Novel glucocorticoid effects on acute ... - Wiley Online Library

Journal of Neurochemistry, 2003, 84, 705–716

doi:10.1046/j.0022-3042.2003.01604.x

Novel glucocorticoid effects on acute inflammation in the CNS Klaus Dinkel, Anna MacPherson and Robert M. Sapolsky Department of Biological Sciences, Stanford University, Stanford, California, USA

Abstract The CNS can mount an inflammatory reaction to excitotoxic insults that contributes to the emerging brain damage. Therefore, anti-inflammatory drugs should be beneficial in neurological insults. In contrast, glucocorticoids (GCs), while known for their anti-inflammatory effects, can exacerbate neurotoxicity in the hippocampus after excitotoxic insults. We investigated the effect of GCs on the inflammatory response after a neurological insult. Intact control (INT; intact stress response GC profile), adrenalectomized/GC-supplemented (ADX; low basal GC profile) and GC-treated (COR; chronically high GC profile) rats were injected with kainic acid into the hippocampal CA3 region. Lesion size was determined 8–72 h later. The inflammatory response was characterized using

immunohistochemistry, RNAse protection assay and ELISA. The INT and COR rats developed larger CA3 lesions than ADX rats. We found that GCs surprisingly caused an increase in relative numbers of inflammatory cells (granulocytes, monocytes/macrophages and microglia). Additionally, mRNA and protein (IL-1b and TNF-a) levels of the pro-inflammatory cytokines IL-1a, IL-1b and TNF-a were elevated in COR rats compared with INT and ADX rats. These data strongly question the traditional view of GCs being uniformly anti-inflammatory and could further explain how GCs worsen the outcome of neurological insults. Keywords: brain injury, cytokines, neuroimmunology, steroids, stress. J. Neurochem. (2003) 84, 705–716.

Brain damage as a consequence of a neurological insult, such as stroke, trauma or seizure, develops from a complex series of pathophysiological events over time. Excessive activation of neurons by excitatory neurotransmitters (e.g. glutamate), which are massively released as a consequence of energy depletion, results in excitotoxic neuron death (Beal 1992; Lipton and Rosenberg 1994; Whetsell 1996). Accumulating evidence during the last decade has shown that brain injury is also accompanied by a marked inflammatory reaction characterized by infiltration of granulocytes and monocytes/ macrophages into the respective brain parenchyma, activation of resident brain cells (e.g. microglia and astrocytes) and expression of pro-inflammatory cytokines, adhesion molecules and other inflammatory mediators (Perry and Gordon 1991; Feuerstein et al. 1998b; Dirnagl et al. 1999). There is considerable evidence that the inflammation contributes significantly to the developing neuronal damage by mechanisms like the release of neurotoxic substances, such as free radicals or cytokines (Barone and Feuerstein 1999; Rothwell et al. 1996; McGeer and McGeer 1999). Accordingly, antiinflammatory drugs should have a beneficial effect in the context of a neurological insult. Glucocorticoids (GCs) are well known for their antiinflammatory and immunosuppressive properties (Marx 1995; Wilckens and De Rijk 1997) and are, therefore,

widely and successfully used in the treatment of autoimmune diseases, chronic inflammation or transplant rejection. Glucocorticoids have been shown to suppress production of pro-inflammatory cytokines (Kern et al. 1988; Lew et al. 1988) and to decrease the cellular inflammatory infiltrate by down-regulation of adhesion molecules (Goulding et al. 1998). They have also been used to treat inflammatory diseases within the CNS, such as edema arising from brain tumors (Barnes and Adcock 1993), bacterial/viral encephalitis (Salaki et al. 1984; Coyle 1999) or to improve the rate of recovery from acute exacerbation in multiple sclerosis patients (Filippini et al. 2000). In the context of various neurological insults, however, it was shown that GCs exacerbated neuron loss (Sapolsky 1985; Sapolsky and

Received September 26, 2002; revised manuscript received October 29, 2002; accepted October 30, 2002. Address correspondence and reprint requests to Dr Klaus Dinkel, Department of Biological Sciences, Gilbert Hall, Stanford University, Stanford, CA 94305-5020, USA. E-mail: [email protected] Abbreviations used: ADX, adrenalectomized; BBB, blood–brain barrier; BSA, bovine serum albumin; COR, corticosteroid treated; CORT, corticosterone; FAST-DAB, FASTTM-diaminobenzidine; GC, glucocorticoid; INT, intact controls; KA, kainic acid; met, metyrapone; mRPA, multiple template RNase protection assay; PBS, phosphate-buffered saline; rt, room temperature.

2003 International Society for Neurochemistry, J. Neurochem. (2003) 84, 705–716

705

706 K. Dinkel et al.

Pulsinelli 1985; Stein-Behrens et al. 1992). The GCs seemed to interfere with neuronal energy metabolism, thereby reducing the survival capability of neurons during brain injury. Hippocampal neurons (McEwen et al. 1986; Sapolsky 1994) have been shown to be especially sensitive to this GC effect. Brain injury is associated with an enhanced release of GCs as a consequence of the activation of the hypothalamicpituitary-adrenal axis by stress (Munck et al. 1984) and inflammatory mediators like IL-1a (Besedovsky et al. 1986; Berkenbosch et al. 1987). Glucocorticoids should ameliorate the damage during brain injury by suppressing the inflammatory reaction but instead have been found to increase neuron loss. Therefore, we investigated in this study the effect of GCs on: (i) lesion size, (ii) cellular inflammatory infiltrate and (iii) cytokine pattern after excitotoxic insult in vivo over the course of time. The glutamate agonist kainic acid (KA) was injected into the CA3 area of the hippocampus of untreated control (INT; intact stress response; GC levels in the normal range of physiological stress response), adrenalectomized/GC-supplemented (ADX; no stress response; low basal GC levels) and corticosterone-treated (COR; chronically high GC levels in the upper stress level) rats.

Materials and methods Animals and materials Male Sprague–Dawley rats (weighing 250–300 g; Simonsen Laboratories, Gilroy, CA, USA) were housed under a 12-h light– dark cycle with free access to food and water. Corticosterone (CORT), KA, metyrapone (met; 2-methyl-1,2-di-3-pyridyl-1-propanone), paraformaldehyde, bovine serum albumin (BSA) and FASTTMdiaminobenzidine (FAST-DAB) were purchased from Sigma (St Louis, MO, USA). All antibodies were purchased from BD PharMingen (San Diego, CA, USA). Glucocorticoid manipulations All experiments were conducted under the guidelines described in the US Public Health Service Policy on Human Care and Use of Laboratory Animals. Four groups of rats (INT, ADX, COR and INT + met) were used in the experiments, each group displaying a different GC level. In the control group (INT) no GC manipulations were performed; thus animals had basal GC levels (approximately 1–10 lg/dL) throughout except for the physiological stress response to surgery and the subsequent KA-induced seizures; this involved an increase of approximately 30 lg/dL above baseline by 30 min after seizure onset and levels 10–15 lg/dL above baseline for the next 6 h before returning to baseline (Stein and Sapolsky 1988). In another group, rats were bilaterally adrenalectomized (ADX) under halothane anesthesia and a 15% CORT pellet was implanted s.c. to generate basal GC levels of approximately 6 lg/dL (Stein-Behrens et al. 1994). Rats were given 3 days for hormone levels to stabilize prior to stereotaxic surgeries. In a subset of experiments, intact rats were treated 1 h before KA injection with the steroid synthesis inhibitor metyrapone (200 mg/kg in 0.9% NaCl, s.c.). In these rats

(INT + met), GC levels were locked to basal levels of approximately 6–7 lg/dL (Stein and Sapolsky 1988). In the fourth group (COR) of rats, chronically high levels of CORT of approximately 28 lg/dL were generated for approximately 20 h/day by injecting rats daily for 3 days with 3 mg CORT/day s.c. in sesame oil (SteinBehrens et al. 1994). These injections continued after microinfusion until the indicated timepoint. As GCs are steroid hormones, they readily penetrate the blood–brain barrier (BBB) and neuronal membranes, then interacting with their intracellular receptors. Stereotaxic microinfusion Rats were anesthetized with an i.p. injection (1 mL/kg bodyweight) of Ôrodent cocktailÕ (76 mg/mL ketamine, 1.5 mg/mL promace and 7.7 mg/mL xylazine) and placed in a stereotaxic frame. The rats were microinfused stereotaxically into the CA3 area of the hippocampus (A/P ) 3.85, L/M ± 3.35 from bregma and D/V ) 2.55 from dura). The CA3 area was defined as the pyramidal cell layer beginning in between the blades of the dentate gyrus, continuing around Ammon’s horn until the thickening of the cell field that characterizes CA2. Animals were injected either with 0.06 lg KA dissolved in phosphate-buffered saline (PBS) or with PBS alone as a control. Quantification of lesion size At indicated times after microinfusion, rats were anesthetized by halothane inhalation and then, under deep anesthesia, perfused intracardially with a 0.1% heparin/0.9% saline solution followed by 3% paraformaldehyde solution. The brains were post-fixed in 3% paraformaldehyde (PFA) for 24 h and cryoprotected with PBS 15% sucrose. On a cryostat microtome, 30-lm coronal sections were cut, dried and stained with cresyl violet using the Nissl method. Lesion size in the CA3 area of the hippocampus was measured using a 10 · 10 optical grid in the ocular of the microscope at 40· and counting the number of pixels with damaged/missing cells. As an internal control, the non-lesioned hemisphere of each section was used to measure the size of the intact CA3 area, which was set to 100%. Counts were converted into area measurements and lesion size was expressed as percentage damage of the intact contralateral CA3 area. Starting from the visible needle track, counts were taken from at least four coronal sections at 0.1 mm increment. All sections were scored blind using a light microscope. This technique produces assessments of lesion size which correlate significantly with more arduous cell counting (Sapolsky and Stein 1989). Immune cell immunohistochemistry At specific times after microinfusion, animals (n ¼ 6 rats/group and time) were killed by inhalation of excess halothane and then decapitated. The brains were removed immediately and quick frozen in 2-methylbutane at )42C for 3 min. Cryostat sections (15 lm) were cut, mounted on slides, dried and kept at )70C until use. Slides were fixed in icecold acetone at )20C for 3 min, treated with 0.03% H2O2 solution for 10 min at room temperature (rt) to block endogenous peroxidase activity and then blocked with 5% normal goat serum or 3% BSA solution for 15 min at rt. Sections were incubated with the respective primary antibody (see Table 1) diluted in PBS 3% BSA (anti-CD11b/c, 3 lg/mL; anti-CD3, 2 lg/mL; anti-CD45RA, 5 lg/mL; anti-granulocyte, 3 lg/mL; anti-ED1-like, 1 lg/mL and anti-NKR-P1A, 3 lg/mL) for 2 h at rt in a humid chamber. After rinsing the slides in three changes of PBS they were


Glucocorticoids and CNS inflammation

Table 1 Antibodies used for immunohistochemical staining of different types of immune cells Antibody

Cell type

Isotype

OX-42 (anti-CD11b/c) G4.18 (anti-CD3) OX-33 (anti-CD45RA) HIS48 1C7 (anti-ED-1 like) 10/78 (anti-NKR-P1A)

Microglia T-cells B-cells Granulocytes Monocytes/macrophages NK-cells

ms ms ms ms ms ms

IgG2a IgG3 IgG1 IgM IgG1 IgG1

incubated with the respective biotinylated secondary antibody for 40 min at 37C in a humid chamber. The slides were rinsed again and then treated with a horseradish peroxidase-streptavidin solution (1 : 400 in PBS 3% BSA) for 45 min at rt. Peroxidase labeling was visualized by incubation with FAST-DAB solution as a substrate for 2–4 min. The details for the primary antibodies employed in this study are given in Table 1. The respective positively stained immune cells in the CA3 area were counted in at least three coronal sections (starting at the needle track) with an increment of 0.1 mm using 40· and 100· magnification. All sections were scored blind and manually to exclude infiltrating cells in the area of the needle track. IL-1b immunohistochemistry Frozen coronal hippocampal sections (15 lm) were fixed in icecold acetone : methanol (3 : 1) at )20C for 10 min, treated with 0.03% H2O2 solution for 10 min at rt to block endogenous peroxidase activity, rinsed in 50 mM Tris 0.4% Triton X-100 for 5 min at rt and then blocked with 10% normal goat serum for 15 min at rt. Sections were then incubated for 48 h at 4C with polyclonal anti-rat IL-1a antibody (Pierce-Endogen, Rockford, IL, USA) diluted 1 : 100 in PBS (0.3% Triton-X100, 5% BSA). After three 5-min washes in PBS (0.1% Triton-X100), sections were incubated with the biotinylated anti-rabbit Ig secondary antibody for 40 min at 37C in a humid chamber. The slides were washed again and immunoreactivity was tested by the avidin-biotin-peroxidase technique (Vectastain ABC kit; Vector Laboratories, Burlingame, CA, USA). Peroxidase labeling was visualized by incubation with FAST-DAB solution as a substrate for 2–4 min. RNase protection assay At indicated times (4, 8, 12 and 24 h after microinfusion), the hippocampus was dissected and quick frozen. Tissue was homogenized/disrupted with a 1.5-mL Pellet Pestle grinder (KimbleKontes, Vineland, NJ, USA) and total RNA was extracted using a RNA isolation kit (BD PharMingen). RNA concentrations were determined by spectrophotometry. Detection and semiquantitation of a variety of rat cytokine mRNAs were performed on 10 lg of total RNA utilizing the Riboquant Multiprobe Rnase Protection Assay System and the rCK-1 template set (BD PharMingen) following the manufacturer’s instructions. Briefly, [a-32P] UTP-labeled antisense RNA probes were synthesized by in vitro transcription of cDNA templates for 11 rat cytokines (IL-1a, IL-1b, TNF-b, IL-3, IL-4, IL-5, IL-6, IL-10, TNF-a, IL-2 and IFN-c) and two housekeeping genes (L32 and GAPDH) as internal controls. DNA templates were degraded by DNaseI digestion and probes were purified by phenol : chloroform extraction and

707

ethanol precipitation with subsequent hybridization to total RNA at 56C overnight. Samples were treated with RNase A+T1 and protected double-stranded RNA was purified by phenol : chloroform extraction and ethanol : salt precipitation. Samples were resuspended in Quickpoint loading dye and resolved on a denaturing 6% polyacrylamide 7M urea gel using the Quickpoint Rapid Nucleic Acid Separation System (Invitrogen, Carlsbad, CA, USA). After overnight exposure at ) 80C to Biomax X-ray films (Kodak, Rochester, NY, USA), bands were quantified by densitometric analysis using the Digital Science 1D software (Kodak). The mRNA levels are expressed as the ratio of the respective cytokine band and the corresponding L32 band in densitometric units. ELISA for IL-1b and TNF-a IL-1b and TNF-a protein levels were determined by commercially available rat-specific ELISA (Pierce-Endogen) according to the manufacturer’s instructions. Animals were killed 8 h after KA or PBS injection, the respective hippocampi were dissected free on ice and immediately snap frozen in liquid nitrogen. Brain samples were placed in sterile icecold PBS containing a protease inhibitor cocktail [0.2 mM aminoethyl-benzenesulfonyl-fluoride (AEBSF), 1 lg/mL aprotinin, 1 mM benzamidine, 1 mM EDTA, 10 lM leupeptin and 10 lg/mL pepstatin] following homogenization using a 1.5-mL handheld homogenizer (Kimble-Kontes). Samples were then centrifuged (12 000 r.p.m., 20 min, 4C), the supernatant fluid removed and divided into 50-lL aliquots that were stored at ) 70C until used. Samples were run in duplicate in the ELISA; protein concentrations of all samples were measured by the bicinchoninic acid (BCA) method (Pierce-Endogen) and cytokine levels expressed as pg/mg protein. Statistical analysis Data are given as mean values ± SD. Analysis of results was performed using linear regression analysis or ANOVA followed by all pairwise multiple comparison post-hoc procedure (Tukey test or SNK test, respectively); p-values < 0.05 were considered statistically significant. For correlation analysis (linear regression), the data (lesion and infiltrate, respectively) of the different treatment groups (ADX, INT and COR) were combined for the comparison of the respective timepoints.

Results

Effect of glucocorticoids on kainic acid-induced hippocampal damage We first investigated whether different GC levels had a significant effect on the lesion size in the CA3 area 8–72 h after KA-induced excitotoxic insult (Fig. 1). Kainic acid was injected locally into the hippocampus. Damage to the CA3 region of the hippocampus was quantified after cresyl violet staining. As would be expected, KA caused significant lesion damage in control rats (INT). Adrenocortical status significantly modified the extent of damage ( p < 0.001, F ¼ 88.6, d.f. ¼ 2/49). In ADX animals, with their reduced GC exposure, the maximum lesion size had developed after 8 h and did not change significantly thereafter ( post-hoc



100 90 80

***

ADX INT

***p 0.05. A significant correlation (*p < 0.05) was found between the extent of damage and the preceding inflammatory infiltration (previous timepoint).

post-hoc test). There was also an increase in relative microglia numbers between the ADX and INT group but it was not statistically significant. Like the neutrophil and macrophage data, these results showed that GCs surprisingly caused an increase instead of a decrease in inflammatory cells in the CA3 area after KA injection. Because of the unexpected nature of these findings, we tested whether the extent of damage at any given time point was significantly correlated with the extent of inflammatory infiltration at that time or at the next time point (i.e. if damage predicted the subsequent extent of inflammatory infiltration). The correlation analysis revealed that the pro-inflammatory effects of GCs were not a mere consequence of them causing more hippocampal damage (Table 2). On the contrary, the extent of damage at any given time correlated significantly with the inflammatory infiltration at the previous timepoint (Table 2), thus indicating that neuronal damage was caused by the preceding inflammation. The increase in neuronal damage and cellular inflammatory infiltrate after kainic acid injection are dependent on steroid synthesis The differences seen between the intact control (INT) and the COR group were definitely caused by GCs alone. However, since removing the adrenal glands removes many more circulating factors than corticosterone, it is possible that the differences seen between adrenalectomized and non-adrenalectomized animals might be due to factors other than GCs. Therefore, we compared intact animals (adrenal glands present and GC levels in the high stress range of 28–32 lg/dL) with intact animals which were treated with metyrapone 1 h before KA insult. Metyrapone is a potent inhibitor of steroid synthesis which locks GC secretion at basal levels of 6–7 lg/dL (INT + met). We found that 40 h after KA administration neuronal damage (Fig. 3a) as well as inflammatory cellular infiltration of neutrophils (Fig. 3b) and macrophages (Fig. 3c) in the hippocampus was significantly reduced in metyrapone-treated animals compared with intact animals (p < 0.01 by SNK post-hoc test). These results clearly showed that the exacerbation of neuronal damage and increase in cellular inflammatory infiltrate was caused by GCs and not by other factors as a consequence of the presence or absence of adrenal glands.

Effect of glucocorticoids on cytokine mRNA pattern In the periphery, GCs are known to suppress inflammation by inhibiting synthesis of pro-inflammatory cytokines. We examined the effect of GCs on the cytokine pattern over the course of time after KA injection using multiple template RNase protection assay (mRPA). Animals were killed at indicated times (4, 8, 12 and 24 h) after KA or PBS, respectively and total RNA was isolated from the hippocampus, pooled (n ¼ 4–6 rats/group and time point) and subjected to mRPA (Fig. 4a shows one of three mRPAs performed using different mRNA pools with similar results; Fig. 4b–e show the densitometric analysis of the cumulative data of the three RPAs performed). The template set included the pro-inflammatory cytokines IL-1a, IL-1b, TNF-b, IL-3, IL-4, IL-5, IL-6, IL-10, TNF-a, IL-2 and IFN-c. Elevated mRNA levels of IL-1a, IL-1b, IL-6 and TNF-a mRNA could be detected 4–12 h after KA injection in all three groups (Fig. 4a, left panel). Quantification of the respective cytokine bands was done by densitometry (Fig. 4b–e) and band intensity was given as the ratio of cytokine : L32 mRNA in densitometric units. Baseline cytokine mRNA synthesis after PBS injection was the same in all three groups at the respective times (Fig. 4a, right panel, representative 8 h PBS controls shown). Adrenocortical status significantly modified respective mRNA cytokine synthesis ( p < 0.001, F ¼ 260.3, d.f. ¼ 2/35 for IL-1a; p < 0.001, F ¼ 357.6, d.f. ¼ 2/35 for IL-1b; p < 0.001, F ¼ 20.8, d.f. ¼ 2/35 for IL-6 and p < 0.001, F ¼ 85.9, d.f. ¼ 2/35 for TNF-a) 4–8 h after KA. IL-1a (Fig. 4b) and IL-1b (Fig. 4c) mRNA levels in the ADX rats were significantly higher than in INT rats ( p < 0.001 for 4 and 8 h), indicating an anti-inflammatory effect of GCs. In agreement with the data regarding cellular inflammation, the COR group (Fig. 4b, 4–8 h) showed significantly higher IL-1a mRNA levels than in INT rats ( p < 0.001 for 4 and 8 h) as well as when compared with ADX rats ( p < 0.001 for 4 and 8 h). The same effects could also be observed for IL1-b (Fig. 4c; p < 0.001 for 4 and 8 h compared with ADX and p < 0.001 for 4 and 8 h compared with INT by post-hoc test) and TNF-a (Fig. 4e; p < 0.001 for 4 h compared with ADX and p < 0.001 for 4, 8 and 12 h compared with INT by posthoc test) mRNA levels. At the indicated time points, the INT


Glucocorticoids and CNS inflammation 711

(a)

70

**

60

% CA3 Damage

50 40 30 20 10 0 INT

(b)

INT+met

50 45

** **p < 0.01

40

Granulocytes

35 30 25 20 15 10 5 0 INT

(c)

INT+met

120

** 100

Macrophages/Monocytes

group always showed lower mRNA levels of the respective cytokine, whereas the COR group showed higher mRNA levels compared with the ADX group. It is noteworthy that the magnitude of the inhibitory effect (65–183% of control) caused by GC levels in the INT group was smaller compared with the enhancing effect (173–413% of control) caused by the GC levels in the COR group. There were no significant differences detected in IL-6 mRNA levels between the ADX and INT group at any time point (Fig. 4d, 4–24 h by posthoc test). The COR group, however, showed increased IL-6 mRNA compared with ADX and INT animals 4 h after KA (p < 0.001 for 4 h as compared with ADX and p < 0.001 for 4 h as compared with INT by post-hoc test). Thus, GCs did not suppress but enhanced IL-6 mRNA synthesis at these times. These results indicate that, depending on the dosage, GCs can have anti- or until now unknown pro-inflammatory effects on cytokine mRNA synthesis after KA injury in the hippocampus.

80 60 40 20 0 INT

INT+met

Fig. 3 Increases in hippocampal damage and cellular inflammatory infiltrate after kainic acid (KA) injection are dependent on steroid synthesis. Rats were left intact (INT) or treated with the steroid synthesis inhibitor metyrapone (met; 200 mg/kg, s.c.) 1 h before intracerebral KA injection. Lesion size was determined after cresyl violet staining of 30-lm coronal brain sections using a superimposed optical grid. (a) Hippocampal injury is expressed as the ratio between lesion size and CA3 size and given as percentage CA3 damaged at 40 h after KA injection. Another set of sections from the same brains was subjected to immunohistochemical staining for (b) neutrophil granulocytes or (c) macrophages. Immunoreactive cells in the CA3 area were counted using a 40· and a 100· magnification. n ¼ 5 rats/group. **p < 0.01; SNK post-hoc. Note that the overall numbers of granulocytes at 40 h after KA are much smaller compared with those shown in Fig. 2(a). The reason for this shift is unknown; however, the two sets of studies were carried out 3 months apart and with different batches of KA.

Effects of glucocorticoids on cytokine protein expression In order to determine if the observed increases/differences in mRNA levels were translated to protein expression, IL-1b (Fig. 5a) and TNF-a (Fig. 5b) protein levels were determined 8 h after KA or PBS injection using a rat-specific ELISA. Compared with the respective control (PBS injection), ADX, INT and COR animals showed an increase in IL-1b and TNF-a expression in response to kainate injection. In line with the mRNA data, KA-induced IL-1b protein levels were much higher (52–122 pg/mg) than TNF-a levels (12–18 pg/mg). Interestingly, the IL-1b levels in the PBSinjected control group were significantly higher ( p < 0.05) in COR animals compared with ADX and INT animals (Fig. 5a), thus pointing towards a pro-inflammatory effect of chronically high GCs even in the absence of an insult. Further confirming the mRNA data, adrenocortical status significantly modified respective cytokine synthesis ( p < 0.001, F ¼ 28.5, d.f. ¼ 5/21 for IL-1b and p < 0.05, F ¼ 11.6, d.f. ¼ 5/21 for TNF-a); 8 h after KA IL-1b mRNA levels in ADX rats were significantly higher than in INT rats ( p < 0.05). The COR group showed significantly higher IL-1b expression than INT rats ( p < 0.001), as well as when compared with ADX rats ( p < 0.001). TNF-a levels were not significantly different between ADX and INT animals as well as between ADX and COR animals. However, there was a statistically significant difference in TNF-a levels between COR and INT animals ( p < 0.02). Thus, the obtained protein data for IL-1b and TNF-a clearly confirm the effects observed at the mRNA level. The most likely source of these cytokine mRNAs and proteins is neurons or glial cells since 4–8 h after KA only a few inflammatory cells could be detected (see Figs 2a–c). This was further supported by immunohistochemical staining of IL-1b in hippocampal sections of COR animals 8 h after KA injection (Figs 6a–e). Compared with PBS control rats



(a)

ADX P

4h

INT

COR

PBS Controls (8h)

8h 12h 24h 4h 8h 12h 24h 4h 8h 12h 24h

P ADX INT COR

IL-1 áα IL-1 β â

IL-6 TNF-α TNF-á

L32

(b)

(c) IL-1β/L32 mRNA (DU)

IL-1α/L32 mRNA (DU)

0.600 0.250 0.200 0.150 0.100 0.050

0.500 0.400 0.300 0.200 0.100 0.000

0.000 ADX

INT

COR

ADX

IL-1α

(d)

(e) TNF-α/L32 mRNA (DU)

IL-6/L32 mRNA (DU)

0.400 0.350 0.300 0.250 0.200 0.150 0.100

INT

COR

IL-1β 0.250 0.200 0.150 0.100 0.050

0.050 0.000

ADX

INT

COR

IL-6

0.000

ADX

INT

COR

TNF-α

Fig. 4 Effect of glucocorticoids (GCs) on the cytokine mRNA pattern over the course of time after kainic acid (KA) injection. At 4, 8, 12 and 24 h after KA injection total RNA was isolated from the hippocampi of adrenalectomized (ADX), intact (INT) and GC-treated (COR) rats, respectively. RNA (10 lg) (pooled from n ¼ 4–6 rats/group and time) was used to perform a rat cytokine (rCK-1 panel) RNase protection assay. RNase-protected fragments were resolved on a 6% polyacrylamide/7M urea gel and visualized by autoradiography. Lane P shows

the unprotected probe template. Densitometric analysis of the respective bands is shown for (b) IL-1a, (c) IL-1b, (d) IL-6 and (e) TNF-a. Results are presented as the ratio of cytokine : L32 mRNA band intensity in densitometric units (DU). (a) shows one of three independent experiments; (b)–(e) show the densitometric analysis of the cumulative data of all three experiments. h and j represent the four different timepoints (4, 8, 12 and 24 h) after KA injection. PBS, Phosphate-buffered saline.

(Figs 6d and e) IL-1b immunoreactivity was enhanced in response to kainate (Figs 6a–c). Intense IL-1b staining was detected in the pyramidal neuron layer of the CA3 region of the hippocampus as well as in the stratum oriens and stratum

lucidum which contain mostly glial cells. The morphology and localization of IL-1b-positive cells suggest that glial cells and neurons are the primary source of this cytokine in response to KA.


Glucocorticoids and CNS inflammation

(a) 160.0

**

IL-1β (pg/mg protein)

140.0 120.0

** p