Results

Glutamine uptake and glutamate secretion increased with glutamine concentration up to 2 mM glutamine

We first determined how HCAECs take up glutamine in both 5.5 mM (normal) and 15 mM (high) glucose by adding 0–10 mM glutamine to the cell culture media. We used low glutamine concentrations to replicate physiological circulating levels and high glutamine concentrations to simulate in vitro cell culture media. HCAECs took up significantly more glutamine as media glutamine concentration increased in both normal and high glucose, as analyzed by one-way ANOVA (p < 0.0001; Fig. 1a, d). In normal glucose, HCAECs took up 5 and 10 times more glutamine, respectively, at 5 and 10 mM glutamine than at the physiological level of 0.5 mM glutamine (p < 0.0001; Fig. 1a). Similarly, in high glucose, HCAEC took up more than 4 times more glutamine in 5 and 10 mM glutamine as compared to 0.5 mM glutamine (p < 0.001; Fig. 1d). Glutamine uptake was not impacted by osmotic effects (Online resource 3). Glutamine uptake was only statistically different between normal and high glucose at 10 mM glutamine, where glutamine uptake in high glucose was around half the uptake in normal glucose (p = 0.0280; Fig. 1g).

Fig. 1: Glutamine uptake and glutamate secretion increased with glutamine concentration; however, glutamate secretion plateaued above 2 mM glutamine. HCAEC were incubated with 0–10 mM glutamine in normal (5.5) and high glucose (15 mM) for 24 h. Glutamine uptake and glutamate secretion were measured in normal (a, b) and high (d, e) glucose using a YSI bioanalyzer. The glutamate: glutamine ratio was determined for each condition in (c) normal and (f) high glucose. g Comparison of glutamine uptake in normal vs. high glucose at 10 mM glutamine. n = 9 biological replicates. h Representative glutaminase-1 (GLS-1) and β-actin Western blots, with quantification of (i) normal glucose and (j) high glucose. n = 9 biological replicates. Data were analyzed using an ordinary one-way ANOVA with a Dunnett post-hoc multiple comparisons test

We then measured secretion of the primary glutamine-derived metabolite, glutamate, into the cell culture media. Glutamate secretion also increased with increasing media glutamine concentration in both normal and high glucose conditions (p < 0.0001 by ANOVA; Fig. 1b, e). In normal glucose, glutamate secretion was about 10 times higher for HCAEC in 0.5 mM as compared to 0 mM glutamine (p = 0.0001) and about 2 times higher for HCAEC in 2, 5, or 10 mM glutamine as compared to 0.5 mM glutamine (p = 0.0004, 0.0010, and < 0.0001, respectively). In both normal and high glucose, HCAEC glutamate secretion did not change above 2 mM glutamine. Glutamate secretion was not impacted by osmotic effects (Online resource 3). There were no statistically significant differences in glutamate secretion for HCAEC in normal and high glucose at any glutamine concentration.

We calculated the ratio of glutamate secreted to glutamine uptake as a measure of glutaminolysis. From 0.2 to 2 mM glutamine, the glutamate: glutamine ratio remained statistically similar at around 0.7 for HCAEC in normal and high glucose. At 5 and 10 mM glutamine, the glutamate: glutamine ratio decreased to around 0.2 for HCAEC in normal glucose (p < 0.0001) and around 0.3 for HCAEC in high glucose (p = 0.0534 at 5 mM and p = 0.0159 for 10 mM glutamine). There were no statistically significant differences in the glutamate: glutamine ratio for HCAEC in normal and high glucose.

Glutaminase 1 (GLS-1) catalyzes glutamine deamidation to glutamate. We therefore determined how extracellular glutamine concentration affected GLS-1 protein by Western blot (Fig. 1h, j). As we did not observe significant changes in glutamate secretion at glutamine concentrations higher than 2 mM, we measured glutaminase protein levels at 0, 0.5 and 2 mM glutamine. HCAEC in normal glucose and 0.5 or 2 mM glutamine had twice as much GLS-1 compared to HCAEC in 0 mM glutamine (p = 0.0006 and p = 0.0015, respectively; Fig. 1i). Similarly, HCAEC in high glucose and 0.5 or 2 mM glutamine had 50% more GLS-1 compared to HCAEC in 0 mM glutamine (p = 0.0088 and p = 0.0289, respectively; Fig. 1j). GLS-1 protein was not higher at 2 mM glutamine as compared to 0.5 mM glutamine for HCAEC in normal or high glucose. These data suggest that ECs take up more glutamine as extracellular glutamine concentration increases but reach a limit of glutamate secretion, possibly due to a lack of additional GLS-1.

Since endothelial cells depend on glycolysis, we assessed glucose uptake and lactate secretion in HCAECs in increasing extracellular glutamine. Glutamine supplementation did not significantly alter glucose uptake (Online resource 1a, c). Lactate secretion increased with extracellular glutamine for HCAEC in both normal and high glucose (p = 0.0007 and p < 0.0001 by ANOVA for normal and high glucose, respectively; Online resource 1b, d). Lactate secretion at 10 mM glutamine was about 25% higher for HCAEC in normal and high glucose as compared to HCAEC in 0.5 mM glutamine. These data show that extracellular glutamine did not significantly change glycolysis, suggesting that the two metabolic pathways remain distinct.

Glutamine increased oxidative respiration and isotope enrichment through the forward TCA cycle with limited impact on reverse carboxylation

Glutamine is a primary carbon source for the endothelial TCA cycle. As we did not measure changes in glutamate secretion above 2 mM glutamine, we measured how 0, 0.5, or 2 mM extracellular glutamine impacted mitochondrial activity (Fig. 2a) using a Seahorse Mito Stress test. HCAEC oxygen consumption rate (OCR), a measure of mitochondrial oxidative respiration, increased with glutamine (p = 0.0016 for basal OCR and p < 0.0001 for all others by ANOVA). Basal OCR was five time higher for cells in 0.5 and 2 mM glutamine compared to 0 mM glutamine (p = 0.0342 and 0.0013, respectively; Fig. 2b). Maximal respiration, which measures mitochondrial ability to meet increased energy demand (Desler et al., 2012), and spare capacity, which is the difference between maximal and basal respiration, more than doubled for cells in 0.5 (p = 0.008) and 2 mM (p < 0.0001) glutamine relative to 0 mM glutamine (Online resource 2a, b). Similarly, cells in 0.5 and 2 mM glutamine had double the non-mitochondrial oxygen consumption of cells in 0 mM glutamine (p = 0.0003 and p < 0.0001, respectively; Online resource 2c), indicating increased cellular oxidase activity. Only spare respiratory capacity was statistically significantly higher for cells in 2 mM as compared to 0.5 mM glutamine (p = 0.0058). We observed a similar OCR response to glutamine in HCAECs cultured in high glucose (Online resource 2d-h). Osmolarity did not affect OCR for HCAECs in low glucose; however, osmolar effects increased OCR for HCAEC in high glucose (Online resource 4). Taken together, these data show that glutamine fuels endothelial mitochondrial activity, although the effect is limited at higher glutamine concentrations.

Fig. 2: Glutamine increased endothelial oxidative respiration and isotope enrichment through the forward TCA cycle; reverse carboxylation also increased with glutamine but was limited overall. HCAECs were incubated for 24 h with 0, 0.5, or 2 mM glutamine in normal (5.5 mM) glucose culture. a Representative oxygen consumption rate (OCR) as measured by Seahorse Mito Stress test. b Basal respiration from three experiments, normalized to the 0 mM glutamine condition. n = 10–16 biological replicates. Data were analyzed using ordinary one-way ANOVA with a Tukey post-hoc multiple comparisons test. (c) HCAECs were cultured with 0, 1, 2, or 5 mM 1-13C- or 5-13 C-glutamine for 24 h, after which cells were collected and the isotope distribution was determined using LC-MS. Isotope labeled fractions for 5-13 C-glutamine metabolites are shown in blue. Isotope labeled fractions for 1-13 C-glutamine metabolites are shown in red with diagonal hatching. 5-13 C-glutamine was used to measure glutamine flux through the forward TCA cycle, since α-KG loses its first carbon as CO2 when it converts to succinate, thereby retaining the M1 label. 1-13 C-glutamine was used to measure reverse carboxylation, since when α-KG is converted to citrate in the reverse TCA cycle, the first labeled carbon is preserved. M0 and M1 refer to the number of 13C atoms incorporated into a metabolite, where M0 has no 13C and M1 has one 13C atom. n = 3 biological replicates. Created in BioRender. Kheradmand-Hajibashi, M. (2026) https://BioRender.com/gom6hqu

Since extracellular glutamine increased endothelial oxidative respiration, we used isotope-assisted tracing to get a more detailed understanding of how glutamine contributes to TCA cycle metabolites. We conducted all labeling experiments in normal glucose, as we observed no differences in TCA cycle activity for HCAEC in normal and high glucose. We used 1 mM glutamine instead of 0.5 mm glutamine to model physiological glutamine concentration due to concerns about our ability to detect isotope labeling at the lower concentration. We used 2 mM glutamine to model a commonly used endothelial cell culture media concentration and 5 mM glutamine to observe changes with additional glutamine. We used 1-13 C-glutamine and 5-13 C-glutamine parallel labeling to understand glutamine entry into forward and reductive carboxylation. In the forward (oxidative) TCA cycle, 1-13 C-glutamine loses its labeled C-1 carbon when α-KG is converted to succinate. However, in the reductive carboxylation pathway, the C-1 carbon from 1-13 C glutamine appears in citrate. Thus, we expect a significant M1 isotopomer abundance for citrate with no significant M1 isotopomer abundances for other TCA cycle metabolites in the 1-13 C glutamine experiment if reductive carboxylation is active. In contrast, 5-13 C-glutamine retains its labeled C-5 carbon in the forward TCA cycle. The C-5 carbon from 5- 13 C glutamine appears in C-1 or C-4 position of succinate (due to molecular symmetry). It then passes to the C-1 or C-4 position of fumarate, malate and oxaloacetate, finally ending up in the C-1 or C-6 position of citrate. In the next turn through the forward TCA cycle, the labeled carbon is lost as CO2. Therefore, we expect substantial M1 isotopomer abundances for all TCA cycle metabolites in the 5-13 C glutamine labeling experiment. Since the cells are likely to concurrently operate both the forward TCA cycle and reductive carboxylation, we expect to observe a superimposition of the labeling scenarios described above.

Our natural abundance corrected LC-MS isotope enrichment data (Fig. 2c) showed that nearly all intracellular glutamine was labeled in HCAEC treated with 1, 2, or 5 mM 1-13C-glutamine or 5-13 C-glutamine. 5-13 C-glutamate enrichment increased from 69% to 76%, and 5-13 C-α-KG enrichment increased from 64% to 71% with increasing extracellular glutamine. Labeled fraction of TCA metabolites in the forward TCA cycle also increased in a concentration dependent manner as extracellular glutamine increased from 1 to 5 mM, including succinate from 4% to 12%, fumarate from 31% to 50%, malate from 40% to 55%, and citrate from 39% to 49%. However, 1-13 C-citrate enrichment was less than 5% across all glutamine concentrations, indicating limited reductive carboxylation. Interestingly, succinate had the highest unlabeled pool among all TCA cycle metabolites, which could indicate that unlabeled material present before the labeling period contained substantial succinate, or that the cells contain a stable intracellular succinate pool or generate succinate from other carbon sources.

Glutamine increased the total abundance of all TCA metabolites except for succinate, which decreased

Next, we analyzed the total abundance of glutamine, glutamate, and TCA metabolites by determining the integrated peak area, which is proportional to the concentration. For HCAEC in 2 or 5 mM glutamine, intracellular glutamine total abundance was 2 and 5 times higher than for HCAEC in 1 mM glutamine (p = 0.0056 and p < 0.0001, respectively; Fig. 3a). Intracellular glutamate total abundance was 20 times higher for HCAEC cultured in 1 mM glutamine as compared to 0 mM glutamine (p < 0.0001) and then an additional 50% higher for HCAEC in 5 mM glutamine as compared to 1 mM glutamine (p < 0.0001; Fig. 3b). The glutamate: glutamine total abundance ratio for HCAEC in 2 and 5 mM glutamine declined by 50% and 75%, respectively, as compared to the ratio for HCAEC in 1 mM glutamine (p = 0.0003, and < 0.0001, respectively; Fig. 3c). These total abundance data confirm that the endothelial cells continued to take up more glutamine as extracellular glutamine concentration increased but did not produce more glutamate, as previously shown (Fig. 1).

Fig. 3: Increasing glutamine increased total abundance of all TCA metabolites except for succinate which decreased. HCAECs were treated with 0, 1, 2, or 5 mM of 1-13 C- or 5-13 C-glutamine for 24 h and then analyzed by LC-MS. a-c Total abundance of L-glutamine, L-glutamate and glutamate: glutamine ratio. Total abundance of TCA cycle metabolites (d) α-KG, e fumarate, f citrate, g malate, and h succinate. n = 6 samples for 1, 2, and 5 mM glutamine and n = 3 for 0 mM glutamine. All data points were normalized to the average value at 0 mM glutamine. Data were analyzed using ordinary one-way ANOVA with a Dunnett post-hoc multiple comparisons test. i Succinate was measured in media by LC-MS after 24 h HCAEC incubation with 0 or 2 mM glutamine. n = 5 biological replicates. j Representative curve and summation of (k) succinate dehydrogenase (SDH) activity for HCAEC cultured in 0 and 5 mM glutamine for 24 h. All data points were normalized to average value at 0 mM glutamine. n = 10 biological replicates. Data were analyzed using Mann Whitney test

Similar to the labeled fraction data, TCA metabolite total abundance increased in the presence of extracellular glutamine, except for succinate, which decreased. α-KG, fumarate, citrate, and malate total abundance increased between 3 and 5 times for HCAEC cultured in 1 mM as compared to 0 mM glutamine (Fig. 3d-g). While α-KG and citrate total abundance did not change at higher glutamine concentrations, malate and fumarate demonstrated small increases with glutamine concentration. Malate total abundance was 35% higher at 2 mM glutamine (p = 0.0024) and 46% higher at 5 mM glutamine (p = 0.0001) as compared to 1 mM glutamine, and fumarate was 29% higher at 2 mM glutamine (p = 0.0426) and 51% higher at 5 mM glutamine (p = 0.0007) as compared to 1 mM glutamine. In contrast to the other TCA metabolites, succinate total abundance decreased by 34%, 44%, and 56% in HCAEC cultured in 1, 2, and 5 mM glutamine compared to 0 mM glutamine (p = 0.0078, 0.0002, and < 0.0001, respectively; Fig. 3h).

We then analyzed several mechanisms by which succinate total abundance could decrease with increasing glutamine. Cells can transport succinate out of the cell via MCT1 and OAT transporters (Bisbach et al., 2022; Huang et al., 2024). However, when we measured succinate in the media by LC-MS after 24 h of HCAEC incubation with 0 or 2 mM glutamine, we did not observe an increase in extracellular succinate (Fig. 3i). Glutamine deprivation has also been shown to cause deSUMOylation of succinate dehydrogenase (SDH), leading to reduced SDH activity and therefore reduced succinate oxidation to fumarate (Xia et al., 2021; Liu et al., 2023). We therefore measured how increasing extracellular glutamine affected SDH activity. SDH activity was 19% higher in HCAEC in 5 mM compared to 0 mM glutamine (p = 0.0117; representative experiment in Fig. 3j, three compiled experiments in Fig. 3k). Thus, increased SDH activity could be one means by which elevated extracellular glutamine reduced intracellular succinate.

Glutamine enriched amino acids and glutathione in addition to TCA metabolites

Since our data showed that ECs took up more glutamine as extracellular concentration increased but did not secrete more glutamate or proportionally increase TCA metabolite labeling, we investigated where the extra glutamine was metabolized using our LC-MS data. A principal component analysis (PCA) score plot of the 5-13 C-glutamine metabolite labeled fractions revealed clear separation of samples along component 1 (PC1 = 51.6%), with the 0 mM glutamine condition clustering distinctly from glutamine-treated groups (Fig. 4a). To identify the metabolites driving the separation observed in the PCA score plot, we next examined the PCA loading plot (Fig. 4b). In addition to glutamine, glutamate, and TCA cycle metabolites, the amino acids L-proline, L-aspartate, and GABA along with the tripeptide glutathione most contributed to the separation of glutamine-treated cells (Fig. 4b). A heatmap of the glutamine-labeled fractions of metabolites that most strongly contributed to the separation further indicated that HCAEC treated with 1, 2, or 5 mM glutamine are metabolically distinct from HCAEC treated with 0 mM glutamine but not highly different from each other (Fig. 4c). The PCA score plot of the 1-13 C-glutamine metabolite labeled fractions also showed that the 0 mM glutamine condition clustered distinctly from glutamine-treated groups (Online resource 5a), and the metabolites identified using the PCA loading plot were similar to those from the 5-13 C-glutamine metabolite (Online resource 5b, c).

Fig. 4: Glutamine carbons were incorporated into amino acids and glutathione in addition to TCA metabolites. HCAECs were cultured with 0, 1, 2, or 5 mM 5-13C-glutamine for 24 h, after which metabolites were extracted and profiled by LC–MS. a PCA score plot of the labeled fraction of each metabolite, showing a separation between cells cultured with versus without glutamine but minimal differences with increasing glutamine concentration. b PCA loading plot of the labeled fraction, with the top 11 loadings labeled. c Heatmap of metabolite labeled fractions

While labeled fraction analysis provides information on isotopologue distribution, it does not capture changes in the overall metabolite pool size. To address this, we performed PCA on total metabolite abundance. The PCA score plot revealed separation along PC1 between the 0 mM glutamine condition and glutamine treated samples (Online resource 6). Notably, the 1 mM glutamine condition formed a distinct cluster from the 5 mM glutamine condition along PC1, indicating that although labeled fraction analysis did not show major differences among glutamine concentrations, total metabolite abundance was sensitive to increased glutamine. These results suggest that glutamine concentration influenced metabolic pool sizes even when isotopologue distributions appeared similar.

Glutathione and proline were enriched with glutamine-derived carbons and increased in total abundance with glutamine; however, arginine and citrulline total abundance decreased with increasing glutamine

HCAEC cultured with glutamine separated from HCAEC cultured without glutamine due to glutamine-derived glutamate contributing carbon to both glutathione and proline. Glutathione isotopic enrichment from 1-13C-glutamine and 5-13C-glutamine increased from no detectable enrichment at 0 mM glutamine to 63% enrichment at 1 mM glutamine, respectively (Fig. 5a). Glutathione isotopic enrichment increased by an additional ~ 5% at 2 and 5 mM glutamine. Total glutathione abundance increased by more than 6 times when glutamine was increased from 0 mM to 1, 2, or 5 mM (p < 0.0001; Fig. 5f); however, no differences in total glutathione abundance were observed among HCAEC cultured in 1, 2, or 5 mM glutamine.

Fig. 5: Glutathione and proline were enriched with glutamine-derived carbons. HCAECs were cultured with 0, 1, 2, or 5 mM 1-13C- or 5-13 C-glutamine for 24 h, after which cells were collected and the isotope distribution and total abundance of metabolites were determined using LC-MS. a-e Glutathione, proline, ornithine, citrulline, and arginine isotope labeled fractions. n = 6 biological replicates. Quantification of total abundance of f-j proline, glutathione, ornithine, citrulline, and arginine. n = 6 biologic replicates for 1,2,and 5 mM glutamine and n = 3 biologic replicates for 0 mM glutamine. Data were analyzed using ordinary one-way ANOVA with a Tukey post-hoc multiple comparisons test. Created in BioRender. Kheradmand-Hajibashi, M. (2026) https://BioRender.com/pb4r3y6

Glutamate can also be metabolized to pyrroline-5-carboxylate (P5C), which can then be directed toward either proline synthesis or the ornithine cycle. Proline isotopic enrichment increased by 36%, 43%, and 48% at 1, 2, and 5 mM 1-13C-glutamine and 5-13C-glutamine, respectively, compared to 0 mM glutamine (Fig. 5b). Proline total abundance doubled in HCAEC cultured in 1 mM glutamine as compared to 0 mM glutamine (Fig. 6g; p = 0.0002). Similar to glutathione, no difference in proline total abundance was observed in HCAEC cultured in 1, 2, and 5 mM glutamine. In contrast, there was no detectable carbon enrichment in ornithine cycle metabolites (Fig. 5c-e). However, total metabolite abundance in this pathway was altered by glutamine (Fig. 5h-j). Increasing extracellular glutamine from 0 to 2 or 5 mM decreased citrulline total abundance by 47% (p = 0.0153) and 53% (p = 0.0039), respectively. Similarly, increasing glutamine decreased arginine total abundance by 39% at 2 mM glutamine (p = 0.0161) and 46% at 5 mM glutamine (p = 0.0050) compared to 0 mM glutamine. Ornithine total abundance remained unchanged across glutamine concentrations. Together, the isotopic enrichment data suggest that glutamine contributes carbon to glutathione and proline but not to ornithine cycle metabolites. Reduced arginine and citrulline abundance suggest that glutamine altered their utilization.

Fig. 6: Glutamine-derived aspartate contributed carbons to UDP-GlcNAc, and glutamine altered fatty acids and one-carbon metabolism. HCAECs were treated with 0, 1, 2, and 5 mM 1-13C- or 5-13 C-glutamine for 24 h, after which cells were collected, and the isotope enrichment and total metabolite abundance was determined using LC-MS. Schematic of glutamine-derived aspartate incorporation into UDP-GlcNAc. Isotope enrichment from 5-13 C-glutamine into (a) aspartate, and (c) UDP-GlcNac. n = 3 biological replicates. Quantification of total abundance of (b) aspartate, and (d) UDP-GlcNAc, (e) eicosapentaenoic acid (EPA) and icosatrienoic acid and (f) one carbon metabolites folate and histidine. n = 6 biologic replicates for 1, 2, and 5 mM glutamine and n = 3 biologic replicates for 0 mM glutamine. Data were analyzed using ordinary one-way ANOVA with a Tukey post-hoc multiple comparisons test. Created in BioRender. Kheradmand-Hajibashi, M. (2026) https://BioRender.com/xjjzmwd

Glutamine contributed carbons to UDP-GlcNAc via aspartate and altered fatty acids and one-carbon metabolism

Since we also observed aspartate isotope enrichment, we next examined how increasing extracellular glutamine concentration impacted pathways downstream of aspartate such as UDP-GlcNAc synthesis. Glutamine-derived α-ketoglutarate becomes oxaloacetate, which is then transaminated to form aspartate. Aspartate is incorporated into orotate, which becomes uridine triphosphate (UTP) and is then incorporated into UDP-GlcNAc (Fig. 6). LC-MS analysis showed increased isotopic enrichment of aspartate from 5-13C-glutamine, with enrichment increasing by 45%, 56%, and 60% at 1, 2, and 5 mM glutamine, respectively, compared to no glutamine (Fig. 6a). Aspartate total abundance increased by 33-, 44-, and 70-fold as glutamine increased from 0 mM to 1, 2, and 5 mM, respectively (p = 0.0003, < 0.0001; Fig. 6b). Isotopic distribution analysis from 5-13C-glutamine revealed a progressive increase in labeled carbon incorporation into UDP-GlcNAc, with enrichment increasing by 7%, 9%, and 16% at 1, 2, and 5 mM glutamine, respectively (Fig. 6c). In parallel, total UDP-GlcNAc abundance increased by around 50% as glutamine concentration increased from 0 to 2 or 5 mM glutamine (p = 0.0276, and 0.0070, respectively; Fig. 6d), with no differences observed between 2 and 5 mM glutamine. Collectively, the isotope enrichment data suggests that increasing glutamine increases its incorporation into UDP-GlcNAc through increased aspartate availability.

We observed that the total abundance of unsaturated fatty acids eicosapentaenoic acid (EPA) and icosatrienoic acid decreased with increasing glutamine concentration. EPA abundance decreased by 70% for HCAEC in 1 mM glutamine as compared to 0 mM glutamine (p < 0.0001, Fig. 6e) and decreased an additional 12% for HCAEC in 5 mM glutamine (p = 0.0101). Similarly, icosatrienoic acid abundance decreased by 48% for HCAEC in 1 mM glutamine as compared to 0 mM glutamine (p = 0.0034, Fig. 6e), with no additional decrease at higher glutamine concentrations.

We finally observed significant alterations in the total abundance of metabolites associated with 1 C metabolism, a process that supports purine biosynthesis, DNA and protein methylation, and redox homeostasis among others (Ducker & Rabinowitz, 2017; Hwang et al., 2023). In the folate cycle, which supports 1 C metabolism, the biologically active form of folate (tetrahydrofolate, THF) and its reduced form (5-methyl-THF) serve as carriers for 1 C units. New 1 C units enter the folate cycle from the amino acids serine, glycine, and histidine among others. In our LC-MS data, increasing glutamine concentration from 0 to 2 mM decreased total folate abundance by 25% (p = 0.0078), while increasing glutamine concentration from 0 to 5 mM decreased total folate abundance by 60% (p = 0.0001, Fig. 6f). Similarly, increasing glutamine concentration from 0 to 1, 2, or 5 mM decreased total histidine abundance by 51%, 61%, or 77% (p < 0.0001 for all). Both glycine and serine total abundance showed smaller changes with increasing glutamine, with glycine 38% lower at 5 mM glutamine compared to 1 mM glutamine (p = 0.01870, online resource. 7). These total abundance data suggest that glutamine altered 1 C metabolism.