoZin-3, AM (Fig. 4E).

FIGURE 4.

Characterization of ZIP6KO cell line. (A) Schematic representation of the existing mutations in both alleles of the Slc39a6 gene in the ZIP6KO cell line generated from a Cas9 stable cell line transfected with guide RNAs targeting exon 3. (B) Real-time PCR was conducted in the in ZIP6KO line for the entire ZIP transporter family, and results compared with those in stable Cas9-expressing line (Cas9-WT line) (n = 3–6). *p < 0.05, **p < 0.01 using t test. (C) Proliferation analysis, counting cells at day 2 and 4 in ZIP6KO line compared with Cas9-WT line. (D) Zinc transport assay using Zinquin, monitored by fluorescence microscopy for 10 min after adding (arrow) 100 μM ZnSO4 in the external solution. (E) Evaluation of zinc content by flow cytometry in cells incubated for 24 h with 0, 10, and 100 μM ZnSO4 and stained with FluoZin-3, AM. Left, Representative histogram of ZIP6KO and Cas9-WT cells incubated with 0 and 100 μM ZnSO4. Right, Quantification at different zinc concentrations. **p < 0.01 compared with basal condition, Bonferroni-corrected ANOVA.

FIGURE 4.

Characterization of ZIP6KO cell line. (A) Schematic representation of the existing mutations in both alleles of the Slc39a6 gene in the ZIP6KO cell line generated from a Cas9 stable cell line transfected with guide RNAs targeting exon 3. (B) Real-time PCR was conducted in the in ZIP6KO line for the entire ZIP transporter family, and results compared with those in stable Cas9-expressing line (Cas9-WT line) (n = 3–6). *p < 0.05, **p < 0.01 using t test. (C) Proliferation analysis, counting cells at day 2 and 4 in ZIP6KO line compared with Cas9-WT line. (D) Zinc transport assay using Zinquin, monitored by fluorescence microscopy for 10 min after adding (arrow) 100 μM ZnSO4 in the external solution. (E) Evaluation of zinc content by flow cytometry in cells incubated for 24 h with 0, 10, and 100 μM ZnSO4 and stained with FluoZin-3, AM. Left, Representative histogram of ZIP6KO and Cas9-WT cells incubated with 0 and 100 μM ZnSO4. Right, Quantification at different zinc concentrations. **p < 0.01 compared with basal condition, Bonferroni-corrected ANOVA.

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We then studied the zinc transport properties of the Zip6KO cell line activated by anti-CD3/anti-CD28. After adding 100 μM ZnSO4 to the external solution of Cas9-WT cells, the transport kinetics was significantly higher from those in the Zip6KO line when monitoring zinc entry using Zinquin (Fig. 5A). In addition, using flow cytometry in activated cells stained with FluoZin-3, AM, we observed that activated T cells had higher zinc content than nonactivated cells. However, zinc content was lower in activated Zip6KO cells than in activated Cas9-WT cells (Fig. 5B). In this scenario, we studied activation markers using real-time PCR at early and late activation time points. The activation program was substantially altered in our Zip6KO line, with IL-2 and IFN-γ being strongly affected at early time points (Fig. 5C, 5D). In contrast, CD25 expression was less affected by Zip6 knockdown (Fig. 5E). We also monitored IL-2 production using flow cytometry under anti-CD3/anti-CD28 activation (Fig. 5F) and observed fewer IL-2–positive cells in the Zip6KO line than in control cells, in line with the previous results. Moreover, we also studied the positive effect of zinc supplementation on T cell activation (Fig. 5G). Our results showed that, unlike control cells, there was no potentiation in the activation of Zip6KO cells with 100 μM Zn2+. We wanted to confirm this result performing an ELISA. Our results showed that although zinc promotes IL-2 secretion in Cas9-WT cells at 10 and 100 μM Zn2+, ZIP6KO cells failed to produce the cytokine after 6 h anti-CD3/anti-CD28 stimulation (Fig. 5H). In this scenario, zinc has been shown to regulate early events of TCR signaling pathway that are translated in different ZAP70 phosphorylation levels (16). However, in our Zip6KO cells we did not observe a diminished ZAP70 phosphorylation compared with control cells upon activation, suggesting that the initial steps of the activation pathway were maintained (Fig. 5I).

FIGURE 5.

Impaired activation in ZIP6KO cell line. (A) Zinc transport assay using Zinquin, monitored by fluorescence microscopy for 10 min after adding (arrow) 100 μM ZnSO4 in the external solution; ZIP6KO and Cas9-WT cells activated for 24 h with anti-CD3/anti-CD28 Abs. Right, Bar graph representing the area under the curve at 10 min (n = 6; n = 140–168 cells). ***p < 0.01 using t test. (B) Evaluation of zinc content by flow cytometry in ZIP6KO and Cas9-WT cells activated for 24 h with anti-CD3/anti-CD28 Abs and stained with FluoZin-3, AM. Left, Representative histogram of ZIP6KO and Cas9-WT cells under basal and activated conditions. Right, Quantification of fluorescence intensity compared with basal Cas9-WT cells (n = 3), arbitrary units. (CE) Real-time PCR to quantify the expression of IL-2 (C), IFN-γ (D), and CD25 (E) in ZIP6KO and Cas9-WT cells activated at 4 and 24 h using anti-CD3/anti-CD28 Abs (n = 3). (F and G) Flow cytometry analysis of IL-2 immunostaining in ZIP6KO and Cas9-WT cells in RPMI + 10% FBS medium (F) (n = 3) or at different zinc concentrations (G) (n = 6, 3 for WT, ZIP6KO, respectively). (H) IL-2 secretion measurement by ELISA of cells activated with anti-CD3/anti-CD28 Abs for 6 h at different extracellular zinc concentration. (A, F, and I) t test. (B–E, G, and H) Bonferroni-corrected ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001 compared with basal conditions, #p < 0.05, ##p < 0.01, ###p < 0.001 between cell lines. (I) Western blot against phospho-ZAP70 (pZAP70) and total ZAP70 in calls activated at 15, 30, and 60 min with anti-CD3/anti-CD28 Abs. Left, Representative Western blot. Right, Bar graph quantification (n = 3–6). *p < 0.05, **p < 0.01, t test compared with nonactivated conditions.

FIGURE 5.

Impaired activation in ZIP6KO cell line. (A) Zinc transport assay using Zinquin, monitored by fluorescence microscopy for 10 min after adding (arrow) 100 μM ZnSO4 in the external solution; ZIP6KO and Cas9-WT cells activated for 24 h with anti-CD3/anti-CD28 Abs. Right, Bar graph representing the area under the curve at 10 min (n = 6; n = 140–168 cells). ***p < 0.01 using t test. (B) Evaluation of zinc content by flow cytometry in ZIP6KO and Cas9-WT cells activated for 24 h with anti-CD3/anti-CD28 Abs and stained with FluoZin-3, AM. Left, Representative histogram of ZIP6KO and Cas9-WT cells under basal and activated conditions. Right, Quantification of fluorescence intensity compared with basal Cas9-WT cells (n = 3), arbitrary units. (CE) Real-time PCR to quantify the expression of IL-2 (C), IFN-γ (D), and CD25 (E) in ZIP6KO and Cas9-WT cells activated at 4 and 24 h using anti-CD3/anti-CD28 Abs (n = 3). (F and G) Flow cytometry analysis of IL-2 immunostaining in ZIP6KO and Cas9-WT cells in RPMI + 10% FBS medium (F) (n = 3) or at different zinc concentrations (G) (n = 6, 3 for WT, ZIP6KO, respectively). (H) IL-2 secretion measurement by ELISA of cells activated with anti-CD3/anti-CD28 Abs for 6 h at different extracellular zinc concentration. (A, F, and I) t test. (B–E, G, and H) Bonferroni-corrected ANOVA, *p < 0.05, **p < 0.01, ***p < 0.001 compared with basal conditions, #p < 0.05, ##p < 0.01, ###p < 0.001 between cell lines. (I) Western blot against phospho-ZAP70 (pZAP70) and total ZAP70 in calls activated at 15, 30, and 60 min with anti-CD3/anti-CD28 Abs. Left, Representative Western blot. Right, Bar graph quantification (n = 3–6). *p < 0.05, **p < 0.01, t test compared with nonactivated conditions.

Close modal

We then wondered whether the impaired activation observed in our Zip6KO cells might be due an artifact related to clone selection. For this reason, we repeated the activation experiments using a different Zip6KO clone (Zip6KO2) with a slightly different frame shift mutation in the Slc39a6 alleles (Supplemental Fig. 1A). We discarded a clonal selection effect because the Zip6KO2 clone also showed lower activation than control cells (Supplemental Fig. 1B–D). We did anti-CD3 and anti-TCR immunostainings in our KO clones to rule out the possibility that they had lost CD3 or TCR components during the selection process (Supplemental Fig. 1E, 1F). Thus, we found that the percentage of TCR-positive cells in M1 was 76.1 ± 0.8 in WT, 83.0 ± 0.3 in Zip6KO, and 98.5 ± 0.2 in Zip6KO2 cells and the percentage of CD3-positive cells in M1 was 82.9 ± 0.8 in WT, 92.1 ± 0.2 in Zip6KO, and 99.5 ± 0.1 in Zip6KO2 cells. The TCR total mean fluorescence intensity was 2991 ± 71 in WT, 1886 ± 27 in Zip6KO, and 3854 ± 130 in Zip6KO2 cells and the CD3 mean fluorescence intensity in arbitrary units was 5158 ± 188 in WT, 3836 ± 125 in Zip6KO, and 7043 ± 43 in Zip6KO2 cells. All data are expressed as mean ± SD. We ruled out the possibility that the activation defect present in our clones was due to different expression of TCR molecular components because Zip6KO2 cells have similar impairment than Zip6KO cells and higher expression of TCR and CD3 than Cas9-WT cells when activating with anti-CD3/anti-CD28 stimulation (Supplemental Fig. 1G). In addition, Zip6KO cells with high TCR expression levels (Zip6KO High) were sorted and activated (Supplemental Fig. 1H). This new population of Zip6KO cells expressing high levels of TCR complex showed as well an impaired production of IL-2 compared with control cells.

We decided to explore the activation profile of the Zip6KO cells independently of TCR signaling events by using PMA/ionomycin stimulation (Fig. 6). Monitoring IL-2 and IFN-γ expression by real-time PCR, we observed lower activation in Zip6KO and Zip6KO2 cell lines compared with Cas9-WT cells (Fig. 6A, 6B). In addition, we explored a dose-dependent stimulation to rescue the Zip6KO line with a stronger stimulus of ionomycin. Contrary to what we observed in Cas9-WT cells, we could not see any improvement in IL-2 production upon 2.5 μM treatment (Fig. 6C). In a similar manner, we also detected an important reduction of secreted IL-2 in the extracellular medium of Zip6KO cells compared with Cas9-WT cells activated with PMA/ionomycin (Fig. 6D).

FIGURE 6.

ZIP6KO affects activation independently of TCR machinery. (A and B) Real-time PCR to quantify the expression of IL-2 (A) and IFN-γ (B) in ZIP6KO, ZIP6KO2, and Cas9-WT cells activated for 4 and 24 h with PMA/ionomycin (n = 3). Bonferroni-corrected ANOVA, *p < 0.05, ***p < 0.001 compared with basal conditions, #p < 0.05, ###p < 0.001 between cell lines. (C) Flow cytometry analysis of IL-2 immunostaining in ZIP6KO and Cas9-WT cells for 6 h activated at 0.1, 1, and 2.5 μM ionomycin plus 10 nM PMA (n = 3). Bonferroni-corrected ANOVA, ***p < 0.001 compared with 0.1 μM condition and ###p < 0.001 between cell lines. (D) IL-2 secretion measurement by ELISA of cells activated for 6 h with PMA/ionomycin (n = 2, 3 for Cas9WT, ZIP6KO, respectively). **p < 0.01, t test. (E) Representative Western blot of phospho-GSK-3β, GSK-3β, and β-actin in ZIP6KO and Cas9-WT cells activated for 1 h using anti-CD3/anti-CD28 Abs or PMA/ionomycin. (F) Flow cytometry analysis of IL-2 immunostaining in ZIP6KO and Cas9-WT cells activated for 6 h with PMA/ionomycin, upon addition of 5 mM of LiCl or KCl (n = 3), *p < 0.05 compared with basal conditions, ###p < 0.001 between cell lines, Bonferroni-corrected ANOVA.

FIGURE 6.

ZIP6KO affects activation independently of TCR machinery. (A and B) Real-time PCR to quantify the expression of IL-2 (A) and IFN-γ (B) in ZIP6KO, ZIP6KO2, and Cas9-WT cells activated for 4 and 24 h with PMA/ionomycin (n = 3). Bonferroni-corrected ANOVA, *p < 0.05, ***p < 0.001 compared with basal conditions, #p < 0.05, ###p < 0.001 between cell lines. (C) Flow cytometry analysis of IL-2 immunostaining in ZIP6KO and Cas9-WT cells for 6 h activated at 0.1, 1, and 2.5 μM ionomycin plus 10 nM PMA (n = 3). Bonferroni-corrected ANOVA, ***p < 0.001 compared with 0.1 μM condition and ###p < 0.001 between cell lines. (D) IL-2 secretion measurement by ELISA of cells activated for 6 h with PMA/ionomycin (n = 2, 3 for Cas9WT, ZIP6KO, respectively). **p < 0.01, t test. (E) Representative Western blot of phospho-GSK-3β, GSK-3β, and β-actin in ZIP6KO and Cas9-WT cells activated for 1 h using anti-CD3/anti-CD28 Abs or PMA/ionomycin. (F) Flow cytometry analysis of IL-2 immunostaining in ZIP6KO and Cas9-WT cells activated for 6 h with PMA/ionomycin, upon addition of 5 mM of LiCl or KCl (n = 3), *p < 0.05 compared with basal conditions, ###p < 0.001 between cell lines, Bonferroni-corrected ANOVA.

Close modal

We also studied the role of the zinc-dependent kinase GSK-3β, which has been found to bind to Zip6 (7, 23). During T cell activation, GSK-3β is inactivated by phosphorylation, allowing NFAT retention in the cell nucleus (24, 25). Therefore, we evaluated GSK-3β phosphorylation in our Zip6KO line using Western blot. We did not observe any major differences in phospho-GSK-3β under activation between Zip6KO and Cas9-WT cells using anti-CD3/anti-CD28 or PMA/ionomycin stimuli (Fig. 6E). Moreover, to test whether GSK-3β activity underlies the activation defects observed in our KO line, we inhibited GSK-3β using LiCl and then repeated IL-2 staining during activation with PMA/ionomycin. Our results showed that LiCl potentiated activation in Cas9-WT cells but did not affect Zip6KO cells (Fig. 6F).

Trace metals have diverse functions in the body, being essential for the structure and activity of proteins and participating in many reactions. Of these, zinc could be considered a special case. In addition to being a structural component of many proteins, there is growing evidence supporting its role as a second messenger, thus sharing a number of features with calcium (2, 26). Therefore, cytosol concentrations of zinc are tightly regulated by the coordinated action of transporters and chelators. Moreover, fast and slow changes in cytosolic zinc levels modify the activity of existing enzymes and promote changes in the transcription program (8). Moreover, zinc, calcium, and redox signaling are highly interconnected, allowing cells to integrate signals and effectively respond to environmental changes. However, although calcium signaling has been studied intensively in recent decades, the zinc signaling machinery is still poorly understood. In this scenario, our work tries to dissect the molecular players involved in one of the most pronounced impacts caused by zinc imbalance, the immune dysfunction.

Early studies of zinc deficiency showed that it is essential for immune system function (2). These studies were further supported by the phenotypic characterization of the genetic disease acrodermatitis enteropathica, which is caused by mutations in Slc39a4, resulting in zinc malabsorption, lymphopenia, and thymus atrophy (27, 28). Zinc deficiency is currently considered a worldwide public health problem that increases childhood morbidity by around 4% (29). However, the cellular and molecular mechanisms that underlay immune dysfunction and the specific role of zinc in immune cells are still poorly understood. Thus, although zinc is known to affect T lymphocyte maturation, differentiation, and cytokine production (1214), there is no consensus on its specific role in T cells. On the one hand, zinc has been shown to be required for the correct TCR signaling (16, 17), and activated T cells are known to have increased zinc content (15, 16). On the other hand, there have also been contrasting results regarding zinc supplementation and T cell activation (16, 18, 20). Our work shows that high doses of zinc could cause cytotoxicity, as has been seen in neurons (30). In contrast, zinc supplementation engages the activation machinery, including NFAT, NF-κB, and AP-1 cascades, when cell activation is triggered using antiCD3/antiCD28 Abs. This beneficial effect is not that evident when using PMA plus ionomycin as stimuli, as previously reported (18), highlighting the requirement of specific zinc entry routes for the positive modulation. Our results show that the NF-κB pathway is the most sensitive to aberrant zinc entry using ionomycin, an ionophore that creates zinc-permeable membrane pores (22). In fact, in innate immune cells, zinc acts as an immunosupressor through its ability to prevent nuclear translocation of NF-κB (9). The triple-reporter line used in this study has allowed us to dissect and better understand the impact of zinc in the T cell activation cascade. Finally, our results provide strong evidences of the positive effects on T cell response upon activation caused by physiological concentrations of zinc and by zinc supplementation (Fig. 1C, 1E). These results, together with the potentiation observed with high levels of zinc to strong stimuli (Fig. 1E), allow to better understand the protection provided by zinc supplements against infections reported in some studies (3).

Our work highlights Zip6 as a key component of the T cell activation machinery. Other transporters have previously been postulated as candidates for the increased cytosolic zinc content present in activated cells (16, 19, 20). We do not discard the notion that Zip8 participates in zinc transport in activated lymphocytes, as previously reported (20). In fact, Zip8 expression is induced in activated Jurkat cells and isolated human lymphocytes but not in mouse T cells. Zip10 is also a plausible candidate considering the regulation observed in human T cells and murine lymphocytes, and also because we found that Zip10 expression depends on the presence or absence of zinc in the extracellular medium. Moreover, Zip10 and Zip6 form heteromers in other cellular systems (7). However, in Jurkat cells, we did not observe upregulation under activation, which argues against the relevance of the heteromer in this cell type. Moreover, the Zip10KO mouse model does not demonstrate any T cell dysfunction (10, 11). Finally, our results do not support a role for Zip12 in activation (19), given the low expression of this transporter in all the models we have studied. In contrast, it would be interesting to further study the role of Zip7. This transporter is activated by extracellular zinc entry and increases zinc levels in the cytosol by transporting zinc out of the endoplasmic reticulum (6). Our experiments showed that Zip7 is upregulated in activated Jurkat cells and mouse T cells.

Our Zip6KO cell lines show markedly impaired activation, although they proliferate normally under basal conditions. The normal zinc transport in nonactivated cells could be explained by some compensatory mechanism, so we cannot discard the involvement of this transporter under nonactivated conditions. Moreover, activated T cells have markedly reduced ability to transport extracellular zinc and have reduced zinc content (Fig. 5A, 5B). These data support the role of Zip6 zinc transport during activation, and the need of specific routes for zinc entry. It is remarkable that high doses of zinc did not produce any positive modulation in cells activated with anti-CD3/anti-CD28 Abs (Fig. 5G, 5H), supporting the idea that Zip6 is the functional gate during activation required to engage the TCR signaling machinery (16, 17). However, it is even more surprising that our Zip6KO cells did not benefit from activation by PMA/ionomycin (Fig. 6C, 6D), as this drug potentially allows unspecific zinc entry and activates T cells downstream of the TCR. Remarkably, the inhibition of GSK-3 kinase could not rescue Zip6KO activation, so we discard that the underlying defect in our KO cells is a dysregulation on this enzyme, a hypothesis based on the interaction between Zip6 and GSK-3β previously described (7, 23) and the required kinase inactivation for the correct activation of T lymphocytes (24, 25). Therefore, the absence of Zip6 seems to chronically alter some as yet unknown molecular players involved in signal transduction during T cell activation. Despite the compensatory mechanisms observed by upregulation of other cell transporters in Zip6KO cell line, those proteins were unable to restore the activation program. Zip6, together with Zip5 and Zip10, belongs to a branch of Zip transporters characterized by the presence of a prion-like ectodomain that has been shown to be relevant in the interactome of Zip6 and its function during cell proliferation and development (7, 31, 32). Whether this specific role of Zip6 on activation might relay on these specific molecular determinants is an attractive hypothesis that requires further studies.

Taken together, our results support the notion that zinc is essential for T lymphocyte physiology, provide new evidence of the how the signaling cascade is modulated during T cell activation, and highlight the Zip6 transporter as an important player during the activation process. However, further work is needed to fully understand the role of this protein in the activation machinery. The ultimate future goal would be to use Zip6 as a therapeutic target for immunomodulation, given that it is an accessible transmembrane protein at the plasma membrane involved in T lymphocyte activation.

This work was supported by a grant from the Spanish Ministry of Economy and Competitiveness (SAF2014-52228-R).

The online version of this article contains supplemental material.

Abbreviations used in this article:

KO

knockout

PI

propidium iodide

RT

room temperature

WT

wild type.

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The authors have no financial conflicts of interest.

Supplementary data