Abstract
Macrophage inflammatory protein (MIP-1α), a member of the CC chemokine subfamily, has been shown to attract T cells and monocytes in vitro and to be expressed at sites of inflammation. Although the in vitro activities of MIP-1α have been well documented, the in vivo biological activities of MIP-1α in humans have not been studied. To address this, we challenged human subjects by intradermal injection with up to 1000 pmol of MIP-1α and performed biopsies 2, 10, and 24 h later. Although no acute cutaneous or systemic reactions were noted, endothelial cell activation, as indicated by the expression of E-selectin, was observed. In agreement with its in vitro activity, monocyte, lymphocyte, and, to a lesser degree, eosinophil infiltration was observed, peaking at 10–24 h. Surprisingly, in contrast to its reported lack of in vitro neutrophil-stimulating activity, a rapid infiltration of neutrophils was observed in vivo. This neutrophil infiltration occurred as early as 2 h, preceding the appearance of other cells, and peaked at 10 h. Interestingly, we found that neutrophils in whole blood, but not after isolation, expressed CCR1 on their cell surface. This CCR1 was thought to be functional as assessed by neutrophil CD11b up-regulation following whole-blood MIP-1α stimulation. These studies substantiate the biological effects of MIP-1α on monocytes and lymphocytes and uncover the previously unrecognized activity of MIP-1α to induce neutrophil infiltration and endothelial cell activation, underscoring the need to evaluate chemokines in vivo in humans.
Chemokines are a family of small proteins with well-documented chemoattractant activity. These proteins are classified into four groups depending on the presence or absence of intervening amino acids between the first two N-terminal conserved cysteines (1, 2). A number of studies have shown the expression of chemokines at sites of inflammation (3). Although chemokines have overlapping functions and target cell specificities, neutralizing Ab studies in animals have demonstrated their lack of redundancy in various disease models (4, 5, 6, 7, 8, 9). One chemokine in the CC family that has been particularly well studied is macrophage inflammatory protein (MIP-1α)3.
MIP-1α has been shown to be elevated in several inflammatory diseases including rheumatoid arthritis, idiopathic pulmonary fibrosis, sarcoidosis, asthma, and the cutaneous disease, lichen planus (10, 11, 12, 13, 14). In animal model systems, neutralizing Abs to MIP-1α have been shown to decrease disease intensity and limit monocyte and lymphocyte recruitment (4, 6, 7, 8). The in vitro profile of activity of MIP-1α in chemotaxis assays includes effects on B lymphocytes, activated T lymphocytes (CD8>CD4), NK cells, basophils, dendritic cells, and eosinophils (15, 16, 17, 18, 19, 20, 21). Many investigators have documented that MIP-1α has no direct effect on human neutrophil chemotaxis in vitro. In one report, Schall and colleagues (22, 23) demonstrated that MIP-1α induced a small but concentration-dependent intracytoplasmic calcium flux in human neutrophils that was pertussis toxin sensitive.
Although in vitro studies illustrate the potent chemoattractant activity of chemokines under static conditions, they cannot absolutely predict chemokine activity at localized sites in vivo where conditions of blood flow require several adhesion steps before migration into tissue sites can occur (24). Furthermore, chemokines have been shown to bind to proteoglycans and to be modified by surface peptidases; both of these events can influence their chemotactic activity in vivo, underscoring the importance of in vivo experimentation (25, 26, 27, 28, 29). In vivo studies in animals in which chemokines have been injected into various anatomical sites indicate significant species differences in the response to some chemokines, making interpretation of this data difficult (30, 31, 32, 33, 34). For example, studies in mice engineered to be deficient in the major MIP-1α receptor CCR1 indicate that MIP-1α/CCR1 are critical for the neutrophilic response to infection with Aspergillus fumigatus and inflammatory insults such as acute lung injury following pancreatitis (35, 36). Since rodents have no CXCR1 homologue (or IL-8 receptor A), but do have abundant CCR1 on their granulocytes, it has been speculated that MIP-1α and CCR1 may serve the same function in rodents as CXCR1 and IL-8 play in humans (35). Thus, studies in rodents and other animal species have their limitations and make it imperative that in vivo responses be assessed in humans.
To address the biological activity of MIP-1α in vivo in humans and better define its role in disease, we injected MIP-1α into the skin of normal and atopic volunteers. We designed the present studies to determine whether the injection of purified human recombinant MIP-1α elicits the selective recruitment of different leukocyte subtypes. We speculated that by studying the responses to this chemokine in a complex multicellular tissue environment, we might uncover novel effects of MIP-1α not predicted by in vitro studies using highly purified leukocyte subtypes. For example, in our recent analysis of the dermal response to RANTES challenge, we detected activation of microvascular endothelial cells (24). Because allergic subjects have primed or activated eosinophils, T lymphocytes, and monocytes, we also wanted to determine whether allergic subjects have a response that was either quantitatively or qualitatively different from nonallergic subjects (37, 38, 39, 40). Our studies confirm in vitro results demonstrating potent effects of MIP-1α on monocytes and uncover an unexpected and potent effect of MIP-1α on neutrophils and endothelial cells in vivo.
Materials and Methods
Subjects
These studies were approved by the Johns Hopkins Bayview Medical Center Institutional Review Board for human research. Nine otherwise healthy allergic human subjects between the ages of 23 and 50 years were enrolled in the phase I study evaluating the effects of escalating doses of MIP-1α (Table I⇓). Subjects were characterized as allergic if they had a history of respiratory allergies and symptoms referable to these allergies within the last year, an elevated total IgE and a positive multiallergen RAST screen (Phadiatop Kabi Pharmacia Diagnostics, Uppsala, Sweden). Use of antihistamines or systemic glucocorticoids was discontinued for at least 2 wk or 2 mo before this study, respectively. An initial group of subjects was divided into three cohorts of three subjects each to evaluate the safety of intradermal injection of escalating doses of MIP-1α. Successive cohorts were injected with progressively higher doses of MIP-1α as follows: first cohort, 10, 50, and 100 pmol; second cohort, 200, 300, and 400 pmol; and third cohort, 600, 800, and 1000 pmol. A sterile saline control was included in all subjects. Each subject underwent two 6-mm skin biopsies 24 h after injection of saline and the highest dose of MIP-1α.
Characteristics of subjects in the study
Nine allergic and nine nonallergic healthy human subjects between the ages of 18 and 53 years were enrolled in the phase II study to evaluate the kinetics of the response to MIP-1α (1000 pmol) injection (Table I⇑). Subjects were characterized as allergic based on the criteria listed above and as nonallergic if they had normal total IgE, negative multiallergen screen, and no history of allergies or symptoms suggestive of allergic rhinitis, asthma, or atopic dermatitis. All subjects were injected with saline at one site and 1000 pmol MIP-1α at three sites. A 6-mm skin biopsy was performed at the sites 2, 10, and 24 h after MIP-1α injections and at one of those time points for the saline-injected sites. Blood pressure, pulse, body temperature, and signs and symptoms at the injection sites were monitored over the course of the study. A complete blood count with differential was done immediately before the first injection and 24 h later to determine whether intradermal injections of MIP-1α had any effects on the number of circulating leukocytes.
The allergic subjects enrolled in phases I and II had significantly elevated total IgE and multiallergen RAST measurements as compared with nonallergic subjects (Table I⇑). The allergic subjects enrolled in phase II had total circulating eosinophil counts which were significantly elevated compared with those of the nonallergic subjects.
MIP-1α
MIP-1α (lot 95351) was purchased from PeproTech (Princeton, NJ) and was shown to be >99% pure by SDS-PAGE and reversed-phase HPLC analysis. It contained <1.25 endotoxin units/mg protein as determined by the Limulus amebocyte lysate assay (BioWhittaker, Walkersville, MD) and was shown to be free of contaminants by laser desorption mass spectroscopy. The lyophilized preparation was first diluted with sterile water to make a stock solution (778.5 μg/ml) and subsequent dilutions were with sterile saline (no carrier protein). Stability studies demonstrated that the stock solution of MIP-1α maintained bioactivity for at least 6 mo at −70°C and several weeks at 4°C. Aliquots of the stock solution were stored at −80°C and thawed only once just before each injection and then discarded. Intradermal injections (0.1 ml) of MIP-1α or saline were given at four separate sites on the volar forearm, and each site was marked with a surgical pen to provide reference for subsequent clinical evaluations and skin biopsies.
Routine histology, immunohistochemistry, and immunofluorescent staining
Skin biopsy specimens were fixed in Formalin and embedded in paraffin. Eosinophil counts were performed on Wright-Giemsa-stained sections. To further characterize the cellular infiltrate, additional sections (5 μm) were stained with a variety of cell-specific Abs (Table II⇓) and the appropriate controls using the Vectastain ABC-AP kit and the Vector Red substrate kit (Vector Laboratories, Burlingame, CA) and, where noted, with the addition of a permeabilization step. Immunofluorescent staining for the eosinophil granule protein, major basic protein (MBP) was performed to further quantify both the intact and degranulated eosinophils. The number of cells/mm2 was determined by counting an area of 4 × 4 reticules, at ×400 magnification using an Olympus BX60 microscope (Olympus, New Hyde Park, NY) for all markers except MBP. MBP staining was quantified using a scoring system of 0–3+ based on the extent and intensity of the fluorescent stain (41). The scores for the endothelial adhesion molecule E-selectin were calculated by dividing the total number of vessels staining for E-selectin by the total number of von Willebrand factor (VWF)-positive vessels in an adjacent tissue section and multiplying by 100. All specimens were scored in a blinded fashion by two independent investigators. The interrater reliability between the two investigators with regard to cell counts ranged from 63% for CD68+ cells to 99% for neutrophil elastase+ cells.
Abs used in immunohistochemical studies
Endothelial cell cultures and flow cytometry
Endothelial cells used in this study were from collagenase digestion of HUVECs or the immortalized human dermal microvascular endothelial cell line (HMEC, 5A12, a generous gift from Dr. Robert Swerlick, Emory University, Atlanta, GA) (42, 43). Endothelial cells were grown to confluence in 6-well plates (Costar, Cambridge, MA) and either first passage (HUVEC) or passages 25–40 (HMEC) cells were stimulated with either recombinant human MIP-1α (10, 50, or 100 ng/ml), IL-1β (IL-1β, 10 ng/ml), or medium alone for 4 and 24 h (37°C, 5% CO244).
RT-PCR analysis of CCR1 expression
Total RNA was isolated from human neutrophils, eosinophils, mononuclear cells, and HUVECs using the RNazol B method (Tel-Test, Friendswood, TX). Human neutrophils were isolated from whole blood of healthy nonallergic donors, collected into EDTA-containing tubes, and purified by density centrifugation on 1.079 g/ml Percoll (Pharmacia, Uppsala, Sweden). Contaminating erythrocytes were removed by hypotonic lysis and neutrophil purity was enhanced by CD9-negative selection using the immunomagnetic bead technique (final purity ≥ 98%). Human eosinophils were purified to 99% purity from allergic donors using the negative selection immunomagnetic bead technique (45). Mononuclear cells were obtained from buffy coats of healthy blood donors using Ficoll-Hypaque (Pharmacia) discontinuous density gradient centrifugation. These mononuclear cell preparations were composed of 20% monocytes and 80% lymphocytes.
First-strand cDNA was prepared from the total RNA using oligo(dT) primers (Boehringer Mannheim, Indianapolis, IN) and Superscript RT enzyme (Life Technologies, Gaithersburg, MD) as follows: aliquots of 1 μg of total RNA in 5 μl diethylpyrocarbonate-treated water were mixed with 4 μl of 5× First-Strand buffer (250 mM Tris, 375 mM KCl, and 15 mM MgCl2; Life Technologies), 2 μl of 0.1 M DTT, 4 μl of 20 mM dNTP mix, and 2 μl of 25 μM oligo(dT) and heated at 70°C for 10 min, then chilled on ice. Subsequently, 2 μl of 10 U/μl RNase inhibitor (Life Technologies) and 1 μl of 200 U/μl RT were added to the tube. The reaction was conducted in a Hybaid Omnigene thermocycler (Hybaid) for 50 min at 42°C and then for 5 min at 99°C. Aliquots of cDNA were amplified by PCR using Taq polymerase (Life Technologies) and specific primers for CCR1 and β-actin in the same tubes according to the method described previously (46). Briefly, 5 μl of the first-strand cDNA was mixed with 3′ and 5′ primers (20 pmol each), 1 U of Taq DNA polymerase, 5 μl 10× PCR buffer (200 mM Tris and 500 mM KCl, pH 8.4), 2 mM MgCl2, 200 μM dNTP mix, and water to bring the final reaction volume to 50 μl. The samples were sealed with a drop of mineral oil and the PCR reaction was performed as follows: first, a 3-min cycle with a denaturing temperature of 94°C, followed by 30 cycles of 94°C for 30 s, 55°C for 1 min, and 72°C for 1 min. A final elongation step at 72°C for 7 min was performed. The cDNA from mononuclear cells and HUVECs were used as positive and negative controls, respectively. The nucleotide sequences of CCR1 and β-actin primers were 5′ primer, ACT CCG TGC CAG AAG GTG AA and 3′ primer, ATG GCA TCA CCA AAA ACC CA; and 5′ primer, TGA CGC GGT CAC CCA CAC TGT GCC CAT CTA and 3′ primer, CTA GAA GCA TTG CGC TGG ACG ATG GAG GG, respectively. At the end of the reaction, PCR products were separated by electrophoresis on a 2.5% agarose gel. The amplified products were identified based on the predicted size of 251 bp by comparison with positive control and a DNA ladder of known m.w.
Human neutrophil CCR1and CCR5 surface expression
Whole blood was obtained by venipuncture and collected in tubes containing EDTA. Surface expression of CCR1 and CCR5 on neutrophils was examined with indirect immunofluorescence and flow cytometry evaluating granulocytic cells based on scatter characteristics. Concomitant CD16 labeling was used to distinguish eosinophils (CD16−) from neutrophils (CD16+2 goat anti-mouse IgG Ab (Tago, Burlingame, CA). RBCs were lysed after the final labeling step using the whole-blood lysing reagent kit (Coulter) following the manufacturer’s instructions. After fixation with 1% paraformaldehyde in PBS, cells were evaluated using an Epics Profile II flow cytometer (Coulter ). Data are expressed as fold control (mean fluorescence intensity(MFI), CCR1 or CCR5, divided by background MFI).
Whole-blood analysis of CD11b up-regulation
The effect of MIP-1α on CD11b expression on monocytes and neutrophils was determined as described previously (47). Briefly, human blood was collected in EDTA from 18 healthy donors. Blood was prewarmed at 37°C for 5 min before adding various concentrations of MIP-1α (0.01, 0.1, 1.0, 10, 100, and 1000 nM). After a 15-min stimulation, the tubes were placed into an ice bath and 1 ml of cold PBS containing 2% FCS and 0.2% sodium azide was added. The tubes were centrifuged at 200 × g for 10 min at 4°C, and cells were stained for the presence of CD11b. CD11b expression on neutrophils was analyzed by direct immunofluorescence using the FITC-conjugated anti-CD11b Ab (Caltag, Burlingame, CA) and a FITC-conjugated human IgG as a control. Monocytes and neutrophils were identified by their forward and side scatter light profiles. The data were then expressed as a fold increase over baseline and calculated as follows: fold increase = (MFI (chemokine) − MFI (autofluorescence))/(MFI (buffer) − MFI (autofluorescence)).
Statistics
Statistical analysis was performed using the Student t test. A p value of < 0.05 was considered to be significant. Correlations among cell counts or between cell counts and E-selectin expression were tested using the Spearman rank nonparametric method. Data were expressed as the means ± SEM.
Results
Clinical evaluation of human subjects
No significant side effects were noted after MIP-1α injection in any of the 27 subjects who participated in this study, and no subjects dropped out of the study for any reason. Blood pressure, pulse, and body temperature were monitored over the course of the study (at 0.5, 1, 1.5, 2, 10, and 24 h after challenge in phase I, and at 2, 10, and 24 h in phase II) and did not change after MIP-1α injections. Additionally, these injections (cumulative dose, 3000 pmol) did not induce a significant change in peripheral blood neutrophils (3609 ± 275 cells/mm3 before vs 4283 ± 283 cells/mm3 after), lymphocytes (2350 ± 150 cells/mm3 before vs 2406 ± 129 cells/mm3 after), monocytes (384 ± 46 cells/mm3 before vs 448 ± 34 cells/mm3 after), or eosinophils (202 ± 39 cells/mm3 before vs 173 ± 33 cells/mm3 after). No immediate cutaneous reactions were noted in any of the subjects. Several mild reactions were noted 10 or more h after MIP-1α injection at the injection sites in 17 subjects (3/9 subjects in phase I and 14/18 subjects in phase II). These localized reactions occurred at the 1000 pmol MIP-1α injection sites and included tenderness (33%), ecchymosis (33%), and erythema larger than 1 cm in diameter (33%). No immediate reactions were noted in any of the subjects.
Leukocyte recruitment following MIP-1α injection
The cellular infiltrates observed in biopsies taken 24 h after injection of MIP-1α at 100-, 400-, and 1000-pmol doses were assessed by Giemsa and immunohistochemical staining (Fig. 1⇓). A parabolic dose-response curve was observed for CD68+ and CD3+ cell recruitment, with maximal recruitment noted at 400 pmol (Fig. 1⇓, A and C). Unexpectedly, neutrophils were recruited at all three doses, although there was no apparent dose-response relationship for the MIP-1α doses tested (Fig. 1⇓B). Eosinophil migration was quite modest, reaching maximal migration at 1000 pmol (Fig. 1⇓D). Despite this modest eosinophil infiltration, the presence of extracellular MBP staining suggested that the eosinophils were activated as demonstrated by their degranulation (Fig. 2⇓).
Characterization of leukocytes in biopsies 24 h after intradermal injection of allergic subjects with increasing doses (100, 400, 1000 pmol) of MIP-1α or saline. A parabolic dose response was seen for monocytes (A, CD68+ cells) and T lymphocytes (C, CD3+ cells). Eosinophil recruitment (D) was seen only at the 1000-pmol dose and neutrophil tissue migration (B) occurred even at the lowest dose tested. These results were not subjected to statistical analysis because of the small sample size (n = 3/dose of MIP-1α). Data presented as means ± SEM of n = 3 for MIP-1α and n = 9 for saline, respectively.
Dose response of cellular and extracellular MBP staining in skin biopsies obtained 24 h after challenge with increasing doses of MIP-1α (100 pmol, 400 pmol, 1000 pmol) in allergic subjects (n = 3 in each dose for MIP-1α and n = 9 for saline). A, MIP-1α challenge induced dose-dependent eosinophil recruitment (cellular) and degranulation (extracellular). Representative photomicrographs of MBP-stained tissue 24 h after saline challenge (B) or after 1000 pmol of MIP-1α (C) are shown (magnification, ×160).
To determine the kinetics of MIP-1α-induced cellular recruitment, we injected 1000 pmol of MIP-1α at three sites and performed skin biopsies 2, 10, and 24 h later. Results of the cellular phenotyping from this study are presented in Fig. 3⇓. Recruitment of neutrophils was apparent as early as 2 h after MIP-1α injection (Fig. 3⇓A). Recruitment of monocytes (CD68+; Fig. 3⇓B), neutrophils, cutaneous lymphocyte Ag (CLA)+ cells (Fig. 3⇓C), and T lymphocytes (CD3+; Fig. 3⇓D) plateaued by 10 h after challenge, whereas recruitment of eosinophils had not plateaued by 24 h (Fig. 3⇓E). The cellular MBP semiquantitative staining score correlated with eosinophil counts from Giemsa-stained sections (r = 0.64, p = 0.0001), and eosinophil degranulation only reached statistical significance at 24 h. Interestingly, the magnitude and kinetics of leukocyte recruitment did not differ between allergic and nonallergic subjects (data not shown). Therefore, all of the cell counts reported in Fig. 3⇓ represent the combined data of 18 (9 allergic and 9 nonallergic) subjects. The order of magnitude of leukocyte recruitment in response to the 1000-pmol dose listed in order of absolute cell counts was: neutrophils > monocytes > lymphocytes > eosinophils. Importantly, the MIP-1α lot used in this study failed to induce chemotaxis of purified human neutrophils in vitro (data not shown). Representative immunohistochemical and Giemsa-stained sections from skin biopsies taken 24 h after MIP-1α injection are illustrated in Fig. 4⇓, A–E. To better characterize the selectivity of the leukocyte migration in response to intradermal MIP-1α challenge, we compared the differential of the tissue-infiltrating leukocytes 10 h after MIP-1α injection to that seen in the peripheral blood of the same subjects before MIP-1α challenge, as shown in Table III⇓. When the migration was evaluated in the context of the circulating pool of leukocytes, the recruitment of monocytes was the most selective; monocytes represented 5 ± 1% of the circulating leukocytes and 41 ± 3% of the tissue leukocytes.
Kinetic analysis of the cellular recruitment in response to intradermal injection of 1000 pmol MIP-1α in allergic and nonallergic subjects. Cell counts from allergic and nonallergic subjects were combined since there were no significant differences between these two groups (n = 18). Leukocyte counts following saline injections at all three time points (2, 10, and 24 h) did not differ and were therefore averaged. The skin biopsies from allergic subjects had slightly higher CLA+ cell counts than nonallergic subjects, although this did not reach statistical significance (data not shown). No immunofluorescent staining for MBP was detected in saline control sites. Data are presented as means ± SEM of n = 18 allergic and nonallergic subjects. ∗, p < 0.05 vs saline).
Representative tissue sections obtained 24 h after cutaneous challenge with 1000 pmol of MIP-1α. A, Cellular staining for CD68 using a mAb. No staining was noted using a control Ab (inset). B, Staining with mAb directed against neutrophil elastase and control Ab (inset). C, Staining with CD3 polyclonal Ab and negative control rabbit serum (inset). D, Staining with CLA mAb and negative control Ab (inset). E, Giemsa-stained section demonstrating tissue eosinophilia. F, Endothelial staining with E-selectin Ab and negative control Ab (inset). All photomicrographs were taken at ×400 original magnification, with the exception of the E-selectin-stained sections which were ×100.
Selectivity of MIP-1α (1000 pmol)-induced leukocyte recruitment
Endothelial E-selectin expression
Skin biopsies were also analyzed for the expression of the endothelial adhesion molecule E-selectin. We noted a dramatic induction of E-selectin expression in dermal endothelial cells as early as 2 h after MIP-1α (1000 pmol) injection, reaching maximal expression 10 h after challenge (Figs. 4⇑F and 5). There was no significant difference in the endothelial E-selectin expression between allergic and nonallergic groups and therefore their values are combined (data not shown). The extent of endothelial E-selectin expression at 24 h correlated with tissue infiltration by monocytes (CD68+), T lymphocytes (CD3+), and neutrophils (r ≥ 0.49, p = 0.0001). The tissue migration of CLA+ cells did not correlate with E-selectin expression, which is surprising since CLA is thought to be an important ligand for E-selectin (48, 49).
Lack of direct effects of MIP-1α on endothelial cells
To determine whether MIP-1α was capable of directly inducing endothelial adhesion molecule expression, we incubated HUVECs and HMECs with MIP-1α (10, 50, and 100 ng/ml) in vitro for 4 or 24 h and performed flow cytometry with Abs to the endothelial adhesion molecules E-selectin, VCAM-1, and ICAM-1 (n = 2). MIP-1α had no effect on the expression of any of these endothelial adhesion molecules at any of the concentrations or stimulation times tested (data not shown). IL-1β (10 ng/ml × 4 h) stimulation was used as a positive control and induced expression of all three adhesion molecules (data not shown).
Human neutrophil CCR1 and CCR5 expression
The influx of neutrophils in response to intradermal MIP-1α challenge (Figs. 1⇑B, 3A, and 4B) suggested that these leukocytes might express one of the MIP-1α receptors CCR1 or CCR5 (15, 50). To investigate this possibility, we analyzed neutrophils for the expression of CCR1 mRNA using RT-PCR and for surface expression of CCR1 and CCR5 using flow cytometry. Fig. 6⇓A shows that highly purified (99%) neutrophils from four donors constitutively express CCR1 mRNA (lanes 4–7). Eosinophils (>99% purity, lane 1) are shown as a positive control, and HUVECs and water (lanes 3 and 8, respectively) as negative controls. To demonstrate that the small number of contaminating eosinophils in the neutrophil samples did not explain our ability to amplify CCR1 mRNA, we simultaneously analyzed mRNA isolated from a CCR1-negative HUVEC sample (lane 3, HUVEC alone) to which we added a number of eosinophils equal to those contaminating the neutrophil preparation (lane 2, HUVEC and 1% eosinophils). Because we did not detect CCR1 mRNA expression under these conditions, we concluded from these studies that neutrophils themselves express CCR1 mRNA.
Cutaneous endothelial E-selectin expression at various time points after MIP-1α (1000 pmol) injection (n = 18, 9 allergic and 9 nonallergic subjects). Saline values from all three time points were combined since there were no differences among the three time points (2, 10, and 24 h; n = 18). The E-selectin values were calculated by dividing the number of vessels staining for E-selectin by the total number of vessels staining with the pan-endothelial marker VWF and multiplying by 100. Data are presented as means ± SEM. ∗, p < 0.05 vs saline).
Human neutrophils express CCR1 as determined by RT-PCR and flow cytometry. A, CCR1 mRNA expression was amplified by RT-PCR using mRNA isolated from neutrophils from four separate donors (lanes 4–7). Eosinophils (>99% purity, lane 1) are shown as a positive control (lane 1), and HUVECs and water (lanes 3 and 8, respectively) as negative controls. To ensure that the signal observed in the neutrophil preparations could not be due to the ∼1% eosinophil contamination, eosinophils were added back to HUVECs to obtain a final concentration of 1% (lane 2). The amplified products were the predicted size of 251 bp, as determined by comparison with a DNA ladder of known m.w. PCR for β-actin was performed to confirm the integrity of the RNA for each sample (data not shown). Surface expression of CCR1 (B) and CD16 (C) on granulocytes from healthy human donors (representative of six donors) is shown. Histograms of flow cytometric analysis with an anti-CCR1 (2D4) or anti-CD16 (3G8) mAb are represented by open histograms and the IgG1 isotype controls are depicted by the filled histograms.
We next determined whether CCR1 and CCR5 were expressed on the surface of leukocytes using flow cytometry. Initial studies performed on purified neutrophils (n = 8) were unable to detect any CCR1 or CCR5 surface expression, and, as expected, these neutrophils did not have a functional response to MIP-1α stimulation (e.g., chemotaxis or up-regulation of CD11b surface expression). Subsequent studies were performed on whole-blood preparations from healthy subjects (n = 6). In these studies, we observed an average MFI of 2.1 ± 0.3-fold control, with 31% positivity for CCR1 and no expression of CCR5 (1.0 ± 0.1-fold control). A representative CCR1 histogram is shown in Fig. 6⇑B. The cells within the granulocyte scatter region were almost entirely neutrophils as determined by their CD16 staining (99.5 ± 0.4% positivity, Fig. 6⇑C). Therefore, the CCR1 surface expression could not be attributed to eosinophils. Interestingly, the neutrophil CCR1 expression was greater than that seen on the cells found within the lymphocyte scatter, which were at the limit of detection (MFI, 1.1 ± 0.1-fold control; 9% positive), but considerably less than that seen on cells within the monocyte scatter (MFI, 14.4 ± 2.9-fold control; 88% positive; data not shown). One interpretation of these findings is that neutrophil CCR1 expression is lost during cell isolation. Next, we determined the effects of MIP-1α on human neutrophils in vitro.
In initial studies, we were unable to demonstrate chemotaxis or up-regulation of surface expression of the integrin subunit CD11b with MIP-1α on isolated neutrophils, in agreement with the work of other investigators (22, 23). In contrast, when freshly collected whole blood was stimulated for 15 min with MIP-1α (10 nM), CD11b up-regulation was observed (n = 18 donors) on both neutrophils (MFI-fold increase, 1.4 ± 0.1) and monocytes (MFI-fold increase, 2.1 ± 0.1) as shown in Fig. 7⇓, A and B, respectively. Although the magnitude of this increase was small, the EC50 for this response, as shown in Fig. 7⇓C, was comparable to that achieved using leukotriene (LT)B4 as the agonist (EC50, 3.8 nM for MIP-1α vs 4.0 nM for LTC4). Whole-blood chemotaxis assays were attempted but were futile, since erythrocytes prevented any leukocyte migration because they plugged the filter in the Boyden microchemotaxis chamber (Neuroprobe, Cabin John, MD).
MIP-1α up-regulates surface expression of CD11b on neutrophils and monocytes. Representation examples (n = 18 donors) demonstrating CD11b up-regulation on whole-blood neutrophils (A) or monocytes (B) after a 15-min, 37°C incubation with 10 nM MIP-1α are shown. Resting CD11b expression is depicted by the filled histograms, and the MIP-1α-stimulated CD11b expression is depicted by the open histograms. There was no change in autofluorescence of either cell type in response to MIP-1α. The EC50 value for this response in neutrophils was comparable to that achieved using LTB4 as the agonist (C, EC50, 3.8 nM for MIP-1α vs 4.0 nM; n = 2–4).
Discussion
MIP-1α is a chemokine thought to be important in several diseases characterized by the recruitment and activation of mononuclear cells. In vitro studies to date have led to the belief that it is not active as a neutrophil chemoattractant (3). However, our studies indicate that MIP-1α, either by direct or indirect effects, is a neutrophil chemoattractant in vivo in humans. We found that intradermal challenge with even low doses of MIP-1α induced a marked influx of neutrophils. We also observed that intradermal injection of MIP-1α led to the activation of microvascular endothelial cells. Both of these findings could have important, unrealized implications for the role of MIP-1α in disease.
The infiltration of lymphocytes and monocytes into intradermal sites following MIP-1α injection is in agreement with in vitro chemotaxis studies. Memory T lymphocyte migration into skin is believed to rely on expression of the CLA Ag (51). However, characterization of the lymphocytes infiltrating the skin after MIP-1α challenge revealed that only 50% of these cells expressed CLA. Furthermore, although CLA+ cells bind to endothelial E-selectin, which is believed to be responsible for their migration to skin (52), in our study there was no correlation between the percentage of vessels staining for E-selectin and the number of CLA+ cells following MIP-1α challenge. It is also interesting to note that when comparing the percentage of infiltrating cells with the number of cells in the periphery of the same individual, lymphocyte recruitment in response to MIP-1α was the least selective as compared with the other cell types (Table III⇑). In contrast, MIP-1α induced the most selective infiltration of monocytes. This supports the hypothesis that MIP-1α may be a critical cytokine in chronic inflammatory conditions characterized by large numbers of tissue monocytes/macrophages, such as multiple sclerosis and rheumatoid arthritis (6, 8).
The magnitude of monocyte and lymphocyte infiltration at 24 h after challenge increased from the 100–400 pmol dose, but was less at the 1000 pM dose. The reason for this parabolic dose-response curve is unclear. One possible explanation is that at higher concentrations, diffusion of MIP-1α into the systemic circulation desensitizes (and internalizes) MIP-1α receptors on circulating leukocytes, resulting in less migration. However, since neutrophil and eosinophil migration was not decreased at higher dose levels, such a systemic receptor desensitization seems unlikely. Alternatively, one could speculate that at higher dose levels, MIP-1α is also activating cells as they infiltrate the skin sites, releasing factors that may feedback and dampen further cell infiltration. Further studies will be necessary to address these possibilities.
The robust infiltration of neutrophils occurring following MIP-1α injection was unexpected based on in vitro results (23, 50, 53, 54). In a similar study with RANTES, another CC chemokine which shares CCR1 and CCR5 with MIP-1α, no neutrophils were observed. Possible explanations for these paradoxical results include the presence of a unique chemokine receptor for MIP-1α on neutrophils, the possibility that RANTES does not stimulate sufficient signaling through their shared receptors (CCR1 or CCR5), or that the neutrophil infiltration was secondary to the release of other factors. Since the kinetics of neutrophil infiltration was rapid, especially as compared with other cell types, and the neutrophil infiltration response was near maximal at a dose of MIP-1α (100 pmol) that was too low to recruit other cell types (Fig. 1⇑), an indirect effect involving other infiltrating cells seems less likely. We have recently made the observation that epithelial cells express CCR1 and CCR5 (S. Shahabuddin, unpublished data). Therefore, we cannot rule out the possibility that MIP-1α may act on other resident cells, such as epithelial cells, to cause the release of inflammatory mediators which in turn recruit neutrophils. Of note, no neutrophil chemotactic activity was observed in supernatants from a resident tissue cell (HUVEC) stimulated with MIP-1α despite the fact that these cells have been shown to express relevant receptors (CCR4 and CCR5) (55).
Although most investigators have found that MIP-1α is not chemotactic for purified human neutrophils, we (and others) have shown that these cells express mRNA for the MIP-1α receptor CCR1, and we demonstrate here the constitutive surface expression of CCR1 by flow cytometry (Fig. 6⇑) (53, 54). Interestingly, we could only detect CCR1 surface expression on neutrophils from whole-blood preparations and could not detect surface expression on cells that had undergone isolation procedures, including density gradient centrifugation and hypotonic RBC lysis (data not shown). In agreement with these results, we detected MIP-1α-dependent CD11b up-regulation on neutrophils from whole blood but not in purified neutrophils. This suggests that the standard isolation techniques may affect neutrophil chemokine receptor surface expression and might help explain the lack of MIP-1α chemotactic activity on purified neutrophil preparations. A possible explanation for this is the release of RANTES or MIP-1α from platelets during the isolation procedure, since platelets are a rich source of these CC chemokines (56). RANTES release could desensitize and internalize CCR1 and thereby prevent responses to CCR1 ligands in vitro (57). Interestingly, Bonecchi et al. (53) have recently demonstrated that IFN-γ up-regulates CCR1 mRNA and surface expression on purified neutrophils, and that these stimulated neutrophils now undergo chemotaxis to MIP-1α and RANTES in vitro. Collectively, this work suggests that MIP-1α may directly or indirectly play a role in the development of tissue neutrophilia, in sharp contrast to other members of the CC chemokine subfamily. In support of this notion, Murch et al. (58) have shown that infants who develop bronchopulmonary dysplasia have increased numbers of neutrophils and elevated MIP-1α in their bronchoalveolar lavage fluid.
In addition to causing leukocyte recruitment, we also observed an effect of MIP-1α on endothelial cell activation in vivo. This suggests that chemokines may recruit leukocytes in vivo in part by inducing endothelial adhesion molecule expression. The up-regulation of selectins on endothelial cells would be expected to facilitate leukocyte rolling under flow and thus prepare them for subsequent steps of integrin-induced firm adherence and chemotaxis into tissue. There was a significant correlation between endothelial E-selectin expression and the influx of all leukocytes, suggesting that the endothelial activation resulting from MIP-1α challenge may have contributed to cell migration. Since we could not detect enhanced expression of ICAM-1, E-selectin, or VCAM-1 on HUVECs and HMECs stimulated with MIP-1α in vitro, it is likely that endothelial activation in vivo occurred indirectly, possibly due to the release of endothelial-activating cytokines (e.g., IL-1 or TNF) by other cell types. Although monocytes have been shown to release TNF-α in response to MIP-1α, a close view of the kinetics of monocyte recruitment and E-selectin expression suggests that the endothelial activation preceded significant monocyte influx. Alternatively, MIP-1α may induce IL-1 or TNF release from other cell types or the ability of MIP-1α to activate endothelium in vivo may require factors not provided in our tissue culture systems (59). The E-selectin expression is not due to endotoxin because the levels detected in the MIP-1α preparation were <1.25 endotoxin units/mg protein (60).
Strikingly different results were observed following the intradermal challenge of RANTES as compared with MIP-1α. Intradermal RANTES challenge led to a selective migration of eosinophils and lymphocytes, whereas MIP-1α recruited neutrophils and monocytes as well as lymphocytes and eosinophils. The recruitment of eosinophils by 500 pmol of RANTES was substantially greater than that seen with 1000 pmol of MIP-1α (112 ± 23 cells/mm2 vs 26 ± 6 cells/mm2, respectively). There were no clear differences in the kinetics of eosinophil tissue infiltration between allergic and nonallergic subjects following MIP-1α challenge, in contrast to the accelerated migration of eosinophils in allergic subjects that occurred with RANTES. Furthermore, the allergic phenotype of enrolled subjects had no influence on the kinetics or magnitude of the monocyte, neutrophil, or lymphocyte response to MIP-1α, suggesting that allergic status does not affect the in vivo response to MIP-1α. A likely explanation for the greater eosinophil response observed in the RANTES study may relate to the ability of RANTES, but not MIP1α, to interact with the eosinophil receptor CCR3.
Although MIP-1α is known best for its effects on cell migration, it is thought to have several other important biological functions which include cell activation, mast cell and/or basophil histamine release, and PG-independent pyrogenic activity (30, 61, 62). Our results suggest that MIP-1α skin challenge can induce eosinophil activation as measured by interstitial MBP staining. We were unable to assess monocyte or neutrophil activation in this model. MIP-1α had no effect on cutaneous mast cell histamine release since no acute wheal-and-flare reactions were seen in any of the 27 subjects injected with MIP-1α in doses ranging from 100 to 1000 pmol. This is in contrast to studies by Alam et al. (63) demonstrating that human MIP-1α induced mouse footpad swelling, which was maximal 30 min after injection and was felt to represent mast cell degranulation based on electron microscopic examination. These differences may be due to differences in chemokine and chemokine receptor specificity between mouse and human. Our study was not designed to test the effects of MIP-1α on basophil mediator release, although it is possible that the erythema seen 10 h after 1000 pmol MIP-1α in approximately one-third of the subjects may have been secondary to basophil recruitment and activation. Even with a cumulative dose of 3000 pmol (24 μg) per subject, we were unable to detect any elevations in body temperature during monitoring of subjects for up to 24 h after their last intradermal injection. These findings suggest either that MIP-1α does not have pyrogenic effects in vivo in humans or that these localized cutaneous injections were not sufficient to develop systemic levels of MIP-1α.
In summary, we demonstrate that MIP-1α is a potent in vivo leukocyte recruiting agent in humans. Relative to the corresponding peripheral cell numbers, MIP-1α is most selective for monocytes. However, we also identified previously unrecognized effects of MIP-1α on endothelial cells and neutrophils in vivo. We have shown that the tissue neutrophilia could be due to a direct effect of MIP-1α on neutrophils but we cannot exclude the possibility that this neutrophilia was due to secondary mediators released by other cells within the tissue. The potent in vivo effect of MIP-1α on tissue neutrophilia raises important questions about the role of MIP-1α and its receptor, CCR1, in neutrophil-mediated diseases. Our results also provide further support for the contention that the human challenge model we have developed will yield new insights into the role of chemokines in human diseases.
Acknowledgments
We thank Drs. Paul Ponath and Walter Newman from LeukoSite Inc. for the anti-CCR1 (2D4) Ab, Dr. Robert Swerlick from Emory University for the HMEC cell line, Joanne Alsruhe for her assistance in histopathology, Jim Plitt for guidance with RT-PCR, Sherry Hudson and Dr. Bruce Bochner for help with the flow cytometry experiments, and Bonnie Hebden for her expert secretarial assistance in the preparation of this manuscript.
Footnotes
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↵1 This work was supported by National Institutes of Health Grants AI01226-03 and AR31891 and by Pfizer, Inc.
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↵2 Address correspondence and reprint requests to Dr. Lisa A. Beck, Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Circle, Baltimore, MD 21224-6801. E-mail address: lab{at}jhmi.edu
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↵3 Abbreviations used in this paper: MIP-1α, macrophage inflammatory protein-1α; MBP, major basic protein; CLA, cutaneous lymphocyte Ag; VWF, von Willebrand factor; HMEC, human microvascular dermal endothelial cell; MFI, mean fluorescence intensity.
- Received August 16, 1999.
- Accepted January 3, 2000.
- Copyright © 2000 by The American Association of Immunologists
References
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