Free heme induces neuroinflammation and cognitive impairment by microglial activation via the TLR4/MyD88/NF-κB signaling pathway

The animal experiments were approved by the Ethics Committee of the First Affiliated Hospital of Zhengzhou University. All animal experiments were carried out in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Six-month-old male Sprague–Dawley rats (500-600 g) were obtained from Shanghai Jihui Laboratory Animal Care Co.,Ltd. Animals were maintained under standard controlled environment at constant temperature of 22 ± 2 °C and humidity of 50 ± 10% with 12-h light/dark cycle. Food and water were available ad libitum. The rats were left to acclimatize to the environment for two weeks prior to inclusion in the experiment.

The rats were randomized into three groups: 1) Control group (n = 10), in which rats received sham operation; 2) Heme group (n = 10), in which rats received an intraperitoneal injection of hemin; and 3) Heme + HPX group (n = 10), in which rats received an intraperitoneal injection of hemin and an intracerebroventricular injection of HPX. On the first day of the experiment (T1), the control group rats received an intraperitoneal injection of 0.5 mL normal saline (NS). The Heme group and Heme + HPX group rats received an intraperitoneal injection of 0.5 ml hemin (25 mg). After 12 h (T2), the control group and Heme group rats received an intracerebroventricular injection of 5 μl 0.1% sodium azide, and the Heme + HPX group rats received an intracerebroventricular injection of 5 μl (5 μg) HPX. Behavior tests were conducted over the next 6 days (T3). Euthanasia was performed by intraperitoneal injection of pentobarbital sodium (75 mg/kg) on day 7 (T4) after the behavioral tests. The brain tissue of rats was taken, half of which was used for immunofluorescence staining. The other half was made into brain tissue homogenate (Fig. 1A).

Hemin (H9039, Sigma, USA) was freshly dissolved in DMSO and diluted with NS to a final concentration of 50 mg/ml in darkness. Rats in the Heme group and Heme + HPX group were intraperitoneally injected with 0.5 ml (25 mg, 50 mg/kg) hemin. The dose was selected based on previous studies [20, 21, 26–29]. Rats in the control group were intraperitoneally injected with 0.5 ml NS. HPX (Abcam-ab198629, UK) was dissolved in 0.1% sodium azide and diluted to 1 μg/μl. Rats in the Heme + HPX group were intracerebroventricularly injected with 5 μl (5 μg) HPX. Rats in the control group and Heme group were intracerebroventricularly injected with 5 μl 0.1% sodium azide. The dose was selected based on previous studies [21, 30].

Rats were anesthetized with isoflurane (2–3% isoflurane in O₂ at 2 L/min). Anesthetized rats were placed on a stereotaxic apparatus and the skull was fixed and secured with four stainless-steel screws. An incision to the scalp exposed the surface of the skull and bregma. A burr hole was drilled into the bone of the right hemisphere with a stainless steel 26-gauge cannula, located 1.5 mm lateral to, and 0.8 mm posterior to the bregma. A 25 μl Hamilton micro syringe was very slowly inserted to 3.5 mm beneath the dural surface to inject 5 μl HPX or vehicle (0.1% sodium azide). The micro syringe was kept in place for 5 min to ensure the effectiveness of injection before withdrawal. After withdrawal, the incision was sutured, and the rats recovered from anesthesia.

After injection of hemin or hemin + HPX, the rats were subjected to the MWM test to measure cognitive function (spatial learning and memory). The MWM test had two sessions: the acquisition session for five consecutive days and the probe trial session on day 6. In the MWM test, a black circular tank (150 cm in diameter and 50 cm in height) filled with water (22 ± 2 °C) was divided into four quadrants. A hidden platform (diameter 10 cm) was placed in the middle of one quadrant 1–1.5 cm below the water. The swimming path of each rat was recorded by video camera mounted directly above the tank. In the acquisition session, the rats were placed into the water facing the wall of the pool in one of the four quadrants. Each rat was allowed 120 s to find and mount the platform. When the rat found the platform, it was kept on the platform for 10 s. If the rat did not find the platform within 120 s, it was guided to the platform and allowed to stay on it for 10 s, and then the escape latency was recorded as 120 s. The escape latency, path length and swimming speed were recorded. In the probe trial session, 24 h after the last trial of the acquisition session, the original platform was removed. Rats were placed in the quadrant opposite to the platform quadrant and allowed to swim for 120 s. The number of platform crossings and time spent in the targeted quadrant were recorded.

The BV2 mouse microglia cell line (FH0355) and HT22 mouse hippocampal neuron cell line (FH1027) was both purchased from Shanghai FuHeng Biology Co., Ltd. in China. On the first day, BV2 cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco) supplemented with 10% fetal bovine serum (FBS) (Sigma–Aldrich, MO, USA) and 1% penicillin/streptomycin (Gibco) in humidified 5% CO₂ air environment at 37 °C. On the second day, the BV2 cell culture medium was replaced, and the cells were divided into 4 groups: 1) the Control group (BV2), in which BV2 cells were incubated with medium only for 24 h; 2) the Heme group (BV2 + Heme) in which BV2 cells were treated with 40 μM hemin (H9039, Sigma, USA) for 24 h; 3) the MyD88 inhibition group (BV2 + Heme + T6167923) in which BV2 cells were treated with 40 μM hemin and 2 μM MyD88 inhibitor (T6167923, #HY-19744, MedChemExpress, USA) for 24 h; and 4) the NF-κB inhibition group (BV2 + Heme + JSH-23), in which BV2 cells were treated with 40 μM hemin and 20 μM NF-κB inhibitor (JSH-23, #S7351, Selleck, USA) for 24 h. Then, the BV2 cells were harvested for subsequent experiments.

The BV2 and HT22 cell coculture system was conducted using Corning Transwell polycarbonate membrane cell inserts (Corning Costar Corp, USA). BV2 cells were placed above the HT22 cell layer. HT22 cells were cultured in 24-well plates, and BV2 cells were seeded onto Transwell inserts. Coculture systems were divided into 4 groups: 1) Neuron group (HT22) in which only HT22 cells were incubated with medium for 24 h; 2) Neuron Heme group (HT22 + Heme) in which HT22 cells were treated with 40 μM hemin (H9039, Sigma, USA) for 24 h; 3) Neuron + microglia group (HT22 + BV2) in which HT22 cells cocultured with BV2 cells were incubated with medium for 24 h; 4) Neuron + microglia Heme group (HT22 + BV2 + Heme) in which HT22 cells cocultured with BV2 cells were treated with 40 μM hemin (H9039, Sigma, USA) for 24 h (Fig. 4A). Then, the coculture systems were harvested for subsequent experiments.

The complex of free heme and HPX could be recognized and cleared by CD91. Therefore, in the rat experiment, we detected the expression of CD91 in brain tissue by FCM. The brain tissue was dissected and minced into small pieces in 0.125% trypsin and 0.02% EDTA for digestion at 37 °C for 25 min. Then, brain tissue suspensions were passed through cell strainer (70 μm, Fisher brand) followed by incubation in culture bottles treated with poly L-lysine in 5% CO₂ at 37 °C for 30 min. When only a few cells adhered to the wall, the cell suspension was centrifuged at 300 g at 4 °C for 5 min and resuspended in medium for counting. After washing with PBS, the cells were resuspended in binding buffer at a density of 1 × 10⁶ cells/ml and subjected to be incubated with a primary antibody (ab92544, Abcam) and then a secondary fluorescent antibody (ab7010, Abcam) in the dark at 4 °C for 30 min. Stained cells were spined down to a pellet and resuspended in the wash buffer for further analysis. In the cell culture experiment, HT22 cell apoptosis was assessed by Annexin V-FITC and propidium iodide (PI) (40302ES20, Yeasen Biotechnology, Shanghai, China). HT22 cells were digested with trypsin and centrifuged at 300 g at 4 °C for 5 min to wash. Cell pellets were resuspended in binding buffer at a density of 1 × 10⁶ cells/ml, and Annexin-FITC and PI were added. After reaction at room temperature for 15 min in the dark, binding buffer was added, and apoptotic cells were analyzed. Cells were analyzed on flow cytometer (FC 500, Beckman Coulter) as soon as possible, and the data were analyzed by FlowJo software (Informer Technologies, USA).

In rat experiment, the dorsal hippocampus was dissected based on the atlas of Paxinos and Watson and homogenized in RIPA lysis buffer followed by centrifugation at 13,000 × g for 10 min at 4 °C. The supernatant was collected, and the protein concentration was determined using bicinchoninic acid (BCA) assay kit (Beyotime Biotechnology, Shanghai, China). Commercially available ELISA kits for measuring TNF-α (CER1393, CRK Pharma, Wuhan, China), IL-1β (CER1094, CRK Pharma, Wuhan, China) and IL-6 (CER0042, CRK Pharma, Wuhan, China) levels were used according to the manufacturers’ instructions. In cell experiment, the culture medium of each group was collected, and the quantities of TNF-α (EK282, MultiSciences, Hangzhou, China), IL-1β (EK201B, MultiSciences, Hangzhou, China), and IL-6 (EK206HS, MultiSciences, Hangzhou, China) were determined using ELISA kits according to the manufacturer’s instructions.

The expression of ionized calcium binding adaptor molecule 1 (Iba-1) was detected in rat experiment. The rat brain was fixed and sectioned. Then the sections were dehydrated with a series of graded ethanol solutions, cleared with distilled water, deparaffinized by xylene, hydrated with gradient alcohol and boiled for antigen retrieval. After blocking nonspecific binding with 3% bovine serum albumin (BSA)/10% normal goat serum for 30 min, the sections were incubated with primary anti-Iba-1 antibody (ab178846, Abcam) at 4 °C overnight. Then, the sections were washed and incubated with the secondary antibody goat anti-rabbit IgG-H&L (Alexa Fluor® 647) (ab150079, 1:1000, Abcam) at room temperature in the dark with slow shaking for 1 h. In the cell coculture experiment, HT22 cells were fixed, permeabilized, blocked, and incubated with primary antibody against neuron-specific nuclear protein (NeuN) (ab104224, Abcam) at 4 °C overnight. After washing, the cells were incubated with goat anti-mouse IgG-H&L (Alexa Fluor®647) (ab150115, 1:1000, Abcam) secondary antibody at room temperature in the dark with slow shaking for 1 h. In the BV2 cell culture experiment, the expression of CD86 was detected. The cells were fixed, permeated, blocked, and incubated with primary antibodies against CD86 (A16805, ABclonal) at 4 °C overnight. After washing, the cells were incubated with secondary antibodies at room temperature in the dark with slow shaking for 1 h. The nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI) and are shown in blue. Images were obtained by laser scanning confocal microscopy (Nikon, Tokyo, Japan), and ImageJ software (Bethesda MD, USA) was used for analysis.

Western blot analysis was performed to detect the possible mechanism of the TLR4/MyD88/NF-κB signaling pathway in neuroinflammation induced by free heme in rat and BV2 cell experiments. Brain tissue from the rats or harvested BV2 cell pellets were homogenized in ice-cold RIPA lysis buffer (Beyotime Biotechnology) with the addition of protease and phosphatase inhibitor cocktail (Beyotime Biotechnology) to obtain total protein. Protein concentrations were quantified by BCA protein assay kit (Beyotime Biotechnology) according to the manufacturer’s instructions. Protein samples were suspended in loading buffer, boiled at 100 °C for 10 min, separated by 8–12% SDS-PAGE and transferred to polyvinylidene fluoride membranes (Millipore). The membranes were blocked with 5% skim milk for 1 h at room temperature and then incubated with primary antibodies at 4 °C overnight. In rat experiment, primary antibodies were as follows: anti-TLR4 (A5258, 1:1000, ABclonal, Wuhan, China); anti-MyD88 (ab219413, 1:1000, Abcam, Cambridge, UK); anti-p-NF-κB p-p65 (#3033, 1:1000, Cell Signaling Technology, Beverly, MA, USA); and anti-NF-κB p65 (#8242, 1:1000, Cell Signaling Technology, Beverly, MA, USA). In BV2 cell experiment, the primary antibodies were as follows: anti-TLR4 (19,811–1-AP, 1:1000, Proteintech, Wuhan, China); anti-MyD88 (ab219413, 1:1000, Abcam, Cambridge, UK); anti-p-NF-κB p-p65 (#3033, 1:1000, Cell Signaling Technology, Beverly, MA, USA); and anti-NF-κB p65 (10745–1-AP, 1:1000, Proteintech, Wuhan, China). The membranes were incubated with species-appropriate horseradish peroxidase (HRP)-conjugated secondary antibodies (1:5000, Cell Signaling Technology) for 1 h at 37 °C. Finally, the protein bands were visualized with enhanced chemiluminescence and quantified using ImageJ software.

Statistical analysis in this study was conducted using SPSS 22.0 software (IBM Corp., Armonk, NY, USA) and GraphPad Prism 9 (GraphPad Software, San Diego, CA). One-way analysis of variance (ANOVA), two-way ANOVA, repeated ANOVA, and unpaired or paired Student’s t test were used to evaluate the differences among the groups, followed by Tukey’s post hoc test or Bonferroni’s post hoc test, as appropriate. Data are presented as means ± SEM. A value of p < 0.05 was set as statistical significance.

The MWM test was conducted to measure the cognitive function (spatial learning and memory) of rats. The swimming path of each rat was recorded (Fig. 1 A, B). In the acquisition session, the control group rats required the shortest time, and the Heme group rats required the longest time to find the target platform from day 1 to day 5. Rats in the control group and Heme + HPX group required less time on day 5 than on day 1 to find the target platform. While in the Heme group, the escape latency was not significantly different between day 5 and day 1. As the training progressed, the rats in the control group exhibited a discernible drop in escape latency. However, the Heme group did not show the same training performance as the control group (Fig. 1 C). No motor dysfunction was detected according to the swimming speed results (Additional file 1: Fig. S1). In the probe trial session, rats in the Heme group and Heme + HPX group crossed the platform less frequently and spent less time (%) in the target quadrant than those in the control group. Compared with the Heme group, the Heme + HPX group rats crossed the platform more frequently and spent more time (%) in the target quadrant (Fig. 1 B, D, E). The MWM test data indicated that Heme had negative effects on the spatial learning and memory abilities of rats and HPX could alleviate these effects.

HPX has high heme affinity, and heme-HPX complexes can be endocytosed by microglia. HPX has neuroprotective effect of keeping free heme at safe levels by preventing unregulated heme uptake [31]. CD91 is receptor protein for heme-HPX complexes. The uptake by CD91 leads to endocytosis and lysosomal degradation of the heme-HPX complexes [32]. So, the number of CD91 + cells represented the elimination condition of free heme. We hypothesized that free heme could enter the brain and activate the heme-HPX-CD91 scavenging system. FCM was performed to analyze the CD91 + cells and the results indicated that on day 7 after administration, CD91 + cells were significantly increased in the Heme group compared with the control group, and the Heme + HPX group had fewer CD91 + cells compared to the Heme group (Fig. 1 F, G, H). CD91 is the receptor to identify the heme-HPX complex. Therefore, the increase of CD91 + cells induced by hemin suggests that, there are still many heme-HPX complexes to be scavenged after a week of hemin administration. In addition, the decrease of CD91 + cells in the Heme + HPX group suggests that, HPX may help the brain clean excessive free heme, and the heme-HPX complexes have been cleaned faster compared to the Heme group within 7 days.

To explore inflammation in rat brain, we took brain homogenates and detected the inflammatory cytokines TNF-α, IL-1β and IL-6. The ELISA analysis showed that, compared with the control group, TNF-α expression was significantly increased in the Heme group. Meanwhile, compared with the Heme group, TNF-α expression was decreased in the Heme + HPX group, suggesting that heme could induce inflammatory reaction in rat brain and that HPX treatment could reduce the reaction (Fig. 2A). Similarly, IL-1β expression was significantly increased in the Heme group compared with the control group and decreased in the Heme + HPX group compared with the Heme group (Fig. 2B). However, IL-6 expression was significantly decreased in the Heme group compared with the control group and increased in the Heme + HPX group compared with the Heme group (Fig. 2C), which might be related to the time of obtaining brain tissue, which was one week after administration.

To investigate the microglial response to free heme and HPX, rat brains were dissected, and immunofluorescence staining was used to assess the expression of Iba-1, which is a calcium binding protein specifically expressed in microglia. The results showed that Iba-1 was widely expressed in the hippocampus of rat brain (Fig. 2 D). We counted the number of Iba-1 + cells and found that compared with the control group, Iba-1 + cells were significantly increased in the Heme group. Meanwhile, the number of Iba-1 + cells was decreased in the Heme + HPX group compared with the Heme group (Fig. 2 E, F), suggesting that free heme could increase the number of microglia in rat brain and that HPX treatment could reduce the number of microglia. These findings indicate that free heme could induce activation of microglia and neuroinflammation in rat brain, but HPX treatment could reduce these effects.

Microglia play an important role in the process of CNS injury and are the main effector cells of neuroinflammation. To investigate the inflammatory response of microglia induced by free heme, we conducted BV2 cell culture in vitro. After hemin treatment, we detected inflammation-related cytokines (TNF-α, IL-1β, and IL-6). ELISA analysis showed that compared with the control group, TNF-α, IL-1β and IL-6 expression was significantly increased in the Heme group of BV2 cells (Fig. 3 A, B, C). Next, FCM was conducted to determine whether hemin could induce neuronal apoptosis in HT22 cells. We found that the percentage of apoptotic cells was significantly increased in the Heme group compared with the control group (Fig. 3 D, E). Additionally, immunofluorescence staining was conducted to observe neurons (Fig. 3 F). These findings indicated that hemin significantly induced microglial inflammation and neuronal apoptosis.

To verify that microglia were involved in neuronal apoptosis, we cocultured BV2 cells and HT22 cells with hemin treatment (Fig. 4 A) and detected neuronal apoptosis in HT22 cells by FCM. We found that in HT22 cells cultured alone or in the HT22 + BV2 cell coculture system, hemin significantly induced neuronal apoptosis. Notably, compared with that in the HT22 Heme group, the apoptosis of neurons in the HT22 + BV2 Heme group was significantly increased, indicating that microglia were involved in and increased neuronal apoptosis (Fig. 4 B, C). Then, immunofluorescence staining was used to observe HT22 cells in four groups (Fig. 4 D). NeuN antibodies were tools for staining live mature neurons. In HT22 cells alone, hemin didn’t reduce the number of NeuN + cells. While in the coculture system, hemin significantly reduced the number of NeuN + cells (Fig. 4 E). Therefore, the results suggests that microglia aggravated free heme-induced neuronal apoptosis. Furthermore, hemin might decrease the number of live mature neurons under the effect of microglia.

We assumed that free heme could cause microglial inflammation and neurocognitive dysfunction by regulating the TLR4/MyD88/NF-κB signaling pathway. In the rat experiment, to verify this hypothesis, we detected TLR4, MyD88, p-P65 (p-NF-κB) and P65 (NF-κB) proteins in rat brain tissue by western blot (Fig. 5 A). The results indicated that, compared with the control group, hemin significantly increased the protein levels of TLR4, MyD88 and p-P65. Notably, compared with Heme group, HPX significantly reduced the protein levels of TLR4 and MyD88 (Fig. 5 B).

In cell culture experiment, to further evaluate the effects of hemin on the regulation of the pathway, BV2 cells were treated with hemin or hemin + inhibitor for 24 h. Western blot analysis results revealed that hemin significantly increased TLR4, MyD88 and p-P65 protein levels compared with the control group. As expected, the MyD88 inhibitor (T6167923) and NF-κB inhibitor (JSH-23) significantly downregulated hemin-induced pathway protein levels. Interestingly, the protein level of TLR4 was decreased by MyD88 inhibitor (T6167923) and NF-κB inhibitor (JSH-23) treatment compared with that in the Heme group, which might be related to the positive feedback regulation of the pathway (Fig. 5 C, D). Next, we hypothesized that inhibiting the pathway could reduce the inflammatory response of microglia. To verify this hypothesis, we detected inflammatory cytokines. ELISA analysis showed that the MyD88 inhibitor (T6167923) and NF-κB inhibitor (JSH-23) significantly decreased hemin-induced TNF-α, IL-1β and IL-6 expression in BV2 cells (Fig. 5 E, F, G).

Then, we observed the expression of the M1 surface marker CD86 in BV2 cells. Consistent with the inflammatory cytokine ELISA results, the CD86 + cells were significantly increased in the Heme group compared with the control group, and MyD88/NF-κB inhibition reversed the increase of CD86 + cells compared to the Heme group (Fig. 5 H, I). Overall, the data demonstrated that free heme could cause neuroinflammation by microglial activation via the TLR4/MyD88/NF-κB signaling pathway.

This study was approved by the Life Science Ethics Committee of Zhengzhou University.

Not applicable.

The authors declare no competing interests.

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.