Background
The prevalence of metabolic syndrome (MetS) in adults is increasing worldwide [
1], largely due to several factors such as ageing of the population, increased life expectancy and chronic overnutrition and physical inactivity [
2]. MetS refers to a cluster of inter-connected metabolic abnormalities involving glucose metabolism (diabetes milletus), lipid metabolism (hypercholesterolaemia and dyslipidaemia), elevated blood pressure and central obesity. In particular, obesity is emerging as a major health concern owing to its key role in MetS and being a well-recognised risk factor for the development of type 2 diabetes (T2D) and related cardiovascular diseases (CVDs) [
3]. Recently, interdisciplinary research in neuroscience and immunology has linked overnutrition to neuroinflammation, particularly in the hypothalamus and hippocampus [
4]. A growing body of evidence suggests that a complex interplay exists between accelerated adiposity, hyperglycemia and cognitive decline [
5]. Indeed, clinical studies demonstrate that patients with T2D are more susceptible to symptoms of neurological impairment, but how T2D may induce neurological deficits remains undetermined.
The recently identified systemic and chronic inflammatory component of MetS is a primary candidate to confer the increased risk of cognitive decline in both obese and hyperglycemic individuals, perhaps by priming the specialised resident macrophage population (microglia) towards an inflammatory state to establish a form of low-grade neuroinflammation [
6]. Chronic hyperglycemia in its own right is known to have deleterious effects on the brain. Meta-analyses of non-obese hyperglycemic type 1 diabetes patients show mild declines in mental speed and flexibility compared to non-diabetic individuals [
7]. Moreover, increased blood-brain barrier permeability, oxidative damage and neuronal apoptosis are all associated with hyperglycemia in experimental animal models of hyperglycemia [
8‐
10]. These conditions are seen similarly in conjunction with obesity, where negative relationships exist between body mass index (BMI) and neuronal viability, myelin integrity and grey matter volumes of the hippocampus in middle-aged adults [
11,
12] However, both clinical and experimental studies have yet to elucidate the specific contributions of either obesity or hyperglycemia to its implicated neuroinflammatory component. Foundational animal studies have primarily investigated the contributions of obesity and hyperglycemia in mice deficient in either the leptin (
ob/ob) or leptin receptor (
db/db) as a result of genetic mutations. While beneficial, findings in these models have often proven difficult to translate to, and do not accurately reflect, the disease aetiology of obesity and diabetes in humans [
13]. Furthermore, these models do not allow for the analysis of individual components of MetS in the development of neuroinflammation.
The present study was designed to dissect the specific contributions of high-fat feeding and hyperglycemia to neuroinflammation. We hypothesise that the systemic inflammatory component as a result of high-fat feeding is required to initially activate the brain’s immune system, which may be worsened by progressive hyperglycemia. To test our hypothesis, we assessed changes to systemic inflammatory cytokines, microglia numbers and astrocytosis in the hippocampus, and immune cell infiltrate in the brains of high-fat diet (HFD)-fed and streptozotocin (STZ)-treated C57BL/6 mice over 9 and 18 weeks. We also used intravital multiphoton microscopy to assess leukocyte-endothelial cell interactions in the cerebral vasculature of these mice in vivo.
Methods
Mice
Five to 6-week-old adult male C57Bl/6 J mice were obtained from the Monash Animal Research Platform and were housed in specific pathogen-free (SPF) conditions with access to food and water ad libitum (Monash Medical Centre Animal Facility, Clayton, VIC, Australia). Mice were housed 2–5 per cage in temperature-controlled rooms (22 °C) under a standard 12 h light-dark cycle. Prior to the start of experiments, mice were acclimatised for a minimum of 7 days before use. Mice were monitored every 3–4 days for specific health monitoring parameters including body weight, alertness, activity, coat health, dehydration, and gait. All procedures were approved by the Monash University Animal Ethics Committee under regulations that comply with the National Institute of Health guidelines for the care and use of laboratory animals.
Induction of hyperglycemia and high-fat feeding
After acclimatisation, all mice were randomly assigned treatment groups. Mice for use in the control group underwent three intraperitoneal (IP) injections of citrate vehicle (CIT, 0.01 M, pH 4.5) 3, 5 and 7 days prior to commencement of control diet (CON; SF09-091, 16% kilojoules from lipids, Speciality Feeds, WA) for 9 or 18 weeks. These mice were designated with the group name “CIT + CON”. To develop a high-fat feeding model, a separate group of mice was treated identically with citrate vehicle and were placed on a high-fat diet (HFD; SF16-104, 44% kilojoules from lipids, Specialty Feeds, WA) for 9 or 18 weeks and designated “CIT + HFD”. To develop a mild hyperglycemia-only model, mice were treated with three separate IP injections of low-dose streptozotocin within 5 min of preparation (STZ; 55 mg/kg in 0.01 M CIT) 3, 5 and 7 days prior to the commencement of control diet for 9 or 18 weeks. These mice were designated with the group name “STZ + CON”. To develop a combination model of both hyperglycemia and high-fat feeding, mice were treated with three separate IP injections of STZ (55 mg/kg in 0.01 M CIT) 3, 5 and 7 days prior to the commencement of a HFD for 9 or 18 weeks. These mice were designated the group name “STZ + HFD”. Low-dose streptozotocin (STZ) treatment was selected as it causes the targeted destruction of the insulin-producing β cells of the pancreas to produce mild hyperglycemia [
14].
Glucose and insulin measurements
Fasting blood glucose was monitored fortnightly in all mice as a health monitoring parameter using a hand-held glucometer (Accu-Chek Performa, Roche) from whole blood collected via saphenous tail vein bleeds. Mice were fasted for 3 h prior to blood collection. Two weeks before end-point, a glucose tolerance test was performed. Baseline fasting blood glucose levels were obtained and mice were then injected IP with 0.8 g/kg glucose solution. Further measurements were taken at 15, 30, 45, 60, 90 and 120 min post-injection via tail vein bleeds. In addition, glycated haemoglobin (HbA1c) was assessed at the experimental endpoint. Mice were anaesthetised with isofluorane prior to transcardiac puncture with a heparinised 26G needle (100 I.U/mL). A small aliquot of whole blood (20 μL) was used for HbA1c analysis, via the Cobas B 101 (Roche Diagnostics). Plasma was collected from the remaining blood and stored at − 80 °C until required. Insulin levels were evaluated from plasma using a mouse ultrasensitive insulin ELISA kit (Alpco, Beijing, China) in single replicates according to the manufacturer’s instructions.
Flow cytometry of brain immune cell isolates
The left cerebral hemisphere was cut into small pieces and digested in 500 μL collagenase digestion buffer at 37 °C for 30 min (collagenase XI (125 U/mL), hyaluronidase (60 U/mL) and collagenase type I-S (450 U/mL) in DPBS (Ca2+/Mg2+ supplemented)). Samples were kept on ice and passed through a 70 μm nylon cell strainer. Single-cell suspensions were subjected to 30% isotonic Percoll, underlaid with 70% Percoll and centrifuged for 20 min at 600g at room temperature. The interphase containing the mononuclear cells was collected, washed and stained with eFluor 506-conjugated anti-mouse CD45 (30-F11, eBioscience), FITC-conjugated anti-mouse CD3 (145-2CC, eBioscience), eFluor 450-conjugated anti-mouse B220 (RA3-6B2, eBioscience), PE-Cy7-conjugated anti-mouse CD11b (M1/70, eBioscience) and PE-conjugated anti-mouse CD68 (FA/11, eBioscience) with Fc-receptor blocker (1:100, 2.4G2, BD Pharmingen) for 15 min on ice in the dark. After staining, the cells were washed, resuspended with counting bead mixture (50 / μL) and 7AAD live-dead viability stain (1:50, 420404, Biolegend) in FACS buffer. Samples were run using a flow cytometer (BD FACSCanto II) and analysed using FlowJo (v10.3.0). Cell numbers were normalised to the left cerebral hemisphere weight of each mouse.
Immunofluorescence
Mice were deeply anaesthetised with a cocktail of ketamine (150 mg/kg) and xylazine (10 mg/kg) prior to cardiac perfusion. An 18G needle was inserted into the left ventricle to perfuse ice-cold PBS and subsequent 4% paraformaldehyde (PFA; in PBS, pH 7.4). After perfusion, the brain was left to fix in PFA for 1 h and cryoprotected in 20% sucrose overnight. Brains were frozen in isopentane for cutting using a cryostat. Tissue anterior to the hippocampus was discarded until the defining bowtie-like structures of Cornu Ammonis (CA1) could be visually identified. Sections were cut at 10 μm and left at room temperature for 30 min prior to storage at − 80 °C until required. Upon use, sections were acclimatised to room temperature prior to rehydration with Tris-buffered saline containing 0.1% Tween 20 (TBST, pH 7.6), thrice, for 5 min. Sections were encircled with hydrophobic ink to maintain a staining well. Sections were permeabilised in 0.5% Triton X-100 (in 1x TBST, pH 7.6) at room temperature for 15 min in a humidifying chamber. Sections were then incubated for 48 h at 4 °C with either APC-conjugated anti-mouse Iba1 (1:1000, NCNP24, Wako Chemicals) to label microglia, anti-mouse GFAP (1:250, 1B4, BD Pharmingen) to label only activated astrocytes or PE-conjugated anti-mouse CD45 (1:200, 30F11, eBioscience) in TBST to label general immune cells. After incubation, excess stain was removed and sections were washed thrice with TBST for 10 min at room temperature, followed by a deionised water wash for 5 min. Sections were cover-slipped using DAPI ProLong Diamond Antifade Mountant (Invitrogen) to stain all nuclei and were left to cure for a minimum of 1 h at 4 °C in the dark. The 18-week cohort was stained with primary mouse anti-Iba1 (1:200, NCNP24, Wako Chemicals) and secondary anti-rabbit IgG (1:200, AlexaFluor-568, A-11011, Life Technologies) according to the manufacturer’s recommended instructions. To perform neuronal counts, the sections were stained with guinea pig anti-mouse NeuN (1:1000, MAB3777, Merck Millipore) and donkey anti-guinea pig IgG (1:400, AlexaFluor-647, Jackson Immunoresearch).
Quantification of neurons and immune cells
Imaging of Immunofluorescence sections was performed on a Nikon C1 confocal laser-scanning microscope (Hamamatsu, Japan) at × 20 magnification with 405 nm (DAPI), 568 nm (PE) and 637 nm (APC) lasers. All microscope settings and laser powers were kept identical between each batch of sections imaged. Cell counts were performed manually, under blinded conditions, selected by appropriate cell body co-localisation of DAPI+ and either CD45+, IBA1+, or NeuN cells using FIJI (v1.51, NIH). Adjustments to brightness and contrast were made to individual files to allow optimal visualisation of CD45+, Iba1+ and NeuN+ cells only. Quantification of GFAP+ activated astrocytes per area stained was performed using identical threshold settings for all samples within each cohort using FIJI.
Cytometric bead array
Plasma cytokine levels were measured by cytometric bead array (CBA) using the BD CBA Mouse Th1/Th2/Th17 Cytokine Kit (BD Biosciences, San Jose, CA, USA). The kit was used for parallel detection of mouse interleukin-2 (IL-2), interleukin-4 (IL-4), interleukin-6 (IL-6), interferon-γ (IFNγ), tumour necrosis factor (TNF), interleukin-17A (IL-17A) and interleukin-10 (IL-10). Standards were reconstituted with 2 mL of assay diluent to a maximum concentration of 5000 pg/mL and serially diluted to a concentration of 2.5 pg/mL. Capture bead mix was prepared by adding 60% of total capture beads to 40% assay diluent as per the kit instructions. Thereafter, 25 μL of bead mix, 25 μL of plasma and 15 μL phycoerythrin (PE) detection reagent were added consecutively to each well and incubated at room temperature for 2 h in the dark. Samples were washed and resuspended in wash buffer. Assay diluent was used as an internal control in place of sample plasma and returned negative values for all cytokines. Samples were measured using the Navios flow cytometer (Beckman Coulter Inc) and analysed using FlowJo (v 10.3.0). Standard curves and interpolations were constructed using GraphPad Prism 7.0 software.
Real-time in vivo imaging of cerebral vasculature
To examine the leukocyte-endothelial cell interactions in the brain at experimental endpoints, intravital multiphoton microscopy of the brain was performed. Mice were anaesthetised by IP injection of an anaesthetic cocktail consisting of 150 mg/kg ketamine hydrochloride and 10 mg/kg xylazine and the tail vein was cannulised to administer fluorescently labelled antibodies and/or additional anaesthetic, if required. Body temperature was maintained using a heat pad. Mice were immobilised in a stereotactic frame, whereby the skin overlying the skull was swabbed with ethanol and a 1 cm vertical incision was made to expose the skull. The skin was retracted, and a 5 mm × 5 mm cranial window was shaved to a thin membrane using a dental drill. PE-conjugated anti-mouse CD31 (390, 1 mg/25 g mouse, eBioscience) and PacBlue-conjugated anti-mouse GR1 (RB6-8C5, 1 mg/25 g mouse, eBioscience,) were injected through the intravenous cannula to label the cerebral brain vasculature and neutrophils/monocytes, respectively. At least three post-capillary venules of approximately 20–40 μm in diameter were chosen per mouse. Images and videos were acquired using a multiphoton microscope (Leica SP5), equipped with a × 20 water-dipping objective (NA 1.0) and a MaiTai pulsed infrared laser (SpectraPhysics) set to an excitation wavelength of 810 nm. A 512 × 512 pixel image was acquired every 1.5 s for 9 min. Adjustments to brightness and contrast were made to individual files to allow optimal visualisation and measurements of blood vessels and cells. Multiple measurements of the width of blood vessels were taken and averaged, while the length of the vessel was measured along its centre, both using the segmented line tool. The number of adherent, intravascular cells was normalised to the vessel surface area and time of recording. Cells were considered adherent if they interacted with the vessel wall for 30 s or more. The measurements from all fields of view from one mouse were averaged with the final data representing neutrophil/monocyte numbers from one mouse.
Statistical analysis
All statistical analyses were performed using GraphPad Prism Software (La Jolla, CA, USA). The normality of all data were first assessed using a Shapiro-Wilk normality test and analysed using a one-way analysis of variance (ANOVA) or Kruskal-Wallis posthoc test if appropriate. Each group was compared to the CIT + CON group. Some data were excluded due to outliers, determined using Grubbs’ test (Prism 7) at P ≤ 0.05. All values are mean ± SEM. A value of P ≤ 0.05 was considered statistically significant.
Discussion
In this study, we dissect the specific contributions of high-fat feeding and hyperglycemia to neuroinflammation in experimental mouse models of MetS. We demonstrate that acute hyperglycemia induces regional-specific effects on the brain by elevating microglial numbers and promotes astrocytosis in the hippocampus. In addition, we demonstrate that chronic hyperglycemia supports the recruitment of peripheral leukocytes, particularly GR1+ granulocytes, to the cerebral microvasculature. Moreover, we provide evidence that these changes are independent of the systemic inflammation associated with high-fat feeding. Taken together, our findings suggest that hyperglycemia rather than high-fat feeding contributes to the development of regional neuroinflammation in animal models of MetS.
MetS in humans is associated with a wide-array of phenotypes. Therefore, the need for clinically relevant and highly reproducible animal models is paramount. Research has primarily centered on using monogenetic deletions or mutations that indirectly induce severe obesity, yet in the case of T2D, obesity is usually second only to environmental influences, not manifestations of genetic mutations [
16]. Thus, the clinical relevance of these models to obesity in MetS and T2D in humans remains controversial. In this study, we used a high-fat diet as a method of inducing weight gain and low-dose STZ-treatment in producing a form of mild hyperglycemia. While a definitive diagnosis of hyperglycemia in T2D can vary depending on the weight, gender and race of an individual, clinical benchmarks constitute mild hyperglycemia as > 7–9% glycosylated haemoglobin [
17]. Our findings indicate that the STZ treatment regime induced a clinically comparable mild hyperglycemia (8–9%), which was sustained over both 9-week and 18-week time points. To induce weight gain, high-fat feeding was selected to reflect the macronutrient composition of the westernised diet, strongly implicated in the development of obesity and insulin resistance in MetS [
18]. We found that high-fat feeding (ad libitum) significantly induced weight gain. Similarly, high-fat feeding was associated with significant impairments to glucose metabolism and hyperinsulinaemia, but not hyperglycemia. Previous studies of HFD in C57Bl/6 J mice note percentage weight gain and insulin resistance consistent with our findings, yet contrastingly, with induced hyperglycemia, which may in part be due to the composition of lipids in our diet (22% fat), compared to these studies (60% fat) [
19]. Our attempt of a combination model (STZ + HFD) resulted in weight gain at 18 weeks, but not 9 weeks. The delayed progression of weight gain in the mice of STZ + HFD group may be attributed to preferential lipid metabolism following impairment of glucose uptake. While only observational, these are critically important findings. Our use of high-fat feeding and STZ-treatment seemingly echo a sounder model of weight gain and hyperglycemia after 18 weeks, reflecting similar conditions and levels relevant to those seen in clinical MetS, without the need of gene deletions or mutations to induce these conditions. Although beyond the scope of this study, it would be important to validate whether our models demonstrate the diverse comorbidities of MetS, such as cardiovascular disease, nephropathy and atherosclerosis. The presence of these complications that arise from MetS patients would lend credence to our models in more accurately representing the more complex physiology and clinical presentation of MetS patients.
Administration of STZ alone to male C57BL/6 mice is commonly associated with retarded weight gain, as we and others routinely observe [
20]. Similar retarded weight gain is also evident in rats subjected to STZ alone [
21], as well as other mouse models where there is also hyperglycemia in the absence of high-fat diet [
22,
23]. This retarded weight gain is a direct result of the degree of hyperglycemia, as it is not evident at a milder hyperglycemia [
21], and is preventable by insulin replacement [
23]. Rather than proposing that high glucose protects against the weight-gaining effect of a HFD, we would hence propose that a HFD protects against the retarded weight gain effect of hyperglycemia in rodent models. This protective effect of HFD on retarded weight gain as a result of impaired insulin availability may reflect a specific rodent phenomenon rather than a potential means of preventing obesity in diabetic humans. Interestingly, however, patients diagnosed with type 1 diabetes (insulin deficiency) exhibit a more severe hyperglycemia than patients diagnosed with type 2 diabetes (insulin resistance), yet it is the latter that are more commonly associated with obesity [
24]. Clinical and experimental evidence has established that chronic hyperglycemia and weight gain in MetS are associated with both localised and systemic inflammation, yet studies struggle to delineate this from the effects of hyperglycemia alone. Our current study is able to shed light on this. By giving a HFD to both CIT- and STZ-treated mice, we examined whether systemic inflammation was driven solely by high-fat feeding, or in conjunction with hyperglycemia. Our findings show that mice in the CIT + HFD and STZ + HFD groups had increased levels of systemic pro-inflammatory IL-6 after 18 weeks, but not STZ + CON mice, suggesting that high-fat feeding is more likely to contribute to systemic inflammation over hyperglycemia. Previous experimental studies of high-fat feeding in C57Bl/6 J mice note significant increases in IL-6 and other pro-inflammatory cytokines after only 8 weeks [
25], while studies involving the use of STZ-treatment in mice vary, noting the presence of IL-1β and TNFα in pancreatic tissue [
26] and IFNy and TNFα systemically [
27]. In accordance with these studies, our results demonstrate that chronic weight gain induced by 18 weeks of high-fat feeding impacts significantly on systemic inflammation.
Inflammation is recognised as a key mechanism of diabetes progression and its multifactorial complications; its likely contribution to diabetes-induced neuropathy is now also clearly emerging [
28]. Indeed, diabetes is regarded as a low-grade, chronic inflammatory disorder [
29,
30]. Systemic inflammation is unmistakably present in diabetic patients [
31,
32] and is considered a contributing mechanism to peripheral disease progression [
33]. CD68 is a protein highly expressed by monocytes and macrophages, as well as microglia. Our observation that a combination of hyperglycemia and high-fat feeding leads to elevated CD11b
+CD68
+ cells is consistent with a diabetes-driven phenotype of increased circulating monocytes and tissue microglia, and suggests that inflammatory signalling is also likely to be implicated in both the systemic and central complications of diabetes, analogous to the known contribution of inflammation to other neurological disorders [
28].
Under steady-state conditions, the number and function of microglia are tightly regulated, comprising the immune-dominant cell type in the brain. In response to an acute immune stimulus, microglia proliferate, increase phagocytosis, and clear the pathogenic insult—a process referred to as priming [
34]. Generally, microglia will undergo apoptosis upon resolving inflammation, but in response to a chronic stimulus, microglia may become notoriously long-lived, contributing to—and thus enhancing—neuro-destructive effects. In our study, mice in the STZ + CON group demonstrated marked increases in the number of hippocampal Iba1
+ microglia after 9 weeks of treatment. This finding is consistent with a previous study that reported hippocampal-specific elevations in Iba1
+ microglia in STZ-treated rats [
35]. Moreover, mice in the STZ + CON group demonstrated a significant increase in activated astrocytes, as indicated by elevated GFAP staining. Intriguingly, these changes were seemingly ameliorated at the 18 weeks endpoint, with no difference found amongst the experimental groups. As neuroinflammatory responses are concomitant with the correct homeostasis functioning of the CNS and are capable of resolving over time, our findings suggest that the STZ-induced changes in microglial numbers and astrocytosis are representative of an acute insult, in this case, hyperglycemia. Chronic hyperglycemia has been associated with cerebral endothelial cell dysfunction and apoptosis, which may provide an entry-point for circulating inflammatory cytokines [
36]. We propose that with more severe weight gain (i.e. obesity) and hyperglycemia, beyond the endpoints examined here, the effects of systemic inflammation and peripheral leukocyte recruitment could establish a form of chronic, non-resolving neuroinflammation in the brain. This would reflect the reasoning by which systemic cytokine increases in obese T2D patients are thought to become evident later in life and support the concept that neurological impairments are primarily seen in aged individuals [
37]. While only conjecture, the notion that hyperglycemia could act as the primary insult in priming microglia prior to the systemic inflammation formed by the secondary insult (obesity and/or ageing) is an interesting prospect that remains to be fully examined.
Having established the presence of systemic inflammation in HFD-fed mice, we characterised whether this was associated with the presence of adherent leukocytes to the cerebral vasculature. The extravasation of leukocytes from the peripheral vasculature into the brain parenchyma is a critical component in both acute and chronic neuroinflammatory disorders. This phenomenon has been predominantly explored in rodent models of multiple sclerosis (MS) and ischaemic stroke in vivo [
38,
39], yet the influences by which this occurs are not fully understood. Here, we observed adherent GR1
+ granulocytes (neutrophils/monocytes) in the surface cerebrovasculature of STZ + CON mice after 18, but not 9 weeks. Interestingly, this adherence was not associated with any increases of IL-6 or other cytokines measured, suggesting that the recruitment of these cells is independent of systemic inflammation and high-fat feeding. Hyperglycemic conditions have been shown to induce endothelial cell dysfunction and promote low-grade inflammation [
40]. Indeed, previous in vitro studies of human brain endothelial cell cultures reveal a high degree of apoptosis under hyperglycemic conditions [
41]. To our knowledge, this is the first study of its kind to demonstrate in vivo granulocyte trafficking along the cerebrovasculature of hyperglycemic mice. Moreover, immunofluorescence of the cortex of STZ + CON mice after 18 weeks demonstrated an increase in the presence of CD45
+ blood-derived leukocytes. Given the lack of changes in inflammatory cells and markers measured using flow cytometry at the level of hemispheric tissue, it is possible that the GR1
+ granulocytes detected with intravital microscopy eventually transmigrated into the brain parenchyma and presented as this CD45
+ population.
Although not specifically examined in this study, the notion of whether high-fat feeding under hyperglycemic conditions seemingly protected against the gliosis we observed at 9 weeks is an interesting prospect. High-fat feeding or the “ketogenic diet” has recently been shown to provide disease-modifying activity in a broad range of neurodegenerative disorders including Alzheimer’s disease [
42]. These observations are supported by studies in animal models and isolated cells that showed ketone bodies, especially β-hydroxybutyrate, to confer neuroprotection against diverse types of cellular injury [
43]. A recent study in young mice highlights that ketogenic diet intervention in mice induces significant increases in cerebral blood flow and P-glycoprotein transports on the blood-brain barrier to facilitate clearance of amyloid-β, a hallmark of Alzheimer’s disease [
44]. Therefore, the absence of hyperglycemia-associated elevation of microglia and astrocytosis in the STZ + HFD group may further suggest a neuroprotective role of high-fat feeding.
Despite the overwhelming evidence linking ageing to a wide array of systemic and neuroinflammatory disorders [
45], it is important to note that the mice used in these experiments were not considered aged. It would be pertinent, therefore, to repeat this investigation in aged mice to examine whether age-associated risk factors could provide the necessary threshold to drive chronic neuroinflammation. In support of this, we observed that both microglial numbers and the degree of astrocytosis in control mice hippocampus increased significantly by four and fivefold, respectively, between 9 and 18 weeks of treatment. Previous studies in aged mice also note increases in CA1-specific gliosis [
46] and that both microglia and astrocytes are more susceptible to inflammatory stimuli [
47]. However, impairments to microglial function have also been observed in aged mice with regard to phagocytosis and motility [
48,
49], which may suggest that this increase in microglial numbers may be a mechanism of compensation for otherwise dysfunctional microglia. Indeed, this age-associated increase in key immune populations of the hippocampus may explain the increased vulnerability to cognitive functions of psychomotor speed and working memory in MetS patients [
50].