Hypothyroidism during the developmental stage induces disruption of hippocampal neurogenesis in later life, as well as inducing oxidative stress in the brain. The present study investigated the preventive effect of co-exposure to an antioxidant on disruptive neurogenesis induced by developmental exposure to anti-thyroid agent in rats. For this purpose, we used two antioxidants, α-glycosyl isoquercitrin (AGIQ) and α-lipoic acid (ALA). Mated female Sprague Dawley rats were either untreated (control) or treated with 12 ppm 6-propyl-2-thiouracil (PTU), an anti-thyroid agent, in drinking water from gestational day 6 to postnatal day (PND) 21, the latter group being subjected to feeding basal diet alone or diet containing AGIQ at 5,000 ppm or ALA at 2,000 ppm during PTU exposure. On PND 21, PTU-exposed offspring showed reductions in a broad range of granule cell lineage subpopulations and a change in the number of GABAergic interneuron subpopulations. Co-exposure of AGIQ or ALA with PTU altered the transcript levels of many genes across multiple functions, suggestive of enhancement of synaptic plasticity and neurogenesis. Nevertheless, immunohistochemical results did not support these changes. PTU exposure and co-exposure of AGIQ or ALA with PTU did not alter the hippocampal lipid peroxidation level. The obtained results suggest a possibility that thyroid hormone depletion itself primarily disrupts neurogenesis and that oxidative stress may not be involved in the disruption during development. Transcript expression changes of many genes caused by antioxidants may be the result of neuroprotective actions of antioxidants rather than their antioxidant activity. However, no preventive effect on neurogenesis suggested impairment of protein synthesis via an effect on mRNA translation due to hypothyroidism.
The hippocampus is an important temporal lobe brain structure involved in cognition, learning, and memory. The hippocampus postnatally generates new neurons within the subgranular zone (SGZ) of the dentate gyrus (DG), which is termed “adult neurogenesis” (Fig. 1)1, 2. This neuronal production consists of multistep processes, including a number of developmental phases, such as self-renewal of neural stem cells, the facilitation of continued division of precursor cells to produce new granule cells, and subsequent differentiation and migration of these new cells into the granule cell layer (GCL)1, 2. In the hilus of the DG, subpopulations of γ-aminobutyric acid-ergic (GABAergic) interneurons innervate granule cell lineage populations to control neurogenesis in the SGZ2, 3. In addition to GABAergic neuronal inputs, various types of neurons outside the SGZ also create a synaptic connection with neurons in the DG, such as glutamatergic neurons in the entorhinal cortex providing axonal projections to the DG4 and cholinergic neurons originating from the septal nucleus and nucleus of the diagonal band of Broca innervating neurons in the hilus of DG4. Glutamatergic inputs to the SGZ are important for maintaining proper proliferation and differentiation of the granule cell lineage subpopulations3.
Thyroid hormones are crucial for brain development during fetal and neonatal periods. They play important roles in neuronal proliferation and migration, neuritogenesis, synaptogenesis, and myelinogenesis5. Previous studies have shown that rat developmental hypothyroidism causes aberrant brain growth involving diverse cellular populations and also impairs inherent brain structures and functions5. Hypothyroidism impairs neuronal migration and results in subcortical band heterotopia in the corpus callosum5, 6, as well as white matter hypoplasia with suppression of both axonal myelination and oligodendrocytic accumulation7. It has been considered that maternal hypothyroidism is associated with autism spectrum disorders (ASD)5. Multiple studies have highlighted the involvement of key processes, such as those including neurogenesis, neurite growth, synaptogenesis, and synaptic plasticity, in the pathophysiology of neurodevelopmental disorders, such as ASD8. Therefore, an experimental induction of developmental hypothyroidism could provide a reasonable model for ASD9.
Oxidative stress is defined as an imbalance between reactive oxygen species (ROS) production and the antioxidant defense system in an organism and is involved in various disorders such as neurodegenerative diseases and malignant tumors. Neural injury in the central and peripheral nervous system caused by some kinds of neurotoxicants is considered to be related to the induction of oxidative stress, but it is unclear how neurotoxicants cause oxidative stress and neurotoxicity. Importantly, SGZ cells in the DG generate ROS, because these cells have a high cellular activity for proliferation and differentiation requiring high oxygen demand10. With regard to the effect of hypothyroidism on brain tissues, induction of oxidative stress has been reported in the rat hippocampus as a result of administration of an anti-thyroid agent during the developmental stage or adult stage11, 12. Furthermore, co-exposure of extracts of a medicinal plant, Nigella sativa, with an anti-thyroid agent, 6-propyl-2-thiouracil (PTU), from the gestation period to adult stage in rats reduces apoptosis in the hippocampal DG, the Cornu Ammonis region (CA) 1 and CA3 areas, as compared with PTU exposure alone13. Therefore, there is a possibility that developmental exposure to an antioxidant may prevent hypothyroidism-related disruption of hippocampal neurogenesis induced by administration with an anti-thyroid agent during development in rats.
The present study was performed to clarify whether developmental exposure to an antioxidant has a potential to prevent hypothyroidism-related disruption of hippocampal neurogenesis induced by administration with PTU during development in rats. For this purpose, we used two antioxidants, α-glycosyl isoquercitrin (AGIQ) and α-lipoic acid (ALA), in the present study. AGIQ, also known as enzymatically modified isoquercitrin, is a flavonol glycoside derived by enzymatic glycosylation of rutin. AGIQ is a mixture of quercetin glycoside, consisting of isoquercitrin and its α-glucosylated derivatives, with 1–10 or more of additional linear glucose moieties and has greater water solubility and bioavailability14. AGIQ has been reported to exert anti-oxidative15, anti-inflammatory16, anti-hypertensive17, anti-allergic18 and tumor suppressive15, 19, 20 properties. ALA, a natural compound that is chemically named 5-(1,2-dithiolan-3-yl)pentanoic acid and is also known as thioctic acid21, is another metabolic antioxidant. In addition to direct antioxidant activity, ALA and its endogenous counterpart dihydrolipoic acid (DHLA), which is rapidly formed after uptake into the body’s cells, contributes to the nonenzymatic regeneration of reduced glutathione, vitamin C, vitamin E, and coenzyme Q10 in vivo22. Moreover, DHLA stimulates glutathione synthesis by enhancing cellular cysteine uptake. As DHLA is a supplier of reducing equivalents for the regeneration of detoxification enzymes, it is capable of supporting repair of oxidative damage22. ALA has been reported to prevent or ameliorate several ailments such as cardiovascular diseases, diabetic complications including retinopathy and neuropathy, and hypertension because of its antioxidant properties23. It is known that once absorbed in the body, AGIQ and ALA can pass the blood-brain barrier and be distributed in the brain24, 25. We have recently found that continuous AGIQ exposure from the developmental stage can facilitate fear extinction learning associated with enhancement of synaptic plasticity at the adult stage in rats26. Experimentally, ALA has been shown to ameliorate brain oxidative injury induced by methionine and choline deficiency27. Therefore, if oxidative stress is involved in aberrant neurogenesis in the hippocampal DG induced by developmental hypothyroidism, there is a possibility that co-exposure to AGIQ or ALA would suppress aberrant neurogenesis in the hippocampal DG.
Materials and Methods
Chemicals and animals
PTU (purity >99%; CAS No. 51-52-5) was purchased from MilliporeSigma (St. Louis, MO, USA). AGIQ (purity >97%) was supplied by San-Ei Gen F.F.I. Inc. (Osaka, Japan). DL-ALA (purity ≥99%; CAS No. 1077-28-7) was purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo, Japan). Fifty mated female Slc:SD rats were purchased from Japan SLC, Inc. (Hamamatsu, Japan) at gestational day (GD) 1, where GD 0 was the day of appearance of a vaginal plug. Mated female rats were individually housed in polycarbonate cages with paper bedding until postnatal day (PND) 21, where PND 0 was defined as the day of delivery. Animals were maintained in an air-conditioned animal room (temperature, 23 ± 2°C; relative humidity, 55 ± 15%) with a 12-h light/dark cycle. Mated female rats were allowed to access to powdered basal diet (CRF-1, Oriental Yeast Co., Ltd., Tokyo, Japan) and tap water ad libitum until the start of developmental exposure to PTU with or without exposure to AGIQ or ALA. Offspring were weaned on PND 21 and thereafter reared three to five animals per cage and provided with powdered basal diet (CRF-1) and tap water ad libitum.
Mated female rats were randomly divided into four groups by stratified randomization according to the body weight on GD 5; 14 rats were fed basal diet and tap water (untreated controls), 12 rats were fed basal diet and water containing 12 ppm PTU (PTU alone), 12 rats were fed diet containing 5,000 ppm AGIQ and water containing 12 ppm PTU (PTU + AGIQ) or 12 rats were fed diet containing 2,000 ppm ALA and water containing 12 ppm PTU (PTU + ALA) (Fig. 2). Animals were treated from GD 6 to day 21 post-delivery with PTU with or without AGIQ or ALA. Based on a previous study that showed apparent aberrations in neuronal development in the hippocampal structure in offspring28, the PTU dose was set at 12 ppm. The chosen dosages of AGIQ and ALA have both been shown to suppress the promotion of hepatic preneoplastic lesions in rats19, 20.
Dams were subjected to measurement of body weight, and food and water consumption, twice a week between GD 6 and PND 21. On PND 4, the litters were randomly culled to preserve 6 or 7 male pups and 1 or 2 female pups per litter. If dams had fewer than 6 male pups, more female pups were included to maintain a total of 8 pups per litter. The offspring were weighed twice a week until PND 21. All dams and oﬀspring were checked for general conditions in terms of appearance of abnormal gait and behaviors at the time of body weight measurement. Dams were euthanized by exsanguination from the abdominal aorta under CO2/O2 anesthesia on PND 21.
In the present study, male offspring were selected for immunohistochemical and gene expression analyses of the hippocampus because neurogenesis is influenced by circulating levels of steroid hormones during the estrous cycle29. On PND 21, 10 male offspring per group (1 pup per dam) were subjected to perfusion fixation for brain immunohistochemistry through the left cardiac ventricle with ice-cold 4% (w/v) paraformaldehyde (PFA) in 0.1 M phosphate buffer (pH 7.4) at a flow rate of 10 mL/min under deep anesthetization with CO2/O2. For transcript expression analysis, 6 male offspring per group (1 pup per dam) were euthanized by exsanguination from the abdominal aorta under CO2/O2 anesthesia and subjected to necropsy, and brains were removed and then fixed in methacarn solution at 4°C for 4 hours. For lipid peroxidation measurement, 6 to 8 male offspring per group (1 pup per dam) were euthanized by exsanguination from the abdominal aorta under CO2/O2 anesthesia and subjected to necropsy, and bilateral hippocampi were removed and stored at –80°C. All female offspring were similarly euthanized under anesthesia and subjected to necropsy, and brain tissues were removed and stored at –80°C. The remaining male offspring were maintained without exposure to PTU, AGIQ, or ALA until PND 77, and body weight, as well as food and water consumption, was measured once a week.
On PND 77, 8 to 10 male offspring per group (1 pup per dam) were subjected to perfusion fixation with ice-cold 4% PFA buffer solution for brain immunohistochemistry at a flow rate of 35 mL/min. For transcript expression analysis, 6 to 8 male offspring per group (1 pup per dam) were subjected to necropsy, and removed brains were fixed in methacarn solution.
The dosing schedule of PTU and necropsy time points of the present study were identical to those in a previous study6, following recommendations in the Organization for Economic Co-operation and Development (OECD) guideline for the testing of chemicals (Test No. 426: Developmental Neurotoxicity Study)30. All procedures in this study were conducted in accordance with the Guidelines for Proper Conduct of Animal Experiments (Science Council of Japan, 1 June 2006) and according to the protocol approved by the Animal Care and Use Committee of Tokyo University of Agriculture and Technology. All efforts were made to minimize animal suffering.
Immunohistochemistry and apoptotic cell detection
After perfusion with 4% PFA buffer solution on PND 21 and PND 77, brains were additionally immersed in the same solution overnight at 4°C. In untreated controls, 3-mm-thick coronal slices were prepared at −3.0 mm from the bregma on PND 21 and at −3.5 mm from the bregma on PND 77. In the PTU-exposure group, brain size was turned out to be small, and coronal slices were prepared at the proportionally similar position to the untreated controls. Brain slices were immersed in 4% PFA buffer solution overnight at 4°C and were routinely processed for paraffin embedding and sectioned into 3-μm-thick slices.
Brain sections from offspring on PND 21 and PND 77 were subjected to immunohistochemistry using primary antibodies against the following antigens: proliferating cell nuclear antigen (PCNA), a cell proliferation marker in the SGZ; glial fibrillary acidic protein (GFAP), which is expressed in type-1 neural stem cells (radial glial cells) in the SGZ and astrocytes2; SRY box 2 (SOX2), which is expressed in type-1 neural stem cells and type-2a progenitor cells in the SGZ1; T-box brain 2 (TBR2), which is expressed in type-2b progenitor cells in the SGZ1; doublecortin (DCX), which is expressed in type-2b and type-3 progenitor cells and immature granule cells in the SGZ and GCL2; neuronal nuclei (NeuN), which is expressed in postmitotic neurons of both immature and mature granule cells in the SGZ and GCL2; and reelin (RELN), parvalbumin (PVALB), calbindin-D-29K (CALB2), and somatostatin (SST), which are expressed in GABAergic interneurons in the DG hilar region3; activity-regulated cytoskeleton associated protein (ARC), Fos proto-oncogene, AP-1 transcription factor subunit (FOS), and cyclooxygenase 2 (COX2), which are members of the immediate-early genes involved in synaptic plasticity31, 32 in the GCL. The respective primary antibodies were applied to brain sections for incubation overnight at 4°C. The primary antibodies are listed in Supplementary Table 1 (online only). One section per animal was subjected to immunohistochemistry of each molecule.
To block endogenous peroxidase, deparaffinized sections were incubated in 0.3% (v/v) H2O2 solution in absolute methanol for 30 min. The antigen retrieval conditions that were applied for some antibodies are listed in Supplementary Table 1 (online only). Immunodetection was conducted using a Vectastain® Elite ABC kit (Vector Laboratories Inc., Burlingame, CA, USA) with 3,3’-diaminobenzidine (DAB)/H2O2 as the chromogen. Hematoxylin counterstaining was then performed, and coverslips were mounted on immunostained sections for microscopic examination.
To evaluate apoptosis in the SGZ of the DG in the offspring, a terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) assay was performed using an ApopTag Peroxidase In Situ Apoptosis Detection Kit (MilliporeSigma) according to the manufacturer’s instructions, with DAB/H2O2 as the chromogen. One section per animal was subjected to a TUNEL assay.
Evaluation of immunoreactive cells and apoptotic cells
Immunoreactive cells, i.e., PCNA+, GFAP+, SOX2+, TBR2+, DCX+, NeuN+, ARC+, FOS+, and COX2+ cells or TUNEL+ apoptotic cells, in the SGZ and/or GCL were bilaterally counted and normalized for the length of the SGZ (Fig. 1). Immunoreactive cells distributed within the hilus of the hippocampal DG, i.e., RELN+, PVALB+, CALB2+, SST+, or NeuN+ cells, were bilaterally counted and normalized per area unit of the hilar area (Fig. 1).
Immunoreactive neurons located inside of the CA3, consisting of large pyramidal neurons that can be morphologically distinguished from relatively small interneurons, were excluded from counting immunoreactive cells in the hilus of the DG. The number of each immunoreactive cellular population (except for NeuN+ cells in the GCL) or TUNEL+ apoptotic cells was manually counted under microscopic observation using a BX53 microscope (Olympus Corporation, Tokyo, Japan). In the case of NeuN+ cells in the GCL, the number of immunoreactive cells for counting was high, and therefore, an image analysis-assisted automatic cell counting method was applied. More specifically, digital photomicrographs at ×200-fold magnification were taken using a DP72 Digital Camera System (Olympus Corporation) attached to a BX53 microscope, and positive cell counting was performed by applying the WinROOF image analysis software package (version 5.7; Mitani Corporation, Fukui, Japan). The length of the SGZ and the hilar area were measured in microscopic images at ×40-fold magnification by applying the cellSens Standard (version 1.9; Olympus Corporation).
Transcript expression analysis
Transcript expression levels in the hippocampal DG were examined using real-time reverse-transcription polymerase chain reaction in offspring on PND 21 and PND 77. Brain tissues were dissected according to the whole-brain fixation method using methacarn solution as previously reported33. In brief, 2-mm-thick coronal cerebral slices were prepared at the position of −3.0 mm from the bregma. Hippocampal DG tissues were collected from the slice using a punch-biopsy device with a pore size of 1 mm in diameter (Kai Industries Co., Ltd., Gifu, Japan). Total RNA was extracted from tissue samples from each group (n=6 per group at both PND 21 and PND 77) using