Introduction
Early brain injury that occurs at the time of bleeding is the leading cause of mortality (30–70%) following subarachnoid hemorrhage (SAH) [
1,
2]. SAH survivors are at risk of developing delayed cerebral vasospasm, delayed cerebral ischemia, or delayed ischemic neurological deficits during the hospital course [
2]. Delayed vasospasm develops in approximately 70% of patients between 3 and 14 days after SAH [
1,
2]. For decades, it has been considered as the single and the most important cause of delayed cerebral ischemia and poor outcome [
3]. Even patients with favorable outcomes are frequently left with significant residual memory, reduced executive functioning, or language deficits [
4]. Cerebral vasospasm following aneurysmal SAH is the leading cause of death and disability after aneurysm rupture [
5]. Cerebral ischemia secondary to vasospasm occurs in 20 to 30% of these patients and has been correlated with a 1.5- to threefold increase in mortality in the first 2 weeks after SAH [
3,
6]. Although cerebral vasospasm associated with SAH has been recognized for more than 50 years, adequate treatment is still illusive. Thence, the pathophysiological mechanism contributing to this form of arterial dysfunction is a topic of intense experimental study.
One of the major consequences of early brain injury is apoptosis, which is known to occur within minutes to 24 h after SAH [
7]. Immune cells, including microglia and astrocytes, release pro-inflammatory factors to induce cell apoptosis and activation of transcription factor, resulting in positive feedbacks. Microglia and astrocytes also mediate neuropathic behavior by modulating the activity of spinal neurons to cause central sensitization, which has been associated with inflammatory neuropathies in autoimmune thyroid diseases, complex regional pain syndrome, osteoarthritis, rheumatoid arthritis, post-operative pain, and SAH [
7,
8].
Hepatoma-derived growth factor (HDGF) is a 240 amino-acid protein isolated from human hepatoma cells [
9]. Surface expressed nucleolin has recently been identified as an HDGF receptor [
10]. During cell development, HDGF stimulates cell proliferation in fibroblasts, endothelial cells, and hepatoma cells [
11]. It is also a growth factor related to tissue organogenesis and is involved in the development and regeneration of the liver [
12‐
14], lungs [
15,
16], kidney [
17], heart [
18], and the vascular system [
19‐
21]. On the other hand, HDGF expression is associated with various malignant cancers, including hepatocellular carcinoma (HCC), gastric cancer, non-small cell lung cancer, pancreatic cancer, and melanoma [
22,
23], to name a few. In addition, HDGF plays important roles in various cellular events, including ribosome biogenesis, RNA processing, DNA damage repair, and transcriptional regulation [
17]. Furthermore, many studies indicate that HDGF is a mitogen with extracellular proliferative effects on hepatoma cells, fibroblasts, vascular smooth muscle cells, and endothelial cells [
16]. However, no reports showed the relationship between HDGF and SAH. Hence, this was the focus of the present investigation.
Materials and Methods
Animal Preparation
The study procedures were executed in accordance with the protocol approved by the Committee of Institutional Animal Research at Kaohsiung Medical University. Male Sprague–Dawley rats were purchased from BioLasco (BioLasco Taiwan Co., Ltd., Taipei, Taiwan, authorized by Charles River Lab). After arriving at the Kaohsiung Medical University vivarium, the rats were acclimated for at least 1 week before being used in the experiment. All rats were housed at a constant temperature (24 °C) and at regular light/dark cycles between 6:00 am and 6:00 pm, with free access to a standard diet. Animals weighing between 350 and 450 g were used in this study.
SAH Induction
The one-shot SAH model was utilized. Briefly, rats were anesthetized by intraperitoneal injection of 40 mg/kg Zoletil 50® containing a mixture of zolazepam and tiletamine hypochloride (Virbac, Carros, France). The head was fixed in a stereotactic apparatus (Stoelting, Wood Dale IL, USA), and a 25-gauge butterfly needle was advanced into the cisterna magna to withdraw 0.3 mL of cerebrospinal fluid (CSF). Fresh, autologous, and non-heparinized blood (0.1 mL/100 g of body weight) drawn from the central tail artery was then slowly instilled into the subarachnoid space through a butterfly needle and tubing. Afterwards, the animals were kept in a ventral recumbent position for at least 30 min to promote ventral blood distribution. The morbidity caused by SAH was 100%, and the mortality was 0% in this study. The respiratory pattern of rats was inspected closely, and mechanical ventilation was provided if necessary. Upon fully awakening, the animals were sent back to the vivarium. In this study, there are five groups: (1) control group; (2) rHDGF ab group, treatment with rHDGF antibody into the subarachnoid space; (3) SAH group; (4) SAH + pre-rHDGF ab group, treatment with rHDGF antibody into the subarachnoid space within 24 h before SAH; and (5) SAH + post-rHDGF ab group, treatment with rHDGF antibody into the subarachnoid space within 24 h after SAH. All rats were used following randomization.
Preparation of HDGF Hyperimmune Serum (Recombinant HDGF Antibody)
HDGF hyperimmune serum was customized by Leadgene Biomedical Inc. (Tainan, Taiwan). Briefly, 6- to 8-week-old BALB/c mice were primed by intraperitoneal injection of 50 μg of recombinant rat HDGF (rHDGF) protein in complete Freund’s adjuvant. Two weeks after the first injection, the mice were given another injection of 25 μg rHDGF antibody in PBS. The procedure was repeated at weeks 4 and 5 after antigen priming. The sera were collected and stored at – 20 °C until use.
Neurological Evaluation
We followed the method of neurobehavioral evaluation from Dr. Huang et al. (2017) [
24]. Neurobehavioral evaluation of animals was performed by assessing the sensorimotor integration of the forelimb and hind-limb activities using a modified limb-placing test that consisted of ambulation as well as placing and stepping reflex [
25]. Motor deficit index (MDI) represented the sum of scores for walking by lower limbs and for placing/stepping response and was determined at 48 h after induction of SAH. Higher MDI values indicated poorer neurological outcomes. In addition, neurological evaluation was used as a double-blind trial.
Tissue Processing
This protocol was conducted in a as previously described in Dr. Huang et al. (2017) [
24]. At the end of experiments, each animal was anesthetized again for perfusion and fixation. The thoracic cage was opened, and the left ventricle was canalled using a No. 16 catheter. After clamping, the descending aorta and puncturing the right atrium, the brain was perfused with 180 mL of 2% paraformaldehyde and then 100 mL of phosphate buffer (0.01 M) under 36 °C and 100 mm Hg perfusion pressure. Gross inspection of harvested brains was performed to confirm the presence of subarachnoid blood clots over the basilar artery (BA), and the specimen was immersed in a fixative solution. The BAs were then separated from the brainstems, and the middle third of each vessel was dissected out. These arterial segments were flat-embedded in paraffin, and BA cross sections were cut into 3-μm sections and were stained with hematoxylin and eosin stain for subsequent analysis.
Morphometric Assessment of BA
This protocol was conducted in a as previously described in Dr. Huang et al. (2017) [
24]. Three cross sections from the middle-third BA in each animal were analyzed by a trained research staff blinded to the experimental groups. The thickness of BA was defined as the largest vertical distance between the inner surface of endothelium and the outer surface of adventitia. The arterial cross-sectional area was calculated using computer-based morphometric analysis (Image J; Universal Imaging Corp., USA). The average area of BA cross sections from each rat was calculated to obtain mean values for the degree of vasospasm at 48 h after SAH.
Immunofluorescence Staining
After deparaffinization and rehydration, paraffin-embedded brain samples were heated by steam for 30 min to retrieve antigen using DAKO antigen retrieval solution (DAKO, Carpenteria, CA). Slides were then washed twice with Tris-buffered saline (TBS) and immersed in a 3% hydrogen peroxide solution for 10 min to inhibit endogenous peroxidase. Upon washing twice in TBS, the sections were incubated with TUNEL kit, mouse anti-NeuN (Merck; MAB377; Germany), mouse anti-GFAP (Sigma; G3893; USA), and rabbit anti-Iba1 (proteintech; 10,904–1-AP; USA) antibodies at room temperature to detect the presence of the cell number of DNA damage, astrocytes, microglia, and neuron cells, respectively. Slides were again washed twice with TBS and subsequently incubated with anti-rabbit antibody (Thermo; A11008; USA) and anti-mouse antibody (Thermo; A10524; USA) for 3 h at room temperature. Afterwards, the slides were washed twice with TBS, stained and mounted within Fluroshield™ with DAPI (Sigma; F6057; USA), and take a picture following LSM 700 confocal microscope.
Western Blot
Rat cortex samples were collected at 48 h after SAH. Tissue extracts were prepared in 1 mL of ice-cold lysis buffer [50 mM Tris–HCl, pH 7.4, 0.25 M NaCl, 0.1% Nonidet P-40, 5 mM EDTA, 50 mM NaF, 1 × cocktail of protease inhibitors (Sigma, St. Louis, MO), 1 mM phenylmethylsulfonyl fluoride, and 1 mg/L aprotinin] and incubated on ice for 30 min. After centrifugation at 8000 g for 20 min at 4 ℃, protein amount in the supernatant was quantified using a protein assay kit from Bio-Rad Laboratories, and 60 μg of protein samples were separated by 12% SDS–polyacrylamide gel electrophoresis. Proteins were then transferred to nitrocellulose membranes. The membranes were blocked in PBS containing 5% fat-free milk for 90 min at room temperature and then incubated with antibodies (HDGF and cleaved caspase-3) for 2 h at room temperature. After washing, the membranes were incubated for 1 h at 25 °C with the appropriate horseradish peroxidase-labeled secondary antibodies, and the bound antibodies were visualized and quantified by chemiluminescence detection. The expression level of proteins of interest was normalized to the densitometric units of β-actin.
ELISA
Serum and cerebrospinal fluid (CSF) samples were collected at 48 h after SAH, centrifuged immediately at 2000 × g for 10 min at 4 °C to remove cells, and were stored below − 15 °C prior to analysis. X These samples were concentrated by passing through C2 columns (Amersham, Nutley, USA), and the levels of TNF-α, IL-1β, and IL-6 were determined using an inflammatory factor ELISA system (Amersham) at 450 nm.
Statistics
The results were analyzed using SPSS version 20.0 (IBM SPSS Statistics, location). Data were presented as mean ± standard deviation (SD). Group results were compared using Student’s t test, Mann–Whitney U test, or one-way analysis of variance (ANOVA). A P value of < 0.05 was considered statistically significant.
Discussion
It has been reported that HDGF stimulates cell proliferation in fibroblasts, endothelial cells, and hepatoma cells [
10] as well as mediates inflammation [
33]. Consistent with these findings, our study showed that treatment with rHDGF ab could decreased the proliferation of microglia and astrocytes as well as reduced the levels of pro-inflammatory factors induced by SAH (Figs.
2,
3, and
4).
Inflammation and cytokines may participate in the pathology of blood–brain barrier (BBB) disruption and brain edema, which are characteristic features for both clinical and experimental SAH [
34,
35]. A variety of inflammatory cytokines, including IL-1β, IL-6, and TNF-α, are strongly associated with brain injury in the rat [
36]. Inhibition of IL-1β has been shown to attenuate early brain injury (EBI) and improve BBB function after SAH [
37]. The present study showed that pre-treatment with rHDGF ab decreased the SAH-induced production of TNF-α, IL-1β, and IL-6.
Many studies have shown that HDGF may play a role in apoptosis. For example, silencing the HDGF gene has been demonstrated to prevent TNF-α/cycloheximide-induced apoptosis [
38]. In addition, HDGF is found to be essential for TNF α-induced release of pro-apoptotic factors from mitochondria [
38]. In contrast, it has been demonstrated that knockdown of HDGF gene induces apoptosis [
39,
40] and cell cycle arrest in several human cancers. HDGF knockdown not only induces expression and de-phosphorylation of the pro-apoptotic protein Bad, but also inactivates ERK and Akt, resulting in activation of the intrinsic apoptotic pathway in cancers [
40‐
42]. However, in our study, blocking HDGF attenuated SAH-induced neuron cell apoptosis in the brain (Fig.
6). Similarly, HDGF knockdown triggers the Fas-mediated extrinsic apoptotic pathway in HepG2 cells through the nuclear factor-κB (NF-κB) signaling [
43]. NF-κB is a nuclear transcription factor that acts as a key regulator of both inflammatory response and cell death [
30,
31]. Our study showed that pre-treatment with rHDGF ab down-regulated inflammatory factors and p-NFκB/NFκB ratio. These data supported the notion that blocking HDGF attenuated apoptosis of neuron cells in the cortex after SAH (Fig.
5D).
Brain-derived neurotrophic factor (BDNF), a molecule that regulates neuronal survival and differentiation, has a critical role in synaptic plasticity [
44]. However, BDNF is initially synthesized as a precursor, proBDNF, which is trafficked to the regulated secretory pathway [
29]. A previous report showed that mature BDNF can protect neurons from amyloid-beta (Aβ)-induced apoptosis by increasing the expression of bcl-2 [
36]. (Irrelevant) Our results showed that the protein levels of proBDNF were significantly elevated in the cortex of SAH rats, and pre-treatment with rHDGF ab decreased proBDNF protein expression following SAH, suggesting that the beneficial effects seen with blocking HDGF may be mediated, at least partially, by the BDNF pathway.
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