Published online before print December 23, 2003
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EA 3442: Laboratoire de Biologie Expérimentale–Immunologie, Faculté des Sciences, Université de Nancy 1, Vandoeuvre, France
1Correspondence: University Henri Poincare–Nancy I, Science Faculty, EA 3442, Boulevard des Aiguillettes BP 239, 54506 Vandoeuvre-les-Nancy, France. E-mail: Rachel.Boxio{at}scbiol.uhp-nancy.fr
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Key Words: isolation method • fMLF stimulation • exocytosis • NADPH oxidase • lifespan
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Understanding cellular functions on the molecular level has been greatly enhanced by the possibility to mutate, suppress, or overexpress individual genes. However, primary human neutrophils are too short-lived for those techniques, and the cell-culture models for neutrophils do not fully differentiate and are difficult to transfect. This has prompted a renewed interest into mouse neutrophils, which could be obtained from genetically modified animals. The number of neutrophils that can be obtained from mouse blood is very limited [5 ]. Therefore, many researchers use bone marrow preparations to study neutrophils in mice. How are the functional characteristics of these cells compared with blood neutrophils? Is it possible to obtain functionally mature neutrophils from mouse bone marrow?
We wanted to improve our knowledge about the physiology of mouse neutrophils and to compare the functional capacities of bone marrow and blood neutrophils. Here, we present evidence that a subpopulation of mouse bone marrow leukocytes consists of functionally mature neutrophils.
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) of the layer was calculated according to the manufacturer’s information (52%, RI 1.347, =1.083 g/ml; 69%, RI 1.349, =1.090 g/ml; 78%, RI 1.350, =1.110 g/ml). The neutrophils from the 69%/78% interface and the upper part of the 78% layer were harvested into BSA 1%-coated tubes, after carefully removing the cells from the upper phases. After one wash with 2 ml HBSS–EDTA + BSA 1%, remaining red cells were eliminated by hypotonic lysis. After a final wash with 2 ml HEPES buffer (140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 10 mM HEPES, pH 7.2), 350,000 ± 40,000 cells were obtained per mouse, 97 ± 1% neutrophils, identified by staining the nuclei with Türks reagent (Merck, Darmstadt, Germany). These cells were suspended in HEPES buffer + 9 mM glucose at 106 cells/ml and used within 6 h. On several occasions, isolated cells were analyzed by cytospin centrifugation followed by Wright-Giemsa coloration according to the manufacturer’s instructions (Hemacolor, Merck). Chemicals were bought from Sigma Chemical Co. (St. Louis, MO) unless indicated otherwise.
Preparation of mouse neutrophils from bone marrow
Mice were killed, the femur and the tibia from both hind legs were removed and freed of soft tissue attachments, and the extreme distal tip of each extremity was cut off [6
]. HBSS–EDTA solution was forced through the bone with a syringe. After dispersing cell clumps, the cell suspension was centrifuged (400 g, 10 min, 4°C) and resuspended in 1 ml HBSS–EDTA. Cells were then treated on a three-layer Percoll gradient exactly as described above for blood cells. We obtained 6 ± 0.6 x 106 cells per mouse, and 94 ± 1% of them were morphologically mature neutrophils (bands and segmented).
Preparation of human neutrophils
Human PMN were isolated from blood by 20% dextran (1:1) sedimentation at room temperature for 40 min. The leukocyte-rich upper fraction was collected, layered on a continuous Histopaque®-1077 gradient, and centrifuged (1000 g, 20 min, room temperature) without braking. The remaining red cells in the neutrophil pellet were eliminated by hypotonic lysis and spun down (300 g, 10 min, 4°C). After a final wash with 2 ml phosphate-buffered saline (PBS), staining the nuclei with Türks reagent identified the neutrophils. Cells (1.5x108) were suspended in 200 µl PBS supplemented with 2% protease inhibitor cocktail. The isolated neutrophils were kept on ice until the experiments were performed within a few hours after the preparation.
Superoxide production
ROIs were quantified by measuring luminol-dependent chemiluminescence at 37°C using a Wallac 1420 multilabel counter (Perkin Elmer, Courtaboeuf, France) at approximately one measurement per well every 6 s. White polypropylene, 96-well plates (Nunc, Roskilde, Denmark) were covered with BSA (1%, 1 h, 37°C). After coating, the plates were washed three times with HEPES buffer. Mouse neutrophils (2x104 cells) were suspended with 10 µg/ml luminol, 5 µg/ml cytochalasin B, 4 U/ml horserardish peroxidase, and HEPES buffer for a final volume of 200 µl per well. The cells were stimulated with formylmethionyl-leucyl-phenylalanine (fMLF), and superoxide anion generation was measured by integrating photon counts for 5 min before and after agonist addition. The solvent control, dimethyl sulfoxide (DMSO), did not stimulate ROI production.
ß-Glucuronidase release
To measure the release of the primary granules, we established a method of detecting the activity of the marker ß-glucuronidase from the supernatants of stimulated mouse neutrophils. Tubes were coated with BSA (1%, 1 h, 37°C). Mouse neutrophils (2x104 cells) were suspended in these tubes in HEPES buffer + glucose for a final volume of 100 µl with 5 µg/ml cytochalasin B (unless otherwise indicated). Cells were prewarmed for 10 min at 37°C before addition of various concentrations of fMLF, 1 µM A23187, 200 nM phorbol 12-myristate 13-acetate (PMA), or solvent (DMSO) alone. After incubation at 37°C for 30 min, secretion was stopped by rapid cooling on ice and addition of 100 µl ice-cold HEPES buffer + glucose and was centrifuged (400 g, 10 min, 4°C). Duplicates of 50 µl supernatant were dispensed in 96-microwell plates, and 50 µl reaction medium [0.1 M sodium acetate, pH 4.0, 0.02% Triton X-100, 5 mM 4-methylumbelliferyl ß-D-glucuronide (Roth, Karlsruhe, Germany)] was added. The reaction was stopped after 2 h at 37°C by adding 50 µl 0.3 M glycine/NaOH, pH 10.4, 10 mM EGTA. Fluorescence was quantified by using a Wallac 1420 multilabel counter with 365 nm excitation and 450 nm emission wavelength. ß-Glucuronidase release from murine PMN is expressed as percent of release compared with the total content detected in supernatants of Triton X-100-lysed PMN. Background release in the presence of solvent was subtracted.
Lactoferrin release
Mouse neutrophils (2x104 cells) were suspended in 100 µl HEPES buffer + glucose with 5 µg/ml cytochalasin B and were prewarmed for 10 min at 37°C before addition of various concentrations of fMLF, 1 µM A23187, 200 nM PMA, or solvent (DMSO) alone. After incubation at 37°C for 30 min, secretion was stopped by rapid cooling on ice and addition of 100 µl ice-cold buffer and was centrifuged (400 g, 10 min, 4°C). Supernatants or a solution of human lactoferrin (10–50 ng per well) were diluted twofold in carbonate buffer (90 mM NaHCO3, 36 mM Na2CO3, pH 9.6) and were added to Nunc Maxisorp F96 immunoplate wells for 3 h at 37°C. Nonspecific binding sites were blocked with PBS–Tween 1%, supplemented with 2% BSA overnight at 4°C. PBS–Tween 1% was used for all subsequent steps, which were conducted at room temperature and separated by several washings. Rabbit anti-human lactoferrin antibody (dilution 1/1000) was added for 1 h, followed by goat peroxidase-conjugated anti-rabbit immunoglobulin G (IgG; dilution 1/10,000) for 1 h. 3.3.5.5'-Tetramethylethylenediamine (5 mg/ml; Interchim, Montlucon, France) in 100 mM acetate buffer, pH 6.0, 1.2% H2O2, was added for 15 min, followed by 750 mM H2SO4. Absorbance was read at 450 nm with a Wallac 1420 microplate reader, and the lactoferrin concentration was calculated using the human lactoferrin calibration curve. As data obtained with mouse samples could not be quantified exactly using standard curves done with human lactoferrin, lactoferrin release from murine PMN is expressed as fold increase compared with the amount detected in supernatants of unstimulated PMN [6
].
Western blotting
The specificity of the anti-human lactoferrin antibody with murine lactoferrin was determined by Western blotting. Human and murine PMN suspensions were divided into two batches. One part was incubated in HEPES buffer with 200 nM PMA at 37°C for 30 min in the presence of 5 µg/ml cytochalasin B. The other part was taken up in lysis buffer (20 mM Tris, 400 mM KCl, 20% glycerol, 2 mM dithiothreitol) for 20 min on ice and lysed by three cycles of freezing in liquid nitrogen/thawing on ice. Both batches were then spun down, and the supernatant was solubilized in 4x concentrated sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer and boiled for 10 min. Samples were run on a 10% SDS-PAGE, electroblotted onto nitrocellulose sheets (Hybond N, Amersham Pharmacia Biotech), blocked with 3% BSA, and probed with rabbit anti-human lactoferrin antibody (dilution 1/500) followed by goat peroxidase-conjugated, anti-rabbit IgG (dilution 1/10,000) in PBS–Tween 1%, BSA 3%. Signal was developed and analyzed using a chemiluminescence substrate (Roche Diagnostics France S.A, Meylan), autoradiographic film (Hyperfilm ECL, Amersham Pharmacia Biotech), and the GelDoc2000 imaging system (BioRad, Hercules, CA).
Viability of neutrophils in culture
Neutrophils from mouse blood and bone marrow were cultured in RPMI-1640 medium supplemented with 10% heat-inactivated fetal bovine serum and 100 U/ml penicillin and streptomycin on 96-well plates (20,000 cells/50 µl/well) for up to 48 h at 5% CO2, 37°C. The bioluminescence of adenosine 5'-triphosphate (ATP) was used as a marker of cell viability using the Cell Titer GloTM luminescent cell viability assay (Promega, Madison, WI) according to the manufacturer’s instructions. The luminescence was read on the Wallac 1420 plate reader and reported to a standard curve for the determination of the number of viable cells.
Analysis and statistics
Data are expressed as mean ± SEM. Statistical significance was tested with unpaired Student’s t-test; P values below 0.05 were considered significant. Sigmoidal and exponential fits were performed in Origin (Microcal, Northampton, MA). The half-life of the cells was determined as t1/2 = 0.693 x 
, with being the time constant of the exponential decay fit.
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Figure 1. Neutrophil isolation from blood and bone marrow. PMN were isolated by centrifugation over a three-layer Percoll® gradient. The photographs show cytospin preparations of cells from the 69%/78% interface stained by Giemsa coloration. A large majority of donut-shaped nuclei was found in bone marrow PMN (A) and blood PMN (B). Original scale bar = 20 µm. (C) Comparison of yield and purity and percent of recovery of blood and bone marrow neutrophil populations isolated as described in Materials and Methods. Mean ± SEM; n = 23–27. Recovery was calculated using the values of Chervenick et al. [8
].
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40% of marrow cells as determined by Wright’s stain and peroxidase staining. Forty-three percent of these marrow neutrophils were bands and segmented cells that are considered to constitute the effective storage compartment of marrow, as they seem to be the only cells that are released to the blood in large numbers under ordinary circumstances [8
]. According to these values, we recovered over 60% of the morphologically mature neutrophils from the bone marrow. The recovery from blood was over 85%, suggesting that mouse blood PMN have a fairly homogenous density. Only few studies have compared the functional characteristics of bone marrow and blood neutrophils, and to our knowledge, none of these studies was done on mice. We concentrated on the bactericidal functions that can be induced by the bacterial peptide fMLF, which has been widely used to characterize human neutrophils. These functions are the generation of ROIs by the reduced nicotinamide adenine dinucleotide phosphate (NADPH) oxidase as well as the release of secondary and primary granules [1 ].
fMLF-induced superoxide production from blood and bone marrow mouse neutrophils
We established a method, which determined the amount of ROIs by luminol-dependent chemiluminescence and which allows a quantification of the production of free radicals on few cells (20,000 cells per well). To investigate the patterns of the respiratory burst in blood and bone marrow neutrophils, cytochalasin B-pretreated cells were stimulated with 3 µM fMLF, and the superoxide production was recorded 5 min before and 5 min after the stimulation (Fig. 2A
). As expected from human neutrophils, fMLF induced a rapid and transient production of ROIs. Compared with peripheral blood neutrophils, the bone marrow cells showed no defect in superoxide release upon stimulation with fMLF.
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Figure 2. fMLF-induced superoxide production from blood and bone marrow neutrophils. ROI production was determined by luminol-enhanced chemiluminescence. (A) Kinetics of production by blood (broken line) and bone marrow (solid line) neutrophil oxidative burst. Cells were incubated at 37°C in the presence of cytochalasin B (5 µg/ml) for 5 min before stimulation with 3 µM fMLF. This figure shows a representative experiment; n  6. (B) Effect of cytochalasin B (cytoB): Bone marrow cells were preincubated at 37°C in the presence (solid line) of or in the absence (dotted line) of cytochalasin B (5 µg/ml) for 5 min before stimulation with 3 µM fMLF. This figure shows a representative experiment; n > 3. (C) Dose-response curve of total ROI production induced by different concentrations of fMLF in the presence of cytochalasin B (5 µg/ml). The EC50 for oxidative burst for blood and bone marrow neutrophils in response to fMLF was 0.52 ± 0.1 and 2.10 ± 0.54 µM, respectively (P<0.05). Results are shown as mean ± SEM. The lines represent the average of the fit curves from individual experiments; n = 4–5.
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For a quantitative comparison of the ROI production from blood and bone marrow PMN, we integrated luminescence signal over 5 min to evaluate the total superoxide release. The response to a wide range of concentrations of fMLF in the presence of cytochalasin B (Fig. 2C) showed a sigmoid dose-response curve. The maximal response was obtained with 3 µM fMLF for blood PMN and 10 µM fMLF for bone marrow PMN. Higher doses up to 100 µM did not further increase but decreased the response of bone marrow cells (data not shown). The EC50 for oxidative burst in the blood and bone marrow PMN in response to fMLF was 0.52 ± 0.1 and 2.10 ± 0.54 µM, respectively (n=4–5). Thus, blood neutrophils responded to fMLF stimulation with somewhat higher affinity than bone marrow cells (statistically significant, P<0.01). The maximal response of bone marrow cells was almost identical to the response of blood cells. Concerning the oxidative burst, these results indicate that bone marrow neutrophils are as functional as blood neutrophils.
Mouse neutrophils release ß-glucuronidase, a marker for primary granules
The enzymatic contents of the granules in human PMN are well-known [11
], and those of the murine neutrophils are much less known. Based on the assumption that ß-glucuronidase, which is known to be present in mouse neutrophils [12
], is localized to the same granules as in their human counterparts, we developed a method for primary granule secretion by measuring the ß-glucuronidase activity using a fluorescent substrate for this enzyme. The assay measures the activity of this enzyme in the supernatant of just 5000 cells. The enzyme release was compared with the total ß-glucuronidase activity from Triton-lysed PMN. Blood and bone marrow PMN had the same total ß-glucuronidase activity (107.6±3.6 vs. 106.4±4.5 arbitrary fluorescence units; n=33 and 18). Addition of fMLF stimulated the release of up to 25% of the primary granules in mouse neutrophils. Similar to the ROI production, the dose-response curve to fMLF for primary granule release was sigmoid with a maximal response at 3 µM and 10 µM fMLF for blood and bone marrow PMN, respectively (Fig. 3A
). The values of EC50 were similar to those obtained for ROI production (0.71±0.11 µM and 1.19±0.09 µM, P<0.01). The maximal response had a tendency to be higher for bone marrow PMN, but this difference was not statistically significant (P=0.28).
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Figure 3. Primary granule secretion. The secretion of the primary granules was measured using the activity of the ß-glucuronidase in the supernatant of stimulated PMN. (A) Dose-response curve of primary granule secretion, induced by different concentrations of fMLF in the presence of cytochalasin B (5 µg/ml) at 37°C, plotted as percentages of the total cellular ß-glucuronidase contents. The EC50 for blood (dotted line) and bone marrow (solid line) neutrophils in response to fMLF was 0.71 and 1.19 µM, respectively (P<0.05). Results are shown as mean ± SEM. The lines represent the average of the fit curves from individual experiments; n 6. (B) Comparison of blood (open bars) or bone marrow cells (solid bars). Left side, Significant effect of cytochalasin B on the primary granule secretion in response to a 3-µM fMLF (P<0.05, n6). Right side, Significant response to PMA (200 nM) and the Ca2+ ionophore A23187 (1 µM) in the presence of cytochalasin B compared with control DMSO (P<0.05, n5). Data are shown as mean ± SEM.
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Taken together, blood and bone marrow mouse PMN can be stimulated to mobilize ß-glucuronidase-containing primary granules in response to the chemoattractant fMLF, to PMA, and to the Ca2+ ionophore A23187 in the presence of cytochalasin B. The affinity for fMLF was similar as for ROI production and about twofold higher for blood PMN compared with bone marrow PMN. The overall response was similar between the two cell populations.
Lactoferrin release by mouse neutrophils
A characteristic feature of neutrophils is the presence of several types of granules with specific physiological roles and partially distinct mechanisms of exocytosis. After having evaluated the secretion of the primary granules, we wanted to know if on the one hand, the secondary granules were liberated in response to the same agonists and on the other hand, how much fMLF was necessary to stimulate the release of these granules. We chose lactoferrin as a marker for secondary granules that can be detected by enzyme-linked immunosorbent assay (ELISA) [6
]. There was no antibody against mouse lactoferrin available; therefore, we tested the specificity of a commercial antibody against human lactoferrin by Western blot. Human lactoferrin purified from human milk (90 kDa, according to the manufacturer), BSA, the supernatant of PMA-stimulated human and murine neutrophils, or lysed human and murine neutrophils were analyzed by SDS-PAGE and Western blot. PMA is known to stimulate secondary granule release from human neutrophils [14
]. Figure 4
shows that the anti-human lactoferrin antibody recognized one single band in each sample, at 91 kDa or 87 kDa in the supernatant and the lysate of human and mouse neutrophils, respectively. The molecular weight of this band and the fact that it appeared in the supernatant of PMA-stimulated PMN confirmed this protein to be lactoferrin. However, we could note a slight difference in migration between human and mouse lactoferrin, despite a similar number of amino acids (711 for human, accession number NP_002334; 707 for mouse, accession number BAA13633; protein sequence homology: 70%) and a similar theoretical, molecular mass (78.3 kDa for human; 77.7 kDa for mouse). This difference in migration might be explained by a difference in the number of sites and extent of glycosylation of this protein. Indeed, the polypeptide chain of human lactoferrin possesses two glycosylation sites with five different glycopeptide structures possible [15
], and the mouse lactoferrin presents only one site of glycosylation [16
].
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Figure 4. The specificity of anti-human lactoferrin antibody mouse lactoferrin. Triton-lysed PMN or the supernatant of PMA-stimulated PMN were analyzed by SDS-PAGE and Western blot. Lanes: 1, Purified milk lactoferrin (Lf), Sigma, 40 ng; 2, BSA, Sigma, 500 ng; 3, supernatant of PMA-stimulated human PMN from 0.64 x 106 cells, 3 µg total protein; 4, lysed human PMN, 0.17 x 106 cells, 1 µg total protein; 5, supernatant of PMA-stimulated mouse PMN from 2.53 x 106 cells, total protein not determined; 6, lysed mouse PMN, 1.11 x 106 cells, 18 µg total protein. Representative of at least three experiments per condition. M.W., Molecular weight.
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Figure 5. Secondary granule secretion. ELISA quantified specific granule exocytosis as lactoferrin release. Data are expressed as fold increase relative to the amount of lactoferrin found in the supernatant of PMN incubated with solvent (DMSO) alone. (A) Specific granule secretion induced by different concentrations of fMLF in the presence of cytochalasin B (5 µg/ml) is plotted as fold increase over the control DMSO (fMLF solvent). The EC50 for blood (dotted line) and bone marrow (solid line) neutrophils in response to fMLF was 0.8 and 0.2 µM, respectively. Results are shown as mean ± SEM; n 4. (B) Left side, The effect of cytochalasin B on secondary granule secretion. Blood (open bars) or bone marrow cells (solid bars) were preincubated at 37°C in the presence or in the absence of cytochalasin B (5 µg/ml) for 5 min before stimulation with 3 µM fMLF. Right side, secondary granule secretion in response to PMA (200 nM) and the Ca2+ ionophore A23187 (1 µM) in the presence of cytochalasin B compared with DMSO solvent control (Student’s test, P<0.05). The data are shown as mean ± SEM; n 3. The response of PMN of marrow to these two stimulators was statistically more important than those of blood (P<0.05).
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Blood neutrophils have a reduced viability
Blood neutrophils are short-lived cells that are constantly replaced by cells from the bone marrow. The lifespan of the cells is a critical determinant for the number of cells in the blood. As our preparation of bone marrow neutrophils seems to consist of terminally differentiated cells, we wanted to compare the lifespan of blood and bone marrow PMN. Neutrophils were kept under standard cell culture conditions for up to 48 h. After 24 h, almost all blood neutrophils were dead, and 25% of the bone marrow PMN survived. The time course of the number of viable cells revealed an exponential decay, which was significantly faster for blood PMN (Fig. 6A
). The half-life of bone marrow PMN in culture, calculated from the exponential fit of the time course, was twice as long as the half-life of blood PMN (P<0.05). Blood and bone marrow neutrophils appeared to enter apoptosis. They showed nuclear condensation like apoptotic human neutrophils [17
] and loss of mitochondrial membrane potential (R. Boxio et al., unpublished data).
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Figure 6. Viability of bone marrow and blood neutrophils in culture. Intracellular ATP was determined to assess the viability of blood and bone marrow neutrophils after different times of culture under identical conditions. (A) Decrease in viability as a function of time of culture. Each point is the mean of a triplicate measurement. The lines represent an exponential fit through the data. The graph shows one example representative of three independent experiments. (B) Mean half-life of blood and bone marrow neutrophils in culture determined by exponential fit as shown in A; n = 3. Data are shown as mean ± SEM; the difference between the two populations is statistically significant (P<0.05).
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Previous studies have shown that bone marrow of humans and mice contains large numbers of myeloid cells with banded or segmented nuclei [4 , 8 ]. Furthermore, surface markers for mature neutrophils have been found on mouse bone marrow cells [19 , 20 ]. Based on the shape of the nuclei, our preparation of neutrophils from mouse bone marrow is indistinguishable from peripheral blood neutrophils. Our goal was to compare the functional qualities of mouse bone marrow neutrophils to blood neutrophils.
The chemotactic peptide fMLF is a major activator of human neutrophils, and recent studies have shown that mouse neutrophils also respond to this agonist with production of reactive oxygen species by the NADPH oxidase [21 ]. For a more detailed characterization, we have determined the EC50 for fMLF, the kinetics, and the extent of the response. We found only a minor difference between blood and bone marrow PMN, whereas Berkow and Dodson [4 ] reported reduced ROI production in human bone marrow PMN.
Another action of fMLF is the stimulation of exocytosis. Neutrophils contain four distinct types of granules and secretory vesicles, which are released differentially [22 , 23 ]. In general, the exocytosis of primary granules requires the strongest stimulation. This may reflect the need for stringent control of these granules, which contain the most aggressive substances. We demonstrate here that mouse neutrophils, like human neutrophils, release primary granules after stimulation with fMLF. Again, only minor differences between bone marrow and blood neutrophils appear. However, the EC50 for primary granule release by fMLF from mouse blood neutrophils is 0.71 µM compared with 0.01 µM in human neutrophils [24 ]. Like human neutrophils, stimulation of primary granule release by fMLF requires cytochalasin B. To our knowledge, this is the first demonstration of primary granule exocytosis from mouse neutrophils.
A similar picture is found for secondary granules. Their release from adherent mouse neutrophils has been shown before [6 ]. As in human PMN, PMA and Ca2+ ionophore are stronger stimuli for secondary granules than for primary granules [25 , 26 ].
For primary granule release and NADPH-oxidase activation, peripheral blood neutrophils responded to fMLF with higher affinity than bone marrow cells. This may reflect the final steps of differentiation that enhance the responsiveness of the blood neutrophil for the activation of highly bactericidal mechanisms. Secondary granule release may be required before the direct contact with bacteria, thus earlier in the life of a neutrophil. This might explain why secondary granule release appeared to be more sensitive to fMLF in bone marrow cells. There might be a hierarchy in the installation of the bactericidal functions during the cellular maturation of the neutrophil.
Human bone marrow contains an important reserve pool of neutrophils [2
]. The most likely role of the functional neutrophils in the bone marrow reserve pool is to replace peripheral neutrophils or supplement at times of increased demand during infection. The size of the reserve pool and the peripheral pool depends on the rate of synthesis, the lifespan of the cells, and the rates of migration into the bloodstream and into tissue. Releasing bone marrow cells with a longer lifespan into the blood is an effective way of increasing the number of circulating neutrophils. In the natural environment of the bone marrow, the difference in the half-life of the two cell populations may be larger than the twofold difference, which we observed under culture conditions. It has recently been suggested that aging mouse PMN may return to the bone marrow and be eliminated there [27
]. This raised the possibility that our bone marrow preparation contained a large number of rapidly dying cells. However, we find that the half-life of these cells in culture is twice as long as that of blood neutrophils. The reported half-life of human blood PMN in culture is approximately 24 h [28
], thus substantially longer than in mice. This difference may contribute to the fact that the portion of neutrophils in the mouse white blood cell count is much lower than in human blood (5–25%, depending on the mouse strain, ref. [10
], vs. 60% of human blood leukocytes), and the total white blood cell count is similar. The literature suggests that neutrophils die by apoptosis, and their lifespan is regulated by the balance between pro- and antiapoptotic proteins under the influence of extracellular signals [3
]. Our data are consistent with the idea that neutrophils receive antiapoptotic signals in the bone marrow, which are less present or absent in the bloodstream. Granulocyte-colony stimulating factor (G-CSF) and granulocyte macrophage (GM)-CSF may be among those signals, although GM-CSF had a much smaller effect on mouse neutrophils than G-CSF [29
]. A mouse of 25 g body weight has 50 x 106 mature neutrophils in its bone marrow [8
] and 2.3 x 106 neutrophils in the blood (volume, 1.9 ml). With a half-life of 6 h, 10 x 106 cells need to be replaced every day. Thus, the bone marrow reserve corresponds to 5 days of neutrophil production under normal conditions and less when more neutrophils are needed to combat an infection. The reserve pool could also serve as a buffer for irregularities in the PMN development in the bone marrow, which takes 10–14 days in humans [7
].
In summary, our findings suggest that mouse bone marrow contains a reserve pool of functionally mature neutrophils that might be readily mobilized when needed during bacterial infections. This pool contains 20 times more neutrophils than the blood. For numerous experiments with mouse neutrophils, the bone marrow may be an attractive and valuable alternative to peripheral blood. Nevertheless, certain studies will require several samples of blood neutrophils from the same animal during the course of the experiment, for example, the analysis of the neutrophil response during infection, drug treatment, or studies on nutrition.
Received July 21, 2003; revised October 29, 2003; accepted November 12, 2003.
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G. Hajishengallis, M. Wang, G. J. Bagby, and S. Nelson Importance of TLR2 in Early Innate Immune Response to Acute Pulmonary Infection with Porphyromonas gingivalis in Mice J. Immunol., September 15, 2008; 181(6): 4141 - 4149. [Abstract] [Full Text] [PDF]
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M. D. Trottier, M. M. Newsted, L. E. King, and P. J. Fraker Natural glucocorticoids induce expansion of all developmental stages of murine bone marrow granulocytes without inhibiting function PNAS, February 12, 2008; 105(6): 2028 - 2033. [Abstract] [Full Text] [PDF]
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S. L. Farnworth, N. C. Henderson, A. C. MacKinnon, K. M. Atkinson, T. Wilkinson, K. Dhaliwal, K. Hayashi, A. J. Simpson, A. G. Rossi, C. Haslett, et al. Galectin-3 Reduces the Severity of Pneumococcal Pneumonia by Augmenting Neutrophil Function Am. J. Pathol., February 1, 2008; 172(2): 395 - 405. [Abstract] [Full Text] [PDF]
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B. T. Kile, A. D. Panopoulos, R. A. Stirzaker, D. F. Hacking, L. H. Tahtamouni, T. A. Willson, L. A. Mielke, K. J. Henley, J.-G. Zhang, I. P. Wicks, et al. Mutations in the cofilin partner Aip1/Wdr1 cause autoinflammatory disease and macrothrombocytopenia Blood, October 1, 2007; 110(7): 2371 - 2380. [Abstract] [Full Text] [PDF]
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L. A. McNamee and A. G. Harmsen Both Influenza-Induced Neutrophil Dysfunction and Neutrophil-Independent Mechanisms Contribute to Increased Susceptibility to a Secondary Streptococcus pneumoniae Infection Infect. Immun., December 1, 2006; 74(12): 6707 - 6721. [Abstract] [Full Text] [PDF]
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M. Nasirikenari, B. H. Segal, J. R. Ostberg, A. Urbasic, and J. T. Lau Altered granulopoietic profile and exaggerated acute neutrophilic inflammation in mice with targeted deficiency in the sialyltransferase ST6Gal I Blood, November 15, 2006; 108(10): 3397 - 3405. [Abstract] [Full Text] [PDF]
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R. G. King, B. R. Herrin, and L. B. Justement Trem-Like Transcript 2 Is Expressed on Cells of the Myeloid/Granuloid and B Lymphoid Lineage and Is Up-Regulated in Response to Inflammation J. Immunol., May 15, 2006; 176(10): 6012 - 6021. [Abstract] [Full Text] [PDF]
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B. Mehrad, S. J. Park, G. Akangire, T. J. Standiford, T. Wu, J. Zhu, and C. Mohan The lupus-susceptibility locus, sle3, mediates enhanced resistance to bacterial infections. J. Immunol., March 1, 2006; 176(5): 3233 - 3239. [Abstract] [Full Text] [PDF]
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A. M. Condliffe, K. Davidson, K. E. Anderson, C. D. Ellson, T. Crabbe, K. Okkenhaug, B. Vanhaesebroeck, M. Turner, L. Webb, M. P. Wymann, et al. Sequential activation of class IB and class IA PI3K is important for the primed respiratory burst of human but not murine neutrophils Blood, August 15, 2005; 106(4): 1432 - 1440. [Abstract] [Full Text] [PDF]
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A. Lennartsson, K. Pieters, K. Vidovic, and U. Gullberg A murine antibacterial ortholog to human bactericidal/permeability-increasing protein (BPI) is expressed in testis, epididymis, and bone marrow J. Leukoc. Biol., March 1, 2005; 77(3): 369 - 377. [Abstract] [Full Text] [PDF]
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C. R. Brown, V. A. Blaho, and C. M. Loiacono Treatment of Mice with the Neutrophil-Depleting Antibody RB6-8C5 Results in Early Development of Experimental Lyme Arthritis via the Recruitment of Gr-1- Polymorphonuclear Leukocyte-Like Cells Infect. Immun., September 1, 2004; 72(9): 4956 - 4965. [Abstract] [Full Text] [PDF]
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