Peripheral blood mononuclear cells (PBMCs) of healthy donors were isolated by leukapheresis. Leukapheresis was approved by the Ethics committee of the University Hospital Regensburg (Ethic Vote Number 13-101-0240; 13-101-0238); all human participants gave written informed consent. Human monocytes were isolated from PBMCs by density gradient centrifugation over Ficoll/Hypaque followed by counterflow centrifugation elutriation (19). Monocytes were seeded on Teflon foils (to allow detachment of cells after cultivation, DuPont) at a concentration of 1×10⁶ cells/mL in RPMI1640 supplemented with 2% AB-serum (heat-inactivated; Bavarian red cross, Germany), penicillin (100 U/mL), streptomycin (100 µg/mL), with or without 2 mM L-glutamine (all from Thermo Fisher Scientific) with 2% pooled human serum for 7 days. Alternatively, macrophages were seeded at a concentration of 2×10⁶ cells per well in a 6-well plate in RPMI (supplements mentioned above). DCs were generated by incubating monocytes in RPMI1640 with 10% FCS (heat-inactivated; Sigma-Aldrich) and 224 IU/ml GM-CSF and 144 IU/ml IL-4 (both PeproTech) in the absence or presence of L‐glutamine for seven days. When indicated, cultures were treated with α-ketoglutarate (Keto; Sigma-Aldrich) at concentration of 1 mM or L-methionine-S-sulfoximine (MSO; Sigma-Aldrich) at a concentration of 1 mM.

Bone marrow-derived macrophages (BMDM) were generated from bone marrow cells of wildtype C57BL/6NCrl (Charles River Breeding Laboratories) in Teflon bags containing DMEM medium (Gibco) supplemented with 10% fetal calf serum (FCS) (Sigma-Aldrich), 5% Horse Serum (Cell Concepts) and supernatant of L929 cells containing M-CSF for 7 days at 10% CO₂ as described earlier (20, 21).

The human AML cell line THP-1 was obtained from the DSMZ (Braunschweig, Germany) and cultured in RPMI1640 (PAN, Aidenbach, Germany) supplemented with 10% fetal calf serum (FCS) (heat-inactivated), penicillin (50 IU/mL) and streptomycin (50 µg/mL), and 2 mM L-glutamine.

Cell proliferation in culture was monitored using the CASY Cell Counter system (OMNI Life Science).

Cytokine secretion was determined in 24 h culture supernatants by commercially available enzyme-linked immunosorbent assays (R&D Systems) according to the manufacturer’s protocol.

To perform [¹³C₅]glutamate tracing in cell culture, human monocyte-derived dendritic cells, macrophages and the AML cell line THP-1 were cultured in glutamine rich media (as described above) for 7 days for primary human cells and 48 h for THP-1. Cells were then washed with glutamine-free medium and re-cultured for 24 h with 2 mM [¹³C₅]glutamate (Cambridge Isotope Labs). After supernatants had been taken off, cells were washed twice in PBS and immediately frozen in liquid nitrogen. Supernatants and cell pellets were kept at −80°C until further analysis. Experiments were performed with cells from 3 different donors, for THP-1 at 3 different time-points each.

For metabolite extraction cell pellets were vortexed with 600 µL of 80% methanol (80:20 methanol: water, v/v) and centrifuged at 10.000 g and 4°C for 5 minutes. The supernatant was collected, and the remaining pellet was washed twice with 200 µL of 80% methanol employing 15.000 g for the last centrifugation step. The combined extracts were dried using an infrared vortex vacuum evaporator (CombiDancer, Hettich AG, Baech, Switzerland) and the residue was reconstituted in 100 µL pure water. Amino acids isotopologues were analyzed by high performance liquid chromatography (HPLC)–electrospray ionization (ESI)–tandem MS (MS/MS) after propyl chloroformate derivatization (22) employing a modified derivatization procedure that uses 17.4% propyl chloroformate and 82.6% isooctane as reagent 2. A 10 µL aliquot of the cell extract or 10 µL of cell culture supernatant were employed for derivatization. The MS/MS method was modified to include transitions for each possible isotopologue. Raw peak areas were corrected for natural stable isotope abundance and tracer impurity using IsoCorrectoR (23). Data are presented as mean ¹³C enrichment.

Samples were lysed in RIPA (Sigma-Aldrich) separated by 12% SDS-PAGE and transferred to PVDF membranes, blocked with 5% milk (Sucofin) in TBS buffer with 0.1% Tween for 1 h, and incubated with primary antibodies overnight (GLUL; rabbit-anti-human/mouse, (Abcam Cat#197024, RRID: AB_10704544) and Actin; rabbit-anti-human/mouse, (Sigma-Aldrich Cat #A2066, RRID: AB_476693)). On the next day, membranes were washed and then incubated with the following secondary antibody for 1 h at RT: goat-anti-rabbit HRP (DakoCytomation Cat #P0448, RRID: AB_2617138). Detection was performed by chemiluminescence (ECL, Amersham Bioscience) and analyzed using the chemiluminescence system Fusion Pulse 6 (Vilber Lourmat).

In order to analyze immune cells, the following markers were used: CD11b FITC (BD Biosciences Cat #561015, RRID: AB_10561676), CD14 FITC (Beckman Coulter Cat# IM0645U, RRID : AB_130992), HLA-DR (BD Biosciences Cat #347400, RRID: AB_2868846), CD86 (BD Biosciences Cat #555657, RRID: AB_396012), HLA-ABC FITC (BD Biosciences Cat #555552, RRID: AB_395935), SIRP1α PE (BioLegend Cat #372104, RRID: AB_2650861), CD1a PE (Beckman Coulter, Cat #6603185), and DC-Sign PE (R and D Systems Cat #FAB161P, RRID: AB_357064). Cells were harvested in FACS tubes and washed with 1 ml of FACS wash buffer (PBS + 2% FCS). After addition of the antibodies, cells were incubated at 4°C for 20 min in the dark. After a washing step with 1 ml FACS wash buffer, cells were resuspended in FACS wash buffer and immediately analyzed by flow cytometry. To determine cell autofluorescence, unstained cells were used. Flow cytometry was performed using a BD FACS Calibur or Celesta. Data were analyzed with the FlowJo software (v10.8.1, Tree Star, RRID: SCR_008520).

Oxygen consumption was monitored non-invasively over time under cell culture conditions using the PreSens technology. 0.3×10⁶ cells were seeded in 1 ml in OD24 OxoDish^® plates (PreSens Precision Sensing GmbH) with pre-calibrated O₂ sensors at the bottom of each well for non-contact reading by the SDR SensorDish^® Reader (PreSens Precision Sensing GmbH) through the transparent material of the plate.

Extraction of RNA, reverse transcription and quantitative real-time PCR for gene expression analysis in myeloid cells was performed as described elsewhere (24). Primer sequences (all from Eurofins MWG Operon, Ebersberg, Germany) were as follows (-5’-3’): 18S rRNA: sense, 5′-ACCGATTGGATGGTTTAGTGAG-3′ and antisense, 5′-CCTACGGAAACCTTGTTACGAC-3′. GLUL: sense, 5′-CATGTATCTCGTGCCTGCTGCC-3′ and antisense, 5′GTTGCTCACCATGTCCATTATCCGT-3′; GLUD1, gene-specific Primer PCR SYBR Green assay (Assay ID qHsaCED0038578) and GLUD2, gene-specific Primer PCR SYBR Green assay (Assay ID qHsaCED0038220).

The siRNAs targeting GLUL and the respective non-target control pool were achieved from Dharmacon and dissolved in PCR-grade H₂O to a final concentration of 1 µg/µL and stored at -20°C. Electroporation with siRNAs was performed with human monocyte-derived macrophages, which were differentiated for 7 days in RPMI1640 supplemented with 2% AB-serum, penicillin (100 U/mL), streptomycin (100 µg/mL), 2 mM L-glutamine and 2% pooled human serum. A total of 1×10⁷ macrophages per electroporation were washed first with RPMI1640 without phenol red and then with 10 mL OptiMEM (Thermo Fisher Scientific). Afterwards, cells were suspended in OptiMem to a final concentration to a final concentration of 1×10⁷ cells/mL. Electroporation with siRNA was performed with 1 µg siRNA/1×10⁷ cells using the Gene Pulser XCell Electroporation System (BioRad). Cells were directly seeded at a concentration of 1×10⁶ cells/mL in pre-warmed cell culture medium with or without 2 mM L‐glutamine.

Statistical parameters reported in the figures and the figure legends include the exact number of experiments (n), the definition of the center, statistical tests applied and statistical significance. Statistical analyses were performed with the GraphPad Prism software (version 9). Two-group comparisons of matched data sets were performed using the Wilcoxon test, as the number of experiments did not allow testing for normality distribution. Multiple comparisons of matched data sets were performed using the one-way ANOVA and post-hoc Tukey’s multiple comparison test or Friedman test and post-hoc Dunn’s or the two-way ANOVA and post-hoc Bonferroni’s multiple comparison test. In figures asterisks denote statistical significance (*p < 0.05, **p < 0.01, ***p < 0.001).

Myeloid cell lines can serve as a model system for human primary monocytes (25). Therefore, we investigated initially the impact of glutamine deficiency on the monocytic leukemia cell line THP-1. After 48 h of culture with glutamine (2 mM) and without glutamine (glutamine free medium plus serum), we determined cell numbers of THP-1 cells. Glutamine deprived culture conditions resulted in reduced proliferation ( Figure 1A ) but cell viability was preserved ( Figures 1B, C ). Next, we investigated the long-term impact of glutamine deprivation. THP-1 cells were counted and reseeded at 300.000 cells/mL every 48 h for 12 days and the theoretical cell yield was calculated. In the absence of glutamine, THP-1 cells showed almost no detectable proliferation, while in the presence of glutamine cells proliferated exponentially and showed consistently high viability ( Figures 1C–E ). These data indicate a strong dependence of THP-1 cells on exogenous glutamine supply, which might in part be related to their strong proliferative activity.

Primary human monocytes, macrophages and DCs do not proliferate, and we were interested whether primary cells would, therefore, be less dependent on glutamine supply. We seeded monocytes in either glutamine-containing (2 mM) or glutamine-free medium in the presence of 10% FCS, GM-CSF and IL-4 to induce monocyte DC differentiation (26). Based on the literature 10% FCS should result in glutamine levels of approximately 0.1 mM in unsupplemented culture medium (27). Successful differentiation was followed by determining DC surface marker expression and LPS-stimulated cytokine secretion in culture supernatants. After 7 days, cell viability, cell numbers and morphology of monocyte-derived DCs were comparable ( Figures 2A–D ). The absence of glutamine did not affect the expression of DC surface markers such as CD1a, DC-SIGN, and HLA-DR ( Figures 2E, F ). Furthermore, production of IL-12, IL‐10, IL-6 and TNF was not altered by glutamine deprivation ( Figures 2G ).

Human blood monocytes can be differentiated into either DCs or monocyte-derived macrophages in vitro. To study the impact of glutamine on differentiation and activation of monocyte-derived macrophages, we cultured monocytes on Teflon foils for 7 days with 2% human serum with and without glutamine. 2% human serum should result in about 0.012 mM glutamine in supplement-free culture medium as human serum contains ~ 0.6 mM glutamine (28–30). Similar to our results with DCs, no major differences were observed regarding cell viability, cell yield and morphology ( Figures 3A–D ). The expression of various surface markers as well as LPS-stimulated cytokine secretion was also not significantly altered ( Figures 3E–G ). We also investigated the impact of glutamine shortage in a short-term setting. Here, by trend IL-6 secretion was lowered and addition of α-ketoglutarate (Keto) partially counteracted the effect ( Figure 3H ).

Furthermore, we investigated whether macrophage polarization and different culture conditions would change glutamine dependency. Monocytes were differentiated into macrophages on plastic plates in the presence of 2% human serum with or without IL-10 or LPS. Again, glutamine deprivation did not lead to macrophage cell death and did not block differentiation-associated morphological changes ( Figures S1A–C ).

Protein expression of glutamine synthetase (GS), the enzyme that catalyzes de novo glutamine synthesis, largely determines glutamine dependency. GS expression varies between cells and is low in auxotrophic (tumor) cells, which leads to glutamine-addiction (31). The conversion of glutamate, ATP and ammonia into glutamine is irreversibly blocked by L-methionine-S-sulfoximine (MSO) (32) (schematic view Figure 4A ). To investigate the observed differences in glutamine dependency, we measured the mRNA expression of glutamine synthetase (GLUL), glutaminase (GLS) and glutamate dehydrogenase (GLUD1) by RT-PCR in THP-1 and human macrophages grown in glutamine-rich (2 mM) and glutamine-poor medium. GLUL mRNA expression was detected in THP-1 and not altered by glutamine absence or presence ( Figure 4B ). In contrast, GS protein expression was low in the presence of glutamine and strongly upregulated under glutamine deprivation ( Figure 4C ). Macrophages showed a strong constitutive expression of GLUL mRNA and GS protein, which was by trend slightly upregulated in the absence of glutamine ( Figures S2A, B ). Surprisingly, GS protein was almost undetectable in murine macrophages, which may also relate to the different culture conditions between human and murine macrophages ( Figure S2B ). Glutaminase mRNA expression was lower in primary cells, while GLUD1 expression did not differ between leukemia cells and macrophages ( Figures S2C, D ).

As THP-1 cells upregulated GS under glutamine-deficient culture conditions, we wondered whether glutamate supplementation could rescue THP-1 proliferation. Indeed, glutamate increased dose-dependently the proliferation of THP-1 cells in absence of glutamine and cell counts returned to control levels upon high glutamate supplementation ( Figure 4D ). Glutamate also reverted the suppression of oxygen consumption in THP-1 cells ( Figure 4E ). Additional MSO treatment reversed the positive effect of glutamate, whereas cells cultured in the presence of glutamine were not affected by MSO (data not shown). To determine the enzyme activity of GS in THP-1, we performed tracing analyses. THP‐1 cells were incubated for 24 h with 2 mM [¹³C]glutamate in the presence or absence of 1 mM MSO. The intracellular and extracellular levels of [¹³C]glutamine and [¹³C]glutamate, resulting from intracellular glutamate metabolism, were determined in cell lysates and supernatants of THP-1 cells by mass spectrometry. As expected, glutamate supplementation resulted in intracellular and extracellular glutamine ¹³C enrichment, which was decreased by MSO treatment ( Figures 4F, G ). Higher enrichment was detected in macrophages ( Figures S2E, F ). Surprisingly MSO did only partially suppress glutamine production in macrophages.

To explore the impact of GS on monocyte differentiation, we cultured monocytes with or without glutamine in the presence or absence of MSO, blocking GS activity. MSO treatment had no impact on DC viability or cell yield in the presence of glutamine but severely affected survival in the absence of glutamine ( Figures 5A–C ). GS was strongly upregulated in DCs cultured without glutamine, notably on protein but not on mRNA level ( Figures 5D, E ). DCs metabolized glutamate to glutamine, which was completely blocked by MSO ( Figure 5F ). MSO also impaired viability, growth and respiration of THP-1 cells under glutamine-restricted culture conditions ( Figures 5G–J ). In contrast, MSO had no effect on macrophage survival ( Figure S3A–C ) in line with our finding that MSO did not completely block GS activity in macrophages. Therefore, we analyzed whether targeting of GLUL protein directly with siRNA would alter macrophage differentiation. In 4 experiments knockdown of GLUL slightly lowered the cell yield in the absence but not in the presence of glutamine ( Figure S3D ). These results demonstrate that GS is essential for myeloid cells under conditions of glutamine shortage, which might be especially important in the tumor environment.