Toll-like receptor 2 deficiency leads to delayed exacerbation of ischemic injury
© Bohacek et al.; licensee BioMed Central Ltd. 2012
Received: 12 January 2012
Accepted: 13 July 2012
Published: 8 August 2012
Using a live imaging approach, we have previously shown that microglia activation after stroke is characterized by marked and long-term induction of the Toll-like receptor (TLR) 2 biophotonic signals. However, the role of TLR2 (and potentially other TLRs) beyond the acute innate immune response and as early neuroprotection against ischemic injury is not well understood.
TLR2−/− mice were subjected to transient middle cerebral artery occlusion followed by different reperfusion times. Analyses assessing microglial activation profile/innate immune response were performed using in situ hybridization, immunohistochemistry analysis, flow cytometry and inflammatory cytokine array. The effects of the TLR2 deficiency on the evolution of ischemic brain injury were analyzed using a cresyl violet staining of brain sections with appropriate lesion size estimation.
Here we report that TLR2 deficiency markedly affects post-stroke immune response resulting in delayed exacerbation of the ischemic injury. The temporal analysis of the microglia/macrophage activation profiles in TLR2−/− mice and age-matched controls revealed reduced microglia/macrophage activation after stroke, reduced capacity of resident microglia to proliferate as well as decreased levels of monocyte chemotactic protein-1 (MCP-1) and consequently lower levels of CD45high/CD11b+ expressing cells as shown by flow cytometry analysis. Importantly, although acute ischemic lesions (24 to 72 h) were smaller in TLR2−/− mice, the observed alterations in innate immune response were more pronounced at later time points (at day 7) after initial stroke, which finally resulted in delayed exacerbation of ischemic lesion leading to larger chronic infarctions as compared with wild-type mice. Moreover, our results revealed that TLR2 deficiency is associated with significant decrease in the levels of neurotrophic/anti-apoptotic factor Insulin-like growth factor-1 (IGF-1), expressed by microglia in the areas both in and around ischemic lesion.
Our results clearly suggest that optimal and timely microglial activation/innate immune response is needed to limit neuronal damage after stroke.
KeywordsApoptosis IGF-1 microglia/macrophages neuroinflammation stroke TLR2−/− mice
Increasing evidence suggests that post-ischemic inflammation plays an important role in the evolution of brain injury after ischemia. However, to what extent inflammatory processes are deleterious and/or beneficial to brain recovery is a matter of debate and controversy [1–5]. One of the key features of the brain inflammatory response following ischemic injury is activation of the resident glial cells, including microglia. Activation of microglial cells after stroke is characterized by marked upregulation of the pattern-recognition receptors, such as Toll like receptors (TLRs) [6–14]. TLRs have the ability to bind two types of ligands: (i) pathogen-associated molecular pattern (PAMP) ligands in response to invading pathogens ; and (ii) endogenous danger-associated molecular pattern (DAMP) ligands, recognized in processes that present a threat to the structural integrity such as ischemic injury [12, 16–19]. Recent evidence suggests that TLRs, especially TLR2, may play a key role in the evolution of brain damage following cerebral ischemia [9–11, 20–23]. However, the exact role of TLR2 in modulation of the post-ischemic inflammatory response remains unclear. While previous studies were primarily focused on the role of TLR2 only in the acute phase 24 to 72 h after experimental stroke, the latter phases of the tissue response to ischemia remained unexplored [9, 10, 20, 21, 24]. Our recent work, using an in vivo imaging approach, revealed that the TLR2 response has a marked chronic component following brain ischemia . Therefore, the aim of the work presented here was to investigate the role of TLR2 signaling and microglial activation beyond the 24 to 72 h time window.
Here, we report that TLR2 deficiency markedly affects neuroinflammatory response to ischemic brain injury resulting in altered post-ischemic inflammatory cell activation profiles, decreased proliferative capacity of resident microglia, reduced levels of MCP-1 and reduced accumulation of CD45high/CD11b+ cells at the site of the ischemic lesion. In addition, reduced levels of the neurotrophic and anti-apoptotic factor IGF-1 expressed by microglial cells were observed in TLR2-deficient mice. Importantly, cumulative effects of the altered post-ischemic inflammatory response resulted in a late increase of neuronal apoptosis and delayed exacerbation of ischemic lesion.
All experiments were carried out on 2 to 4 month-old male TLR2−/− mice obtained from Jackson Laboratories (Bar Harbor, ME, USA), and wild-type (WT) C57BL/6 as a control group. Experimental procedures were approved by the Laval University Animal Care Ethics Committee and are in accordance with The Guide to the Care and Use of Experimental Animals of the Canadian Council on Animal Care.
As previously described [3, 11, 25, 26], unilateral transient focal cerebral ischemia was induced by intraluminal filament middle cerebral artery occlusion (MCAO) for 1 h followed by reperfusion periods of 1, 3, 4, 7 or 14 days. Briefly, all animals were anesthetized with 2% isoflurane, and body temperature was regularly checked and maintained at 37°C with a heating pad in order to avoid hypothermia. After midline neck incision, the left common carotid artery and ipsilateral internal and external carotid artery were exposed and isolated from surrounding tissue. A 12 mm long 6–0 silicon-coated monofilament suture (Doccol, CA, USA) was inserted via the proximal external carotid artery into the internal carotid artery and then into the circle of Willis, thus occluding the middle cerebral artery. After the occlusion period of 1 h the monofilament suture was removed, followed by different reperfusion periods. In our experiments we did not observe any increase in mortality in different experimental groups and, in general, mortality was very low. The whole operational procedure was conducted under the operating microscope. All animals were allowed ad libitum access to water and food before and after surgery.
After different reperfusion periods, all animals were anesthetized with an intraperitoneal injection of chloral hydrate (150 mg/kg, 300–350 μl solution, concentration 10 mg/ml), and transcardially perfused with 30 ml 0.9% saline, followed by ice-cold borax buffered 4% paraformaldehyde (PFA) at pH 9.5 (for in situ hybridization), or PBS buffered 4% PFA at pH 7.4 (for immunohistochemistry, immunofluorescent labeling and cresyl-violet staining). Tissue samples were then post-fixed overnight in 4% PFA and equilibrated in PBS buffered 30% sucrose for 48 h. Brains were embedded in Tissue-Tek (O.C.T. compound, Sakura, CA ,USA), frozen at −20°C, cut into 35 μm thick coronal sections using cryostat, and stored at −20°C.
In order to prepare samples for flow cytometric and cytokine antibody array analysis, animals were anesthetized with chloral hydrate and transcardially perfused with ice-cold 0.9% saline to remove all blood from the nervous tissue. Brain samples were surgically removed and immersed in Hibernate-A medium (for flow cytometric analysis), or immediately frozen in liquid nitrogen (for cytokine antibody array analysis).
In situ hybridization
The expression and localization of TLR2 mRNA was detected using S35 labeled riboprobes. Protocols for probe synthesis and in situ hybridization were previously described by Lalancette-Hebert and colleagues [3, 11].
Immunofluorescent labeling was performed on the brain sections of TLR2−/− mice and WT C57BL6 mice, collected 3 and 7 days after the MCAO procedure (all with visible stroke region, n = 4 animals/group/stage), according to the previously described procedure . Briefly, brain sections were blocked for 30 minutes in PBS containing 10% goat serum and 0.25% Triton X-100. Sections were then incubated overnight at room temperature with primary antibody (1:250 mouse monoclonal anti-TLR2 (eBioscience, Burlington, Ontario, Canada), 1:500 rabbit polyclonal anti-Iba1 (Wako, Richmond, VA, USA), 1:2000 rat monoclonal anti-CD68 (AbD Serotec, Oxford, UK), 1:2000 rat polyclonal anti-BrdU (Axyll, Westbury, NY, USA), 1:400 mouse monoclonal anti-BrdU Alexa Fluor 488 conjugated (Molecular Probes, Eugene, OR, USA), 1:500 rat anti-Mac2 (American Type Culture Collection, Manassas, VA, USA), 1:400 rabbit polyclonal anti-cleaved caspase-3 (Cell Signaling, Danvers, MA, USA), 1:300 mouse monoclonal anti-NeuN (Millipore, Billerica, MA, USA), 1:1000 mouse monoclonal anti-glial fibrilary acidic protein (GFAP) (Millipore Billerica, MA, USA), 1: 50 rat monoclonal anti-CD11b (AbD Serotec) and 1:50 mouse monoclonal anti-IGF-1 (Millipore)). Afterwards, sections were incubated for 2 h at room temperature in corresponding secondary antibody 1:500 (Invitrogen, Eugene, OR, USA). Each of the steps above was followed by four 5 minute rinses in PBS-0.25% Triton X-100. At the end of the procedure, sections were coverslipped with Fluoromount G (Electron Microscopy Sciences, Fort Washington, PA, USA) and dried overnight.
Immunohistochemistry was performed according to the previously described protocol [3, 11]. Brain sections of TLR2−/− and WT mice were collected 3, 7 and 14 days after the MCAO procedure (all the stroke region, n = 4 brains/group). After an initial wash in PBS to remove O.C.T., endogenous peroxidase activity was attenuated with 0.6% hydrogen peroxide in PBS1X for 15 minutes. Afterwards, sections were blocked for 1 h in PBS1X containing 10% goat serum and 0.25% Triton X-100, and incubated overnight with 1:400 rabbit anti-cleaved caspase-3 antibody (Cell Signaling) in PBS1x containing 0.25% Triton X-100 and 5% goat serum. Sections were then incubated for 2 h with 1:500 goat anti-rabbit biotinylated secondary antibodies (Jackson ImmunoResearch, West Grove, PA, USA). For the amplification of the positive signal, incubation in avidin-biotin solution was performed (Vectastain ABC kit, Vector Laboratories, Burlingame, CA, USA). Staining was developed with nickel-DAB solution (0.3%) (Vector Laboratories). Each of the steps above was followed by four 5 minute rinses in PBS-0.25% Triton X-100. At the end of the procedure, sections were dehydrated and coverslipped with DPX mounting medium (Electron Microscopy Sciences).
In order to visualize proliferating cells, mice were injected intraperitoneally with 5-bromo-2′-deoxyuridine (BrdU) (50 μg/g of mouse weight in saline) immediately before the MCAO procedure and again 2, 24 and 48 h after surgery. At 72 h after surgery, mice were anesthetized with an intraperitoneal injection of chloral hydrate (150 mg/kg, 300–350 μl solution, concentration 10 mg/ml) and transcardially perfused with 0.9% saline and PBS buffered 4% PFA. Brains were isolated, post-fixed overnight in 4% PFA, equilibrated in PBS buffered 30% sucrose for 48 h, and cut into 35 μm thick coronal sections. Afterwards, pretreatment was performed with 2.0 N HCl for protein hydrolysis (30 minutes, 37.0°C) with further neutralization of HCl with 0.1 M sodium borate (10 minutes), all diluted with KPBS. Immunofluorescent labeling was later performed on brain samples according to the standardized protocol described above.
Flow cytometric analysis
In order to analyze the number of mononuclear cells and their proliferation properties after ischemia, flow cytometric analysis was performed on WT and TLR2−/− mice at 3 and 7 days after MCAO. As previously described [27, 28], mice were injected intraperitoneally with BrdU 12, 24 and 36 h prior to sacrifice, according to BrdU Flow Kit protocol (BD, Franklin Lake, NJ, USA, USA). Collected brain samples were prepared for flow cytometric analysis according to previously described procedures [27, 28]. Briefly, tissue was mechanically dissociated with a Teflon homogenizer and incubated with 1X HBSS containing papain (2 mg/ml) and DNase I (0.025 U/ml) for 30 minutes at 37°C. Afterwards, cell suspension was passed through 70 μm nylon cell strainer (BD). In order to isolate the mononuclear cell fraction, cell suspension was loaded in 30-37-70% isotonic Percoll gradient (GE Healthcare, Uppsala, Sweden) and centrifuged for 40 minutes at 500 g. After centrifugation, an interphase containing mononuclear cells was collected, washed thoroughly, and stained with anti-CD45 conjugated with V500 (BD Horizon), and anti-CD11b conjugated with allophycocyanin (BD). Further, for proliferation analysis, cells were stained with anti-BrdU conjugated with fluorescin isothiocyanate antibody, following BrdU Flow Kit procedure (BD). Samples were afterwards analyzed on a flow cytometer (BD LSRII) immediately after the end of the staining protocol by a ‘blinded’ individual. Cells were gated using front scatter and side scatter to eliminate unviable cells. Further gating was performed using side scatter and CD45 in order to select the inflammatory cell population. Afterwards, two-color analysis with the use of anti-CD11b and anti-CD45 antibodies was performed in order to delineate CD45low/CD11b+ (that is, microglial) and CD45high/CD11b+ (that is, macrophage-like) cell populations [27, 28]. Proliferation analysis was performed using two-color analysis of anti-CD11b and anti-BrdU antibodies. Non-stroked, BrdU-treated animals were used as negative controls for proliferation analysis.
The expression profile of inflammatory cytokines was performed with a Mouse Cytokine Antibody Array (Raybio Mouse Inflammation Antibody Array 1 (Cat#AAM-INF-1), RayBiotech, Norcross, GA, USA). As previously described [3, 29] , protein samples were obtained by homogenization of TLR2−/− and WT brains 1 and 4 days after MCAO (n = 3/group) in 1x cell lysis buffer that was included in the RayBiotech kit (Raybio®Mouse Inflammation Antibody Array 1.1, Ray Biotech, #AAM-INF-1 L). Protein lysates were obtained by homogenization of brain of control and transgenic mice in 1X Cell Lysis Buffer (included in the Ray Biotech kit) with Protease inhibitor cocktail (Sigma #P8340, St. Louis, MO, USA). After extraction, samples were spun down and supernatant was used for the experiment. The protein concentration was determined for each sample and diluted at 300 μg total protein in 1x blocking buffer. For each group (three mice per group) samples were pooled together and incubated with the array membrane overnight at 4°C. After washing in the washing buffer (included in the RayBiotech kit), membranes were incubated with biotin-conjugated antibodies overnight. After washing, membranes were incubated with horseradish peroxidase conjugated streptavidin diluted in the blocking buffer for 2 h. Signal detection was performed according to RayBiotech protocol, by exposing membranes to x-ray film (Biomax MR1; #8701302; Kodak, Rochester, NY, USA), and the obtained results analyzed with ImageJ software.
Assessment of the infarct area
Every sixth coronal cryostat section of TLR2−/− and WT mouse brains collected after reperfusion periods of 3, 7 and 14 days after MCAO (n = 10/group) was stained with cresyl-violet according to the standardized protocol, coverslipped with DPX mounting medium (Electron Microscopy Sciences) and afterwards digitized. The area of infarction (‘direct stroke area’) was quantified with the ImageJ program (version 1.42q for Windows, National Institutes of Health, USA), and the infarct volume was calculated and expressed in mm3. As previously described , correction for edema was applied for brains reperfused for 3 days by calculating the brain swelling. First, the ‘indirect stroke area’ was calculated as contralateral (non-stroked) hemisphere area minus non-stroked area of the ipsilateral (infarcted) hemisphere. A difference between ‘direct’ and ‘indirect’ stroke area represented brain swelling. The correction of ‘direct stroke area’ was performed by subtracting brain swelling volume from ‘direct stroke area’. In addition, ‘indirect stroke area’ represents the volume of healthy tissue that correlates to the volume of ischemic tissue not affected by post-ischemic shrinkage. Thus, ‘direct stroke area’ reflects lesion consolidation (shrinkage) and glial scar formation and ‘indirect stroke area’ represents lesion progression (that is, how does altered inflammatory response reflect on surrounding-cell survival and lesion dynamics).
Quantification and statistical analysis
For the quantification of the immunofluorescent signal (Iba1, CD68, BrdU, Mac2, IGF-1), four fields of view per section (every sixth section, all stroke area), ten sections per animal were acquired on a fluorescent microscope (Leica DM5000B, Solms, Germany) using a 20x Plan Fluotar objective. Immunoreactivity was quantified with the ImageJ software by measuring the integrated optical density (intensity of fluorescence per unit of surface area), and results were expressed in arbitrary units as previously described . The data were averaged and analyzed by a two-tailed unpaired Student’s t-test.
Every sixth sample section through the brain was used for the immunohistochemical labeling with anti-cleaved caspase-3 antibody. The number of positively stained cells was estimated by the optical fractionator method using a Nikon Eclipse 80i microscope equipped with Stereo Investigator software. The ipsilateral (ischemic) hemisphere was traced using a 4 Plan Apochromat and sampled using 60 Plan Apochromat objectives (Nikon, El Segundo, CA, USA). The counting parameters for the experiments were the counting frame size (width and height 100 × 100 μm), sampling grid (500 × 500 μm), dissector height (10 μm) and the guard-zone thickness (1.5 μm). Counts were performed on the entire ischemic area of the ipsilateral (ischemic) hemisphere. Results were expressed as number of cells per mm3 of ischemic tissue. The data were averaged and statistically analyzed by a two-tailed unpaired Student’s t-test.
Cytokine expression assays were quantified by measuring the optical density of each cytokine spot on the membrane with ImageJ software, as previously described [3, 29]. Cytokines were presented on membranes as duplicates and the analysis was performed twice. The background values were subtracted from cytokine expression values. Data were expressed in arbitrary units relative to appropriate positive control. Further, data were averaged and analyzed by a two-tailed unpaired Student’s t-test.
Microglial cells upregulate Toll-like receptor 2 in response to transient ischemic brain injury
Toll-like receptor 2 deficiency alters microglia/macrophage activation profiles
Decreased levels of monocyte chemotactic protein-1 (MCP-1) and CD45high/CD11b+ cells in ischemic brains of Toll-like receptor 2−/− mice
Reduced proliferation of resident microglial cells in ischemic brains of Toll-like receptor 2 deficient mice
A highly characteristic feature of the injury-induced microglial activation is massive, but usually transient, expansion of microglial cell population, peaking 48 to 72 h after initial insult [3, 31]. Our results suggest a decrease in resident microglia numbers. However, at present, it is unclear to what extent the ischemic injury-induced microglial proliferation is affected in the context of TLR2 deficiency. To address this issue, the TLR2−/− and WT mice were injected daily with BrdU. The BrdU-positive/proliferating cells were quantified 3 and 7 days after stroke. In agreement with previous results, quantitative analysis of the BrdU staining (Figure 4A,B) revealed a robust increase in BrdU immunoreactivity in WT mice. As revealed in Figure 4B, we observed a 24.55% reduction in cell proliferation in TLR2−/− mice (control: WT, 0.03 × 108 ± 0.01 × 108, n = 4; TLR2−/−, 0.04 × 108 ± 0.01 × 108, n = 4, P = 0.278; 3 days: WT, 4,97 × 108 ± 0,13 × 108; TLR2−/−, 3,75 × 108 ± 0,42 × 108, n = 4, P = 0.048). To further confirm that BrdU is indeed expressed in proliferating microglial cells, double immunofluorescence for BrdU and Iba1 (markers of cell proliferation and microglia/macrophages, respectively) 3 days after transient MCAO was performed. As expected, results demonstrated that the majority of BrdU-positive cells were also positive for Iba1 staining (Figure 4C). To increase the resolution of the results obtained by BrdU signal intensity measurement the microglial proliferative capacity in TLR2−/− mice was assessed by flow cytometry measurement. Proliferation analysis was carried out on the population of CD45low/CD11b+ cells (microglial population) [27, 28] obtained from TLR2−/− and WT animals injected with BrdU (Figure 4D). The flow cytometric analysis 3 days after stroke revealed a robust, 10-fold increase in the number of CD45low/CD11b+/BrDU+ cells in WT mice (Figure 4D,E). As further shown in Figure 4E, quantitative analysis revealed a significant (55.91%) reduction in the number of proliferating CD45low/CD11b+ cells in TLR2−/− compared with WT mice (control (contralateral): WT, 1.72 ± 0.78%, n = 4; TLR2−/−, 1.39 ± 0.63%, n = 4, P = 0.762; 3 days: WT, 18.51 ± 1.41%; TLR2−/−, 8.16 ± 2.14%, n = 4, P = 0.014). The number of proliferating cells declined by day 7 and there were no significant differences in the numbers of CD45low/CD11b+/BrDU+ cells between the two experimental groups (Figure 4E).
Previously we have shown that the injury-activated resident microglial cells in proliferation upregulate galectin-3 (a Mac-2 marker of activated microglia) . Further characterization of the proliferating cells in our experimental model revealed that approximately 80% of BrdU positive cells co-localize with Mac-2 (Figure 4F). Interestingly, analysis of the Mac-2 immunoreactivity revealed a significant 40.52% reduction in the signal in TLR2−/− compared with WT mice (Figure 4G,H) (control (contralateral): WT, 0.06 × 109 ± 0.03 × 109, n = 4; TLR2−/−, 0.03 × 109 ± 0.01 × 109, n = 4, P = 0.448; 3 days: WT, 4,17 × 109 ± 0,20 × 109; TLR2−/−, 2,48 × 109 ± 0,75 × 109, n = 4, P = 0.019). Altogether, the obtained results indicate that TLR2 deficiency may affect ischemic injury-induced proliferation of the resident microglial cells.
Toll-like receptor 2 deficiency increases delayed neuronal apoptosis and exacerbates ischemic injury
To our surprise, 7 days after transient MCAO, the size of direct stroke area in TLR2−/− mice increased and was significantly larger (31.87%) when compared with controls. Due to a post-ischemic consolidation of the ischemic brain tissue and glial scar formation in the later phase (14 days after transient MCAO), direct stroke area of WT and TLR2−/− mice were reduced compared with 3 and 7 days after MCAO. Still, direct stroke area remained significantly larger in TLR2−/− mice when compared with the WT control group (27.99%) (3 days: WT, 57.92 ± 4.98, n = 10; TLR2−/−, 36.47 ± 3.58, n = 10, P = 0.003; 7 days: WT, 39.00 ± 2.78, n = 10; TLR2−/− 51.43 ± 3.47, n = 10, P = 0.012; 14 days: WT, 22.15 ± 0.96, n = 9; TLR2−/− 28.35 ± 2.09, n = 6, P = 0.010). Quantitative assessment of indirect stroke area (volume of healthy tissue that correlates to volume of damaged, ischemic tissue not affected by post-ischemic consolidation and glial scar formation) 3 days after transient MCAO revealed a significant reduction of indirect stroke area (37.58%) in TLR2−/− mice compared with the control WT group (Figure 5A,C). In addition, 7 days after transient MCAO, indirect stroke area was significantly larger (26.41%) in the group of TLR2−/− mice compared with controls. In the later phase (14 days after transient MCAO), evolution of the ischemic lesion stagnated, and indirect stroke area in TLR2−/− mice remained significantly larger (29.07%) compared with the WT control group (3 days: WT, 57.82 ± 5.04, n = 10; TLR2−/−, 36.09 ± 4.04, n = 10, P = 0.004; 7 days: WT, 42.29 ± 3.24, n = 10; TLR2−/− 53.46 ± 2.19, n = 10, P = 0.010; 14 days: WT, 44.10 ± 3.08, n = 9; TLR2−/− 56.92 ± 5.43, n = 6, P = 0.049). Altogether, these results indicate that TLR2 deficiency induces delayed exacerbation in the size of the ischemic lesion.
Reduced levels of Insulin-like growth factor 1 in brains of Toll-like receptor 2 deficient mice after transient ischemia
Microglial activation and innate immune response are key features of the brain inflammatory response to ischemic injury. Using a live imaging approach we have previously shown that microglial activation after stroke is characterized by a marked long-term induction of the TLR2 signals, thus suggesting an important role of TLR2 signaling in brain ischemia. In the present study, we report altered microglia activation profiles and delayed exacerbation of ischemic injury in the mouse model lacking functional TLR2 receptors. Namely, the TLR2 deficiency resulted in: (1) reduced microglia/macrophage activation after stroke; (2) reduced capacity of resident microglia to proliferate; and (3) decreased levels of MCP-1 and consequently lower levels of CD45high/CD11b+ expressing cells. Importantly, although acute ischemic lesions (24 to 72 h) were smaller in TLR2−/− mice, the observed alterations in the innate immune response were more pronounced at later time points (at day 7) after initial stroke, which finally resulted in delayed exacerbation of the ischemic lesion leading to larger chronic infarctions as compared with WT mice. Finally, reduced microglial activation/proliferation resulted in a significant decrease in expression of IGF-1 in microglial cells after stroke. Altogether, our results clearly suggest that optimal and timely microglial activation/innate immune response is needed to limit the extent of neuronal damage after stroke.
Upon activation, microglial cells start to express TLRs on their surface [11, 34, 35]. The functional role of TLRs is the signal transduction into the cell via TLR domain-containing adaptor proteins and MyD88 adaptor protein resulting in degradation of IκB proteins and translocation of transcription factor NFκB into the nucleus, which induces expression of proinflammatory cytokines in response to PAMPs and DAMPs [5, 36, 37]. Among diverse TLRs, it has been reported that, after transient cerebral ischemia, induction of TLR2 predominates among TLRs, followed by TLR4 and TLR9 . After brain injury and/or pathogen exposition, TLR2 is mainly expressed in the population of microglial cells [11, 20, 38, 39]. In addition, some reports indicate expression of TLR2 in astrocytes, oligodendrocytes and ependymal cells, as well as neurons and neuronal progenitor cells [12, 18, 40, 41]. Our results revealed a marked induction of TLR2 (at the mRNA and protein level) that was restricted to ischemic lesion and peri-infarct zones. No signal was observed contralaterally, that is on the non-stroke side. Further, our analysis revealed that, in early and late phases of the brain response to transient ischemia, upregulation of TLR2 occurred almost exclusively on activated microglial cells (see Figure 1C,D,E). Minor expression was found in a limited subpopulation of neurons situated in the ischemic core area and we did not observe any co-localization between TLR2 and GFAP-stained astrocytes. These results are in agreement with previous reports on TLR2 induction following brain injury from ours  and other laboratories [9, 20, 21, 23, 40, 42].
As discussed, previous studies reported initial neuroprotection against ischemia in TLR2−/− mice [9, 20, 21]. In keeping with previous work, the results of our study revealed and confirmed that, in TLR2−/− mice, the early post-stroke period (up to 3 days after transient MCAO) is characterized by smaller infarcts (see Figure 5). Interestingly, however, initial neuroprotection was not extended to later time points and we observed progressive and delayed exacerbation of neuronal damage in TLR2-deficient mice. Thus, in TLR2-deficient mice an early reduction in microglial response was associated with initially smaller lesions; a long-term decrease in microglia/macrophage activation and proliferation led to increased neuronal apoptosis and delayed exacerbation of ischemic lesion (Figures 2 3 4 5 and 6). Previous work suggests that some inflammatory molecules may have time-dependent functions ; however, the role of TLR2 (and potentially other TLRs) beyond the acute innate immune response in ischemic injury is not well understood. The results presented in this study suggest that TLR2 has a role in injury-induced monocyte/macrophage recruitment as well as in colony proliferation. Increased activation and accumulation of the myeloid cells on the site of the lesion after stroke may occur either due to massive proliferation of resident microglia, which normally peaks 48 to 72 h after initial activation, or due to increased monocyte recruitment from the circulation to the lesion site [3, 31, 44, 45]. Our data suggest that both processes may have been affected in TLR2−/− mice. We detected a selective decrease in MCP-1 levels 1 day after stroke and consequently lower numbers of CD45high/CD11b+ expressing cells at both 3 and 7 days after the onset of ischemia. In addition, TLR2 deficiency was associated with the reduced capacity of resident microglial cells to proliferate. Reduced microglial proliferation was observed in stroked brains of TLR2−/− mice 3 days after MCAO. Although at present we cannot exclude the possibility that the detected decrease in microglial proliferation may be due to initially smaller infarcts, it is important to mention that the size of infarction did increase over time in TLR2−/− mice; however, this was not accompanied by an increase in microglial proliferative response and/or activation. As shown by flow cytometric analysis, the proliferative response stayed low in TLR2−/− mice 7 days after stroke (see Figure 4). On the other hand, a decrease in CD45high/CD11b+ expressing cells observed at days 3 and 7, and a selective decrease in MCP-1 observed in TLR2−/− mice, suggest that TLR2 signaling is needed for adequate recruitment of circulating monocytes. Taken together, these results support in part the hypothesis established on the axonal injury model which suggests that, in the acute phase, TLR2 may modulate proliferation rather than the entry of circulating monocytes [23, 31, 46–48].
The question that arises here is how to correlate deficient monocyte/microglia responses observed in TLR2−/− mice with delayed exacerbation of the ischemic injury. One of the possibilities is that, as previously reported, rapid phagocytic removal of dead or dying cells prevents the release of proinflammatory intracellular components and contributes to the resolution of inflammation . Reduced CD68 signal intensity may suggest a reduced number of phagosomes present on the lesion site in TLR2−/− mice, thus implying that the engulfment of cellular debris is altered with a consequently prolonged inflammation. It is noteworthy that altered microglia/macrophage activation profiles observed in TLR2−/− mice were more pronounced at day 7 post-stroke, which coincided with the initial enlargement of ischemic lesions in TLR2-deficient mice. An additional explanation as suggested by our results is that a defective innate immune response in the brain caused by TLR2 deficiency results in deficient microglia proliferation and consequently reduced levels of IGF-1, which may lead to an increase in delayed neuronal death and exacerbation of ischemic injury.
Based on our results, we may conclude that reduction in numbers as well as aberrant profiles of the activated microglia/macrophages may over time result in a microenvironment that is detrimental for neuronal survival and causes exacerbation of ischemic lesions.
Taken together, our data suggest that functional TLR2 signaling is required for modulation of microglia/macrophage responses in the injured brain. Moreover, our data also indicate that TLR2 induction may exert temporally differential effects. However, the ischemic injury-induced TLR2 response in the brain is instrumental for timely resolution of the post-ischemic inflammation, finally limiting exacerbation and/or propagation of the initial ischemia-induced neuronal damage.
danger associated molecular pattern
glial fibrilary acidic protein
Insulin-like growth factor −1
middle cerebral artery occlusion
Monocyte chemotactic protein-1
pathogen-associated molecular pattern
Toll like receptor
tumor necrosis factor
This work was supported by The Canadian Institutes of Health Research (CIHR) Operating Grant to JK. JK is recipient of the Fonds de la Recherché en Santé Québec (FRSQ) Senior Scholarship. IB, DG and SG received funding from Unity through Knowledge Fund (UKF) grant 35/08. The authors also wish to thank Ms Genevieve Soucy for her technical assistance.
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