Four dairy heifers with body weights of 280–310 kg from the University Austral herd were used in all experiments. Heifers were maintained on an ad libitum grass diet with grain supplementation. All experiments were conducted in strict accordance with protocols approved by the ethical committee of the Universidad Austral de Chile (permit number: 217/2015) and according to the current Chilean Animal Protect Laws.

Primary BUVEC was isolated from bovine umbilical cords, as previously shown (43). Umbilical cords obtained from calves born by cesarean section were kept at 4°C in 0.9% Hank’s Balanced Salt Solution–HEPES (HBSS–HEPES) buffer (pH 7.4, Gibco, Thermo Fisher Scientific, Waltham, MA, USA), supplemented with 500 U/ml penicillin (Sigma-Aldrich, St. Louis, MO, USA) and 50 mg/ml streptomycin (Sigma-Aldrich, St. Louis, MO, USA) for 1 h. After clamping one end of the umbilical cord veins, 0.025% type II collagenase (Worthington Biochemicals Corporation, Lakewood, NJ, USA) in HBSS (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) was infused into the vein lumens. The opened ends of the veins were closed and then incubated at 37°C and 5% CO₂ for 20 min. Thereafter, the veins were gently massaged and the collagenase solutions were collected in 50 ml plastic tubes (Nunc, Thermo Fisher Scientific, Waltham, MA, USA) containing 1 ml FCS (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) to inactivate the collagenase. The venous lumens were washed two times with RPMI 1640 medium (Gibco, Thermo Fisher Scientific, Waltham, MA, USA). The washes were pooled and centrifuged at 400 × g for 10 min. The cell pellets were resuspended in complete ECM (Sciencell Research Laboratories, Carlsbad, CA, USA), cells were plated in 25 cm² plastic tissue culture flasks (Nunc Thermo Fisher Scientific, Waltham, MA, USA) and incubated at 37°C and 5% CO₂ until reaching confluence.

Blood was collected by jugular vein puncture, and neutrophils were isolated as described previously (44). Briefly, following collection into acid citrate dextrose-lined collection tubes (Becton Dickinson, USA), blood specimens were gently rocked for 5 min (Nutator, Becton Dickinson, Franklin Lakes, NJ, USA) and then centrifuged at 1,000 × g at 20°C for 20 min. The plasma and buffy coat layers were aspirated, and remaining red blood cells and neutrophil pellets were suspended in HBSS. Red blood cells were removed by flash hypotonic lysis using a cold phosphate-buffered water solution (5.5 mM NaH₂PO₄, 8.4 mM HK₂PO₄, pH 7.2). Once returned to isotonicity using a hypertonic phosphate buffer solution (5.5 mM NaH₂PO₄, 8.4 mM HK₂PO₄, 0.46 M NaCl, pH 7.2), the samples were centrifuged at 600 × g at 20°C for 10 min. The neutrophil pellets were washed thrice with HBSS. Neutrophil viabilities were determined by trypan blue assays and were never less than 97% by light microscopy. Purity was at least 94%, as assessed by flow cytometry analyses using forward-scatter vs side-scatter dot plot.

Bovine umbilical vein endothelial cell was cultivated on coated Thermanox^® coverslips (22 mm × 60 mm; Nunc, Thermo Fisher Scientific, Waltham, MA, USA) with 10 µg/ml bovine fibronectin (#F1141 Sigma-Aldrich, St. Louis, MO, USA) until reaching confluence. BUVEC monolayers on Thermanox^® coverslips were mounted in a parallel plate flow chamber allowing constant and uniform laminar flow fields, as previously described (45, 46). Pre-warmed (37°C) pH buffer (140 mM NaCl, 10 mM glucose, 1 mM KCl, 1 mM CaCl₂, 1 mM MgCl₂, and 20 mM HEPES) was perfused over the BUVEC for 2–3 min to remove soluble factors and cell debris. Subsequently, neutrophil suspensions were perfused into the systems at flow rates that resulted in a physiological constant wall shear stress of 1.0 dyn/cm² (syringe pump sp 100i; World Precision Instruments, Berlin, Germany), corresponding to the flow rates of small venous vessels. Interactions between BUVEC and neutrophils were visualized using a phase-contrast microscope (Olympus CKX41, Miami, FL, USA) and video (Optikam PRO 5 Digital camera, Ponteranica, BG, Italy) using AMCap v 9.21 software and an Xvid MPEG-4 codec video compressor. After 10 min, quantification of neutrophil adhesion was performed microscopically by determining the number of adherent cells in 10–20 randomly selected vision fields with modifications as published by Maksimov et al. (47). In each adhesion assay, neutrophils of least four different animals were used. For all experiments, primary BUVEC isolates had been passaged at least three, but no more than seven times, as described in Hermosilla et al. (48). In order to discard a possible effect of pH [5.7 in the media with 5 mM d(−) lactic acid], we used HCl.

Neutrophil extracellular traps were quantitated by measuring the amount of extracellular DNA that was stained by the cell-permeant fluorescent dye PicoGreen^® (Invitrogen, Carlsbad, CA, USA). 1 × 10⁶ neutrophils were incubated with different concentrations of d(−) lactic acid (0.1, 0.7, 2.0, and 5.0 mM), 1 µM of ionomycin or vehicle for 30 min. Micrococcal nucleases were added (5 U/well, New England Biolabs, Ipswich, MA, USA) (15 min, 37°C). Samples were centrifuged (300 × g, 5 min). The supernatants were transferred (100 ml per 96-well) and PicoGreen^® (50 ml/well, diluted 1:200 in 10 mM Tris/1 mM EDTA) was added. NET formation was determined by spectrofluorometric analysis (484 nm excitation/520 nm emission) using an automated reader (Varioskan Flash; ThermoFisher Scientific, Waltham, MA, USA) as described elsewhere (10).

A total of 1 × 10⁶ neutrophils were incubated with 1 µM Ar-c155858, 200 µM of Cl-amidine or vehicle (0.01% dimethyl sulfoxide) for 60 min at 37°C. Next, cells were stimulated with 5 mM of d(−) lactic acid with 90 U DNase I (PanReac AppliChem, Darmstadt, Germany) or with vehicle (0.01% ethanol) for 30 min at 37°C. Next, neutrophils were fixed with 2% paraformaldehyde solution for 20 min at room temperature (RT). The cells were treated with blocking buffer (4% of non-fat milk, 1% BSA, and 0.05% Tween-20 in PBS) for 2 h at RT and incubated with either anti-CD11b antibody (#MCA1425, ABD Serotec, Raleigh, NC, USA), or anti-H₄ citrullinated 3 antibody (#07-596, Merck Millipore, Darmstadt, Germany) overnight at 4°C. Neutrophils were washed with PBS three times and incubated with goat anti-rabbit IgG-Alexa 405 (#A31556, Thermo Scientific, Waltham, MA, USA), or goat anti-mouse IgG-Alexa 635 (#A31575, Thermo Fisher Scientific, Waltham, MA, USA) for 2 h at RT. Nuclei were visualized with PicoGreen^® 1:200 dilution (#P7589, Thermo Fisher Scientific, Waltham, MA, USA). The quantification of NETs was performed using the formula (Nets for field/total Neutrophils) × 100 (49) and depicted as fold of control.

We treated 250 × 10³ neutrophils with 5 mM of d(−) lactic acid or vehicle for 30 min. Cells were fixed in 2.5% glutaraldehyde in 0.1 M cacodylate buffer for 15 min and washed gently with 0.1 M cacodylate buffer (all Merck, Darmstadt, Germany). The cells were then post-fixed in 1% osmium tetroxide (Merck, Darmstadt, Germany) diluted in 0.1 M cacodylate buffer, washed three times in distilled water, dehydrated in ascending ethanol concentrations, critical point dried by CO₂ treatment and sputtered with gold. Specimens were examined using a Philips XL30 scanning electron microscope at the Institute of Anatomy and Cell Biology, Justus Liebig University Giessen, Germany.

Total RNA specimens were isolated from 10 × 10⁶ neutrophils per subject using EZNA^® Total RNA Kits (E.Z.N.A.; Promega, Madison, WI, USA). Samples were treated with Turbo DNase-Free^® (Thermofisher Scientific, Waltham, MA, USA). For the cDNA synthesis reaction, 700 ng of total RNA was reverse transcribed using Affinity Script^® QPCR cDNA Synthesis Kits (Agilent Technologies, Cedar Creek, TX, USA). Real-time PCR assays were performed using SYBR Green^® qPCR Master mix reagents (Fermentas Life Sciences, Waltham, MA, USA) and primers specific for bovine monocarboxylate transporter 1 (MCT1), MCT2, MCT3, MCT4, and housekeeping ribonucleoprotein s9 (RPS9). The primers used for the PCR were as follows: MCT1 forward 5′CGCCGCGAGCCGCGTATAA 3′ reverse 5′CCTCCAACTGCTGGTGGCATTGT3′; MCT2 forward 5′CCACCCAGTGCCGGAGA CCA 3′, reverse 5′TCCCGTGTCTAAGGTTGCCCAGG 3′; MCT3 forward 5′GAGGCT GTGGCTGTGCTCATCG 3′ reverse 5′GATCTCGTAGTTCTTGAGCGCGT CC 3′; MCT4 forward 5′ATCCAGCAAGCCCTCCCTTCCC 3′, reverse 5′CCATGGCCAGGAG GGCTGATTC T3′, RSP9 forward 5′GCTGACGCTG GATGAGAAAGACCC 3′, and reverse 5′ATCCAGCACCCCGATAC GGACG 3′. We used the following conditions on a Mx3000P qPCR System (Agilent Technologies, Santa Clara, CA, USA): (1) 1 cycle of 95°C for 10 min, (2) 35 cycles of 95°C for 30 s, 60°C for 60 s, and 72°C for 60 s, and (3; dissociation curve) 95°C for 1 min, 60°C for 30 s, and 95°C for 30 s.

For immunofluorescence visualization of MCTs, 2.5 × 10⁵ neutrophils were incubated in a 3% paraformaldehyde/0.19 M sucrose solution for 15 min and then transferred to slides using cytospin. Slides were washed with PBS and then neutrophils were permeabilized with 100 µM digitonin (Sigma-Aldrich, St. Louis, MO, USA) for 10 min. Slides were washed with PBS, treated with blocking buffer for 1 h, and incubated with anti-MCT1- (#bs-1569R, BIOSSUSA, Woburn, MA, USA), anti-MCT4- (#bs-2698R, BIOSSUSA, Woburn, MA, USA), and anti-CD147- (EMMPRIN H-200: sc-13976, Santa Cruz Biotechnology, Dallas, TX, USA) specific antibodies, overnight. Next, the slides were incubated with goat anti-rabbit IgG-Alexa Fluor 488 (#A11034, Thermofisher Scientific, Waltham, MA, USA) for 2 h. To visualize nuclei, slides were Hoechst 33342 (#H3570, Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA) stained at 2.5 µg/ml for 20 min. Images were acquired with an OLYMPUS^® Fluoview 1000 confocal microscope.

10 × 10⁶ neutrophils were lysed in lysis buffer (50 mM Tris–HCl, pH 8.0; 150 mM NaCl, 1% of NP-40, and 10 µg/ml of protease inhibitors: leupeptin, aprotinin, and pepstatin). Total protein (80 µg) were separated on 12% SDS-PAGE gels and transferred onto nitrocellulose membranes. Membranes were then blocked in TBS buffer (0.1% Tween-20 and 5% non-fat dry milk) for 2 h at RT, washed with TBS-Tween 0.1% (10 mM Tris–HCl, 68 mM NaCl, and 0.1% Tween-20), and incubated with MCT1 (#bs-1569R, BIOSSUSA, Woburn, MA, USA), MCT2 (#ab182178, Abcam, Cambridge, UK), MCT3 (#ab60333, Abcam, Cambridge, UK), MCT4 (#bs-2698R, BIOSSUSA, Woburn, MA, USA), and CD147 (EMMPRIN H-200: sc-13976, Santa Cruz Biotechnology, Dallas, TX, USA) antibodies overnight at a 1:2,000 dilution at 4°C. Membranes were incubated with HRP-conjugated secondary antibody (#sc-2770, Santa Cruz Biotechnology, Dallas, TX, USA) for 2 h at RT, and visualized using an enhanced chemoluminescence system (Thermofisher Scientific, Waltham, MA, USA). Molecular weights of proteins were determined based on the mobility of pre-stained standards of known molecular weight. For control experiments, MCT1, MCT4, and CD147 antibodies were removed by incubation with a stripping solution (100 mM 2-mercaptoethanol; 2% SDS; 62.5 mM Tris–HCl, pH 6.7) for 1 h at 50°C with agitation, followed by several washes with TBS-Tween 0.1%. Membranes were then incubated with anti-β-actin antibodies (#A1978, Sigma-Aldrich, St. Louis, MO, USA).

We treated 5 × 10⁶ neutrophils with 1 µM of Ar-c1555858 (MCT1 inhibitor) or vehicle (0.01% DMSO) for 60 min at 37°C and later stimulated either with 2 mM of l(+) lactic acid or 2 mM of d(−) lactic acid for 30 min at 37°C. Cells were centrifuged at 800 × g for 6 min. Cell pellets were lysed with liquid nitrogen and resuspended in 500 µl of 1 mM CuSO₄ (mobile phase). Lysed cells were centrifuged at 800 × g for 6 min and supernatants (500 µl) were loaded into Amicon^® Ultra-4 3K (Merck Millipore, Darmstadt, Germany) tubes and centrifuged at 75,000 × g for 40 min at 4°C (50). The filtrates were concentrated in a SpeedVac Concentrator (Savant^® SPD131DDA, Thermo Fisher) for 90 min at 45°C and 1.5 atmospheres of pressure. Finally, the concentrates were resuspended in 200 µl of 1 mM CuSO₄, as reported elsewhere (51). For calibration curves, 2–400 µM aliquots of l(+) and d(−) lactic acid were used. Twenty-microliter samples were analyzed by HPLC using a cationic exchange column, Astec CLC-D (15 cm × 4.6 mm, Sigma-Aldrich, St. Louis, MO, USA), at a 1.2 ml/min flow rate at 30°C. The detection wavelength was set at 254 nm (52) using diode array HPLC, Elite la Chrom (VWR-Hitachi, PA, USA).

Primary BUVEC was seeded onto 96-well plates at a final concentration of 20,000 cells/ml. Briefly, BUVECs were treated with 5 mM of d(−) lactic acid, vehicle (pH buffer), or with supernatants from neutrophils exposed to 5 mM of d(−) lactic acid, DNase I, or 0.01% vehicle (ethanol) for 30 min at 37°C. As a positive control for cell death, cells were treated with 0.2% Triton X-100 diluted in pH buffer at 37°C for 30 min. Cells were incubated with 5 µM propidium iodide (PI; Invitrogen, Carlsbad, CA, USA) diluted in pH buffer for 15 min. The PI signals were detected with a fluorescence multiplate reader (Varioskan^®, Thermo Scientific) at 530 nm excitation/620 nm emission wavelengths.

All results presented here in bar graphs or dot plots illustrate mean ± S.E.M. of five independent experiments obtained from five heifers. One-way analysis of variance calculations were performed, and Dunnett’s multiple comparison tests were applied, with significance considered for P-values less than 0.05. Statistical analyses were performed using GraphPad Prism^® (v 5.0; Graphpad Software, USA).

We previously demonstrated that d(−) lactic acid increased CD11b expression and induced L-selectin shedding in bovine neutrophils (42), suggesting that d(−) lactic acid induced neutrophil adhesion to endothelial cells. Here, this was evaluated with neutrophil adhesion assays onto BUVEC monolayers under physiological flow conditions in parallel plate flow chamber systems. Neutrophils were stimulated with increasing concentrations of d(−) lactic acid (0.7–5.0 mM) for 30 min and then cells were perfused onto BUVEC monolayers. d(−) lactic acid at 5 mM concentrations significantly increased neutrophil adhesions onto vascular endothelial sheets (Figures 1A,B). TNFα (10 ng/ml) stimulation of neutrophils (30 min) and BUVEC monolayers (24 h) served as positive controls, as previously described (48). We discard an effect of pH (5.7) using HCl, which was not able to induce bovine neutrophil adhesion (Figure S1 in Supplementary Material). Next, neutrophils were treated with d(−) lactic acid concomitant with anti-CD11a- and anti-CD11b-specific antibodies (Figure 1C). When co-stimulated with anti-CD11b antibodies, significant reductions of neutrophil adhesions onto BUVEC monolayers were observed. Conversely, co-stimulation with anti-CD11a antibodies did not alter lactic acid-induced neutrophil adhesions. CD11b has been demonstrated to promote amphotericin-induced adhesion, independent of CD11a (53). Furthermore, when neutrophils were co-stimulated with 5 mM d(−) lactic acid and anti-CD11b antibodies and then perfused onto anti-ICAM-1-blocked endothelial sheets, neutrophil adhesion decreased more than when neutrophils were only treated with 5 mM d(−) lactic acid and isotype antibodies (Figure 1D). Together, these results suggested that d(−) lactic induced significant neutrophil adhesion reaction onto vascular endothelium, which was a CD11b/ICAM-1-dependent mechanism.

The mechanism of NET formation was originally described by Brinkmann et al. (5). In this mechanism, neutrophils release granules composed of nuclear DNA fibers coated with histones (H₁, H_2A/H_2B, H₃, H₄) and proteolytic enzymes. These fibers ensnare microbes and, thus, control microbial infection of the host (54). Bovine neutrophils stimulated with vital Eimeria bovis sporozoites efficiently trigger NET formation and increased CD11b surface expression (10). Consistent with these findings, bovine neutrophils exposed to 5 mM d(−) lactic acid for 30 min resulted in extrusion of NET-like structures (Figure 2C) that were positive for H₄ citrullinated (Figure 2A) and CD11b (Figure 2B). The presence of extracellular NET structures were confirmed by SEM analyses in which thin fibers released from lactic acid-treated neutrophils were detected (Figure 2D). Quantification of NET by DNA release showed that stimulation with 5 mM of d(−) lactic acid for 30 min significantly induced the release of extracellular DNA (Figure 2E).

Monocarboxylates, such us pyruvate, lactate, and ketone bodies (d-beta-hydroxybutyrate and acetoacetate), are transported across the plasma membrane through MCTs [e.g., SLC16 (55, 56)]. The monocarboxylate family has 14 members, but only four of them (MCT1–4) bi-directionally transport monocarboxylates coupled to protons (57). In human immune cells, such as leukocytes, MCT1, MCT2, and MCT4, proteins have been detected in granulocytes, lymphocytes, and monocytes (58). These transporters were expressed concomitantly with ancillary cell surface glycoproteins such as CD147 (also known as OX-47, EMMPRIN, HT7, and/or basignin). CD147 is a chaperone protein that facilitates cell surface expression of MCT1 and 4 (59). In bovine neutrophils, RPS9, MCT1, and MCT4 mRNAs were expressed, while MCT2 and MCT3 were absent (Figure S2 in Supplementary Material). MCT4 expression levels were 1,000-fold lower than MCT1 levels (Figure 3A). Expressions of MCT1 and MCT4 gene products in bovine neutrophils were corroborated by western blot and immunofluorescence analyses (Figure 3B). Presence of the chaperone protein CD147 on neutrophils was also observed (Figure 3B). Monocarboxylate transport inside neutrophils was measured by assaying intracellular contents with HPLC analyses. Neutrophils were stimulated with either 2 mM of l(+) or d(−) lactic acid or vehicle (HBSS) for 30 min. Stimulation with either enantiomer increased intracellular monocarboxylate concentrations when compared with vehicle controls. However, uptake of l(+) or d(−) lactic acid was reduced by pre-treatment of neutrophils with Ar-c155858 [a selective and specific pharmacological inhibitor of MCT1 (60); Figure 3C (top and bottom panels)]. Together, these results demonstrated that MCT1 and MCT4, and CD147 gene products were expressed in bovine neutrophils. The data suggested that MCT1 facilitated l(+) or d(−) lactate intracellular transport into these cells.

The role of MCT1 activity in NET formation by neutrophils was assessed. Neutrophils were incubated with 1 µM of Ar-c155858 to inhibit endogenous MCT1 and then stimulated with 5 mM d(−) lactic acid. Co-treatment with these agents resulted in significant reduction of NET formation in comparison to d(−) lactic acid treatment alone (Figure 4A). Similarly, treatment with a PAD4 inhibitor (200 µM, Cl-amidine) and DNase I (90 U) reduced neutrophil NET formation, suggesting that d(−) lactic acid-induced NET formation was PAD4- and MCT1-dependant (Figures 4A,B). Furthermore, the role of MCT1 on neutrophil adhesion under continuous flow conditions onto endothelium was assessed. Bovine neutrophils treated with Ar-c155858 to inhibit MCT1 significantly decreased neutrophil adhesion of cells that were previously exposed to 2 and 5 mM d(−) lactic acid (Figure 4C). TNFα is a pro-inflammatory cytokine that has been shown to stimulate adhesion of neutrophils to endothelium through activation of the β-integrin, CD11b/CD18 (61), and NET formation (15). TNFα-induced effects on neutrophils are thought to be mediated through activation of TNF receptors (62) and not through MCT transporter mechanisms. To determine whether Ar-c155858 specifically inhibited d(−) lactic acid-induced neutrophil adhesion, the role of MCT in TNFα-induced neutrophil adhesion was evaluated. Interestingly, Ar-c155858 treatment did not inhibit TNFα-mediated neutrophil adhesion on bovine endothelial sheets in parallel flow chamber assays (Figure 4C). Stimulation of BUVEC monolayers with TNFα for 24 h was used as a positive control. Co-stimulation with DNase I and d(−) lactic acid treatments also significantly decreased neutrophil adhesions onto bovine endothelial sheets. Conversely, DNase I treatments did not alter TNFα-induced neutrophil adhesions onto BUVEC monolayers (Figure 4D).

The role of PAD4 in d(−) lactic acid-induced neutrophil adhesion was evaluated. Adhesion was significantly inhibited after Cl-amidine pre-treatment, prior to stimulation with d(−) lactic acid (Figure 4E). To evaluate whether d(−) lactic acid BUVEC stimulation increased neutrophil adhesions, untreated neutrophils were perfused onto endothelial sheets that had been previously treated with Ar-c155858 or corresponding vehicle (0.01% DMSO), and then cells were stimulated with 5 mM of d(−) lactic acid. Neutrophil adhesions onto endothelial sheets were not observed when compared to positive controls (i.e., TNFα; Figure 4F). Together these results suggested that neutrophil adhesions onto bovine endothelial sheets were dependent on d(−) lactic acid uptake through MCT1 and that NET formations were mediated by PAD4. Neutrophils stimulated with d(−) lactic acid increased adhesions to endothelia that were not recapitulated by direct endothelial stimulations. Moreover, DNase I and Ar-c155858 treatments specifically inhibited d(−) lactic acid-induced neutrophil adhesions.

Previously, d(−) lactic acid stimulation of bovine neutrophils resulted in the release of NET, suggesting that NET may participate in neutrophil adhesions onto activated endothelial cell sheets. cf-DNA may function to activate endothelial cells, which could promote firm neutrophil adhesions. To test this, supernatants from neutrophils previously stimulated with 5 mM d(−) lactic acid or vehicle were collected in the presence or absence of DNase I. Supernatants were perfused on bovine endothelial monolayers in parallel plate flow chambers for 10 min. Next, untreated neutrophils were perfused into pre-treated chambers and adhesion assays were performed. Adhesions of freshly infused, unstimulated neutrophils were significantly increased when endothelial cell sheets were exposed to supernatants collected from 5 mM d(−) lactic-stimulated neutrophils (Figure 5A). Conversely, decreased neutrophil–endothelium adhesions were stimulated by supernatant collected from neutrophils treated with both 5 mM d(−) lactic acid and anti-CD11b antibodies. Endothelial cell sheets were previously blocked with anti-ICAM-1 antibodies or control antibody isotypes (Figure 5B). Together, these data suggested that d(−) lactic acid neutrophil stimulation mediated release of soluble NET structures. These soluble components enabled neutrophil adhesions onto endothelia through a CD11b/ICAM-1-dependent mechanism. Neither supernatant affected endothelial cell viabilities (Figure 5C).