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Abstract
Objectives To investigate the effect of the L-arginine metabolism on arthritis and inflammation-mediated bone loss.
Methods L-arginine was applied to three arthritis models (collagen-induced arthritis, serum-induced arthritis and human TNF transgenic mice). Inflammation was assessed clinically and histologically, while bone changes were quantified by μCT and histomorphometry. In vitro, effects of L-arginine on osteoclast differentiation were analysed by RNA-seq and mass spectrometry (MS). Seahorse, Single Cell ENergetIc metabolism by profilIng Translation inHibition and transmission electron microscopy were used for detecting metabolic changes in osteoclasts. Moreover, arginine-associated metabolites were measured in the serum of rheumatoid arthritis (RA) and pre-RA patients.
Results L-arginine inhibited arthritis and bone loss in all three models and directly blocked TNFα-induced murine and human osteoclastogenesis. RNA-seq and MS analyses indicated that L-arginine switched glycolysis to oxidative phosphorylation in inflammatory osteoclasts leading to increased ATP production, purine metabolism and elevated inosine and hypoxanthine levels. Adenosine deaminase inhibitors blocking inosine and hypoxanthine production abolished the inhibition of L-arginine on osteoclastogenesis in vitro and in vivo. Altered arginine levels were also found in RA and pre-RA patients.
Conclusion Our study demonstrated that L-arginine ameliorates arthritis and bone erosion through metabolic reprogramming and perturbation of purine metabolism in osteoclasts.
- Inflammation
- Arthritis
- Osteoporosis
Data availability statement
Data are available on reasonable request.
This is an open access article distributed in accordance with the Creative Commons Attribution Non Commercial (CC BY-NC 4.0) license, which permits others to distribute, remix, adapt, build upon this work non-commercially, and license their derivative works on different terms, provided the original work is properly cited, appropriate credit is given, any changes made indicated, and the use is non-commercial. See: http://creativecommons.org/licenses/by-nc/4.0/.
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WHAT IS ALREADY KNOWN ON THIS TOPIC
Change in intracellular metabolism can influence the functional state of monocytes and contribute to inflammation and bone loss. Osteoclasts are metabolically active cells that preferentially produce energy by glycolysis.
WHAT THIS STUDY ADDS
This study shows that the amino acid L-arginine inhibits arthritis, blocks osteoclast formation and protects from inflammatory bone loss erosion via switching energy metabolism from glycolysis to oxidative phosphorylation, leading to metabolic reprogramming of osteoclasts. Blockade of osteoclast differentiation by L-arginine is mediated via perturbation of purine metabolism, with increased ATP production and elevated inosine and hypoxanthine levels.
HOW THIS STUDY MIGHT AFFECT RESEARCH, PRACTICE OR POLICY
L-arginine is a nutritional supplement and could, therefore, be useful to inhibit joint inflammation and bone destruction.
Introduction
Rheumatoid arthritis (RA) is a chronic inflammatory disease characterised by joint pain and swelling as well as cartilage and bone destruction. RA affects 1% of the population worldwide and constitutes a critical contributor to disability.1 2 Inflammation is a key precipitator for bone loss in RA, resulting in enhanced bone erosion, premature osteoporosis and increased fracture risk.3 Thereby, chronic inflammation perturbates bone homoeostasis with increased bone resorption over bone formation. TNFα is a key proinflammatory cytokine and a well-known trigger of excessive osteoclast differentiation.4 Signalling pathways downstream of TNFα stimulation, such as NF-κB and MAPK-AP1, have been described as being activated in osteoclasts.5
Osteoclasts require high amounts of energy for their differentiation. Oxidative phosphorylation contributes to the differentiation of osteoclasts,6 while glycolysis influences osteoclast resorptive activity.7 8 Furthermore, mitochondrial pathways are involved in osteoclast function, since lack of PGC-1α, a master regulator of mitochondrial biogenesis and respiration, reduces resorptive activity of osteoclasts.9 Inflammation may alter the functional state of osteoclasts by altering their metabolism. However, to date, no study has investigated the metabolic properties of osteoclasts under inflammatory conditions.
We hypothesised that changes in amino acid metabolism in osteoclasts occur during inflammation. Amino acids have shown to regulate osteoclastogenesis.10–12 L-arginine is a semiessential amino acid built after protein conversion.13 It is used as a nutritional supplement since many years. Physiologically, L-arginine serves as a building block in protein synthesis and participates in deamination through the urea cycle.14 L-arginine can be metabolised by various enzymes such as nitric oxide synthetase (NOS) and arginase (ARG).15 Notably, L-arginine preserves bone mass in mice under non-inflammatory conditions.16 17 Based on the observations, L-arginine could serve as an important link between inflammation and bone loss by reprogramming the metabolic state of osteoclasts. We, therefore, tested the hypothesis that L-arginine may inhibit inflammatory bone loss by altering the metabolic state and function of osteoclasts.
Methods
Mice
Seven-week-old male C57BL/6 mice were purchased from Charles River (Germany). Seven-week-old male DBA/1 mice were purchased from Janvier (Germany). Mice were co-housed for 1 week prior to the start of experiments. The hTNFtg mice (strain Tg197 on C57BL/6 background) have been previously described.18 Evaluation of arthritis in hTNFtg mice initiated 6 weeks after birth and were performed twice a week. The generation of Jun-floxed and LysM-Cre mice has been previously described.19 The mice were bred and maintained on a 129/C57BL/6 background. The generation of Arg-floxed and Tie2-Cre mice on a B6 background has been previously described.20 Arg-floxed Ctsk-Cre mice on a B6 background were obtained by independent backcrossing of Ctsk-Cre and Arg-floxed mice with C57BL/6 mice for 12 generations and intercrossing thereafter. Littermate mice were used as controls. Mice were age and sex matched and used at 7–12 weeks of age. All mice were housed in a temperature-controlled and humidity-controlled facility with free access to food and water.
RNA-seq and bioinformatics
For RNA-seq, 1×106 cells from day 0 were cultured in 1 mL/well osteoclast medium with 20 ng/mL M-CSF and 20 ng/mL RANKL in 24-well plates at 37°C and 5.5% CO2. On day 2 of culture, 18 hours after the stimulation with or without 40 ng/mL TNFα, supplemented or not with 10 mM L-arginine, total RNA was harvested and isolated from the four groups and purified with the RNeasy Mini Kit (QIAGEN) according to the manufacturer’s instructions. RNA-seq was carried out by Novogene. A total amount of 1 µg RNA per sample was used as input material for the RNA sample preparations. Sequencing libraries were generated using the NEBNext Ultra RNALibrary Prep Kit for Illumina (New England Biolabs). Clustering of the index-coded samples was completed on a cBot Cluster Generation System using PE Cluster Kit cBot-HS (Illumina), according to the manufacturer’s instructions. The library preparations were sequenced on an Illumina HiSeq platform and paired-end reads were generated. The filtered reads were aligned to the Mus musculus genome (Mus_musculus. GRCm 38) with HISAT2 (V.2.0.5). HTSeq (V.0.6.1) was used to count the read numbers mapped of each gene, including known and novel genes.
Raw paired-end reads were aligned to the reference genome using Rsubread (V.2.6.4) within R programming language (V.4.1.1).21 Quantification was performed using featureCounts function from RSubread. Differential expression analysis was performed following the previously published pipeline using DESeq2 (V.1.32.0).22 23 Gene Set Enrichment Analysis (GSEA) was performed using Broad Institutes java application.24 25 Heatmaps of normalised counts were plotted using pheatmap (V.1.0.12). Transcription factors were enriched with NetworkAnalyst 3.0.26
Extracellular flux assay using the Seahorse platform
For metabolic analysis, 4×105 cells from day 0 were seeded on Seahorse XF96 Cell Culture Microplates (Agilent) in 200 µL/well osteoclast medium with 20 ng/mL M-CSF and 20 ng/mL RANKL. On day 2 of culture, 4 hours after the stimulation of 40 ng/mL TNFα, and 10 mM L-arginine, mitochondrial respiration (Seahorse XF Cell Mito Stress Test) successively adding final concentrations of 2 µM oligomycin, 2 µM carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP), 1 µM antimycin A and 1 µM rotenone (all from Millipore Sigma) and glycolytic capacity (Seahorse XF Cell Glyco Stress Test) successively adding final concentrations of 10 mM glucose, 2 µM Oligomycin, 50 mM 2-Deoxy-d-glucose (2-DG) (all from Millipore Sigma) were measured in preosteoclasts, according to the manufacturer’s instructions. Oxygen consumption rate and extracellular acidification rate were measured in a Seahorse XFe96 Analyzer (Agilent) and the values were normalised to the protein concentration within the single wells, detected with the DC protein assay (Bio-Rad). Data were analysed with the Seahorse XF Report Generator for Mito Stress Test and Glyco Stress Test. ATP rate assay was performed with Seahorse XF Real-Time ATP Rate Assay Kit, according to the manufacturer’s instructions.
Single Cell ENergetIc metabolism by profilIng Translation inHibition
The Single Cell ENergetIc metabolism by profilIng Translation inHibition (SCENITH) assay was performed as previously described.27 Briefly, 1×106 cells from day 0 were cultured in 1 mL/well osteoclast medium with 20 ng/mL M-CSF and 20 ng/mL RANKL in 24-well plates at 37°C and 5.5% CO2. On day 2 of culture, 10 hours after the stimulation of 40 ng/mL TNFα, and 10 mM L-arginine, preosteoclasts from the four groups were incubated with 2-DG (final concentration: 100 mM) for 10 min and/or followed by Oligomycin (final concentration: 100 mM) for 5 min. Puromycin at the final concentration of 10 µg/mL was supplemented into the medium and the plates were incubated for 1 hour at 37°C. Afterwards, the cells were washed with a cold FACS buffer and detached. The cells were then transferred to a V-BOTTOM 96-well plate (Greiner) and centrifuged at 500 g for 2 min. The cell suspension was blocked with a final concentration of 0.5 µg/mL anti-mouse CD16/32 for 10 min at 4°C and subsequently stained with the surface markers of preosteoclasts for 30 min at 4°C. The gating strategy was illustrated in online supplemental figure S5B. The details of FACS antibodies were shown in online supplemental table S2. The following intracellular staining was carried out using the intracellular Foxp3 Fixation/permeabilisation solution (eBioscience, Thermo Fisher) according to the manufacturer’s instructions and the staining of puromycin was carried out with a directly labelled anti-puromycin antibody-FITC (Millipore Sigma). Samples were analysed using the CytoFLEX S Flow Cytometer (Beckman Coulter, Krefeld, Germany).
Supplemental material
Supplemental material
Mass spectrometry
For mass spectrometry (MS), 5×106 cells from day 0 were cultured in 5 mL/well osteoclast medium with 20 ng/mL M-CSF and 20 ng/mL RANKL in 6-well plates at 37°C and 5.5% CO2. For the 13C6-L-arginine tracing experiment, SILAC medium (Thermo, #88368) supplemented with 0.4 mM L-lysine (Roth, # 39665-12-8) and 0.5 mM 13C6-L-arginine (Sigma, 643440) was used for the control and TNFα-activated group. Additional 10 mM 13C6-L-arginine was supplemented in the L-arginine supplemented group and TNFα+L-arginine (TNFα and L-arginine double treated group). Mediums with 12C6-L-arginine were used as unlabelled control. On day 2 of culture, medium was replaced while supplemented or not with 10 mM L-arginine, stimulated or not with 40 ng/mL TNFα. Mature osteoclasts were then harvested at day 3 from the four groups: Control; L-arginine supplemented group; TNFα activated group; TNFα+L-arginine (TNFα and L-arginine double treated group). And cells pooled from two wells with the same treatments were counted as one sample.
Mature osteoclasts were then lysed in 1 mL 80% MeOH containing reference standards. The mixtures were vortexed, centrifuged (5 min, 16 000 rpm, 4°C) and from each sample two aliquots were separated (350 µL) for further sample preparation. Furthermore, a pooled sample was prepared with aliquots (230 µL) taken from each sample. This pooled mix was split into aliquots of 350 µL, which represented quality controls (QCs) and followed the same SOP as the individual samples.
The samples and QCs were dried under nitrogen at room temperature and stored at −80°C until metabolomics analysis. Prior to the analysis, the samples were reconstituted in 200 µL eluent containing internal standards, vortexed rigorously and transferred to HPLC insert vials.
The samples were analysed using an ultra-high-performance liquid chromatography (UPLC) (Ultimate 3000, Thermo Fisher Scientific) hyphenated to a high-resolution mass spectrometer (QExactive Orbitrap Focus, Thermo Fisher Scientific). All samples were analysed in both, hydrophilic interaction chromatography (HILIC) and reverse phase (RP) chromatography. For HILIC, an ACQUITY UPLC BEH Amide column (1.7 µm, 2.1×100 mm, Waters) was installed. The mobile phase consisted of eluent A (10 mM ammonium formate, 0.1% formic acid; FA) and eluent B (10 mM ammonium formate, 95% acetonitrile, 0.1% FA). The gradient programme started with 100% B (0–2 min), which was decreased to 30% over 12 min. This concentration was hold (14–16.5 min) and returned to the starting conditions over 1 min (17.5 min). The system was equilibrated for 10 min. For RP, an ACQUITY UPLC BEH C18 column (1.7 µm, 2.1×100 mm, Waters) was installed. Chromatographic separation was achieved using a gradient elution, whereby eluent A consisted of water with 0.1% (vol/vol) FA and eluent B of MeOH with 0.1% (vol/vol) FA. The gradient programme started with 10% B. After injection, the percentage of B increased from 10%–98% over 9 min. From 9 min to 11 min, the B was maintained at 98%. Finally, the starting conditions (10% B) were reconditioned from 11 min to 11.50 min. Conditions were maintained till 15 min to re-equilibrate the column. For both, HILIC and RP, the flow rate was 0.35 mL/min, the column heater was maintained at 50°C and the autosampler temperature was 4°C. The injection volume was 2 µL.
The mass spectrometer was operated in full MS/dd-MS2 (discovery) mode with a resolution of 17,500, a scan range of 70–800 and an automatic gain control target of 1e6. Sheath gas, aux gas and sweep gas flow rated were set to 60, 20 and 0, respectively. The spray voltage of the HESI source was 4.00 kV. All samples were analysed in both, positive and negative mode for ionisation.
Recording, processing and evaluation of raw data were accomplished using Trace Finder Software V.4.1 (Thermo Fisher Scientific). Further data processing including spectral alignment, descriptive statistic and metabolite annotations was achieved using the Compound Discoverer Software V.3.1 (Thermo Fisher Scientific). Metabolic pathway enrichment was accomplished via MetaboAnalyst V.5.0.28
Human subjects
Serum samples of RA patients were collected from the Department of Rheumatology, Renji Hospital, Shanghai Jiao Tong University between March 2017 and July 2017, according to the 2010 RA classification criteria by American College of Rheumatology/European League Against Rheumatism.29 A total of 23 RA cases were finally recruited. Two naive patients had no treatment history. The other patients were treated with low dose of glucocorticoids and/or disease-modifying antirheumatic drugs (DMARDs). Patients treated with high-dose glucocorticoids (>10 mg/day) or biological agents within the past 6 months were excluded from the study. Serums of 29 age, gender matched healthy individuals were collected as controls. The general information of all individuals and associated laboratory examinations of RA patients were collected and documented (online supplemental table S1). For pre-RA data, serum samples were drawn from 39 patients who had been referred to the Rheumatology outpatient clinic in Linköping, Sweden, due to musculoskeletal pain and a positive test for anti-citrullinated protein antibodies (anti-CCP2). None of the patients had arthritis on clinical examination at baseline, but developed at least one after median 6 months (IQR 3–24 months). None were treated with corticosteroids or DMARDs at the time of sampling. Mean age was 55 years and 82% were female.
Statistical analyses
GraphPad Prism V.9.0 (GraphPad Software) was used for statistical treatment. Experimental data was shown as mean±SEM. Two-tailed unpaired Student’s t-test for two groups or analysis of variance (ANOVA) with multiple comparisons tests were used as indicated in respective experiments. For kinetic measurements, repeated measures two-way ANOVA (two-way RM ANOVA) was used in which the time factors are matched, to account for the nestedness of the data.
More detailed experimental methods and procedures are provided in online supplemental materials.
Results
L-arginine inhibits arthritis and inflammatory bone loss
To investigate whether L-arginine could protect against inflammation and systemic inflammatory bone loss, we used the serum-induced arthritis (SIA) model and supplemented the mice orally with L-arginine at day 0 or day 4 after serum injection (figure 1A). L-arginine led to significant reduction of clinical arthritis (figure 1B). Synovitis, bone erosions and osteoclast numbers were decreased (online supplemental figure S1A, B), accompanied with reduced expression of Ctsk and Mmp9 in the arthritic joints (online supplemental figure S1C). We also analysed tibial bones of SIA mice and did not find significant differences in bone mass between L-arginine and vehicle treatment (figure 1C,D and online supplemental figure S1D, E). Nonetheless, L-arginine reduced tibial osteoclast number and size (figure 1E,F), while not altering RANKL/OPG ratio (online supplemental figure S1F).
L-arginine protects from bone loss in three murine arthritis models. (A) Arthritis was induced by K/BxN serum transfer into C57BL/6 J mice and 40 g/L L-arginine was supplemented in drinking water, either simultaneously with the K/BxN serum transfer (SIA/L-Arginine day 0) or 4 days after serum transfer (SIA/L-arginine day 4). Mice were sacrificed at day 10 after serum transfer; (B) Arthritis score and quantification of area under the curve (AUC) of arthritis of SIA, SIA/L-arginine day 0 and SIA/L-arginine day 4 groups (n=15–25/group); (C, D) Representative tibial μCT images (C) and quantification of bone volume/total volume (BV/TV), bone density trabecular numbers (Tb.) (N) trabecular thickness (Tb. Th) and trabecular separation (Tb. Sp) (D) in control, control/L-arginine, SIA, SIA/L-arginine day 0 and SIA/L-arginine day 4 groups (n=5–9/group). Scale bars: 1 mm; (E, F) Representative images of tartrate-resistant acid phosphatase (TRAP) staining (E) and quantification of osteoclast surface/bone surface (Oc.S/BS), osteoclast number/tissue area (N.Oc/T.Ar) and osteoclast number/bone perimeter (N.Oc./B.Pm.) (F) in control, control/L-arginine, SIA, SIA/L-arginine day 0 and SIA/L-arginine day 4 groups (n=5–9/group). Scale bars: 500 µm; (G) Collagen-induced arthritis (CIA) was triggered by immunisation with an emulsion of complete Freund’s adjuvant and type II collagen (CII) into DBA/1 J mice, supplemented with 40 g/L L-arginine, after the onset of arthritis (day 23). Mice were sacrificed at day 39 after first immunisation; (H) Arthritis scores and quantification of AUC of CIA and CIA/L-arginine groups (n=7–9/group); (I, J) Representative tibial μCT images (I) and quantification of BV/TV, Tb. N, Tb. Th and Tb. Sp (J) in controls, control/L-Arginine, CIA and CIA/L-arginine groups (n=4–9/group). Scale bars: 1 mm; (K, L) Representative images of tartrate-resistant acid phosphatase (TRAP) staining (K) and quantification of Oc.S/BS, N.Oc/T.Ar and N.Oc./B.Pm. (L) in controls, control/L-arginine, CIA and CIA/L-arginine groups (n=4–9/group). Scale bars: 500 µm; (M) Human TNF-transgenic (hTNFtg) mice developed spontaneously erosive arthritis aggravated with age. 40 g/L L-arginine was supplemented within drinking water 6 weeks after birth. Mice were sacrificed 12 weeks after birth; (N, O) Kinetic arthritis score (N) and measurements of grip strength (O) in hTNFtg−/−, hTNFtg+/−, hTNFtg−/−/L-Arginine and hTNFtg+/−/L-Arginine groups (n=5–10/group) during the course of spontaneous arthritis; (P, Q) Representative tibial μCT images (P) and quantification of tibial BV/TV, Tb. N, Tb. Th and Tb. Sp (Q) in hTNFtg−/−, hTNFtg+/−, hTNFtg−/−/L-arginine and hTNFtg+/−/L-arginine groups (n=5–10/group). Scale bars: 1 mm; (R, S) Representative images of tartrate-resistant acid phosphatase (TRAP) staining (R) and quantification of tibial Oc.S/BS, N.Oc/T.Ar and N.Oc./B.Pm. (S) in hTNFtg−/−, hTNFtg+/−, hTNFtg−/−/L-arginine and hTNFtg+/−/L-arginine groups (n=5–10/group). Scale bars: 500 µm. Graph points indicate individual mice. Data are shown as mean±SE.e.m. Asterisks mark statistically significant difference (*p<0.05; **p<0.01; ***p<0.001; ****p<0.0001), Student’s t-test in (H) (AUC result) and one-way ANOVA in (B) (AUC test), (D, F, J, L, Q, S), two-way repeated measures ANOVA in (B, H, N, O). See also in online supplemental figures S1 and S2. ANOVA, analysis of variance.
We hypothesised that longer-term L-arginine treatment may have a better outcome. Therefore, we treated DBA/1J mice with collagen-induced arthritis (CIA) with L-arginine starting at day 23 post first immunisation (figure 1G). Like in the SIA model, joint inflammation was significantly ameliorated by L-arginine, as shown by reduced arthritic scores (figure 1H). Synovitis, bone erosion and osteoclast numbers were also reduced by L-arginine (online supplemental figure S2A, B), despite no difference in Ctsk and Mmp9 expression in the joints (online supplemental figure S2C). μCT analysis of tibial and vertebral bones showed that L-arginine treatment ameliorated bone loss with significant increases in bone volume and trabecular numbers (figure 1I,J, online supplemental figure S2D, E). Histomorphometry analyses of osteoclasts using TRAP staining showed a significant suppression of osteoclast numbers in L-arginine-treated CIA mice (figure 1K,L). Interestingly, TNFα expression and RANKL/OPG ratio in bone marrow supernatant were not altered (online supplemental figure S2F, G), suggesting that L-arginine might ameliorate bone loss despite the presence of proinflammatory and pro-osteoclastogenic cytokines like TNFα and RANKL, respectively.
Finally, the beneficial role of L-arginine on arthritis was also demonstrated in hTNFtg mice that represent a spontaneous model of inflammatory arthritis.30 Six weeks after birth, L-arginine was supplemented in drinking water (figure 1M). Like the other two experimental models, joint swelling was diminished in L-arginine treated mice accompanied with an improved grip strength (figure 1N,O) and reduced inflammatory area (online supplemental figure S2H, I). Furthermore, μCT and TRAP staining results showed that L-arginine mitigated tibial and vertebral bone destruction with increased bone volume and decreased osteoclast numbers (online supplemental figure S1P–S and J, K). Taken together, L-arginine inhibits arthritis and inflammatory bone loss by reducing osteoclast numbers.
L-arginine mitigates TNFα-induced osteoclastogenesis
To delineate whether L-arginine acts directly on osteoclasts, we isolated bone marrow cells from hTNFtg+/− and littermate control mice, and performed MCSF/RANKL-mediated osteoclast differentiation assays with or without L-arginine (online supplemental figure S3A). A 0.5–10 mM L-arginine did not influence cell viability (online supplemental figure S3B). Osteoclastogenesis was significantly inhibited by L-arginine in a dose-dependent manner in WT and hTNFtg+/− cells (online supplemental figure S3C, D). Bone resorption activity was also reduced by L-arginine in WT and hTNFtg+/− cells (online supplemental figure S3E). The inhibitory role of L-arginine on osteoclast differentiation was further confirmed by a lower expression of several osteoclast-specific genes, such as Traf6, Acp5, Nfatc1, Ctsk and Atp6v0d2, particularly in cells isolated from hTNFtg+/− mice (online supplemental figure S3F).
Next, we performed osteoclast differentiation assays with WT bone marrow cells, which were stimulated with TNFα to mimic the proinflammatory microenvironment. Cultures were supplemented with L-arginine during the whole differentiation process or only at the later phase (figure 2A). The concentrations of L-arginine used did not have any toxic effects (figure 2B). As expected, TNFα stimulation (40 ng/mL) enhanced osteoclast numbers, whereas 10 mM L-arginine profoundly reduced osteoclast differentiation (figure 2C,D). Interestingly, L-arginine inhibited osteoclastogenesis more efficiently when supplemented at the later phase than during the whole procedure of differentiation (figure 2C,D). TNFα stimulation promoted F-actin ring formation in osteoclasts, which was disrupted by L-arginine (figure 2E). Bone resorption activity was also reduced by L-arginine, also in the cells stimulated with TNFα (figure 2F,G). Kinetic analysis of osteoclast-associated genes illustrated their variation along the differentiation process. Their expression was increased after TNFα stimulation, but was significantly diminished after L-arginine treatment (figure 2H). Furthermore, to explore the role of L-arginine on pathological osteoclastogenesis, we isolated synovial inflammatory macrophages with the markers CD45+Cx3cr1+ from the knee and ankle joints of C57BL/6 mice as shown in a previous study.31 Then, we performed osteoclast differentiation assay with these cells and added L-arginine, with or without TNFα stimulation (online supplemental figure S4A, B). As shown in online supplemental figure S4C, we observed that TNFα stimulation could significantly elevate osteoclast differentiation whereas L-arginine again reduced the osteoclast formation. Next, we checked whether osteoclastogenesis is impaired in the complete lack of arginine, with or without TNFα stimulation. As expected, osteoclastogenesis was completely inhibited in Arg-free medium (online supplemental figure S5).
L-arginine reduces TNFα-induced osteoclastogenesis and resorption activity. (A) Schematic diagram of osteoclastogenesis from the bone marrow cells of C57BL/6 J mice; supplementation of L-arginine (10 mM) was done during the whole procedure (days 0–3) or only at the later phase (days 2–3), with or without stimulation with 40 ng/mL TNFα; (B) Cell viability on day 2 of WT osteoclasts exposed to different doses of L-arginine supplementation (0.5 mM, 1 mM, 3 mM, 5 mM, 10 mM) (n=3/group, representative of three independent experiments); (C, D) Quantification of TRAP+ osteoclasts (C) and representative TRAP staining images (D) of mature osteoclasts at day 3 from different treatments (n=5–10/group, representative of three independent experiments). Scale bars: 100 µm; (E) Representative F-actin staining images of mature osteoclasts at day 3, supplementation of L-arginine (10 mM) was done at the later phase, with or without stimulation with 40 ng/mL TNFα. Scale bars: 50 µm; (F, G) Representative images of pit formation assay (F) and quantification of bone resorption area (G) at day 3, supplementation of L-arginine (10 mM) was done at the later phase, with or without stimulation with 40 ng/mL TNFα (n=3/group, representative of three independent experiments). Scale bars: 100 µm. (H) Expression of osteoclast-associated genes at day 0, day 1, day 2 and day 3 of culture; supplementation of L-arginine (10 mM) was done at the later phase, with or without stimulation with 40 ng/mL TNFα (n=3/group, representative of three independent experiments) (I, J) KEGG pathway enrichment results from RNA sequencing data in preosteoclasts supplemented or not with 10 mM L-arginine at the later phase, with or without the stimulation with 40 ng/mL TNFα (n=3/group), comparing L-arginine versus control (I) and TNFα/L-arginine versus TNFα (J); (K) Gene set enrichment analysis (GSEA) results indicating significant difference in oxidative phosphorylation (OXPHOS), comparing TNFα/L-arginine versus TNFα; (L) Heatmap showing the differentially expressed genes61 linked to OXPHOS pathway in control, L-arginine, TNFα and TNFα/L-arginine groups. Genes with an adjusted p value (p-adj) less than 0.05 are assigned as differentially expressed. Pathways with p-adj value less than 0.05 are considered significantly enriched. Data are shown as mean±SE. Asterisks mark statistically significant difference (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001), one-way ANOVA in (B) and two-way ANOVA in (C, F). See also in online supplemental figures S3–S6. ANOVA, analysis of variance.
To delineate the most important downstream metabolites of L-arginine on osteoclast differentiation, particularly after TNFα stimulation, we supplemented the cells with either putrescine, ornithine or spermidine, the metabolites directly produced by arginase catalysis in presence or absence of TNFα. Interestingly, putrescine could significantly diminish osteoclast differentiation and suppress the expression of OC-related genes, especially after TNFα stimulation (online supplemental figure S6A, D, G). At a higher concentration ornithine (25 mM) could also reduce osteoclastogenesis in TNFα-stimulated condition only (online supplemental figure S6B, E, H), whereas spermidine did not alter osteoclast numbers (online supplemental figure S6C, F, I).
Collectively, L-arginine inhibits osteoclast differentiation in vitro, especially in inflammatory conditions.
L-arginine promotes oxidative phosphorylation and ATP production
To delineate L-arginine function, RNA-sequencing was performed in L-arginine-treated osteoclasts that were stimulated with TNFα or were left non-stimulated. Only few genes and related pathways were enriched in cells treated with L-arginine in non-stimulated cells (figure 2I). However, when the cells were stimulated with TNFα, KEGG pathway analysis indicated strong enrichment of oxidative phosphorylation pathway genes after L-arginine (figure 2J). GSEA further demonstrated a significant enrichment of oxidative phosphorylation in the TNFα/L-arginine treated compared with TNFα-stimulated cells (figure 2K). Heatmap analysis also highlighted genes linked to oxidative phosphorylation, which were significantly upregulated in the TNFα/L-arginine treated cells (figure 2L).
To further confirm the boost of oxidative phosphorylation in TNFα/L-arginine treated cells, extracellular flux assay was performed. Glycolysis was unaltered in WT and hTNFtg+/− preosteoclasts (online supplemental figure S7A, B). Oxidative phosphorylation was lower in hTNFtg+/− compared with to WT cells, but was rescued by L-arginine (online supplemental figure S7C). In addition, maximal respiration, non-mitochondrial oxygen consumption, ATP production and proton leak were increased after L-arginine (online supplemental figure S7D). At RNA level, expression of genes involved in oxidative phosphorylation was increased after L-arginine (online supplemental figure S7E). Similarly, TNFα stimulation promoted glycolysis in WT cells, which was not affected by L-arginine (figure 3A). At the same time, TNFα suppressed oxidative phosphorylation and boosted ATP production, which was reversed by L-arginine (10 mM) (figure 3B,C). ATP production by glycolysis increased, while ATP production by oxidative phosphorylation decreased with differentiation into osteoclasts (online supplemental figure S8A). After TNFα stimulation, glycolytic ATP production increased but was switched into oxidative phosphorylation by L-arginine (figure 3D,E).
L-arginine boosts oxidative phosphorylation (OXPHOS) and ATP production of osteoclast under inflammatory conditions. (A) Extracellular acidification rate (ECAR) analysed by extracellular flux assay in preosteoclasts from WT cells; supplementation with L-arginine (10 mM) was done at the later phase, with or without stimulation with 40 ng/mL TNFα (n=9–14/group, representative of three independent experiments); (B, C) Oxygen consumption rate (OCR) (B) and associated mitochondrial ATP production (C) analysed by extracellular flux assay in preosteoclasts from WT cells; supplementation of L-arginine (10 mM) was done at the later phase, with or without stimulation with 40 ng/mL TNFα (n=9–14/group, representative of three independent experiments); (D, E) ATP rate (D) and percentage of ATP produced by glycolysis or OXPHOS (E) analysed by real-time ATP rate assay in preosteoclasts from WT cells; supplementation of L-arginine (10 mM) was done at the later phase, with or without stimulation with 40 ng/mL TNFα (n=6–12/group, representative of three independent experiments); (F, G) Representative histogram of flow cytometry analysis with the treatment of DMSO, 2-DG, Oligomycin and 2-DG/Oligomycin (F) and percentage of mitochondria dependence (G) via Single Cell ENergetIc metabolism by profilIng Translation inHibition (SCENITH) in preosteoclasts (CD45+CD11b+Ly6G- Mcsfr+RANK+ cells) from WT cells; supplementation of L-arginine (10 mM) was done at the later phase, with or without 40 ng/mL TNFα stimulation (n=3–6/group, representative of three independent experiments); (H) Glucose consumption in the cell supernatant of mature osteoclasts at day 3; supplementation of L-arginine (10 mM) was done at the later phase, with or without 40 ng/mL TNFα stimulation (n=2–3/group, pooled from three independent experiments); (I, J) Representative images of JC-1 staining (I) and ratio of JC-1 Aggregate (red) / JC-1 monomeric (green) fluorescence (J) in mature osteoclasts at day 3; supplementation of L-arginine (10 mM) was done at the later phase, with or without 40 ng/mL TNFα stimulation (n=2–3/group, pooled from two independent experiments). Scale bars: 50 µm; (K–N) Quantification of TRAP+ osteoclasts (K, M) and representative TRAP staining images (L, N) of mature osteoclasts at day 3 in control versus L-arginine treated groups (K, L) and TNFα versus TNFα/L-arginine groups (M, N), combined with different concentrations (5 nM, 10 nM, 20 nM) of OXPHOS inhibitors Rotenone and Antimycin. Scale bars: 100 µm. Data are shown as mean±SEM. Asterisks mark statistically significant difference (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001), one-way ANOVA in (C, E, K, M) and two-way ANOVA in (A, B, G, H, I). For A and B, black asterisk means group TNFα versus control and purple asterisk means group TNFα/L-arginine versus TNFα. See also in online supplemental figures S7 and S8. ANOVA, analysis of variance.
To investigate the energy metabolism at single cell level, we performed SCENITH assay in preosteoclasts isolated from the four treatment groups. Arginine increased mitochondrial dependence and decreased glycolytic capacity in TNFα-stimulated cells (figure 3F,G, online supplemental figure S8B–D). No difference of glucose dependence or fatty acid and amino acid oxidation capacity were found (online supplemental figure S8C, D). Furthermore, increased oxidative phosphorylation by L-arginine was not associated with increased glucose consumption (figure 3H). Mitochondrial ROS levels were slightly decreased with L-arginine treatment (online supplemental figure S8E,F). When analysing mitochondrial morphology by transmission electron microscopy, no significant differences could be detected between the four stimulations (online supplemental figure SG). Therefore, we explored possible changes in mitochondrial membrane potential. JC-1 staining demonstrated an impaired mitochondrial membrane potential after TNFα stimulation, while L-arginine restored the membrane potential (figure 3I,J). Finally, when blocking oxidative phosphorylation by rotenone and antimycin, effects of L-arginine were abolished in dose-dependent manner exclusively in TNFα-stimulated condition (figure 3K–N).
Induction of arginase-1 mediates osteoclast inhibition by L-arginine
To investigate the transcriptional regulations in our model, we performed transcription factor enrichment analyses based on our RNA-sequencing data. Interestingly, c-Jun was found enriched in all three transcription factor databases from the DEGs (figure 4A). Since c-Jun is a known transcription factor controlling osteoclast differentiation32 and its expression was negatively related to the expression of arginase-1 (Arg-1) in macrophages,33 we pursued the characterisation of c-Jun at protein level in our four culture conditions. C-Jun expression was enhanced by TNFα, but significantly decreased after L-arginine, especially when L-arginine was supplemented at the later phase of osteoclast differentiation (figure 4B). Next, we analysed the role of c-Jun in osteoclastogenesis with or without TNFα stimulation. To do so, we isolated bone marrow cells from cJunΔLysM and littermate control mice. Lower osteoclast numbers in c-Jun deficient cells were observed, even after TNFα stimulation (figure 4C,D). Interestingly, the expression of Arg1 was profoundly elevated in c-Jun deficient osteoclasts after TNFα stimulation, whereas Arg2 was unchanged (figure 4E, online supplemental figure S9A). We, therefore, investigated the role of Arg-1 in L-arginine-mediated inhibition of osteoclasts. Arginase activity was elevated in the paw tissue lysates of L-arginine treated mice with CIA, while urea level was unchanged and nitrite level decreased (figure 4F, online supplemental figure S9B). These data imply that c-Jun controls Arg1 expression mediating the inhibitory role of L-arginine on osteoclast differentiation.
Induction of Arginase-1 is mediating osteoclast inhibition by L-arginine. (A) Transcription factors enriched from three databases with the differentially expressed genes in RNA-seq data comparing TNFα/L-arginine versus TNFα treated groups; (B) Protein levels of c-Jun in mature osteoclasts at day 3 (normalised to b-actin levels), with supplementation of L-arginine (10 mM) during the whole culture or only at the later phase, with or without 40 ng/mL TNFα stimulation; (C, D) Schematic diagram of the osteoclastogenesis with bone marrow cells from cJun ∆LysM mice or littermates (C, above); representative TRAP staining images (C, bottom) and quantification of TRAP+ osteoclasts (D) from c-Jun-deficient cells or littermate controls at day 3, with or without 40 ng/mL TNFα stimulation (n=3–5/group, representative of three independent experiments). Scale bars: 100 µm; (E) Arginase (Arg)-1 expression in mature osteoclasts from c-Jun-deficient cells or littermate controls at day 3, with or without 40 ng/mL TNFα stimulation (n=3–6/group, representative of three independent experiments); (F) Normalised arginase activity in the paw lysate of mice from controls, control/L-arginine, CIA and CIA/L-arginine groups (n=4–9/group); (G) Putative binding site of c-Jun on the promoter sites of Arg-1 predicted by JASPAR; Binding of c-Jun or IgG on the promoter sites of Arg-1 in preosteoclasts at day 2, stimulated with TNFα for 4 hours (n=4/group); (H) Binding of H3me3K4, H3me3K27 or IgG on the promoter sites of Arg-1 in preosteoclasts at day 2, stimulated with TNFα for 4 hours (n=6/group); (I–J) Schematic diagram of the osteoclastogenesis with bone marrow cells from Arg-1 ∆Tie2 mice or littermates, representative TRAP staining images (I) and quantification of TRAP+ osteoclasts (J) from Arg-1-deficient cells or littermate controls at day 3; supplementation of L-arginine (10 mM) was done at the later phase, with or without 40 ng/mL TNFα stimulation (n=3/group, representative of three independent experiments). Scale bars: 100 µm; (K, L) Schematic diagram of the osteoclastogenesis procedure with bone marrow cells from Arg-1 ∆Ctsk mice or littermates, representative TRAP staining images (K) and quantification of TRAP+ osteoclasts (L) from Arg-1-deficient cells or littermate controls at day 3, supplementation of L-arginine (10 mM) was done at the later phase, with or without 40 ng/mL TNFα stimulation (n=4/group, representative of three independent experiments). Scale bars: 100 µm. Data are shown as mean±SE. Asterisks mark statistically significant difference (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001), one-way ANOVA in (F, G, H) and two-way ANOVA in (D, E, J, L). See also in online supplemental figure S9. ANOVA, analysis of variance; CIA, collagen-induced arthritis.
To verify whether c-Jun regulates Arg1 expression at the transcriptional level especially on TNFα stimulation, we included 5000 bp upstream of the Arg1 transcription start site (TSS) and predicted eight putative cJun binding sites on Arg1 promoter in silico with JASPAR database34 (figure 4G). ChIP-qPCR was then performed with an anti-c-Jun antibody in preosteoclasts stimulated with TNFα. Indeed, c-Jun could bind to the promoter sites 3, 4, 5 and 8 in steady state, and to sites 1, 2, 4, 6 and 8 on TNFα stimulation (figure 4G). Interestingly, the chromatin remodelling markers indicated an inhibitory pattern of c-Jun binding sites, with enhanced H3me3K27 methylation of Arg1 promoter after TNFα stimulation, whereas no change was detected for H3me3K4 methylation (figure 4H). These results indicate that TNFα downregulates Arg1 expression by c-Jun.
Next, we analysed bone marrow-derived osteoclast from two strains of Arg-1 conditional knockout mice. As expected, L-arginine could diminish osteoclast differentiation in WT cells, but not in Arg-1 deficient cells after TNFα stimulation (figure 4I–L).
Collectively, these results suggested that Arg-1 is essential for allowing L-arginine to inhibit osteoclastogenesis after TNFα stimulation.
L-arginine reprogrammes purine metabolism on inflammatory stimulation
To investigate the metabolic flux after L-arginine supplementation, we first performed the metabolic tracing of L-arginine using 13C-isotopologue labelling. As shown in figure 5A, the downstream metabolites labelled with 13C-isotopologue after 13C6-L-arginine supplementation were ornithine, spermine, N-acetyl-spermidine, N-acetyl-citrulline/citrulline, D-proline, pyrroline and pyrrolidine. Interestingly, as shown in the isotopologue distribution chart of the individual metabolite, TNFα stimulation increased the consumption of 13C6-L-arginine, while 13C6-L-arginine supplementation enhanced its cellular concentration. In addition, 13C6-L-arginine supplementation promoted the production of 13C4-ornithine, 13C5-D-proline, 13C4-spermine, 13C4-N-acetyl-spermidine, 13C4-pyrroline and 13C4-pyrrolidine. However, 13C6-L-arginine supplementation reduced the production of 13C5-ornithine with TNFα stimulation, implying enhanced ornithine consumption exclusively in inflammatory conditions. Collectively, these data indicate that elevated L-arginine bioavailability indeed skews the metabolic flux towards Arg-1 direction.
L-arginine perturbates purine metabolism in TNFα stimulated cells. Mature osteoclasts were harvested at day 3; supplementation of L-arginine (10 mM) or 13C6-L-arginine (10 mM) was done at the later phase, with or without 40 ng/mL TNFα stimulation. Untargeted metabolic tracing of 13C-isotopologue or untargeted mass spectrometry (MS) of L-arginine was performed within the following four groups: control, L-arginine, TNFα and TNFα/L-arginine (three replicates per group). (A) Schematic diagram delineating metabolic flux of 13C-isotopologue labelled L-arginine and the isotopologue distribution chart of the detected metabolites (Created with BioRender.com); (B) Metabolic pathway enrichment results showing affected pathways, comparing the groups L-arginine versus control and TNFα/L-arginine versus TNFα; (C) Schematic diagram combining the differentially expressed metabolites from MS data and expression of enzymes from RNA-seq results associated with purine metabolism pathway, comparing TNFα/L-arginine versus TNFα groups; (D) Expression of metabolites from MS results affected in purine metabolism among the four groups; (E) GO pathway enrichment results indicating significant difference in purine metabolism, comparing TNFα/L-arginine versus TNFα groups; (F) Expression of enzymes from RNA-seq results connected to purine metabolism among the four groups. Genes with an adjusted p value (p-adj) less than 0.05 were assigned as differentially expressed. Data are shown as mean±SE. Asterisks mark statistically significant difference (*p<0.05; **p<0.01), two-way ANOVA in (D). See also in online supplemental figure S10. ANOVA, analysis of variance.
Next, to further address the intracellular metabolic changes on L-arginine treatment and clarify the downstream metabolic pathways, we performed untargeted metabolomics analyses of osteoclasts stimulated with or without L-arginine and TNFα. As expected, pathway enrichment from metabolomic data confirmed an increase of arginine and proline metabolism in L-arginine treated osteoclasts (figure 5B). Most interestingly, the purine metabolism was enriched in TNFα+L-arginine treated cells when compared with TNFα stimulated cells (figure 5B). Production of hypoxanthine and inosine were increased, likely resulting from the boosted ATP production (figure 5C,D). Adenosine and adenine production were significantly decreased in TNFα+L-arginine treated cells. In alignment with the metabolomic results, RNA-seq data also demonstrated that purine metabolism associated pathways were significantly enriched in Gene Ontology database in TNFα+L-arginine treated cells (figure 5E). Genes coding enzymes associated with purine metabolism were also found changed accordingly (figure 5F). Especially, adenosine deaminase (ADA) and Nt5c1a were exclusively up-regulated by L-arginine treatment of TNFα-stimulated cells (figure 5F). Genes encoding enzymes in purine metabolism were also analysed on either putrescine, ornithine, or spermidine treatment, with or without TNFα stimulation (online supplemental figure S10A). TNFα stimulation promoted the expression of Ampd, Nt5cla, Pnp, Hprt and Xdh, while putrescine suppressed their expression. However, despite no effect on osteoclastogenesis at the low concentration (5 µm), spermidine enhanced the expression of Ampd, Nt5cla, Ada, Pnp and Xdh especially after TNFα stimulation (online supplemental figure S10B). Ornithine also enhanced the expression of Ampd and Ada after TNFα stimulation (online supplemental figure S10B). Collectively, these results revealed that L-arginine reprograms purine metabolism under inflammatory stimulation.
To further analyse the purine metabolism in osteoclastogenesis, we induced osteoclast differentiation and treated the cells with either inosine or hypoxanthine. Both inosine and hypoxanthine could suppress in vitro osteoclastogenesis especially after TNFα stimulation (figure 6A,B). By preventing inosine and hypoxanthine production with the ADA inhibitors (pentostatin or erythro-9-(2-hydroxy-3-nonyl) adenine), we could reverse the inhibitory role of L-arginine on osteoclast differentiation in the inflammatory milieu (figure 6C–E).
L-arginine attenuates inflammatory bone loss through controlling adenosine deaminase. (A) Schematic diagram of osteoclastogenesis with WT bone marrow cells, supplemented with 20 mM inosine at the later phase, with or without 40 ng/mL TNFα stimulation (above); Representative TRAP staining images (bottom left) and quantification of TRAP+ osteoclasts (bottom right) at day 3 (n=6/group, representative of three independent experiments). Scale bars: 100 µm; (B) Schematic diagram of osteoclastogenesis with WT bone marrow cells, supplemented with 1 mM hypoxanthine at the later phase, with or without 40 ng/mL TNFα stimulation (above); Representative TRAP staining images (bottom left) and quantification of TRAP+ osteoclasts (bottom right) at day 3 (n=5/group, representative of three independent experiments). Scale bars: 100 µm; (C–E) Schematic diagram of osteoclastogenesis with WT bone marrow cells (C), quantification of TRAP+ osteoclasts (D) and representative TRAP staining images (E) at day 3, supplemented with ADA inhibitors (20/40 µM Pentostatin or 2.5/5 µM erythro-9-(2-hydroxy-3-nonyl) adenine (EHNA)) at the later phase, with or without the treatment of 10 mM L-arginine at the later phase, stimulated by 40 ng/mL TNFα (n=3–6/group, representative of three independent experiments). Scale bars: 100 µm; (F) Arthritis induced by immunisation with an emulsion of complete Freund’s adjuvant and type II collagen (CII) into DBA/1 J mice, supplemented with 40 g/L L-arginine in drinking water after onset of arthritis (day 23). ADA inhibitor pentostatin (2 mg/g) was administrated intraperitoneally into the mice at an interval of 24 hours for successive 3 days every 2 weeks starting from day 23. Mice were sacrificed at day 43 after first immunisation; (G) Arthritis score in controls, CIA, CIA/L-arginine, pentostatin, CIA/pentostatin and CIA/L-arginine/pentostatin groups (n=4–5/group); (H, I) Representative images of TRAP staining (H) and quantification of tibial BV/TV, Tb. N, Tb. Th, Tb. Sp, Oc.S/BS and N.Oc./B.Pm. (I) in controls, CIA, CIA/L-arginine, pentostatin, CIA/pentostatin and CIA/L-arginine/pentostatin groups (n=4–5/group). Scale bars: 500 µm; Data are shown as mean±SEM. Asterisks mark statistically significant difference (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001), one-way ANOVA in (D), two-way repeated measures ANOVA in (G), two-way ANOVA in (A, B, I). ADA, adenosine deaminase; ANOVA, analysis of variance; CIA, collagen-induced arthritis.
Next, we analysed the effect of ADA inhibition in vivo in CIA mice supplemented with L-arginine (figure 6F). As previously, L-arginine significantly ameliorated arthritis, however pentostatin injection fully restored arthritis (figure 6G). Histomorphometry analysis of tibial bone indicated that L-arginine supplementation reduced osteoclasts and protected from inflammatory bone loss. However, ADA inhibition reverse the effects of L-arginine (figure 6H,I). Taken together, L-arginine induced purine metabolism mediates inhibition on inflammatory osteoclastogenesis in vitro and in vivo.
Altered arginine levels in RA patients and inhibition of human osteoclastogenesis by L-arginine
To investigate whether arginine levels are changed in RA patients, serum samples from 23 RA patients and 29 age-matched and gender-matched healthy controls were analysed by UPLC-mass spectroscopy (UPLC-MS), to determine the levels of amino acids involved in the urea cycle. Clinical characteristics from RA patients were documented in online supplemental table S1. Compared with healthy controls, arginine levels were significantly increased in the serum of RA patients, while ornithine was decreased, and citrulline and proline levels were unchanged (figure 7A). Moreover, in RA patients with mild to moderate disease activity (DAS28 score <5.2 units), arginine levels were significantly correlated with disease severity (figure 7B). We also tested these amino acid levels in another cohort of pre-RA patients.35 Interestingly, in pre-RA patients subsequently developing arthritis (at-risk progressors), D-arginine levels were enhanced compared with healthy controls, while ornithine and D-proline were significantly decreased, and citrulline levels were unaltered (figure 7C). Collectively, arginine expression was significantly elevated in RA and pre-RA patients, implying a dysregulation of arginine metabolism even at the early stage of the disease.
Altered arginine metabolism in rheumatoid arthritis (RA) patients and L-arginine inhibits human osteoclastogenesis. (A) Levels of arginine, ornithine, proline and citrulline in the serum collected from RA patients and healthy individuals (n=23 for RA; n=29 for healthy controls); (B) Correlation between each amino acid and DAS28 scores in mild to moderate RA patients (2.6<DAS28 score<5.2); (C) Expression level of arginine, ornithine, proline and citrulline in the serum collected from pre-RA patients subsequently developing arthritis (at-risk progressors) and healthy individuals (n=25 for Control; n=39 for At-risk progressors); (D) Schematic diagram of the osteoclastogenesis procedure with peripheral blood mononuclear cell (PBMC) isolated from healthy donors, supplemented or not with 10 mM L-arginine, with or without stimulation with 20 pg/mL TNFα; (E–F) Quantification of TRAP+osteoclasts (E) and representative TRAP staining images (F) in mature human osteoclast at day 7, supplemented or not with 10 mM L-arginine, with or without stimulation with 20 pg/mL TNFα. Scale bars: 100 µm; (G) Expression of osteoclast-associated genes in human PBMC-derived osteoclasts at day 7, supplemented with 10 mM L-arginine, with or without 20 pg/mL human TNFα stimulation. Data are shown as mean±SEM. Asterisks mark statistically significant difference (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001), Student’s t-test in (A, C), two-way ANOVA in (E, G). ANOVA, analysis of variance.
Finally, the role of L-arginine in osteoclasts was also investigated in human cells. Peripheral blood mononuclear cell (PBMC) derived-osteoclasts were treated with 10 mM L-arginine and stimulated with 20 pg/mL TNFα (figure 7D). A significant reduction of osteoclast numbers and osteoclast marker expression was observed after L-arginine treatment, both in cells that were stimulated with MCSF and RANKL and in those that were additionally challenged with TNFα (figure 7E–G). Hence, L-arginine also suppresses human osteoclastogenesis.
Discussion
In the present study, we demonstrated that L-arginine not only dampened inflammation in experimental arthritis, but also reduced bone loss. L-arginine especially suppressed osteoclastogenesis enhanced by proinflammatory factors such as TNFα. Metabolically, TNFα promoted glycolysis and blocked oxidative phosphorylation, resulting in accelerated osteoclastogenesis, whereas L-arginine restored oxidative phosphorylation and promoted ATP production. Subsequently, L-arginine reprogrammed purine metabolism by elevating hypoxanthine and inosine levels, which essentially mediated the inhibitory function of L-arginine on both systemic inflammation and also inflammatory osteoclastogenesis in vitro and in vivo. We also demonstrated that arginase-1 mediates the effects of L-arginine on osteoclast differentiation, which was transcriptionally regulated by c-Jun (figure 8). Taken together, we uncovered a so far unrecognised function of L-arginine in perturbing osteoclast metabolism and inhibiting the function of these bone-resorbing cells.
In inflammatory condition, TNFα promoted c-Jun expression, enhanced glycolysis and blocked oxidative phosphorylation, leading to accelerated osteoclastogenesis. In L-arginine supplementation, L-arginine inhibited c-Jun expression, restored oxidative phosphorylation and promoted ATP production. L-arginine also elevated hypoxanthine and inosine levels by reprogramming purine metabolism, which further mediate the inhibitory role of L-arginine in inflammatory osteoclastogenesis. ADA, adenosine deaminase.
L-arginine is a natural amino acid that is enriched in nuts, seeds and whole grains, food components, which have been shown to beneficially influence arthritis.36 In our study, L-arginine demonstrated a reduction in joint inflammation in three different arthritis models. Apart from inflammation, L-arginine potently inhibited inflammatory osteoclastogenesis in vivo and in vitro. L-arginine treatment led to a decrease of bone destruction and osteoclast numbers in three different arthritis models. Moreover, L-arginine reduced osteoclastogenesis in cells from WT and hTNFtg mice, as well as from synovial inflammatory macrophages. Despite that L-arginine supplementation elicits broad suppression on osteoclast formation, bone marrow derived monocytes cannot differentiate into osteoclasts in Arg-free medium, with or without TNFα. These results indicated that the effect of L-arginine on osteoclastogenesis might be due to an autoregulatory mechanism of L-arginine to determine osteoclast fate,37 where the affinity and the catalytic efficiency of the key enzymes, arginases and NOSs, depend on the concentration of L-arginine.38–40
L-arginine mitigated osteoclastogenesis induced by TNFα, which boosts RANKL-induced NF-κB and MAPK-AP-1 activation and thereby promotes NFATc1-mediated osteoclastogenesis.5 41 In macrophages and osteoclasts, L-arginine and TNFα signalling seems interconnected. Indeed, in alternatively-activated macrophages (AAM), TNFα selectively diminished Arg-1 expression in a time-dependent manner.42–44 TNFα also selectively reduces AAM gene transcription via induction of JunB and c-Jun.42 C-Jun was previously shown to downregulate Arg1 in macrophages.33 We could confirm that TNFα promoted c-Jun protein expression in osteoclasts, which transcriptionally restrained Arg-1 expression by modifying chromatin methylation. L-arginine was sufficient to interfere with the TNFα-cJun-Arg1 axis, resulting in reduced c-Jun production and increased Arg1 expression, hence decreasing osteoclastogenesis. Arg-1 is known as negative regulator of osteoclast differentiation.45 Accordingly, treatment with pegylated recombinant arginase-1 (pegARG1) reduced joint swelling and inhibited osteoclastogenesis in arthritic mice.46 In the present study, we also observed that arginase activity was elevated in vivo after L-arginine supplementation, contributing to the suppression of osteoclastogenesis and bone loss. Moreover, with the two strains of Arg-1 deficient cells, we demonstrated that Arg-1 mediates the inhibition on osteoclastogenesis by L-arginine.
Reprogramming of the energy metabolism of immune cells influences cell fate and function of immune cells. Glycolysis is mostly associated with proinflammatory functions, whereas oxidative phosphorylation often mediates repression of inflammation.47 48 Previous work has suggested that proinflammatory cytokines such as TNFα and IL-1β profoundly suppress oxidative phosphorylation in macrophages.43 49 In accordance, we could as well show that TNFα facilitated glycolysis but blocked oxidative phosphorylation in osteoclast progenitors. Moreover, the induction of oxidative phosphorylation by L-arginine was previously shown in T cells where L-arginine skewed human activated CD4+ T cells from glycolysis towards oxidative phosphorylation.50 In the present study, L-arginine reversed the blockade of TNFα on oxidative phosphorylation, leading to reduced osteoclastogenesis. Interestingly, glucose consumption is highly decreased after L-arginine supplementation, whereas the level of glycolysis, OXPHOS, and ATP production remain stable. L-arginine is converted into 2-oxoglutarate (also known as α-ketoglutarate) as a glucose precursor, which then directly participates in the energy metabolism through OXPHOS.51 Moreover, it is likely that inosine that is altered in our models, might be an alternative carbon source and energy provider for osteoclasts.52
We also observed that L-arginine significantly increased the efficiency of ATP synthesis in osteoclasts by switching the respiratory dependence from glycolysis to oxidative phosphorylation. Increased ATP production might contribute to the suppression of osteoclastogenesis through accelerated actin dynamics, which destroys the proper F-actin aggregation during osteoclast differentiation and its resorption activity.53 Moreover, we observed that L-arginine could inhibit osteoclastogenesis in non-inflammatory conditions, while the OXPHOS and ATP production were not significantly increased as in inflammatory stimulation. It is possible that L-arginine alters osteoclastogenesis via different pathways under non-inflammatory conditions. For instance, L-arginine along with its downstream metabolites may control apoptosis and efferocytosis, interfering with the cell fate.54 55 Nevertheless, it would require further studies to analyse L-arginine effect on osteoclast in non-inflammatory conditions.
Intriguingly, we observed that L-arginine altered purine metabolism, substantially increasing inosine and hypoxanthine and decreasing adenosine after TNFα stimulation. To date, only a few reports have interrogated the role of purine metabolism on bone.56 Elevated adenosine was shown to increase osteoclastogenesis while ADA that catalyses the conversion of adenosine to inosine significantly suppressed osteoclast differentiation.57 Likewise, we used ADA inhibitors in our study to block adenosine metabolisation into inosine/hypoxanthine on L-arginine treatment. ADA inhibitors reversed the effect of L-arginine on osteoclastogenesis even in the context of TNFα stimulation in vitro and in vivo. Moreover, we also observed that the metabolites produced by arginase might have different functions on osteoclast formation and purine metabolism. Putrescine significantly diminished osteoclast differentiation but did not affect purine metabolism, while spermidine promoted purine metabolism especially after TNFα stimulation, but did not affect osteoclasts. Hence, the specific effects of L-arginine on osteoclast formation and purine metabolism might result from the overall combination of Arg-1 catalytic downstream metabolites.
Several studies have shown that systemic arginine levels were decreased in RA patients,58 59 while others presented elevated arginine levels in the serum or local synovial fluid of RA patients.46 60 In our study, we detected an increased arginine levels in RA patients. Elevated arginine levels correlated with disease severity in mild to moderate patients but not in severely active RA patients, suggesting a compensatory arginine elevation in mild RA patients but a dysregulated arginine metabolism in severe RA. In accordance, levels of arginine were found significantly increased in Steinbrockers classs I–III RA patients, but low in Steinbrockers class IV.58 Due to the technical limitation and consistent with previous studies,58–60 only arginine, but not the distinct arginine isomers were detected in RA patients. In our pre-RA patient cohort, an enhanced D-arginine was observed while L-arginine levels were undetected, suggesting a dysregulation of arginine metabolism at the early stage of the disease. D-arginine is a competitor for L-arginine bioavailability, which could inhibit L-arginine uptake.50 58 Thus, L-arginine supplementation may help to enhance the L-arginine bioavailability and function as a therapeutic strategy against RA inflammation and bone destruction. Future investigations are currently investigating the isomers and their actual uptake in RA patients.
In conclusion, our observations show that L-arginine inhibits arthritis, inflammatory osteoclastogenesis and inflammation-induced bone loss by reprogramming amino acid metabolism of osteoclasts. Hence, therapeutic supplementation of L-arginine may be a dietary strategy to inhibit inflammation in arthritis and protect from inflammatory bone loss.
Data availability statement
Data are available on reasonable request.
Ethics statements
Patient consent for publication
Ethics approval
This study involves human participants and was approved by Ethics Committee of Renji Hospital and Swedish Ethical Review Authority. Participants gave informed consent to participate in the study before taking part. All animal experiments were discussed and approved by the University of Friedrich-Alexander-Universität Erlangen-Nürnberg ethics committee and carried out in accordance with protocols approved by the German law.
Acknowledgments
We thank Wolfgang Baum for providing hTNFtg+/– mice and the generation of K/BxN serum. We also thank Daniela Weidner for analysing μCT samples. We would also like to thank Christine Zech, Barbara Happich, Nicole Berndt, Franceska Jelas and Jule Lindörfer for their excellent technical assistance. We thank Uwe Appelt and Markus Mroz of the Core Unit Cell Sorting and Immunomonitoring facility for their technical support in synovial cell sorting.
References
Supplementary materials
Supplementary Data
This web only file has been produced by the BMJ Publishing Group from an electronic file supplied by the author(s) and has not been edited for content.
Footnotes
Handling editor Thomas Pap
SC and YL contributed equally.
Correction notice This article has been corrected since it published Online First. Reference 53 has been corrected. In December 2023, this paper was resupplied as open access.
Contributors SC and AB designed the study. SC and YL performed the in vitro experiments. SC, RS, XiaM and JW performed in vivo experiments. XinM, YL and XC contributed to patient recruitment and data collection. MF, CL and MK contributed to the bioinformatic analysis. US-S contributed to the TEM analysis. VT, CL and AG contributed to the metabolomic analysis. Xianyi M and KK helped with metabolic assays. MMZ and AK contributed to pre-RA data. SC and AB wrote the manuscript. GS, US and AB supervised the study and edited the manuscript. AB is the author responsible for the overall content, and had access to all data and controlled the decision to publish.
Funding This study was supported by the Interdisciplinary Center for Clinical Research grant J90; J76; and A77, the German Research Foundation grant BO-3811/5-1; BO-3811/6-1; FOR2886 TP02, the Collaborative Research Centre 1181 project A01, and the European Research Council Consolidator Grant LS4-ODE and the Synergy Grant 4D Nanoscope. Part of the data from this project has been presented previously at EULAR 2020 and EULAR 2022. The authors have declared that no conflict of interest exists.
Competing interests None declared.
Patient and public involvement Patients and/or the public were not involved in the design, or conduct, or reporting, or dissemination plans of this research.
Provenance and peer review Not commissioned; externally peer reviewed.
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