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Journal of Animal Science - Animal Physiology

Technical note: A procedure to estimate glucose requirements of an activated immune system in steers

 

This article in JAS

  1. Vol. 94 No. 11, p. 4591-4599
     
    Received: June 29, 2016
    Accepted: Aug 10, 2016
    Published: October 7, 2016


    1 Corresponding author(s): baumgard@iastate.edu
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doi:10.2527/jas.2016-0765
  1. S. K. Kvideraa,
  2. E. A. Horsta,
  3. M. Abuajamieha,
  4. E. J. Mayorgaa,
  5. M. V. Sanz Fernandeza and
  6. L. H. Baumgard 1a
  1. a Department of Animal Science, Iowa State University, Ames 50011

Abstract

Infection and inflammation impede efficient animal productivity. The activated immune system ostensibly requires large amounts of energy and nutrients otherwise destined for synthesis of agriculturally relevant products. Accurately determining the immune system’s in vivo energy needs is difficult, but a better understanding may facilitate developing nutritional strategies to maximize productivity. The study objective was to estimate immune system glucose requirements following an i.v. lipopolysaccharide (LPS) challenge. Holstein steers (148 ± 9 kg; n = 15) were jugular catheterized bilaterally and assigned to 1 of 3 i.v. treatments: control (CON; 3 mL saline; n = 5), LPS-administered controls (LPS-C; E. coli 055:B5; 1.5 mg/kg BW; n = 5), and LPS + euglycemic clamp (LPS-Eu; 1.5 mg/kg BW; 50% dextrose infusion to maintain euglycemia; n = 5). In LPS-Eu steers, postbolus blood samples were analyzed for glucose every 10 min. Dextrose infusion rates were adjusted to maintain euglycemia for 720 min. All steers were fasted during the challenge. Samples for later analysis were obtained at 180, 360, 540, and 720 min relative to LPS administration. Rectal temperature was increased ∼0.5°C in both LPS treatments relative to CON steers (P = 0.01). Steers in both LPS treatments were hyperglycemic for ∼3 h postbolus; thereafter, blood glucose was markedly decreased (30%; P < 0.01) in LPS-C relative to both CON and LPS-Eu steers. A total of 516 ± 65 g of infused glucose was required to maintain continuous euglycemia in LPS-Eu steers. Circulating insulin increased in LPS-C and LPS-Eu steers relative to CON (∼70% and ∼20 fold, respectively; P < 0.01). Circulating NEFA increased similarly with time for both CON and LPS-C compared to LPS-Eu steers (∼43%; P < 0.01). Plasma L-lactate and LPS binding protein increased (∼198 and ∼90%, respectively; P < 0.01) and ionized calcium decreased (18%; P < 0.01) in both LPS treatments relative to CON steers. Circulating white blood cells decreased initially in LPS-Eu and LPS-C relative to controls (180 min; 85%) followed by a progressive increase with time (P = 0.02). Blood neutrophils followed the same pattern; however, at 720 min, neutrophils were decreased in LPS-Eu compared to LPS-C, resulting in a decreased neutrophil-to-lymphocyte ratio (54%; P = 0.03). The large amount of glucose needed to maintain euglycemia indicates extensive repartitioning of nutrients away from growth and the importance of glucose as a fuel for the immune system.



INTRODUCTION

Infection and inflammation have negative economic consequences on animal agriculture due to decreased production, inefficient feed utilization, poor reproduction, and increased health care costs. An activated immune system demands a large amount of energy and nutrients (Lochmiller and Deerenberg, 2000; Johnson, 2012), which reprioritizes the hierarchy of nutrient partitioning away from productive purposes. For instance, glucose homeostasis is markedly disrupted during an endotoxin (e.g., lipopolysaccharide, LPS) challenge, and hypoglycemia and hyperlactemia are characteristic hallmarks (Filkins, 1978; McGuinness, 2005; Michaeli et al., 2012). In addition, in vitro experiments suggest activated immune cells experience a substantial increase in glucose consumption and utilize glucose as their primary source for the generation of energy, biosynthetic precursors, and signaling intermediates (Calder et al., 2007; Palsson-McDermott and O’Neill, 2013). The extent of immune cell glucose consumption in vivo is difficult to assess due to the ubiquitous and fluctuating distribution of immune cells and organ-specific changes in insulin sensitivity; however, better understanding the impact on energy status has practical implications to animal agriculture as glucose is obviously an important fuel for productive purposes.

Despite the increase in glucose requirements, an activated immune response is often accompanied by anorexia and thus decreased diet-derived glucose or glucose precursors. To ensure an adequate nutrient supply to the immune system, hepatic glucose output can increase via both glycogenolysis and gluconeogenesis (Filkins, 1978; McGuinness, 1994). Concurrently, there is an increase in peripheral insulin resistance leading to decreased glucose uptake by skeletal muscle and adipose tissue (Lang et al., 1990; Song et al., 2006). Despite the homeorhetic efforts to spare glucose for the immune system, hypoglycemia often develops following a LPS challenge, likely because the immune system’s rate of glucose utilization exceeds the synchronized capacity of the liver to export glucose and insulin-sensitive tissues to reduce glucose disposal (McGuinness, 2005). Therefore, the experimental objective was to estimate the amount of glucose needed to maintain euglycemia following a LPS challenge as a proxy for the amount of glucose required to fuel an acute immune response.


MATERIALS AND METHODS

All procedures were approved by the Iowa State University Institutional Animal Care and Use Committee. Fifteen Holstein steers (148 ± 9 kg) were randomly assigned to individual pens at the Iowa State University Zumwalt Climatic Research Station (Ames, IA). Steers were allowed 5 d to acclimate during which they were implanted with bilateral jugular catheters and fed ad libitum once daily (0600 h) a diet formulated to meet or exceed the predicted requirements (NRC, 2001) of energy, protein, minerals, and vitamins. Steers were randomly assigned to 1 of 3 i.v. bolus treatments: control (CON; 3 mL sterile saline; n = 5), LPS-administered controls in which hypoglycemia was allowed to develop (LPS-C; 1.5 μg/kg BW LPS; n = 5), and LPS-administered in which euglycemia was maintained (LPS-Eu; 1.5 μg/kg BW LPS; n = 5). The LPS dose used was selected based on Waggoner et al. (2009) and their observation of a 30% decrease in circulating glucose 4 h post-LPS. Lipopolysaccharide (Escherichia coli O55:B5; Sigma Aldrich, St. Louis, MO) was dissolved in sterile saline at a concentration of 75 μg/mL and passed through a 0.2 μm sterile syringe filter (Thermo Scientific, Waltham, MA). The total volume of LPS solution administered was approximately 3 mL. In the LPS-Eu treatment, we performed a euglycemic clamp where 50% dextrose (VetOne, Boise, ID) was i.v. infused at a known and adjustable rate utilizing a modular pump (Deltec 3000; Deltec Inc., St. Paul, MN) to maintain the pre-LPS infusion blood glucose levels.

Feed was removed ∼1 h prior to treatment administration and animals remained fasted during the 720 min data collection period. Baseline blood samples were obtained -30, -20, and 0 min relative to bolus administration to establish baseline glucose levels. Each respective treatment bolus was administered immediately following the 0 min blood sample collection. For LPS-Eu steers, postbolus blood samples (1 mL) were collected every 10 min and immediately analyzed for glucose (TRUEbalance glucometer; McKesson, San Francisco, CA). Dextrose infusion began when blood glucose concentration declined below baseline levels, and its rate of infusion was adjusted as necessary to maintain blood glucose concentration at baseline levels (± 10%). The rate of 50% dextrose infusion (mL/h) was transformed to rate of glucose infusion (ROGI; g/h). The total glucose infused for each steer was calculated using the ROGI for each 10-min interval (72 intervals in total) according to the following equation:

Blood glucose was measured every 30 min in CON and LPS-C steers for the first 300 min and every 60 min thereafter. Additional serum and plasma samples (∼10 mL each) for further analysis were collected from all treatments at -30, 0, 180, 360, 540, and 720 min relative to LPS administration. Rectal temperatures were obtained -30 and 0 min relative to bolus administration, every 30 min for the first 300 min, and every 60 min thereafter using a digital thermometer (GLA M700, San Luis Obispo, CA).

Insulin, glucose, NEFA, β-hydroxy butyrate (BHB), LPS binding protein (LBP), L-lactate, serum amyloid A (SAA), haptoglobin, and blood urea nitrogen (BUN) concentrations were determined using commercially available kits according to manufacturers’ instructions (insulin: Mercodia AB, Uppsala, Sweden; glucose: Wako Chemicals USA Inc., Richmond, VA; NEFA: Wako Chemicals USA, Richmond, VA; BHB: Pointe Scientific Inc., Canton, MI; LBP: Hycult Biotech, Uden, Netherlands; L-lactate: Biomedical Research Service Center, Buffalo, NY; SAA: Tridelta Development Ltd., Kildare, Ireland; haptoglobin: Immunology Consultants Laboratory Inc., Portland, OR; BUN: Teco Diagnostics Anaheim, CA). The inter- and intra-assay coefficients for haptoglobin, insulin, NEFA, BHB, L-lactate, SAA, LBP, and BUN assays were 14.0% and 10.2%, 12.6% and 7.0%, 9.9% and 3.9%, 3.8% and 2.3%, 7.1% and 2.9%, 14.2% and 6.2%, 33.0% and 2.6%, and 12.2% and 5.2%, respectively. Ionized blood calcium was measured using an i-STAT handheld machine and cartridge (CG8+; Abbott Point of Care, Princeton, NJ). For white blood cell (WBC) count, a 3-mL blood sample was collected (K2EDTA; BD Franklin Lakes, NJ) and stored at 4°C for 12 h before submission to the Iowa State Department of Veterinary Pathology for complete blood count analysis.

Postbolus blood glucose was divided into 2 phases: a hyperglycemic (0–180 min) and hypoglycemic (180–720 min) phase which were statistically analyzed separately. Rectal temperature and ROGI were analyzed for the entire postbolus period. Remaining parameters were analyzed for the hypoglycemic phase (180–720 min, during which blood samples were obtained). Each animal’s respective parameter was analyzed using repeated measures with an autoregressive covariance structure for blood parameters and spatial power law for rectal temperature and blood glucose. The repeated effect was minute after bolus administration. Each specific variable’s prebolus value served as a covariate. Effects of treatment, time, and treatment by time interaction (except for ROGI, where only the effect of time within the LPS-Eu treatment was analyzed) were assessed as a completely randomized design using PROC MIXED (SAS Inst. Inc., Cary, NC). Preformed contrasts were used to estimate differences between CON and LPS-infused steers (LPS-C and LPS-Eu) as well as between the two LPS-infused treatments (LPS-C vs. LPS-Eu). Data are reported as LSmeans and considered significant if P ≤ 0.05 and a tendency if 0.05 < P ≤ 0.10.


RESULTS

Rectal temperature was increased ∼0.5°C in both LPS treatments relative to CON steers (P = 0.01; Table 1). Both LPS-infused treatments (LPS-C and LPS-Eu) had ∼100% increased circulating glucose for 180 min following LPS administration relative to CON steers (P = 0.04; Fig. 1A). From 180 to 720 min, there was a 30% decrease in blood glucose in LPS-C steers relative to CON and LPS-Eu steers (P < 0.01; Fig. 1A). No differences in blood glucose were observed between CON and LPS-Eu treatments (P = 0.59) following the 180th min, indicating euglycemia was successfully maintained. Glucose infusion began 178 ± 17 min post-LPS administration (range 150–230 min). The ROGI increased with time (P < 0.01) and 516 ± 65 g of glucose were infused to maintain euglycemia (Fig. 1B) from the 180th to 720th min.


View Full Table | Close Full ViewTable 1.

Blood parameters in steers given a bolus of saline (CON), lipopolysaccharide (LPS-C), or lipopolysaccharide accompanied with a euglycemic clamp (LPS-Eu) during the hypoglycemic phase 180–720 min post-LPS administration

 
Treatment1
P-value
Contrasts
Parameter CON LPS-C LPS-Eu SEM Treatment Time Treatment x Time CON v LPS2 LPS-C v LPS-Eu
Rectal Temperature, °C 38.7a 39.2b 39.2b 0.1 0.02 < 0.01 0.34 0.01 0.95
Metabolites
    NEFA, mEq/L 340a 352a 197b 28 < 0.01 < 0.01 < 0.01 0.08 < 0.01
    BHB,3 mg/dL 4.3a 3.8a 2.7b 0.3 0.01 0.09 0.19 0.02 0.03
    Blood urea N, mg/dL 6.7 10.4 10.1 1.1 0.08 0.01 0.34 0.03 0.84
Haptoglobin, μg/mL 136 170 152 65 0.95 0.19 0.41 0.75 0.87
LBP,4 μg/mL 6.9a 14.0b 12.2b 1.0 < 0.01 < 0.01 < 0.01 < 0.01 0.27
SAA,5 μg/mL 87 117 103 11 0.22 < 0.01 0.32 0.10 0.47
Ionized Calcium, mmol/L 1.40a 1.16b 1.19b 0.02 < 0.01 0.01 0.02 < 0.01 0.38
Complete Blood Count
    Neutrophils, x103/μL 4.4 3.8 1.6 1.1 0.25 0.01 0.08 0.21 0.21
    Lymphocytes, x103/μL 6.0a 2.9b 2.8b 0.6 0.01 < 0.01 0.01 < 0.01 0.89
    Monocytes, x103/μL 0.57a 0.20b 0.18b 0.10 0.03 0.02 0.06 0.01 0.83
    Eosinophils, x103/μL 0.29 0.56 0.46 0.06 0.06 0.05 0.17 0.03 0.27
    Basophils, x103/μL 0.13 0.08 0.07 0.02 0.07 < 0.01 0.04 0.03 0.53
    Red blood cells, x106/μL 9.0 9.4 9.6 0.2 0.26 0.02 0.03 0.11 0.65
    Platelets, x103/μL 553a 274b 256b 31 < 0.01 0.15 0.68 < 0.01 0.71
    NLR6 0.8 1.2 0.6 0.2 0.07 0.12 0.24 0.81 0.03
a,bMeans with different letters differ (P ≤ 0.05).
1CON = saline bolus; LPS-C = LPS bolus; LPS-Eu = LPS bolus and euglycemic clamp.
2LPS-C and LPS-Eu treatments.
3β-hydroxy butyrate.
4Lipopolysaccharide binding protein.
5Serum amyloid A.
6Neutrophil-to-lymphocyte ratio.
Figure 1.
Figure 1.

(A) Blood glucose levels in steers administered a bolus of saline (CON), lipopolysaccharide (LPS-C), or lipopolysaccharide accompanied with a euglycemic clamp (LPS-Eu) during both hyperglycemic (0–150 min) and hypoglycemic (180–720 min) phases; (B) the average rate of glucose infusion in LPS-Eu steers; and (C) circulating insulin levels in CON, LPS-C, and LPS-Eu steers during the hypoglycemic phase (180–720 min).

 

Insulin increased 25.5-fold in LPS-Eu steers compared to CON steers (P < 0.01; Fig. 1C). Ad hoc analysis also indicated that LPS-C steers had a 70% increase (P < 0.05) in circulating insulin compared to controls (data not shown). Circulating NEFA and BHB were reduced (43% and 33%, respectively) in LPS-Eu versus CON and LPS-C steers (P ≤ 0.01; Table 1). Compared to CON, steers in both LPS treatments had a similar increase in circulating BUN (53%; P = 0.03; Table 1), L-lactate (199%; P = 0.01; Fig. 2A), LBP (90%; P < 0.01; Fig. 2B), and a tendency for increased SAA (26%; P = 0.10; Table 1). Ionized calcium decreased 16% in both LPS treatments relative to CON steers (P < 0.01; Table 1).

Figure 2.
Figure 2.

Circulating (A) L-lactate, (B) lipopolysaccharide binding protein (LBP), and (C) white blood cell count (WBC) during the hypoglycemic period (180–720 min) in steers given a bolus of saline (CON), lipopolysaccharide (LPS-C), or lipopolysaccharide accompanied with a euglycemic clamp (LPS-Eu).

 

There was a treatment by time interaction (P = 0.02; Fig. 2C) for WBC counts as LPS steers experienced an initial decrease in cell number (180 min; 85%) followed by a progressive increase with time and WBC counts were similar to CON levels by 720 min. Cell types primarily contributing to the WBC temporal changes were neutrophils, lymphocytes, monocytes, and basophils which were initially reduced (9%, 85%, 91%, and 79%, respectively; P ≤ 0.08) at 180 min post-LPS and gradually increased thereafter (Table 1). The progressive increase in neutrophils tended to be reduced in LPS-Eu steers (P = 0.08; Table 1), resulting in a 63% decrease in neutrophil number by min 720 compared to LPS-C steers. This resulted in a 100% decrease in the neutrophil-to-lymphocyte ratio (NLR) in LPS-Eu relative to LPS-C steers (P = 0.03; Table 1). Eosinophils increased 76% and platelets decreased 52% in both LPS treatments relative to CON steers (P < 0.05; Table 1).


DISCUSSION

Animal productivity is suboptimal during infection and inflammation due to increased energy requirements and subsequent nutrient partitioning toward the immune system and away from agriculturally important products (e.g., skeletal muscle, milk, fetus, etc.). For example, skeletal muscle proteolysis is an important source of amino acids needed for acute phase protein synthesis, and this has been studied extensively (Klasing and Austic, 1984a,b; Reeds et al., 1994; Grimble and Grimble, 1998; Reeds and Jahoor, 2001; Li et al., 2007; Johnson, 2012). However, quantifying nutrient requirements of the immune system is difficult due to its dynamic and ubiquitous distribution throughout tissues. In terms of bioenergetics, both whole body energy expenditure and glucose utilization are estimated to increase approximately 50% during an infection (Lang and Dobrescu, 1991; Lang et al., 1993; Plank et al., 1998). This is likely because immune cells become substantial glucose consumers to support the energetic and substrate needs of activation (Calder et al., 2007; Maratou et al., 2007; MacIver et al., 2008; Palsson-McDermott and O’Neill, 2013; Kelly and O’Neill, 2015). For instance, in tissues with a large immune compartment (spleen, liver, lung, and ileum) LPS increases glucose utilization which is enhanced by euglycemia (Lang et al., 1993). Determining energetic requirements of the immune system in vivo is arduous as immune cells are present in virtually every tissue. Furthermore, the distribution of immune cells throughout different tissues is dynamic and can change rapidly as demonstrated by leukopenia observed in this and other studies (Griel et al., 1975; Bieniek et al., 1998). Despite its obvious difficulty, accurately estimating energetic and nutrient requirements of an activated immune response is essential for developing strategies to optimize productivity in immune-challenged animals. Herein, we propose using a euglycemic clamp after LPS administration to estimate the amount of glucose utilized during an intense immune response.

In the current study, we successfully induced an immune response by LPS injection as indicated by pyrexia and increased circulating levels of acute phase proteins LBP and SAA (90% and 26%, respectively; Table 1 and Fig. 2B). While the pattern of increase in acute phase proteins is consistent with other ruminant literature, there is substantial variation in baseline values and magnitudes of change in response to LPS (Schroedl et al., 2001; Carroll et al., 2009; Graugnard et al., 2013; Plessers et al., 2015). Variation in baseline values have even been reported within labs (Khafipour et al., 2009a,b), and the differences in the magnitude of responses are likely due to dissimilar experimental models, route of LPS administration, and/or laboratory procedures. Severe hypoglycemia (30% decrease; Fig. 1A) began ∼180 min postbolus and continued through 720 min. Hyperlactemia (∼200% increase; Fig. 2A) was also evident in LPS-infused steers, especially at 180 min, and declined slightly with time. Additionally, circulating ionized calcium decreased (18%) which agrees with others (Griel et al., 1975; Carlstedt et al., 2000; Waldron et al., 2003b) and may be due to calcium’s involvement with immune system activation (Hendy and Canaff, 2016). The aforementioned observations are indicative of an aggressive and sustained immune response and are similar to other models of endotoxemia (Filkins, 1978; Lang et al., 1993; Waldron et al., 2003b; Michaeli et al., 2012). Dextrose infusion initiation, coinciding with the development of hypoglycemia, was fairly consistent between steers (178 ± 17 min post-LPS; range 150–230 min). The ROGI increased steadily over time before plateauing at ∼650 min post-LPS. From the accumulated ROGI, we estimate the activated immune system uses approximately 516 g of glucose in a 720-min period. However, as further discussed below, this calculation is likely underestimated as it does not incorporate glucose use by immune cells during the hyperglycemic phase.

Nevertheless, the current experimental design has some limitations. First, the extent of peripheral tissue glucose consumption limits our capacity to accurately estimate immune system’s glucose utilization. However, many studies demonstrate reduced insulin sensitivity and/or glucose utilization in both muscle and adipose tissue during endotoxemia both in vivo (Raymond et al., 1981; Ling et al., 1994; Poggi et al., 2007; Mulligan et al., 2012) and in vitro (Song et al., 2006; Liang et al., 2013). Lang et al. (1993) obtained similar results in a rodent model of endotoxemia; however, when providing glucose to maintain euglycemia, glucose utilization increased in skeletal muscle and adipose tissue. Nonetheless, in tissues with a large immune compartment (spleen, liver, lung, and ileum) glucose utilization was increased in LPS-hypoglycemic rats and the increase was even more pronounced in LPS-euglycemic conditions, acknowledging the relative importance of activated immune cells to whole-body glucose consumption. Additionally, endotoxemia likely causes macrophage infiltration into adipose and muscle (Caesar et al., 2012; Pillon et al., 2013), highlighting the difficulty to pinpoint glucose consumption by the immune system using 2-deoxyglucose, a non-metabolizable marker of glucose uptake, as immune cells are located in essentially every tissue. Immune cell distribution radically changes after LPS administration as demonstrated by leukopenia observed in this and other studies (Fig. 2C; Griel et al., 1975; Bieniek et al., 1998), which is likely due to extravasation of leukocytes into tissues, especially those with important immune functions such as liver, kidney, spleen, and lung (Mészáros et al., 1991; Lang and Dobrescu, 1991). In agreement, Mészáros et al. (1991) examined different cell fractions within the liver after an i.v. LPS challenge and demonstrated glucose uptake did not change in parenchymal cells but markedly increased in Kupffer cells and neutrophils. A potential refinement to the current experimental design would be to incorporate tracers to measure glucose uptake by individual tissues during sepsis. However, an accurate measurement would still require isolating and studying different cell types within a given tissue. The aforementioned studies demonstrate glucose incorporation into immune cells increases while extra-immune cells decrease glucose utilization during LPS infusion and supports our assumption that infused glucose was primarily utilized by immune cells rather than peripheral tissue.

A second limitation to our experimental design is the lack of hepatic glucose output measurements which prevents us from estimating the liver’s contribution to the circulating glucose pool. However, increased glycogenolysis and gluconeogenesis following endotoxin administration has been reported in ruminants (Waldron et al., 2003a) and other species (Wolfe et al., 1977; Filkins, 1978; Spitzer et al., 1985; Lang et al., 1985; McGuinness, 1994; McGuinness, 2005). Hepatic glycogenolysis is likely a large contributor to the hyperglycemic period post-LPS bolus in the current study (Fig. 1A), and we are unable to calculate the amount of endogenous glucose utilized by the immune system prior (∼180 min) to dextrose infusion. The hyperglycemic phase is a consequence of orchestrated systemic changes including peripheral insulin resistance and increased hepatic glucose output which successfully provides glucose in excess of immune cell utilization. The LPS-induced hypoglycemia represents the inability of glucose sparing mechanisms (reduced glucose uptake by insulin sensitive tissues and increased hepatic rates of glycogenolysis and gluconeogenesis) to keep pace with the activated immune system’s glucose utilization. If increased rates of hepatic glycogenolysis and gluconeogenesis described in ruminant and non-ruminant models hold true in the current model, then we are underestimating the amount of glucose entering the circulating pool and subsequently the total amount of glucose utilized by the activated immune system.

Interestingly, differences between the LPS-C and LPS-Eu steers were mostly related to metabolism rather than immunity. Particularly, the magnitude of insulin increase in LPS-Eu steers is remarkable, but not unprecedented, as similar results have been described in dogs (Blackard et al., 1976). This suggests an interaction between LPS and glucose infusion as recently proposed (Baumgard et al., 2016). Further, severe hyperinsulinemia likely contributes to the decrease in plasma NEFA and BHB observed in LPS-Eu cows, as insulin is a potent antilipolytic hormone (Vernon, 1992). Regardless of exogenous glucose infusion, hyperinsulinemia following an LPS challenge has been previously observed in cattle (Waldron et al., 2003a,b; Burdick Sanchez et al., 2013), and might be explained by LPS’s amplifying role in glucose-stimulated insulin release reported in various in vitro experiments (Vives-Pi et al., 2003; Bhat et al., 2014). Furthermore, insulin becomes important for immune cell glucose uptake and development during activation (Shimizu et al., 1983; Helderman, 1984; Calder et al., 2007; Maratou et al., 2007).

Another hallmark of endotoxemia is hyperlactemia (Wolfe et al., 1977; Michaeli et al., 2012), which is likely due to the increase glucose utilization via aerobic glycolysis of immune cells after activation (Palsson-McDermott and O’Neill, 2013). In agreement, in the current study, LPS-Eu steers experienced a numerical 48% increase in plasma L-lactate compared to LPS-C steers, which likely represents an increase in glucose utilization by immune cells when glucose supply is not a limiting factor. Other potential sources include skeletal muscle, which may export L-lactate as an oxidative fuel for nonimmune cells in an attempt to “spare” glucose for the immune system, a process akin to the Warburg Effect (Tannahill and O’Neill, 2011). Other adaptations to support the innate immune response demand for energy and biosynthetic precursors include muscle proteolysis (Klasing and Austic, 1984a; Doyle et al., 2011; Michaeli et al., 2012, Johnson, 2012), which also likely occurred in the current experiment based on the increase in circulating BUN in LPS-infused steers. Interestingly, insulin normally inhibits skeletal muscle proteolysis (Allen, 1988), but insulin’s inhibitory effect on muscle proteolysis is compromised during sepsis (Hasselgren et al., 1987) and may be due to LPS-induced insulin resistance and increased glucocorticoids taking primary control of muscle catabolism (Hall-Angerås et al., 1991).

Regarding immune parameters, leukopenia was evident at 180 min post-LPS, and WBC count gradually increased to baseline levels by 720 min. Leukopenia has been observed in other endotoxemia studies (Griel et al., 1975; Bieniek et al., 1998) and is likely due to leukocyte extravasation into tissues. The NLR was increased in LPS-C steers and similar results were observed in LPS-infused rodents (Rose et al., 2007). The NLR is a marker of systemic inflammation as well as a predictor of mortality in various diseases involving the liver and cardiovascular system (Núñez et al., 2008; Leithead et al., 2015). Infusing glucose in LPS-Eu steers attenuated the increase in NLR and perhaps demonstrates a benefit of supplemental glucose to the innate immune system’s ability to detoxify LPS.

In conclusion, our experiment demonstrates the induction of acute endotoxemia causes hypoglycemia within 3 h, and this likely results from the immune system’s rate of glucose utilization exceeding whole-body glucose sparing mechanisms. From the ROGI, we estimate the activated immune system uses approximately 516 g of glucose in a 720-min period. This is ostensibly underestimated because we are unable to account for immune cell glucose utilization during the acute hyperglycemic phase and the liver’s increased contribution to the circulating pool as described in ruminant and nonruminant models following LPS infusion. Regardless, on a metabolic BW basis, the requirement for glucose is approximately 1.0 g/kg BW0.75/h which is comparable with data we have generated in other LPS-euglycemic clamp experiments in growing pigs and lactating cows (1.1 and 0.66 g/kg BW0.75/h, respectively; Kvidera and Baumgard, unpublished data). The consistency in calculated glucose utilization despite different ages, physiological status, and species suggests the reprioritization and extent of fuel utilization by immune cells upon severe activation might be a conserved response. Whether glucose can become a limiting factor in immune response and the benefit of supplemental glucose is not clear. However, from an animal production perspective, infection and inflammation redirect resources toward the immune system and away from utilization and development of economically relevant tissues. Having a better understanding of the energetic and nutrient requirements of the immune response is critical to develop strategies to minimize productivity losses when physiological states or environmental conditions activate the immune system.

 

References

Footnotes


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