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. 2025 Sep 24;11(5):e70601. doi: 10.1002/vms3.70601

Glutaminase Expression in Canine Large‐Cell Alimentary Lymphoma Cells and Effects of Glutaminase Inhibition by CB‐839

Kosei Sakai 1,✉, Masaki Hirao 2, Satoshi Kameshima 1, Yasuhiko Okamura 3, Takuya Mizuno 4, Shunsuke Shimamura 2
PMCID: PMC12457854  PMID: 40988645

ABSTRACT

Glutamine metabolism plays a crucial role in tumour progression, making glutaminase a promising therapeutic target in various human cancers. However, its role in canine large‐cell alimentary lymphoma (AL) remains unclear. This study investigated glutaminase expression and the effects of a glutaminase inhibitor (CB‐839) on canine large‐cell AL cell lines. Western blotting analysed glutaminase expression in three canine large‐cell AL cell lines (CLC, Ema and Nody‐1) and peripheral blood mononuclear cells (PBMCs) isolated from eight clinically healthy dogs. Cell viability was determined in each cell line after treatment with varying concentrations (0–10 µM) of CB‐839. Flow cytometry was used to analyse the cell cycle and assess annexin assays in each cell line following treatment with 1 µM of CB‐839 or a vehicle control. Additionally, metabolome analysis was performed in Nody‐1 cells after treatment with 1 µM of CB‐839 or a vehicle control. Glutaminase expression was significantly higher in cell lines than in PBMCs. CB‐839 suppressed cell proliferation in a dose‐dependent manner, with CLC and Nody‐1 cells exhibiting greater susceptibility than Ema cells. Flow cytometric analysis revealed that CB‐839 induced G0/G1 phase arrest and apoptosis in susceptible cell lines. Metabolomic analysis revealed that CB‐839 led to glutamine accumulation and depletion of key tricarboxylic acid cycle intermediates in Nody‐1 cells. These findings indicate that glutamine metabolism is upregulated in canine large‐cell AL and plays a crucial role in tumour cell growth and survival. Inhibiting glutaminase could serve as a promising therapeutic strategy for this disease.

Keywords: alimentary lymphoma, CB‐839, dog, glutaminase, glutamine, tumour metabolism

Glutamine metabolism is upregulated in canine large‐cell alimentary lymphoma cells and plays a crucial role in tumour cell growth and survival. A glutaminase inhibitor led to metabolic disruption via glutamine accumulation and tricarboxylic acid cycle intermediate depletion in the cells, possibly serving as a promising therapeutic strategy for this disease.

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1. Introduction

Lymphoma is one of the most frequently diagnosed neoplasms in dogs, accounting for approximately 7%–24% of all canine neoplasms and 83% of all canine haematopoietic malignancies (Vail et al. 2013). Canine lymphoma manifests in various anatomical forms, including multicentric, mediastinal, abdominal (alimentary, hepatic, splenic and renal), cutaneous, ocular, central nervous system and pulmonary lymphomas. Alimentary lymphoma (AL) primarily infiltrates the gastrointestinal tract and mesenteric lymph nodes with neoplastic lymphocytes, comprising approximately 5%–7% of all canine lymphomas (Munday et al. 2017; Vail et al. 2020). Based on their cell morphologies, canine AL can be classified into two major types: large‐cell or small‐cell. Although medical therapy remains the mainstay treatment for both, large‐cell AL is particularly aggressive and highly resistant to treatment, resulting in a poor prognosis with a median survival time of only a few months (Lai et al. 2024; Nakagawa et al. 2022; Rassnick et al. 2009; Sogame et al. 2018). Therefore, there is an urgent need to better understand the pathogenesis of canine large‐cell AL and to identify novel therapeutic targets.

Our previous study demonstrated that plasma glutamine concentrations were lower in dogs with AL (predominantly large‐cell type) than in those with chronic enteropathy (Sakai et al. 2023). Additionally, another study reported that dogs with chronic enteropathy exhibited lower plasma glutamine concentrations than clinically healthy dogs (Tamura et al. 2019). These findings suggest that the plasma glutamine concentrations are lower in dogs with AL than in both clinically healthy dogs and those with chronic enteropathy. However, this remains a largely phenomenological observation, and the underlying mechanisms of decreased plasma glutamine concentrations in dogs with AL have yet to be elucidated.

Glutamine is a nonessential/conditionally essential amino acid that plays a pivotal role in clinical illness and stress conditions (Jin et al. 2023; Puurunen et al. 2018). The upregulation of glutamine metabolism has recently been recognised as a critical feature of human cancer cells, increasing their demand for glutamine to support proliferation and survival (Anthony et al. 2024). This metabolic alteration can be linked to decreased blood glutamine concentrations in human cancer patients (Muranaka et al. 2024). A key enzyme in glutamine metabolism is glutaminase, which catalyses the conversion of glutamine to glutamic acid and ammonia. This process plays a pivotal role in fuelling the tricarboxylic acid (TCA) cycle and supporting the biosynthetic pathways essential for tumour growth, including nucleotide, amino acid and lipid synthesis. There are two forms of glutaminase: glutaminase 1 (GLS1) and glutaminase 2 (GLS2). GLS1 is overexpressed in various human cancers, and this phenotype is associated with a worse disease stage and poor prognosis (Lee et al. 2016; Xiang et al. 2019). The heightened activity of GLS1 in cancer cells is considered a hallmark of cancer metabolism and has been proposed as a therapeutic target (Anthony et al. 2024; Jin et al. 2023). Given these findings, the decreased plasma glutamine concentrations in dogs with AL may be associated with an increased demand for glutamine in cancer cells; however, no data currently support this hypothesis. This study aimed to elucidate the role of glutamine metabolism in the pathogenesis of canine large‐cell AL, with a special focus on GLS1, to provide novel insights into potential therapeutic interventions.

2. Materials and Methods

2.1. Reagents and Antibodies

Telaglenastat (CB‐839), which inhibits GLS1 but not GLS2, was obtained from Selleck Chemicals (Houston, TX, USA). This agent was dissolved in dimethyl sulfoxide.

For western blotting, rabbit polyclonal antibodies against GLS1, including splice variants of kidney‐type glutaminase (KGA) and glutaminase C (GAC), were procured from Proteintech Group, Inc. (Munich, Germany). Additionally, a mouse monoclonal antibody against β‐actin was acquired from Sigma‐Aldrich (Saint Louis, MO, USA). The antibody against β‐actin was validated for use in dogs based on the manufacturer's specifications. Horseradish peroxidase‐conjugated goat anti‐rabbit and anti‐mouse immunoglobulin G secondary antibodies were purchased from Bio‐Rad Laboratories, Inc. (Hercules, CA, USA). All the antibodies were diluted according to the manufacturer's recommendations.

2.2. Cell Line Validation and Culture Conditions

Three canine large‐cell AL cell lines (CLC, Ema and Nody‐1) established in our previous study were used (Umeki et al. 2013). Mycoplasma contamination in these cell lines tested negative using the Takara PCR Mycoplasma Detection Set (Takara Bio Inc., Kusatsu, Japan). The cells were maintained in RPMI 1640 medium (FUJIFILM Wako Pure Chemical Corporation, Tokyo, Japan) supplemented with 10% foetal bovine serum (Gibco, Carlsbad, CA, USA) at 37°C with 5% CO2. The human lung cancer cell line (A549) was purchased from RIKEN BRC (Tsukuba, Japan). The cells were maintained in DMEM (Gibco) supplemented with 10% foetal bovine serum at 37°C with 5% CO2.

2.3. Western Blotting

CLC, Ema and Nody‐1 cells were cultured in petri dishes. When the cells reached 70%–80% confluence, they were washed with Hanks' balanced salt solution (HBSS; Sigma‐Aldrich) and incubated with RIPA buffer (Cell Signalling, Danvers, MA, USA) containing protease and phosphatase inhibitor cocktail tablets (Roche Diagnostics, Mannheim, Germany) on ice for 5 min. The cell lysates were centrifuged at 14,000 rpm at 4°C for 10 min, and the supernatants were collected.

Whole blood samples were collected from eight clinically healthy dogs, and peripheral blood mononuclear cells (PBMCs) were isolated based on a previous study (Goto‐Koshino et al. 2011). Proteins were extracted from PBMCs using the same method described above. All experimental procedures involving dogs were approved by the Animal Care Committee of Kitasato University (authorisation number: 24–046).

Total protein concentration in all samples was measured using a BCA Protein Assay kit (Thermo Fisher Scientific, Waltham, MA, USA). Equal sample amounts (12 µg) were loaded onto 10% Mini‐PROTEAN TGX Precast Protein Gels (Bio‐Rad Laboratories Inc.) for electrophoresis. After separation, proteins were transferred to Trans‐Blot Turbo Mini PVDF (Bio‐Rad Laboratories Inc.) and were blocked in EzBlock Chemi (Atto Corp., Tokyo, Japan) at 25°C for 1 h. Primary antibodies were incubated at 4°C for 12 h. Subsequently, the membranes were washed using Tris‐buffered saline with 0.2% Tween 20 (Sigma‐Aldrich) and were incubated with the appropriate secondary antibodies at 23°C for 1 h. Proteins were visualised with Clarity Western ECL Substrate (Bio‐Rad Laboratories Inc.), and the signals were detected using the Mi‐II 600CB (BioTools, Gunma, Japan). β‐actin expression was used as a protein loading control, and the A549 cell lysate served as a positive control. Using an image analysis software (ImageJ, U.S. National Institutes of Health, Bethesda, MD, USA) (Schneider et al. 2012), the density of each band was quantified by K.S.

2.4. Water‐Soluble Tetrazolium Salt Assay

Cell viability was determined utilising a Cell Counting Kit‐8 (Dojindo Laboratories, Kumamoto, Japan) according to the manufacturer's instructions. Briefly, CLC, Ema and Nody‐1 cells were seeded in 96‐well plates with 2.5 × 103, 2.5 × 104 or 1.4 × 104 cells per well, respectively. Cells were treated with varying concentrations (0–10 µM) of CB‐839 for 72 h, followed by the addition of Cell Counting Kit‐8 to each well. The survival fraction was determined based on the absorbance at 450 nm using a Multiskan FC (Thermo Fisher Scientific, MA, USA). All samples were examined in quintuplicate, and three independent experiments were performed.

2.5. Cell Cycle Analysis

CLC, Ema and Nody‐1 cells were seeded in six‐well plates with 1.5 × 105 cells per well for CLC or 2.0 × 105 cells per well for Ema and Nody‐1. The cells were treated with 1 µM of CB‐839 or a vehicle control for 48 h. Subsequently, the cells were washed with HBSS, centrifuged at 1000 rpm for 5 min, fixed in 75% alcohol, and maintained at −30°C until the analysis. The samples were then washed with HBSS and incubated with a staining solution containing 50 µg/mL propidium iodide (PI, Sigma‐Aldrich), 0.1 mg/mL RNase A (NIPPON GENE CO. LTD., Tokyo, Japan), and 0.05% Triton X‐100 (Sigma‐Aldrich) at 37°C for 40 min in the dark. The deoxyribonucleic acid content of the cells was evaluated using CytoFLEX S (Beckman Coulter, Brea, CA, USA). All samples were examined in triplicate, and three independent experiments were performed.

2.6. Apoptosis Assay

To detect apoptotic cells, the MEBCYTO Apoptosis Kit (MBL, Nagoya, Japan) was used according to the manufacturer's instructions. Briefly, CLC, Ema and Nody‐1 cells were seeded in six‐well plates with 1.5 × 105 cells per well for CLC or 2.0 × 105 cells per well for Ema and Nody‐1. The cells were treated with 1 µM of CB‐839 or a vehicle control for 48 h. Subsequently, the cells were washed with HBSS and centrifuged at 1000 rpm for 5 min. The samples were then washed with HBSS and incubated with a staining solution containing FITC‐labelled annexin V (ANXV) and PI at 23°C for 15 min in the dark. The events of early apoptosis (ANXV‐positive/PI‐negative), late apoptosis (double‐positive) and live cells (double‐negative) were counted employing CytoFLEX S. All samples were examined in triplicate, and three independent experiments were performed.

2.7. Metabolome Analysis

Metabolite extraction was performed according to the manual of Human Metabolome Technologies (HMT; Tsuruoka, Japan). Briefly, approximately 1.0 × 107 Nody‐1 cells after treatment with CB‐839 (1 µM) or a vehicle control for 24 h were collected and washed with 10 mL of 5% (w/w) mannitol solution. The cells were then treated with 800 µL of methanol and vortexed for 30 s. Subsequently, the cell extract was treated with 550 µL of ultrapure water containing 10 µM of internal standard solution 1 (HMT) and vortexed for 30 s. The extract was centrifuged at 2300 × g at 4°C for 5 min, and 350 µL of the supernatant was centrifugally filtered through a 5‐kDa cutoff filter (ULTRAFREE MC PLHCC, HMT) (9100 × g, 4°C, 4 h). The filtrate was maintained at −80°C until the analysis. All samples were examined in triplicate.

Metabolome analysis was conducted using the C‐SCOPE package of HMT with capillary electrophoresis time‐of‐flight mass spectrometry for cation analysis and capillary electrophoresis‐tandem mass spectrometry for anion analysis. Intracellular levels of 116 metabolites (52 cations and 64 anions) that play major roles in glycolysis, the pentose phosphate pathway, the citric acid cycle, the urea cycle, the polyamine creatine metabolism pathway, the purine metabolism pathway, the glutathione degradation pathway, the nicotinamide biosynthetic pathway, the choline metabolism pathway and various amino acid metabolism pathways were analysed (Table S1).

2.8. Statistical Analysis

Welch's t‐test was used for comparison between two groups. Prism software version 8.4.2 (GraphPad Software, Boston, MA, USA) was utilised for statistical analysis. Principal component analysis and hierarchical cluster analysis were performed using software developed by HMT. Statistical significance was set at p < 0.05.

3. Results

3.1. Glutaminase Protein Expression

Western blot analysis revealed distinct bands corresponding to two glutaminase isoforms (KGA and GAC) in the positive control, A549 (Figure 1A). Similar bands were observed at approximately the same positions in the canine samples. KGA expression tended to be higher in canine large‐cell AL cell lines than in PBMCs isolated from clinically healthy dogs (Figure 1B). Additionally, the expression levels of GAC and the sum of KGA and GAC were significantly higher in the cell lines than in PBMCs (Figure 1C,D). In the canine large‐cell AL cell lines, the band density of KGA was more distinct in CLC and Nody‐1 cells than in Ema cells, whereas no major differences were observed in the band density of GAC among the cell lines.

FIGURE 1.

FIGURE 1

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(A) Western blot analysis of kidney‐type glutaminase (KGA) and glutaminase C (GAC) in three canine large‐cell alimentary lymphoma cell lines (CLC, Ema and Nody‐1) and peripheral blood mononuclear cells (PBMCs) isolated from eight clinically healthy dogs. The human lung cancer cell line (A549) was used as a positive control. Expression of β‐actin was used as a protein loading control. (B–D) Comparison of the expression levels of KGA and GAC and their sum between canine large‐cell AL cell lines and PBMCs. The expression levels of KGA and GAC are normalised to β‐actin in each sample. Data are presented as relative values when the mean protein level in PBMCs was set to 1.00.

3.2. Effects of CB‐839 on Cell Proliferation

Cell viability of all canine large‐cell AL cell lines reduced in a CB‐839 dose‐dependent manner compared to the vehicle control (Figure 2). The 50% inhibitory concentrations of CB‐839 in CLC, Ema and Nody‐1 were 2.5 × 10−1 µM, > 10 µM and 3.0 × 10−2 µM, respectively.

FIGURE 2.

FIGURE 2

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Cell viabilities of CLC, Ema and Nody‐1 cells after treatment with various concentrations (0–10 µM) of CB‐839 for 72 h. The experiment was performed in quadruplicate. Data are presented as mean ± standard error.

3.3. Effects of CB‐839 on Cell Cycle and Cell Death

As shown in Figure 3, cell cycle analysis using flow cytometry revealed a significant increase in the G0/G1 phase fraction in CLC and Nody‐1 cells after treatment with 1 µM of CB‐839, compared to the vehicle control, while the S and G2/M phase fractions were significantly reduced. There were no significant differences in the sub‐G1 phase fraction between the two groups. In contrast, the G0/G1 phase fraction tended to increase in Ema cells after treatment with 1 µM of CB‐839 compared to those in the vehicle control. However, no significant differences were observed in any phase fraction between the two groups.

FIGURE 3.

FIGURE 3

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(A) Representative data on cell cycle analysis of CLC, Ema and Nody‐1 cells after treatment with CB‐839 (1 µM) or the vehicle control for 48 h. (B) Comparing the percentages of each phase fraction in CLC, Ema and Nody‐1 cells after treatment with CB‐839 or the vehicle control. This experiment was performed in triplicate. Data are presented as mean ± standard error.

As shown in Figure 4, the apoptosis assay showed that the percentage of live cells significantly reduced in CLC and Nody‐1 cells following treatment with 1 µM of CB‐839 compared to those in the vehicle control, while the proportion of late apoptotic cells significantly increased. There were no significant differences in the percentage of early apoptotic cells between the two groups. In contrast, no significant differences in the percentages of any condition were observed between Ema cells treated with 1 µM of CB‐839 and those treated with the vehicle control.

FIGURE 4.

FIGURE 4

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(A) Representative flow cytometry data for the classification of CLC, Ema and Nody‐1 cells after staining with annexin V (ANXV) and propidium iodide (PI). The cells were treated with CB‐839 (1 µM) or the vehicle control for 48 h and then classified as ANXV‐negative/PI‐negative (live cells), ANXV‐positive/PI‐positive (early apoptotic cells) or ANXV‐positive/PI‐positive (late apoptotic cells) using flow cytometry. (B) Comparing the percentages of live, early apoptotic and late apoptotic CLC, Ema and Nody‐1 cells after treatment with CB‐839 or the vehicle control. This experiment was performed in triplicate. Data are presented as mean ± standard error.

3.4. Effects of CB‐839 on Intracellular Metabolite Concentrations

Principal component analysis plots showed clear separation between CB‐839–treated Nody‐1 cells and those treated with the vehicle control (Figure 5). Hierarchical clustering of differentially altered metabolites also demonstrated clear separation between the two groups (Figure 6). Focusing on individual metabolites, intracellular glutamine concentrations were significantly increased in Nody‐1 cells after treatment with 1 µM of CB‐839 compared with the vehicle control, while intracellular glutamic acid concentrations and glutamic acid/glutamine ratios were significantly reduced (Figure 7). Among the TCA metabolites, intracellular concentrations of succinic acid, fumaric acid, malic acid, citric acid and cis‐aconitic acid were significantly reduced in CB‐839–treated Nody‐1 cells compared to those in the vehicle control, while isocitric acid levels showed a reducing trend (Figure 8). No significant differences in the intracellular 2‐oxoglutaric acid concentrations were observed between the two groups.

FIGURE 5.

FIGURE 5

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Principal component analysis plots of Nody‐1 cells after treatment with 1 µM of CB‐839 (black) or the vehicle control (white). This experiment was performed in triplicate. NAD+, oxidised nicotinamide adenine dinucleotide; NADH, reduced nicotinamide adenine dinucleotide; NADP+, oxidised nicotinamide adenine dinucleotide phosphate; NADPH, reduced nicotinamide adenine dinucleotide phosphate.

FIGURE 6.

FIGURE 6

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Hierarchical clustering of 89 differentially altered metabolites in Nody‐1 cells after treatment with 1 µM of CB‐839 (black) and those with the vehicle control (white). This experiment was performed in triplicate. Metabolites indicated in green are downregulated, whereas metabolites indicated in red are upregulated.

FIGURE 7.

FIGURE 7

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Intracellular concentrations of glutamine and glutamic acid and glutamic acid/glutamine ratios in Nody‐1 cells after treatment with CB‐839 (1 µM) or the vehicle control for 24 h. This experiment was performed in triplicate. Data are presented as mean ± standard error.

FIGURE 8.

FIGURE 8

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Intracellular concentrations of tricarboxylic acid cycle metabolites in Nody‐1 cells after treatment with CB‐839 (1 µM) or the vehicle control for 24 h. This experiment was performed in triplicate. Data are presented as mean ± standard error.

4. Discussion

In this study, the expression levels of GLS1, including its splice variants KGA and GAC, were higher in canine large‐cell AL cell lines than in PBMCs isolated from clinically healthy dogs. These findings suggest that GLS1 expression is upregulated in canine large‐cell AL. A similar result has been observed in human diffuse large B‐cell lymphoma (DLBCL), where human DLBCL cell lines showed high GLS1 expression compared to human primary B cells (Solsona et al. 2023). To date, the mechanisms underlying GLS1 expression in human DLBCL remain unclear. A previous study reported that c‐Myc transcriptionally represses miR‐23a/b, resulting in greater expression of mitochondrial GLS1 in a human Burkitt lymphoma cell line, P‐493‐6 (Gao et al. 2009). In contrast, another study reported no correlation between c‐Myc and GLS1 expression status in 18 human DLBCL cell lines (Solsona et al. 2023). Additionally, reduction of c‐Myc expression using an inhibitor of bromodomain and extra‐terminal proteins did not alter GLS1 protein levels in various human DLBCL cell lines (Solsona et al. 2023). Therefore, other transcription factors are likely to be responsible for GLS1 expression in human DLBCL. Further studies are needed to clarify the mechanisms underlying the upregulation of GLS1 expression in both human DLBLCs and canine large‐cell AL.

Metabolomic analysis revealed that CB‐839 treatment induced significant alterations in the intracellular metabolite concentrations in Nody‐1 cells. The increased intracellular glutamine concentrations, as well as reduced intracellular glutamic acid concentrations and glutamic acid/glutamine ratios, indicated effective inhibition of glutaminase activity. To our knowledge, this is the first study demonstrating the cross‐species efficacy of CB‐839 in dogs. Additionally, depletion of key TCA cycle intermediates, including succinic acid, fumaric acid, malic acid, citric acid, cis‐aconitic acid and isocitric acid, suggests impaired mitochondrial metabolism. This metabolic disruption is consistent with the findings in various human cancer cell lines, indicating that CB‐839 suppresses the enzymatic activity of GLS1, thereby limiting the influx of glutamine derivatives into the TCA cycle (Gross et al. 2014; Koch et al. 2020; Santos‐Jiménez et al. 2023; Solsona et al. 2023).

CB‐839 treatment reduced the viability of canine large‐cell AL cells in a dose‐dependent manner. Notably, CLC and Nody‐1 cells were more susceptible to CB‐839 treatment than Ema cells. This difference may be attributed to variations in glutaminase dependency, compensatory metabolic pathways such as glycolysis and fatty acid oxidation, or inherent resistance mechanisms in Ema cells. Because CLC and Nody‐1 cells showed higher KGA expression than Ema cells, the differential expression of KGA may indicate variable susceptibility to CB‐839 among different lymphoma subtypes. Future studies should focus on elucidating the molecular mechanisms underlying CB‐839 resistance in Ema cells and identifying potential combination therapies to enhance treatment efficacy. For example, endogenously produced pyruvate secreted by human breast cancer cell lines acts in a paracrine manner to decrease the susceptibility of recipient cells to CB‐839, and suppression of pyruvate secretion using a monocarboxylate transporter 1 inhibitor antagonises this paracrine effect and increases CB‐839 activity (Singleton et al. 2020). Additionally, a recent study reported that the inhibitory effects of combining CB‐839 with metformin, the first‐line medication for the treatment of Type 2 diabetes, in a human KRAS‐mutant ovarian cancer NOD‐SCID mouse model were stronger than those in the drug‐alone group (Wu et al. 2025).

Flow cytometric analysis revealed increased percentages of the G0/G1 phase fraction and late apoptotic cells in CLC and Nody‐1 cells, which showed high susceptibility to CB‐839. This indicates that CB‐839 suppresses cell proliferation via cell cycle arrest and apoptosis in canine large‐cell AL cell lines. The mechanism underlying the anti‐tumour effects of CB‐839 varies from that reported previously. In human glioma cell lines, CB‐839 causes G0/G1 phase arrest without inducing apoptosis (Koch et al. 2020). In contrast, CB‐839 induces apoptosis but not cell cycle arrest in human lung cancer cell lines (Jamshidi‐Parsian et al. 2024). In human DLBCL cell lines, CB‐839 induces both G0/G1 phase arrest and apoptosis (Solsona et al. 2023).

Canine large‐cell AL shares features including cell morphology, immunophenotype, lesion localisation, biological behaviour and prognosis with human enteropathy‐associated T‐cell lymphoma (EATL) (Matsumoto et al. 2019), which is an aggressive peripheral T‐cell lymphoma. No validated and standardised treatment protocols for human EATL exist due to the rarity of this disease (Somali et al. 2021). Therefore, there is an urgent need to identify novel therapeutic strategies. Because the role of glutamine metabolism in human EATL remains unclear, this study improves our understanding of glutamine metabolism in canine large‐cell AL, with possible applications to human EATL.

In conclusion, this study provides the first evidence that GLS1 expression is upregulated in canine large‐cell AL and that glutaminase inhibition by CB‐839 disrupts glutamine metabolism and suppresses cell proliferation via cell cycle arrest and apoptosis in a subset of canine large‐cell AL cell lines. These findings suggest that glutamine metabolism plays a crucial role in tumour growth and survival in canine large‐cell AL, supporting the potential of glutamine metabolism as a novel therapeutic target for this disease. However, the variability in CB‐839 sensitivity underscores the need for further studies to identify biomarkers predictive of response and explore combination therapies for enhanced efficacy.

Author Contributions

Kosei Sakai: conceptualisation, methodology, data curation, investigation, funding acquisition, project administration, resources, writing – original draft, writing – review and editing. Masaki Hirao: investigation, writing – review and editing. Satoshi Kameshima: resources, writing – review and editing. Yasuhiko Okamura: supervision, resources, writing – review and editing. Takuya Mizuno: supervision, resources, writing – review and editing. Shunsuke Shimamura: supervision, resources, writing – review and editing.

Ethics Statement

The authors confirm that the ethical policies of the journal, as noted on the journal's author guidelines page, have been adhered to and the appropriate ethical review committee approval has been received. All experimental procedures involving dogs were approved by the Animal Care Committee of Kitasato University (authorisation number: 24–046).

Conflicts of Interest

The authors declare no conflicts of interest.

Peer Review

The peer review history for this article is available at https://www.webofscience.com/api/gateway/wos/peer‐review/10.1002/vms3.70601.

Supporting information

Table S1. One hundred sixteen metabolites were analysed in this study.

VMS3-11-e70601-s001.xlsx (10.8KB, xlsx)

Acknowledgements

This work was supported by JSPS KAKENHI Grant Number JP23K14087 and the 2023 Osaka Metropolitan University (OMU) Strategic Research Promotion Project (Young Researcher).

Sakai, K. , Hirao M., Kameshima S., Okamura Y., Mizuno T., and Shimamura S.. 2025. “Glutaminase Expression in Canine Large‐Cell Alimentary Lymphoma Cells and Effects of Glutaminase Inhibition by CB‐839.” Veterinary Medicine and Science 11, no. 5: e70601. 10.1002/vms3.70601

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

References

  1. Anthony, J. , Varalakshmi S., Sekar A. K., Devarajan N., Janakiraman B., and Peramaiyan R.. 2024. “Glutaminase—A Potential Target for Cancer Treatment.” Biomedicine 14, no. 2: 29–37. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Gao, P. , Tchernyshyov I., Chang T., et al. 2009. “ c‐Myc Suppression of miR‐23a/b Enhances Mitochondrial Glutaminase Expression and Glutamine Metabolism.” Nature 458, no. 7239: 762–765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Goto‐Koshino, Y. , Ohno K., Nakajima M., Mochizuki H., Kanemoto H., and Tsujimoto H.. 2011. “A Rapid and Simple Method to Obtain Canine Peripheral Blood‐Derived Macrophages.” Journal of Veterinary Medical Science 73, no. 6: 773–778. [DOI] [PubMed] [Google Scholar]
  4. Gross, M. I. , Demo S. D., Dennison J. B., et al. 2014. “Antitumor Activity of the Glutaminase Inhibitor CB‐839 in Triple‐Negative Breast Cancer.” Molecular Cancer Therapeutics 13, no. 4: 890–901. [DOI] [PubMed] [Google Scholar]
  5. Jamshidi‐Parsian, A. , Jenkins S. V., Tran A., et al. 2024. “CB‐839 Induces Reversible Dormancy in Lung Tumor‐Cells.” European Journal of Pharmacology 982: 176912. [DOI] [PubMed] [Google Scholar]
  6. Jin, J. , Byun J., Choi Y., and Park K.. 2023. “Targeting Glutamine Metabolism as a Therapeutic Strategy for Cancer.” Experimental & Molecular Medicine 55, no. 4: 706–715. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Koch, K. , Hartmann R., Tsiampali J., et al. 2020. “A Comparative Pharmaco‐Metabolomic Study of Glutaminase Inhibitors in Glioma Stem‐Like Cells Confirms Biological Effectiveness but Reveals Differences in Target‐specificity.” Cell Death Discovery 6: 20. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Lai, Y. Y. , Horta R. D. S., Almendros A., Ha P. W. Y., and Giuliano A.. 2024. “L‐LOP/LOPP for the Treatment of Canine Gastrointestinal/Hepatosplenic Lymphoma.” Frontiers in Veterinary Science 11: 1373180. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Lee, J. , Kang J. H., Lee S., et al. 2016. “Dual Targeting of Glutaminase 1 and Thymidylate Synthase Elicits Death Synergistically in NSCLC.” Cell Death & Disease 7, no. 12: e2511. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Matsumoto, I. , Nakashima K., Goto‐Koshino Y., et al. 2019. “Immunohistochemical Profiling of Canine Intestinal T‐Cell Lymphomas.” Veterinary Pathology 56, no. 1: 50–60. [DOI] [PubMed] [Google Scholar]
  11. Munday, S. , John L. V., and Christiane K. M. 2017. “Tumors of the Alimentary Tract.” In Tumors in Domestic Animals. 5th ed., edited by Donald M. J., 499–601. John Wiley & Sons. [Google Scholar]
  12. Muranaka, H. , Akinsola R., Billet S., et al. 2024. “Glutamine Supplementation as an Anticancer Strategy: A Potential Therapeutic Alternative to the Convention.” Cancers 16, no. 5: 1057. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Nakagawa, T. , Kojima M., Ohno K., et al. 2022. “Efficacy and Adverse Events of Continuous l‐Asparaginase Administration for Canine Large Cell Lymphoma of Presumed Gastrointestinal Origin.” Veterinary and Comparative Oncology 20, no. 1: 102–108. [DOI] [PubMed] [Google Scholar]
  14. Puurunen, J. , Tiira K., Vapalahti K., Lehtonen M., Hanhineva K., and Lohi H.. 2018. “Fearful Dogs Have Increased Plasma Glutamine and γ‐Glutamyl Glutamine.” Scientific Reports 8, no. 1: 15976. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Rassnick, K. M. , Moore A. S., Collister K. E., et al. 2009. “Efficacy of Combination Chemotherapy for Treatment of Gastrointestinal Lymphoma in Dogs.” Journal of Veterinary Internal Medicine 23, no. 2: 317–322. [DOI] [PubMed] [Google Scholar]
  16. Sakai, K. , Furuya M., Hatoya S., Kanegi R., Nabetani T., and Shimamura S.. 2023. “A Pilot Study for the Development of a Novel Diagnostic Index Based on Plasma Amino Acid Concentrations to Differentiate Chronic Enteropathy and Alimentary Lymphoma in Dogs.” Journal of Animal Physiology and Animal Nutrition 26, no. 1: 8–18. [Google Scholar]
  17. Santos‐Jiménez, J. D. L. , Rosales T., and Ko B., et al. 2023. “Metabolic Adjustments Following Glutaminase Inhibition by CB‐839 in Glioblastoma Cell Lines.” Cancers 15, no. 2: 531. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Schneider, C. A. , Rasband W. S., and Eliceiri K. W. 2012. “NIH Image to ImageJ: 25 Years of Image Analysis.” Nature Methods 9: 671–675. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Singleton, D. C. , Dechaume A., Murray P. M., Katt W. P., Baguley B. C., and Leung E. Y. 2020. “Pyruvate Anaplerosis Is a Mechanism of Resistance to Pharmacological Glutaminase Inhibition in Triple‐Receptor Negative Breast Cancer.” BMC Cancer 20, no. 1: 470. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Sogame, N. , Risbon R., and Burgess K. E. 2018. “Intestinal Lymphoma in Dogs: 84 Cases (1997–2012).” Journal of the American Veterinary Medical Association 252, no. 4: 440–447. [DOI] [PubMed] [Google Scholar]
  21. Solsona, B. G. , Horn H., Schmitt A., et al. 2023. “Inhibition of Glutaminase‐1 in DLBCL Potentiates Venetoclax‐Induced Antitumor Activity by Promoting Oxidative Stress.” Blood Advances 7, no. 24: 7433–7444. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Somali, Z. A. , Hamadani M., Kharfan‐Dabaja M., Sureda A., Fakih R. E., and Aljurf M.. 2021. “Enteropathy‐Associated T Cell Lymphoma.” Current Hematologic Malignancy Reports 16, no. 2: 140–147. [DOI] [PubMed] [Google Scholar]
  23. Tamura, Y. , Ohta H., Kagawa Y., et al. 2019. “Plasma Amino Acid Profiles in Dogs With Inflammatory Bowel Disease.” Journal of Veterinary Internal Medicine 33: 1602–1607. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Umeki, S. , Ema Y., Suzuki R., et al. 2013. “Establishment of Five Canine Lymphoma Cell Lines and Tumor Formation in a Xenotransplantation Model.” Journal of Veterinary Medical Science 75: 467–474. [DOI] [PubMed] [Google Scholar]
  25. Vail, D. M. , Pinkerton M. E., and Young K. M. 2013. “Hematopoietic Tumors.” In Withrow & MacEwen's Small Animal Clinical Oncology. 5th ed., edited by Withrow S. J., and Vail D. M., 608–678. Elsevier. [Google Scholar]
  26. Vail, D. M. , David Pinkerton M., and Karen Y. M. 2020. “Canine Lymphoma and Lymphocytic Leukemias.” In Withrow & MacEwen's Small Animal Clinical Oncology. 6th ed., edited by David V. M., Douglas T. H., and Julius L. M., 688–715. Elsevier. [Google Scholar]
  27. Wu, H. , Zhang J., Wang Q., Li Z., Li L., and Xie Y.. 2025. “Metformin Combined With CB‐839 Specifically Inhibits KRAS‐Mutant Ovarian Cancer.” Scientific Reports 15, no. 1: 6072. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Xiang, L. , Mou J., Shao B., et al. 2019. “Glutaminase 1 Expression in Colorectal Cancer Cells Is Induced by Hypoxia and Required for Tumor Growth, Invasion, and Metastatic Colonization.” Cell Death & Disease 10, no. 2: 40. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Table S1. One hundred sixteen metabolites were analysed in this study.

VMS3-11-e70601-s001.xlsx (10.8KB, xlsx)

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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