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Am J Physiol Heart Circ Physiol 291: H694-H704, 2006. First published March 24, 2006; doi:10.1152/ajpheart.01271.2005
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Immune complexes alter cerebral microvessel permeability: roles of complement and leukocyte adhesion

Karyn J. Lister and Michael J. Hickey

Department of Medicine and Monash Institute of Medical Research, Monash University, Victoria, Australia

Submitted 1 December 2005 ; accepted in final form 20 March 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Immune complexes (ICs) are potent inflammatory mediators in peripheral tissues. However, very few studies have examined the ability of ICs to induce inflammatory responses in the brain. Therefore, using preformed ICs or the reverse passive Arthus (RPA) model to localize ICs to the pial microvasculature of mice, we aimed to investigate the ability of ICs to induce an inflammatory response in the cerebral (pial) microvasculature. Application of preformed ICs immediately increased pial microvascular permeability, with a minimal change in leukocyte adhesion in pial postcapillary venules. In contrast, initiation of the RPA response in the pial microvasculature induced changes in cerebral microvascular permeability and increased leukocyte adhesion in pial postcapillary venules. The RPA response induced deposition of C3 in perivascular regions adjacent to sites of IC formation. Depletion of C3 abrogated RPA-induced microvascular permeability and leukocyte adhesion, indicating that the complement pathway was critical for this response. Inhibition of leukocyte adhesion via CD18 blockade also reduced IC-induced microvascular permeability. However, this did not require intercellular adhesion molecule-1, inasmuch as blockade of intercellular adhesion molecule-1 did not alter RPA-induced microvascular permeability and adhesion. These findings demonstrate that ICs are capable of rapidly inducing inflammatory responses in the cerebral microvasculature, with the complement pathway and leukocyte recruitment playing critical roles in microvascular dysfunction.

inflammation; cell trafficking; adhesion molecules

IMMUNE COMPLEXES (ICs) are potent proinflammatory mediators that are believed to contribute to inappropriate tissue inflammation in several autoimmune diseases, particularly systemic lupus erythematosus (SLE) (29). The mechanisms of IC-mediated inflammation in peripheral microvasculatures, such as skin and muscle, have been extensively characterized (39, 40, 51). In these tissues, ICs induce rapid changes in microvascular physiology, whereby microvascular permeability and leukocyte-endothelial cell interactions are increased, culminating in leukocyte entry into the tissue. However, it is not known whether ICs are capable of inducing similar changes in the cerebral microvasculature. This is of interest, inasmuch as the brain is among the most commonly affected organs in SLE patients, with autoantibodies and ICs being regularly detected in the cerebral spinal fluid (CSF) and the choroid plexus of the central nervous system (CNS) (7, 29, 56). Furthermore, ICs have been detected in the CNS in association with neurological disorders, such as multiple sclerosis, cerebral malaria, and viral encephalitis (1, 18, 55), as well as other diseases, such as acquired immunodeficiency syndrome, liver cirrhosis, and hypertension (15, 41, 42). However, despite the well-documented proinflammatory abilities of ICs in the peripheral microvasculature, very few studies have investigated the ability of ICs to induce inflammatory responses in the unique microvasculature of the brain.

There is a growing body of evidence that the mechanisms of the inflammatory response in the brain are distinct from those in the periphery. The CNS is considered an immunoprivileged site because of the presence of the blood-brain barrier (BBB) (32), a structural and functional barrier within the microvasculature that restricts the entry of macromolecules and immune cells into the CNS (17, 43). Furthermore, chemoattractants or proinflammatory mediators such as N-formyl-methionyl-leucyl-phenylalanine and TNF, which induce inflammation in peripheral tissues, fail to induce leukocyte recruitment when injected directly into the CNS (4). Physiological assessment of the cerebral microvasculature has provided some explanation for these observations. First, in contrast to many peripheral tissues, constitutive leukocyte rolling is absent or extremely rare in the pial microvasculature (10, 24). Also, the ability of cerebral microvessels to upregulate expression of the adhesion molecules P-selectin, E-selectin, and vascular cell adhesion molecule-1 is markedly restricted relative to tissues such as skin and intestine (10, 19). Additionally, direct examination of the CNS has shown that activated T cells can become adherent in uninflamed CNS white matter microvessels without first undergoing the rolling interactions normally considered a prerequisite for leukocyte adhesion on the endothelial surface (53). Taken together, these observations indicate that the cerebral microvasculature responds to inflammatory stimulation in a highly unique manner. Therefore, it cannot be assumed that the mechanisms of IC-mediated inflammation in the brain are similar to those already elucidated in other tissues.

Despite the refractory nature of the brain to inflammatory stimulation, some observations raise the possibility that ICs could induce inflammation in the cerebral microvasculature. Complement proteins, which are central to IC-mediated responses, are synthesized by CNS-resident brain cells (54). Furthermore, cerebral expression of complement receptors has been shown to be important in the development of brain pathology in experimental autoimmune encephalomyelitis (9, 36, 54). Additionally, Fc receptors, which are essential in facilitating cellular responses in IC-mediated inflammation, are expressed on CNS-resident immune cells (48). These findings demonstrate that many of the cellular and molecular requirements for IC-mediated responses are present in the brain. Therefore, the aim of this study was to determine whether ICs are capable of inducing responses in the cerebral microvasculature. To achieve this aim, we developed a modified model of IC-mediated inflammation in which ICs are localized to the cerebral microvasculature and assessed local alterations in microvascular permeability and leukocyte-endothelial cell interactions. These results reveal that ICs are capable of rapidly inducing an inflammatory response in the cerebral microvasculature.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. Male C57BL/6J wild-type mice (6–12 wk, 20–30 g body wt) were bred in-house at Monash University. The Monash Medical Centre Animal Ethics Committee approved all protocols involving animal use.

Antibodies. Polyclonal rabbit anti-ovalbumin IgG (anti-OVA) and rabbit IgG were obtained from Sigma Chemical (St. Louis, MO). 2E6 (a MAb against murine CD18) and YN1/1.7.4 [a MAb against murine intercellular adhesion molecule-1 (ICAM-1)] were purified from hybridoma supernatant. To assess the role of CD18, the mice were treated with 2E6 (4 mg/kg iv) before initiation of the reverse passive Arthus (RPA) protocol. For assessment of the role of ICAM-1 as a possible ligand for beta2-integrins, the mice were treated with YN1/1.7.4 (2 mg/kg iv) before initiation of the RPA protocol.

Cranial window preparation. A closed cranial window technique was performed as previously described (23, 33). The animals were anesthetized by intraperitoneal injection of a mixture of ketamine hydrochloride (150 mg/kg; Caringbah, NSW, Australia) and xylazine (10 mg/kg; Bayer Pharmaceuticals, Pymble, NSW, Australia). The lateral tail vein was catheterized for administration of additional anesthetic and reagents. The left femoral artery was catheterized for periodic arterial blood sampling. The animal’s head was held in a stereotaxic board, and core temperature was maintained at 37°C with a thermocontrolled heating pad. The skull was exposed by a longitudinal skin incision, and the periosteum over the parietal bone was removed. A craniotomy was performed in the right parietal bone with a high-speed microdrill (Fine Science Tools, North Vancouver, BC, Canada). A stainless steel superfusion chamber (2 mm high, 7 mm ID) was designed with an outer rim extending horizontally from the chamber (15 mm diameter, 0.4 mm wide) and modified to include a concavity (8 mm radium) shaped to the mouse skull. The chamber contained two ports: one for attachment of polyethylene tubing for superfusion of the brain surface with artificial CSF (aCSF) and the other for collection of aCSF samples after superfusion. The chamber was sealed to the skull with bone wax (Lukens Medical, Albuquerque, NM) and secured in place with rapid adhesive (catalog no. 406, Loctite, Caringbah, NSW, Australia). Before exposure of the pial surface, the chamber was filled with aCSF (in mM: 132 NaCl, 24.6 NaHCO3, 2.95 KCl, 1.4 MgSO4, 6.7 urea, 3.71 glucose, and 1.71 CaCl2, pH 7.4, 37°C), which was continuously gassed with 12% O2-5% CO2-83% N2. The bone cap and the underlying dural membrane were removed to expose the pial vessels. The chamber was then sealed using a coverslip held in place with vacuum grease, and aCSF infusion was maintained at 0.3 ml/min. The outlet tubing was positioned 10 cm above the mouse brain to maintain intracranial pressure at 5–8 mmHg throughout the experiment (33).

Cerebral microvascular permeability. Microvascular permeability of the pial vessels was assessed using a modification of a previously published technique (22, 38). The mice were anesthetized, and the closed cranial window was prepared as already described. At the start of the experiment, the animals received a bolus dose (125 mg/kg iv) of FITC-70-kDa dextran (50 mg/ml; Sigma Chemical). aCSF was continuously superfused across the cranial window at 0.3 ml/min for 60 min and collected for the last 2 min of every 5-min period. Clearance of FITC-70-kDa dextran into the aCSF (nl/s) was measured to assess permeability of the pial microcirculation. To determine plasma FITC-dextran concentration, arterial blood samples (50 µl) were taken at 20-min intervals after initiation of the RPA protocol. The concentration of FITC-derived fluorescence in the aCSF and the plasma samples was measured on a 96-well microplate reader (485-nm excitation wavelength and 520-nm emission wavelength; FLUOstar Optima, BMG Labtechnologies, Mt. Eliza, Victoria, Australia), and concentrations were determined by reference to a standard curve. Clearance was determined as the ratio of fluorescence in aCSF to fluorescence in plasma, multiplied by the flow rate of aCSF across the brain (22, 38).

Intravital microscopy. Intravital microscopy of the pial microvasculature was performed as previously described (22). The pial microvasculature was visualized using an intravital microscope (Axioplan 2 Imaging, Carl Zeiss) with a x40 water immersion objective lens (Achroplan x40/0.80 numerical aperature (NA), Carl Zeiss). Images were visualized using a silicon intensifier target video camera (Dage-MTI VE-1000, Sci Tech) to project the images onto a calibrated monitor (model PVM-20N5E, Sony) and recorded for playback analysis using a video-cassette recorder (model NV-HS950, Panasonic, Klapp Electronics, Prahran, Victoria, Australia). Leukocytes were visualized by intravenous injection of 50 µl of 0.05% rhodamine 6G (Sigma Chemical) immediately before each recording. Epifluorescence microscopy (510–560 nm excitation, 590 nm emission) was used to visualize leukocyte-endothelial cell interactions. In each experiment, three postcapillary venules (25–50 µm diameter) were chosen, and the same section was recorded at each time point. Leukocyte rolling flux, rolling velocity, and adhesion were analyzed off-line during video playback analysis. Rolling leukocytes were defined as those moving at a velocity less than that of an erythrocyte within a given vessel. Leukocyte rolling velocity was measured as the time required for a leukocyte to roll along a 100-µm length of the venule. Adherent leukocytes were defined as those that remained stationary for ≥30 s and are shown as the number of cells per square millimeter of pial postcapillary venule surface area (calculated assuming cylindrical vessel geometry). Red blood cell velocity within pial postcapillary venules was determined by analysis of the velocity of 1-µm-diameter fluorescent polystyrene microspheres (FluoSpheres-yellow/green, Molecular Probes, Eugene, OR) injected intravenously before recording, as previously described (22). The microspheres were visualized by epifluorescence (450- to 490-nm excitation and 515 nm emission; Carl Zeiss filter set 09). Intravital microscopy video images were digitized to sequences. After calibration appropriate to the magnification, red blood cell velocity was determined by measurement of the velocity of 20 randomly selected microspheres in two to three postcapillary venules with ImageJ 1.34s software (http://rsb.info.nih.gov/ig/; National Institutes of Health, Bethesda, MD).

Preparation and administration of ICs in pial microvasculature. Soluble ICs were generated using a modification of a previously published technique (49). On the basis of a point of equivalence determined using a precipitin reaction (6, 30), soluble ICs were produced by combination of OVA (Sigma Chemical) and anti-OVA at 10 times antigen excess: 75 µg of OVA and 150 µg of anti-OVA in 1 ml of aCSF for 1 h at 37°C. ICs were then applied to the cerebral microvasculature by superfusion over the pial surface for 10 min. Superfusion was restored to aCSF for the subsequent 60 min. For control mice, OVA was combined with nonspecific rabbit IgG under identical conditions, and the resultant mixture was applied to the pial microvasculature. Leukocyte recruitment and microvascular permeability responses were assessed at various times after administration of preformed ICs. In experiments aimed at examining the response of a peripheral microvasculature to preformed ICs, ICs generated as described above were injected intrascrotally (in 200 µl of saline) into wild-type mice. After 4 h, the mice were anesthetized, and the cremaster muscle was prepared for intravital microscopy exactly as described previously (39, 40). Leukocyte trafficking parameters in cremasteric postcapillary venules were assessed as previously described (39, 40).

RPA reaction in pial microvasculature. The RPA reaction was used as an alternative model of IC-induced inflammation (40). The RPA response was induced by intravenous injection of 1,000 µg of OVA (Sigma Chemical, 200 µl of 5 mg/ml, in sterile saline), followed by superfusion of anti-OVA over the pial surface at 5 or 50 µg/ml in aCSF. Control animals also received OVA intravenously but were superfused with nonspecific rabbit IgG at the same concentration. Leukocyte recruitment and microvascular permeability responses were assessed at various times after initiation of the RPA response.

Depletion of complement. To deplete animals of serum complement, we administered purified cobra venom factor (CVF; Venom Supplies, Tanunda, South Australia, Australia) derived from Naja melanoleuca venom intraperitoneally at 25 µg/animal 24 h before RPA experiments. Similar doses have previously been shown to reduce C3 in plasma to <3% of normal levels (14, 31). To confirm the ability of the brain to respond to C3-independent stimuli after CVF treatment, pial permeability was assessed after 5 min of superfusion with 100 µM bradykinin in aCSF (Auspep, Parkville, Victoria, Australia).

Location of OVA and anti-OVA during cerebral RPA reaction. To determine the location and timing of IC formation during the RPA response in the cerebral microvasculature, a protocol similar to that previously described for the cremaster muscle was used (40). To examine the location of OVA and anti-OVA during the first 10 min of the RPA response, the RPA procedure was performed with fluorochrome (Alexa Fluor 488, Molecular Probes)-conjugated OVA (OVA-488) and nonfluorescent anti-OVA or, conversely, Alexa Fluor 488-conjugated anti-OVA (anti-OVA-488) and nonfluorescent OVA. At 10 min after initiation of the response, brain tissue within the cranial window was removed and snap frozen in OCT embedding medium, and 10-µm coronal sections were prepared on a cryostat for examination by confocal microscopy. The location of blood vessels was determined by staining with the endothelium-specific isolectin IB4 (from Griffonia simplicifolia) conjugated to Alexa Fluor 594 (Molecular Probes) (44). Sections were stained for 30 min at 10 µg/ml, washed in PBS, mounted in antifade fluorescent mounting medium (Dako, NSW, Australia), and examined using confocal microscopy. Confocal images were collected using an inverted microscope (model FV1000, Olympus) with x60 (NA 1.25) and x100 (NA 1.4) oil immersion lenses. Alexa Fluor 488 was imaged by excitation at 488 nm (Ar laser) and Alexa Fluor 594 by excitation at 543 nm (HeNe laser). Images were collated using Soft Imaging System software (SIS, Münster, Germany).

In addition, fluorescent intravital microscopy was used to determine the distribution of anti-OVA during the RPA response over 60 min by a modification of a previously published technique (40). Alexa 594-conjugated anti-OVA (anti-OVA-594) was superfused across the surface of pial vessels in the closed cranial chamber in the presence of intravenously injected OVA-488. The location of anti-OVA-594 during the RPA response was determined using a 530- to 585-nm excitation filter with a 615-nm emission filter (Carl Zeiss filter set 00), and OVA-488 was determined by epi-illumination at 450–490 nm with a 515-nm emission filter (filter set 09, Carl Zeiss). Additional experiments were performed to analyze the distribution of anti-OVA-594 in the absence of OVA. Images were captured (PC2-Vision capture board, Coreco Imaging, Billerica, MA) directly from the silicon intensifier target video camera. Scion Image analysis software (Scion, Frederick, MD) was used to determine the mean fluorescence intensity of anti-OVA-594 in a region of interest positioned immediately adjacent to the vessel. Fluorescence intensity was derived by subtraction of background fluorescence intensity, which was determined before administration of anti-OVA-594, and expressed as mean intensity units.

To determine whether OVA colocalized with anti-OVA during the RPA response, brain tissue was removed from experiments involving OVA-488 and anti-OVA-594. Brain tissue was snap frozen in OCT embedding medium, and 10-µm coronal cryostat sections were prepared for confocal microscopy, as described previously.

Immunohistochemical identification of C3 deposition. The section of brain underlying the cranial window was removed, fixed in periodate-lysine-paraformaldehyde, cryoprotected in 20% sucrose-PBS, and frozen over liquid nitrogen (40, 45). Cryostat sections (10 µm) were prepared and stained for C3 deposition with FITC-conjugated goat anti-mouse C3 (Cappel Laboratories/ICN Biomedical Australasia, Seven Hills, NSW, Australia), as previously described (40). The sections were preincubated with 10% goat serum in 5% BSA-PBS for 10 min and then stained with anti-mouse C3 diluted 1:400 in 1% BSA-PBS for 30 min. The slides were washed in PBS, mounted in aqueous mounting medium, and examined using epifluorescence microscopy (40, 45).

Circulating leukocyte counts. At the conclusion of each experiment analyzing leukocyte recruitment, whole blood was obtained by cardiac puncture. Total leukocyte counts were determined by using a Neubauer hemocytometer (U-Lab, Eltham, Australia).

Statistical analysis. Values are means ± SE. For comparisons involving only two groups, Student’s t-tests were used. For comparisons involving three groups, ANOVA followed by Scheffé’s post hoc tests or Student’s t-test followed by Bonferroni’s multiple comparisons test was used. Statistical significance was set at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effects of extravascular ICs on cerebral microvasculature. ICs generated in vitro are commonly used to examine the effect of ICs in vivo (12, 30, 49). However, in most studies, ICs were administered into the circulation. Our aim was to administer preformed ICs directly to the brain. Therefore, before applying this method to the brain, we sought to determine the response of a peripheral tissue to extravascular IC deposition. Similar to our recent experiments with the RPA model in the cremaster muscle (40), local application of in vitro-generated ICs induced a reduction in leukocyte rolling velocity (data not shown) and increases in leukocyte adhesion and emigration in cremasteric postcapillary venules (Fig. 1).


Figure 1
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  		Fig. 1. Leukocyte recruitment in cremasteric postcapillary venules induced by local application of preformed immune complexes (ICs). ICs [ovalbumin (OVA) and anti-OVA] generated in vitro were injected intrascrotally adjacent to the cremaster muscle, and the muscle was examined via intravital microscopy 4 h later. Responses were compared with those of mice treated with OVA and nonspecific IgG. A: leukocyte adhesion. B: leukocyte emigration. Values are means ± SE of 6 mice per group. *P < 0.05 vs. OVA/rabbit IgG.

 
We next examined responses in the pial microvasculature. In untreated mice, clearance of FITC-70-kDa dextran from the pial circulation was minimal (Fig. 2A), illustrating that BBB integrity was maintained in the closed cranial window preparation, consistent with previous studies (23, 33). Similarly, leukocyte rolling and adhesion were rarely observed in pial postcapillary venules (data not shown), in accordance with previous observations (10). We next examined the responses to preformed ICs administered directly to the pial microvasculature. Although superfusion with OVA-rabbit IgG did not alter permeability, FITC-70-kDa dextran clearance was significantly elevated within 15 min after superfusion with preformed ICs (P < 0.05; Fig. 2B). The response was short-lived, returning to basal levels within 30 min (Fig. 2B). The increase in microvascular permeability in response to preformed ICs was accompanied by an increase in leukocyte adhesion in pial postcapillary venules (Fig. 2C). However, the level of adhesion in OVA-rabbit IgG-treated mice did not differ from that in preformed IC-treated mice (Fig. 2C), indicating that this adhesion was not attributable to IC deposition. Superfusion of preformed ICs did not affect circulating leukocyte counts (see Table 2). These observations indicate that the cerebral microvasculature is capable of rapidly responding to ICs in extravascular tissue.


Figure 2
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  		Fig. 2. Responses of pial microvasculature to superfusion with artificial cerebrospinal fluid (aCSF), preformed ICs (OVA and anti-OVA), or OVA and nonspecific IgG (control) over the surface of the brain. A: clearance of FITC-70-kDa dextran in untreated mice. B: alterations in pial microvascular permeability after superfusion of preformed ICs for 10 min. C: leukocyte adhesion in pial postcapillary venules. Values are means ± SE for untreated mice (n = 7), preformed IC-treated mice (n = 6), and control mice (OVA/rabbit IgG, n = 6). *P < 0.05 vs. OVA/rabbit IgG.

 

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  		Table 2. Circulating leukocyte counts

 
Effect of the RPA reaction on cerebral microvasculature. Extravascular application of ICs allows potential involvement of nonvascular cells in the IC-mediated response. However, in IC-mediated vasculitis, ICs are commonly found in and around blood vessels (5, 29). Therefore, we next examined the response of the cerebral microvasculature to a model of perivascular IC formation, the RPA reaction. In response to 5 and 50 µg/ml anti-OVA, microvascular permeability was elevated above levels in control-treated mice within minutes of initiation (Figs. 3A and 4A). In RPA-treated mice treated with 5 µg/ml anti-OVA, FITC-70-kDa dextran clearance was significantly elevated within 20 min relative to that in rabbit IgG control-treated mice (P < 0.05; Fig. 3A). This immediate increase in permeability in RPA-treated mice occurred in the absence of a simultaneous change in leukocyte rolling flux, rolling velocity, and adhesion in pial postcapillary venules, although a small increase in adhesion occurred by 60 min (P < 0.05; Fig. 3B, Table 1). Induction of the RPA response with 50 µg/ml anti-OVA elevated the clearance of FITC-70-kDa dextran within 15 min and caused a larger increase in vascular permeability than that observed at the lower concentration (Fig. 4A). In addition, leukocyte rolling flux in RPA-treated animals was significantly increased relative to rabbit IgG-treated animals at 60 min: 1.6 ± 0.7 vs. 0.14 ± 0.1 cells/min (Table 1). In these animals, leukocyte adhesion in pial postcapillary venules was also significantly elevated within 30 min and remained significantly elevated above rabbit IgG control at 60 min: 109.5 ± 34.1 and 17.8 ± 9.3 cells/mm2, respectively (P < 0.05; Fig. 4, B–D). These alterations occurred in the absence of changes in microvascular blood flow, as assessed by measurement of microsphere velocity in pial postcapillary venules (Fig. 4E). Circulating leukocyte counts were not affected by these treatments (Table 2). Inasmuch as higher levels of permeability and leukocyte adhesion were observed after treatment with 50 µg/ml anti-OVA, all subsequent experiments were performed at this dose.


Figure 3
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  		Fig. 3. Response of pial microvasculature to intravenous administration of OVA and superfusion of 5 µg/ml anti-OVA [reverse passive Arthus (RPA)] or nonspecific IgG (control). A: alterations in pial microvascular permeability shown as clearance of FITC-70-kDa dextran. B: leukocyte adhesion in pial postcapillary venules. Values are means ± SE for 6 mice in each group. *P < 0.05 vs. OVA/rabbit IgG.

 

Figure 4
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  		Fig. 4. RPA response of pial microvasculature to intravenous administration of OVA and superfusion of 50 µg/ml anti-OVA (RPA) or the same amount of nonspecific IgG (control). A: alterations in pial microvascular permeability shown as clearance of FITC-70-kDa dextran. B: leukocyte adhesion in pial postcapillary venules. C and D: fluorescent intravital microscopy images of pial microvasculature illustrating leukocyte adhesion during the RPA response: untreated pial postcapillary venule with no leukocytes interacting with the endothelial surface (C) and pial postcapillary venule during the RPA response, showing several leukocytes interacting with the endothelial surface (D). E: analysis of microsphere velocity [as an index of red blood cell velocity (VRBC)] in pial postcapillary venules of RPA-treated (n = 3) and control mice (n = 3). Values are means ± SE for 6–7 mice in each group. *P < 0.05 vs. OVA/rabbit IgG.

 

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  		Table 1. Leukocyte rolling flux and rolling velocity

 
Location of IC formation during the cerebral RPA response. In the RPA model, inflammation is initiated when antigen administered intravenously binds to antibody administered locally to the tissue of interest, resulting in IC formation. The mechanism for this response in the cerebral RPA model was unclear. Therefore, we used confocal and fluorescent microscopy to study the localization of OVA and anti-OVA during the RPA response (Figs. 5 and 6). First, confocal microscopy was used to determine the location of anti-OVA-488 relative to the position of the pial microvasculature identified using the endothelium-specific lectin IB4. Within 10 min of the RPA response, anti-OVA localized immediately adjacent to the pial endothelium (Fig. 5A). These findings were supported by intravital microscopy experiments, which showed preferential localization of anti-OVA-594 in perivascular sites within minutes during the RPA response. In contrast, in the absence of intravenous OVA, anti-OVA was distributed more diffusely (Fig. 5, B and C). Image analysis of perivascular fluorescence 60 min after initiation of the RPA response demonstrated that accumulation of anti-OVA-594 immediately adjacent to cerebral microvessels was significantly higher in mice undergoing the RPA reaction than in mice treated with anti-OVA-594 alone (Fig. 5D). We also examined the location and distribution of OVA during the RPA response to determine whether the IC-mediated response was associated with the exit of OVA from the circulation. Within 10 min of initiation of the RPA reaction, OVA-488 was detectable in extravascular tissue, adjacent to lectin IB4-labeled pial vessels (Fig. 6, A–C).


Figure 5
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  		Fig. 5. Localization of anti-OVA in pial vasculature during the RPA response. A: confocal microscopy image showing location of Alexa 488-conjugated anti-OVA (green, arrowhead) adjacent to pial vessels (red), which were identified using Alexa 594-conjugated isolectin IB4, 10 min after initiation of the RPA response (OVA nonfluorescent). Original magnification x100. B and C: intravital microscopy images of pial vessels showing diffuse distribution of Alexa 594-conjugated anti-OVA when superfused over the brain in the absence of intravenous OVA at 15 min (B) and distribution of Alexa 594-conjugated anti-OVA in mice treated with intravenous OVA (RPA response) (C). Preferential localization of anti-OVA is detectable in perivascular regions (arrowheads) of RPA-treated mice from as early as 15 min. D: image analysis of perivascular fluorescence intensity in indicated region of pial vessels of mice treated with anti-OVA-594 alone (n = 3) or anti-OVA-594 + OVA (n = 4) 60 min after initiation of the response. *P < 0.05 vs. anti-OVA-594 alone.

 

Figure 6
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  		Fig. 6. Distribution of Alexa 488-conjugated OVA, colocalization of Alexa 488-conjugated OVA and Alexa 594-conjugated anti-OVA, and complement deposition during the RPA response in pial microvasculature. A–C: confocal microscopy showing location of intravenously administered Alexa 488-conjugated OVA 10 min after initiation of the RPA response. A: distribution of Alexa 488-conjugated OVA (green). B: distribution of Alexa 594-conjugated isolectin IB4 (red) showing location of vascular endothelial cells. C: merged images demonstrating location of OVA outside pial vessels (arrowhead). Original magnification x60. D–F: confocal microscopy of pial microvasculature 60 min after initiation of the RPA response. D: distribution of Alexa 488-conjugated OVA (green). E: distribution of Alexa 594-conjugated anti-OVA (red). F: merged images. Yellow staining indicates colocalization of OVA and anti-OVA. Original magnification x60. G–I: C3 deposition in pial microvasculature: untreated brain (G), 60 min after RPA (H), and 60 min after RPA in cobra venom factor (CVF)-pretreated mouse.

 
Additional experiments were performed using confocal microscopy to confirm colocalization of OVA-488 and anti-OVA-594 and, therefore, IC formation in this model (Fig. 6, D–F). Colocalization of OVA and anti-OVA was detectable in perivascular regions near the brain surface (Fig. 6F). To provide further evidence of IC formation, we examined deposition of C3, which has been shown repeatedly to be generated in response to IC formation (40, 49). In RPA-treated brains, C3 staining was markedly increased, predominantly localized to perivascular regions, in a pattern similar to the distribution of anti-OVA during the RPA response (Fig. 6, G–I). Taken together, these observations suggest that, during the cerebral RPA response, the location of anti-OVA immediately adjacent to the pial endothelium allows for rapid colocalization of OVA and anti-OVA adjacent to pial microvessels.

Cerebral RPA response requires C3. It has previously been demonstrated that complement activation contributes to the development of IC-mediated inflammation in other tissues (40, 49). Therefore, in the next series of experiments, we examined the role of C3 in alteration of permeability and leukocyte-endothelial cell interactions after IC formation. It has been established that CVF depletes C3 (16, 26, 31). In the present study, this was confirmed by the absence of C3 staining in the brains of CVF-treated mice after the RPA response (Fig. 6I). To confirm that CVF did not affect the ability of the pial microvasculature to respond to vasoactive agents, we assessed the response to bradykinin (38). The increase in permeability induced by bradykinin in CVF-treated mice was indistinguishable from that in animals treated with bradykinin alone (Fig. 7A). In RPA-treated mice, C3 depletion suppressed the increase in permeability, returning clearance to levels in OVA-rabbit IgG-treated control mice. CVF also nearly eliminated RPA-induced leukocyte adhesion (Fig. 7, B and C).


Figure 7
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  		Fig. 7. Effect of CVF-induced complement depletion on cerebral microvasculature after bradykinin superfusion and RPA response. A: bradykinin (100 µM, 5 min)-induced alterations in pial microvascular permeability in untreated and CVF-pretreated mice. B: leukocyte adhesion in pial postcapillary venules 60 min after initiation of the RPA response. C: pial microvascular permeability, shown as clearance of FITC-70-kDa dextran. Values are means ± SE for bradykinin (n = 7), bradykinin + CVF (n = 4), RPA (n = 6–7), and RPA + CVF (n = 5). *P < 0.05 vs. RPA.

 
Inhibition of leukocyte adhesion reduces the RPA response in cerebral microvasculature. In previous studies in peripheral tissues, anti-CD18 and anti-ICAM-1 blockade was used to demonstrate a role for leukocytes in mediating IC-induced microvascular injury (35, 37, 52). Therefore, we next assessed the role of leukocyte adhesion in the increased microvascular permeability during the cerebral RPA response. Treatment with anti-CD18 significantly reduced leukocyte adhesion 60 min after induction of the RPA response relative to levels in RPA-treated animals: 15.04 ± 7.8 vs. 86.9 ± 30.3 cells/mm2 (P < 0.05; Fig. 8A). Treatment with this antibody did not affect circulating leukocyte counts relative to those in normal RPA-treated mice (Table 2). In the first 20 min of the RPA response, CD18 blockade had no effect on pial vessel clearance of FITC-70-kDa dextran (Fig. 8B). However, after 55 min, FITC-70-kDa dextran clearance was significantly attenuated relative to that in RPA-treated animals. In contrast, treatment with anti-ICAM-1 did not reduce RPA-induced microvascular permeability or leukocyte adhesion (Fig. 8, A and B).


Figure 8
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  		Fig. 8. Effect of CD18 and intercellular adhesion molecule-1 (ICAM-1) blockade on cerebral RPA response. A: leukocyte adhesion in pial postcapillary venules 60 min after initiation of the RPA response. B: pial microvascular permeability, shown as clearance of FITC-70-kDa dextran. Values are means ± SE for RPA (n = 6–7), RPA + anti-CD18 (n = 6) and RPA + anti-ICAM-1 (n = 4). *P < 0.05 vs. RPA.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Several unique characteristics of the cerebral microvasculature affect the nature of inflammatory responses in the CNS. The BBB restricts macromolecular leakage into the CNS (17, 32). Leukocyte-endothelial interactions and endothelial adhesion molecule expression are also markedly reduced in the CNS relative to peripheral tissues (10). Finally, intracerebral injection of mediators that induce recruitment in other tissues fails to induce recruitment of leukocytes to the CNS parenchyma (3, 4). Taken together, these findings demonstrate that the response of the brain to many inflammatory mediators is markedly suppressed relative to that of most other tissues. Therefore, despite the recognized ability of ICs to generate responses in other tissues, it was unclear whether ICs would be able to generate a response in the brain microvasculature. Here we show that preformed ICs applied extravascularly and ICs formed adjacent to blood vessels (RPA model) rapidly increase microvascular permeability in cerebral microvessels. Furthermore, in the model of perivascular IC formation, larger amounts of IC formation induced substantial leukocyte adhesion in cerebral postcapillary venules, contributing to the later stages of microvascular dysfunction. These studies show that ICs are capable of inducing leukocyte adhesion and microvascular permeability in the cerebral microvasculature.

In initial experiments to test the responsiveness of the brain to ICs, preformed ICs were applied to the surface of the brain, external to the vasculature. This approach is relevant to IC-mediated diseases, such as SLE, in which ICs are commonly detected outside the vasculature, particularly in choroid plexus and skin (29). In addition, previous studies have shown that preformed ICs are capable of producing inflammatory responses comparable to those induced by the classical Arthus model of IC-mediated inflammation (11, 49). The present studies show that ICs applied to extravascular tissue in the brain are capable of increasing microvascular permeability. However, in contrast to the significant increases in leukocyte adhesion and emigration observed when preformed ICs were applied to the extravascular tissue of the cremaster muscle, the response was relatively mild and short-lived and did not involve a marked increase in leukocyte recruitment. This indicated that the cerebral response to extravascular deposition of ICs was reduced relative to the response in the periphery.

However, in diseases such as IC-mediated vasculitis, ICs are commonly deposited perivascularly (5, 29). Therefore, as an alternative approach to examine the effects of ICs in the cerebral microvasculature, the conventional RPA model of perivascular IC formation was examined. This model induced an increase in cerebral microvascular permeability and an increase in leukocyte adhesion within cerebral microvessels, demonstrating that perivascular IC formation in the brain is capable of altering the cerebral microvasculature. Furthermore, it suggests that, in the brain, the location of IC formation is an important determinant of the level of inflammation and microvascular dysfunction.

To examine the mechanism of IC formation in the cerebral RPA model, we used confocal microscopy and fluorescent intravital microscopy to analyze the location of OVA and anti-OVA. Anti-OVA was detected immediately adjacent to microvascular endothelial cells (identified by staining with lectin IB4) shortly after it was applied to the surface of the brain. Presumably, location of anti-OVA at this site allowed for binding with OVA within the vasculature or in the immediate perivascular area. Furthermore, within 10 min of initiation of the RPA response, substantial amounts of OVA had escaped from pial vessels, allowing further IC formation, as demonstrated by confocal microscopy colocalization studies. Deposition of C3 in similar locations further supported the contention that substantial IC formation occurred in this cerebral RPA model.

Previous experiments showed that complement activation and Fc receptors play important roles in the initiation of IC-mediated inflammation (40, 49, 50). The molecular components of each of these pathways are expressed in the CNS, where they have the potential to contribute to inflammatory responses (46–48, 54). Complement receptors on CNS-resident cells have been demonstrated to contribute to the development of immune-mediated cerebral inflammation (9). Furthermore, activation of the complement pathway contributes to brain injury in a range of inflammatory disorders (13, 34, 46). Collectively, these observations indicate that the molecular requirements for IC-mediated responses are present and potentially functional in the brain. The present observations further support this contention. C3 depletion achieved by pretreatment with CVF resulted in a significant attenuation of permeability and leukocyte adhesion in pial postcapillary venules during the RPA response. These findings indicate that the complement pathway plays a key role in the cerebral response to IC formation, similar to its role in many other tissues (8, 27, 40, 49).

In RPA-treated animals, the nature of the microvascular response differed according to the concentration of anti-OVA used. At the lower concentration (5 µg/ml), microvascular permeability increased, but leukocyte adhesion remained minimal, indicating that, at this lower concentration, permeability increased independently of recruited leukocytes. In contrast, the higher concentration of anti-OVA (50 µg/ml) induced an increase in leukocyte adhesion on the cerebral endothelium as well as a larger and more rapid increase in permeability. At this higher concentration, prevention of leukocyte adhesion also reduced permeability, but only after the first 30 min, a time point at which a significant level of leukocyte adhesion was observed. Taken together, these data indicate that, in response to larger amounts of IC formation, one of the mechanisms of complement-mediated vascular injury in cerebral microvessels involves induction of leukocyte adhesion, which amplifies microvascular dysfunction. This is similar to the observed role of leukocytes in mediating IC-mediated microvascular injury in other tissues (28, 35, 37, 51). It is noteworthy that leukocyte adhesion was inhibited by antibodies to beta2-integrins, but not antibodies to ICAM-1. This indicates that leukocyte adhesion in this model requires a beta2-integrin ligand other than ICAM-1.

In this study, we used macromolecular clearance of the pial microvasculature as an index of BBB function. It is conceivable that the pial microvasculature has barrier properties different from those of CNS parenchymal vessels. Furthermore, the additional barrier of the arachnoid epithelium may also affect macromolecular leakage from pial vessels (32). However, studies have shown that pial and CNS microvessels share many features, such as tight junctions and high transendothelial resistance, and the barrier properties of these vessels have also been observed to be similar (2). Nevertheless, in this study of IC-mediated inflammation, we cannot exclude the possibility that the observed response to ICs is a unique feature of the pial microvasculature.

Previous studies have repeatedly demonstrated that the level of constitutive leukocyte rolling in uninflamed cerebral postcapillary venules is extremely low (10, 21, 22). This is explained in part by studies in which expression of the endothelial adhesion molecules most likely to be mediating rolling, P- and E-selectin, was negligible in the uninflamed brain (10, 24). Nevertheless, the brain has been observed to be capable of supporting rapid (within minutes) increases in leukocyte adhesion in response to specific inflammatory stimuli, e.g., nicotine (57). In this study, leukocyte rolling after initiation of perivascular IC formation in pial postcapillary venules was very low (<2 cells/min). This level of rolling is substantially lower than that observed in pial venules in other models of cerebral inflammation (20, 24, 25). However, despite this extremely low level of rolling, adhesion increased significantly after IC formation. Furthermore, direct arrest of leukocytes on the endothelial surface was not observed without prior rolling. This suggests that, during the RPA response, the cerebral endothelium is highly efficient in promoting the transition from rolling to adhesion.

In conclusion, this study demonstrates that the unique microvasculature of the brain is sensitive to IC formation, responding via alterations in microvascular permeability and leukocyte adhesion. These observations raise the possibility that, in diseases in which ICs are present in the CNS, one of the potential actions of ICs is the promotion of microvascular dysfunction and leukocyte recruitment.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
These studies were supported by the National Health and Medical Research Council (NHMRC), Australia, Program Grant 334067. M. J. Hickey is an NHMRC R. D. Wright Fellow.


   ACKNOWLEDGMENTS
 
The authors thank Dr. Ian Harper and Stephen Firth (Monash Micro Imaging, Monash University) for technical assistance with confocal microscopy.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. J. Hickey, Centre for Inflammatory Diseases, Monash Univ. Dept. of Medicine, Monash Medical Centre, 246 Clayton Rd., Clayton, Victoria 3168, Australia (e-mail: michael.hickey{at}med.monash.edu.au)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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