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Department of Animal and Avian Sciences, University of Maryland, College Park, MD 20742
* To whom correspondence should be addressed. E-mail: bhumphre{at}umd.edu.
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ABSTRACT |
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 TOP ABSTRACT IntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Introduction |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Alterations in energy status at the whole-animal level affect lymphocyte function, proliferation, and development (11–13). Cellular anabolic processes associated with lymphocyte proliferation, such as protein synthesis, nucleotide synthesis, and cation transport, are ATP demanding (14), and it has been shown that protein synthesis and nucleotide synthesis are most sensitive to ATP supply (15). Glycolysis is the major ATP-generating pathway in proliferating lymphocytes (16,17) and inhibiting ATP generation via glycolysis adversely affects lymphocyte proliferation (18). Glutaminolysis and ß-oxidation of fatty acids also provide energy in proliferating lymphocytes (16). Because protein synthesis is an energy-consuming process, depleted energy status in lymphocytes results in reduced membrane-bound IL-2 receptor expression and soluble IL-2 receptor secretion (19). This highlights the need for lymphocytes to maintain adequate energy supplies to sustain development and function when metabolic perturbations occur at the organism level.
In chickens, energy metabolism is altered at the whole-animal level during the embryonic to posthatch transition. During embryogenesis, lipid serves as the primary energy source, whereas at hatch, feeding a carbohydrate-rich grain-based diet induces metabolic adaptations over the first 2 wk posthatch that promote carbohydrate utilization (20–26). Development of the adaptive immune system is also critical during this period, because chicken lymphocyte development in primary immune tissues is initiated during embryogenesis and continues for several weeks after hatch (27,28). Developing B lymphocytes proliferate extensively in the bursa during the first few weeks after hatch and migrate to peripheral tissues to form a self-renewing B cell pool (27). In contrast, T cell progenitors enter the thymus in 3 waves during the embryonic days (e)2 of life (e6.5, e12, and e18) where they proliferate up to 3 wk within the thymic cortex prior to their emigration to peripheral tissues (28,29). The effect of altered primary energy-substrate metabolism, at the whole-animal level, on developing B and T lymphocyte metabolism is unknown. Therefore, the objective of these experiments was to determine the ability of developing chicken B and T lymphocytes to acquire and metabolize glucose, glutamine, and fatty acids to generate energy. Three experiments were performed to measure: 1) the mRNA abundance for genes involved in glucose, glutamine, and fatty acid metabolism in developing B and T lymphocytes; 2) the activity of enzymes that catalyze glycolysis, glutaminolysis, and ß-oxidation in developing B and T lymphocytes; 3) serum glucose, glutamine, and nonesterified fatty acid (NEFA) concentrations; and 4) glucose uptake by developing B and T lymphocytes.
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Materials and Methods |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Experimental design. Three experiments were designed to characterize glucose, glutamine, and fatty acid metabolism in bursacytes and thymocytes from embryonic and neonatal chicks. On the day of incubation, eggs were randomly assigned to each sampling time point to achieve similar average egg weights. In all experiments, cells were isolated from the bursa and thymus at selected time points from e17 to neonatal day (d) 14. In Expt. 1, mRNA abundance of genes involved in glucose, glutamine, and fatty acid metabolism in bursacytes and thymocytes were measured on e17, e20, d1, d7, and d14 (n = 5 per time point). In Expt. 2, mRNA abundance and enzyme activity in bursacytes and thymocytes were measured on e20, d1, d3, and d7 (n = 6 per time point). In Expt. 3, glucose uptake by bursacytes and thymocytes was measured on d1, d7, and d14 (n = 6 per time point).
Lymphocyte isolation.
The bursa and the thymus were isolated from each chick and minced with forceps in RPMI 1640 and strained through a 70-µm nylon cell strainer (BD Falcon). Lymphocyte purity of bursacyte and thymocyte populations was analyzed using a FACSCalibur flow cytometer (BD Biosciences). FITC-conjugated mouse anti-chicken 
chain and FITC-conjugated mouse anti-chicken ChT1 (SouthernBiotech) were used as markers for B and T lymphocytes, respectively. Lymphocyte viability was determined by propidium iodide staining. Bursacyte and thymocyte aliquots (2 x 106 cells in Expt. 1; 5 x 106 cells in Expt. 2) were immediately snap-frozen in liquid nitrogen and stored at –80°C until further analysis. In Expt. 3, bursacytes and thymocytes (1 x 106) were cultured in RPMI 1640 to measure glucose uptake.
Quantitation of mRNA by real-time PCR.
In Expts. 1 and 2, total RNA was isolated from bursacytes and thymocytes using the NucleoSpin RNA II Total RNA Isolation kit (Macherey-Nagel) and optical density at 260 nm was used to determine RNA concentrations. Total RNA (0.5 µg) was reverse transcribed as described (22). Intron-spanning gene-specific primers were designed to either cloned or predicted chicken genes (Supplemental Table 1) using the Beacon Designer 4 software (Premier Biosoft International). PCR products were separated by agarose gel electrophoresis, gel purified using the MinElute gel extraction kit (Qiagen), and sequenced (Gene Gateway) to ensure amplification specificity. Quantitative real-time PCR was performed using the iQ SYBR Green Supermix (Bio-Rad). Duplicate reactions for each sample contained 1 µL of the 1:2 diluted reverse transcription reaction and 0.3 µmol/L of each gene-specific primer in a total reaction volume of 12.5 µL. Amplification and detection of specific PCR products were performed using the iCycler iQ Multicolor Real-Time PCR Detection system (Bio-Rad). Thermal cycling conditions for real-time PCR were 95°C for 3 min followed by 40 cycles of denaturing at 95°C for 15 s, annealing for 30 s, and extension at 72°C for 30 s. The optimum annealing temperature for each primer pair is provided in Supplemental Table 1. After 40 cycles, the specificity of each PCR reaction was confirmed by melt-curve analysis. Thermal cycling conditions for melt-curve analysis were 95°C for 1 min, 55°C for 1 min, followed by a linear increase in temperature of 0.5°C/10 s up to 95°C while continuously monitoring fluorescence. The change in mRNA abundance from e17 was calculated using the 
- equation (31) with certain modifications. The amplification efficiency of each reaction was obtained using the LinRegPCR software (32). Target gene mRNA abundance was normalized by geometric averaging of hypoxanthine phosphoribosyltransferase-1, glyceraldehyde phosphate dehydrogenase, TATA box binding protein, ß2-microglobulin, ß-actin, acetyl CoA carboxylase, and fatty acid translocase raw-non-normalized values using the geNorm software (33). Data are presented as the normalized fold-change in mRNA abundance of a gene relative to its mRNA abundance on e17 or e20 for Expt. 1 or 2, respectively.
Enzyme assays. In Expt. 2, bursacytes and thymocytes (5 x 106) were lysed using a cell-lysis buffer with nonionic detergent (1% v:v NP-40 and 0.9% w:v NaCl in PBS). Cell lysate protein content was measured using the Quick Start Bradford Protein Assay kit (Bio-Rad). HK activity was measured as described (34). Carnitine palmitoyl transferase-1 (CPT-1) activity was measured as described (35) with minor modifications. Assay conditions were: 116 mmol/L Tris, pH 8.0, 1.1 mmol/L EDTA, 0.05 mmol/L palmitoyl-CoA, 7.5 mmol/L carnitine, 0.12 mmol/L 5,5'-dithiobis (2-nitrobenzoic acid) (DTNB), 50 µL cell lysate in a reaction volume of 200 µL. Glutaminase (GA) activity was measured as described (36) with minor modifications. Assay conditions were: 80 mmol/L glutamine, 0.5 mmol/L potassium phosphate, 10 µL of cell lysate, 100 µL Tris-hydrazine buffer, pH 8.5 (100 mmol/L Tris-base, 5 mmol/L EDTA, 10 mmol/L MgCl2, 400 µmol/L hydrazine dihydrochloride), 28 mmol/L NAD, and 3 units of glutamate dehydrogenase in a reaction volume of 150 µL. Results of all enzyme assays are presented as nmol of substrate produced per mg protein per min.
Serum substrate concentration. In Expt. 2, blood was collected by cardiac puncture on e20, d1, d3, and d7. Blood was allowed to clot at 25°C for 90 min, centrifuged at 4000 x g; 5 min at 4°C, and then serum was collected and stored at –80°C until further analysis. Serum glucose was measured using the GAHK-20 glucose assay kit (Sigma). Serum glutamine was measured as described (37). Serum NEFA concentration was measured using the NEFA C assay kit (Wako Chemicals).
Glucose uptake.
In Expt. 3, glucose uptake by bursacytes and thymocytes was measured using 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino]-2-deoxy-D-glucose (2-NBDG; Molecular probes). Bursacytes and thymocytes (106) were cultured in RPMI 1640 + 10% fetal bovine serum with or without 20 µmol/L 2-NBDG for 0, 30, and 60 min at 37°C with 5% CO2. Samples incubated without 2-NBDG were used to measure background fluorescence. B and T lymphocyte glucose uptake was determined by measuring fluorescence of chain and ChT1 positive cells, respectively. Mean fluorescence intensity (MFI) due to 2-NBDG uptake was measured for 10,000 gated events for each replicate using a FACSCalibur flow cytometer (BD Biosciences). Background fluorescence was subtracted from MFI of each sample. Data are presented as background-corrected MFI.
Statistical analysis. Dependent variables were analyzed by general linear model procedure (JMP) using a 1-way ANOVA. Prior to analysis, data on mRNA abundance was assessed for homogeneity of variance by Levene's test and was log-transformed when significant (P < 0.05). Data are reported as nontransformed means and pooled SEM. When main effects were significant (P < 0.05), means were compared by Tukey's means comparison. Data on glucose uptake within an incubation time point were compared by least-squares means contrasts. Pearson correlation coefficients between selected dependent variables were calculated by the multivariate procedure using JMP.
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Results |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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chain and ChT1 antigens were used as lineage-specific markers for developing B and T cells, respectively (38,39). Flow cytometric analysis of bursacyte and thymocyte populations showed that 90% of bursacytes were chain positive and 95% of thymocytes were ChT1 positive. The viability of both cell populations exceeded 98%. Gene expression. The mRNA abundance of genes involved in glucose, glutamine, and fatty acid metabolism changed in both bursacytes and thymocytes during the first 2 wk after hatch. Bursacyte glucose transporter-3 (Glut-3) mRNA abundance increased 1.5-fold from d1 to d14, while Glut-1 mRNA abundance decreased 50% from e17 to e20 (P < 0.05; Fig. 1A). Bursacyte hexokinase-1 (HK-1) mRNA abundance reached maximum levels on e20 (Fig. 1B). Thymocyte Glut-1 and Glut-3 mRNA abundance increased after hatch (P < 0.05; Fig. 1C), and HK-1 mRNA abundance increased 2.5-fold from d7 to d14 (P < 0.05; Fig. 1D). Bursacyte sodium coupled neutral amino acid transporter-2 (SNAT-2) mRNA abundance increased 1.5-fold from e20 to d7 (P < 0.05; Fig. 2A) and GA mRNA abundance increased 2.5-fold from e17 to d7 (P < 0.05; Fig. 2B). However, bursacyte SNAT-1 mRNA abundance did not change over time. Thymocyte SNAT-2 mRNA abundance increased 4-fold from d1 to d7 (P < 0.05), SNAT-1 mRNA abundance increased 2-fold from e20 to d7 (P < 0.05), and GA mRNA abundance increased 4-fold from e17 to d7 (P < 0.05; Fig. 2C,D). Bursacyte CPT-1 mRNA abundance decreased 80% from e17 to e20 (P < 0.05; Fig. 3A). Thymocyte CPT-1 mRNA abundance increased 4-fold from e17 to d1 and decreased 50% from d1 to d7 (P < 0.05; Fig. 3B). In Expt. 2, the mRNA abundance of genes (same as in Expt. 1) did not change significantly from e20 to d7 in either bursacytes or thymocytes (data not shown).
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Glucose uptake. Glucose uptake by bursacytes and thymocytes increased with each d to maximum levels on d14 (P < 0.05). In bursacytes, glucose uptake on d14 was higher than uptake on d1 and d7 at 30 and 60 min of incubation (P < 0.05; Fig. 4A). In thymocytes, glucose uptake was higher on d7 than on d1 at either 30 or 60 min of incubation (P < 0.05; Fig. 4B). Similarly, glucose uptake by thymocytes on d14 was higher than on d7 at either 30 or 60 min of incubation (P < 0.05).
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Discussion |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Developing B and T lymphocytes showed increased transcription of glucose transporter genes beginning at the time of hatch. Glut-3 mRNA abundance, an inducible isoform in chickens (41), increased from d1 to d14 in B and T lymphocytes. Glut-3 protein levels were not determined in these experiments, because an antibody is not yet available to this chicken protein, but the increase in glucose uptake and Glut-3 mRNA abundance with age suggests that Glut-3 may be the primary transport protein responsible for increasing glucose uptake in developing B and T lymphocytes after hatch. In developing T lymphocytes, however, Glut-1 mRNA abundance also increased after hatch and this isoform is also likely to play an important role in developing T lymphocyte glucose metabolism. The increase in transcription of both Glut-1 and Glut-3, 2 high-affinity glucose transporters (42), suggests that developing T lymphocytes may have an increased metabolic need for glucose beginning at hatch. In the thymus, high Glut expression and glucose uptake is characteristic of proliferating CD4+CD8+ thymocytes (43,44). Therefore, the apparent increased metabolic need for glucose by developing B and T lymphocytes after hatch suggests increased rates of cell proliferation.
HK-1 mRNA abundance and total HK activity increased in both developing B and T lymphocytes following hatch. Upon entry into the cell, glucose can enter into 1 of several alternative metabolic pathways and this is regulated by different HK isozymes. HK-2 provides glucose-6-phosphate for glycolysis, while HK-2 and HK-3 provide glucose-6-phosphate for either glycogen synthesis or the pentose phosphate pathway (45). There are several lines of evidence to suggest that increased HK activity in developing lymphocytes is associated with increased rates of glycolysis in developing chicken lymphocytes. First, lymphocytes have low activities of glycogen synthase, suggesting low to no internal glycogen stores and, consequently, incorporation of glucose carbon into glycogen in this cell type is minimal (46). Second, malic enzyme provides the majority of reducing equivalents required for fatty acid synthesis in chickens due to an inactive pentose phosphate pathway at hatch (47). Taken together, it is likely that the increased HK activity in developing B and T lymphocytes after hatch reflects an increased flux of glucose carbon through the glycolytic pathway for energy generation.
Some of the metabolic markers for glucose transport and metabolism in developing lymphocytes were correlated with the serum glucose concentration. Glucose availability is sensed by cells via kinases that regulate the transcription of glycolytic and lipogenic genes (48). Glucose sensing is well studied in lipogenic tissues, such as liver and adipose, where increased glucose availability results in the activation of carbohydrate response element binding protein (ChREBP), a transcription factor, which induces the transcription of lipogenic genes (49). The mRNA abundance of acetyl CoA carboxylase-
and sterol regulatory element binding protein-1, 2 lipogenic genes that are transcriptional targets of ChREBP, did not change in developing B and T lymphocytes (data not shown). This suggests that ChREBP is unlikely to integrate the increased serum glucose concentration to glucose metabolism in developing lymphocytes. Similar to ChREBP, AMP-activated protein kinase (AMPK) is responsive to glucose availability and regulates glucose metabolism in mammals (50) and AMPK subunit genes are expressed in developing chicken B and T lymphocytes (data not shown). It remains to be determined whether the AMPK has a role in glucose-mediated adaptation in glycolytic metabolism of developing chicken B and T lymphocytes.
Glutamine is a preferred substrate for system A-encoding isoforms SNAT-1 and SNAT-2 (51) and the level of activity of these transporters is high in proliferating cells. During embryogenesis and for several weeks posthatch, developing B and T lymphocytes proliferate extensively within their respective primary immune tissue, doubling approximately twice daily (29,52). In this study, SNAT-2 mRNA abundance increased in developing B lymphocytes while both SNAT-1 and SNAT-2 mRNA abundance increased in developing T lymphocytes, suggesting heightened metabolic activity for both populations (53,54). Consequently, the increase in transcription of SNAT isoforms and the direct correlation of SNAT-2 mRNA abundance with system A activity (55) suggests that developing B and T lymphocytes increase their ability to acquire glutamine and other neutral amino acids after hatch.
GA catalyzes glutamine to glutamate and ammonia and the activity of this enzyme is correlated with glutamine utilization in lymphocytes (56). Glutamate is partially oxidized via the citric acid cycle to provide energy or is metabolized to provide precursors for protein, fatty acid, and nucleic acid biosynthesis (57). About 40% of glutamate produced by GA is oxidized to CO2 via the tricarboxylic acid cycle in resting and proliferating lymphocytes (46,58). Increased GA enzyme activity in developing B lymphocytes suggests that glutamine is an important energy source for chicken B lymphocytes during the first week after hatch. Indeed, feeding neonatal chicks diets supplemented with glutamine enhances bursa maturation (59). In contrast, developing T lymphocyte GA activity did not change despite an increase in GA mRNA abundance. Glutamine has been shown to be an important energy substrate for T lymphocytes (46,58), so perhaps glutaminolysis in this cell type has already reached maximum by hatch.
In these experiments, there were no significant changes in the transcription of lipogenic and lipolytic genes, nor did CPT-1 enzyme activity change in either lymphocyte population. However, serum NEFA concentrations increased 3-fold by 1 wk posthatch and is likely due to the rapid absorption of lipids from both the diet and yolk sac remnant (60). Though the NEFA concentration increased in serum, lymphocytes did not appear to increase their ability to utilize lipid as an energy source, suggesting that increased ß-oxidation is not associated with developing lymphocyte metabolism posthatch. This is similar to fatty acid metabolism in mammalian proliferating lymphocytes, because these cell types do not increase rates of ß-oxidation despite an elevated serum NEFA concentration caused by fasting (16). However, activated mammalian lymphocytes increase fatty acid uptake (16) and this is related to an increased NEFA demand for plasma membrane synthesis (61). In these experiments, fatty acid transporter mRNA abundance did not change (data not shown) and markers associated with plasma membrane synthesis were not analyzed, so the utilization of NEFA for plasma membrane synthesis could not be evaluated.
Several hematopoietic cytokines and developmental ligands associated with lymphocyte development regulate their energy metabolism, particularly glucose metabolism. For example, IL-7 and Notch-1 increase glucose metabolism in differentiating thymocytes (3,10). This suggests that changes in either IL-7, Notch-1, or other cytokines and ligands may be contributing to the changes in developing B and T lymphocyte energy metabolism. Indeed, IL-7 receptor mRNA abundance increases after hatch in developing lymphocytes (data not shown), indicating that these cells may be more sensitive to IL-7 availability during this period of increased glucose metabolism. In addition, endocrine factors, such as insulin, also regulate lymphocyte metabolism (62) and their levels are altered during the embryonic to posthatch transition (63,64). Therefore, it is likely that cytokines, developmental ligands, and endocrine factors are contributing to altered energy metabolism in developing lymphocytes after hatch.
Developing lymphocytes increase their glucose transport and metabolism to generate energy during the first 2 wk posthatch. Similarly, developing nonimmune tissues adapt to oxidize carbohydrates by increasing hexose transport (24,65) and the transcription of genes involved in carbohydrate metabolism (22,66). This suggests that metabolic responses in B and T lymphocytes are similar to those occurring at the whole-animal level during this life-stage transition. Consequently, factors that regulate metabolic adaptation at the whole-animal level may also regulate developing lymphocyte energy metabolism. Identifying these regulatory mechanisms may help to promote lymphocyte development and maturation, which is vital for adaptive immune responses to pathogens in neonatal chicks.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Abbreviations used: AMPK, AMP-activated protein kinase; ChREBP, carbohydrate response element binding protein; CPT-1, carnitine palmitoyl transferase-1; d, neonatal day; e, embryonic day; Glut-1 and -3, glucose transporter-1 and -3; GA, glutaminase; HK-1, hexokinase-1; MFI, mean fluorescence intensity; 2-NBDG, 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino]-2-deoxy-D-glucose; NEFA, nonesterified fatty acid; SNAT-1 and -2, sodium coupled neutral amino acid transporter-1 and -2.
Manuscript received 18 September 2006. Initial review completed 23 October 2006. Revision accepted 14 November 2006.
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