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Endocrinology, doi:10.1210/en.2006-0766
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Endocrinology Vol. 147, No. 11 5314-5324
Copyright © 2006 by The Endocrine Society

Reduction in the Number of Astrocytes and Their Projections Is Associated with Increased Synaptic Protein Density in the Hypothalamus of Poorly Controlled Diabetic Rats

Alfonso M. Lechuga-Sancho, Ana I. Arroba, Laura M. Frago, Cristina García-Cáceres, Arancha Delgado-Rubín de Célix, Jesús Argente and Julie A. Chowen

Department of Endocrinology (A.M.L.-S., A.I.A., L.M.F., A.D.-R.d.C., J.A., J.A.C.), Hospital Infantil Universitario Niño Jesús, and Department of Pediatrics (L.M.F., C.G.-C., A.D.-R.d.C., J.A.), Universidad Autónoma de Madrid, 28009 Madrid, Spain

Address all correspondence and requests for reprints to: Dr. Julie A. Chowen, Hospital Infantil Universitario Niño Jesús, Departamento de Endocrinología, Avenida Menéndez Pelayo, 65, 28009 Madrid, Spain. E-mail: jachowen{at}telefonica.net.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Processes under hypothalamic control, such as thermogenesis, feeding behavior, and pituitary hormone secretion, are disrupted in poorly controlled diabetes, but the underlying mechanisms are poorly understood. Because glial cells regulate neurosecretory neurons through modulation of synaptic inputs and function, we investigated the changes in hypothalamic glia in rats with streptozotocin-induced diabetes mellitus. Hypothalamic glial fibrillary acidic protein (GFAP) levels decreased significantly 6 wk after diabetes onset. This was coincident with decreased GFAP immunoreactive surface area, astrocyte number, and the extension of GFAP immunoreactive processes/astrocyte in the arcuate nucleus. Cell death, analyzed by terminal deoxyuridine 5-triphosphate nick-end labeling and ELISA, increased significantly at 4 wk of diabetes. Proliferation, measured by Western blot for proliferating cell nuclear antigen and immunostaining for phosphorylated histone H-3, decreased in the hypothalamus of diabetic rats throughout the study, becoming significantly reduced by 8 wk. Both proliferation and death affected astroctyes because both phosphorylated histone H-3- and terminal deoxyuridine 5-triphosphate nick-end labeling-labeled cells were GFAP positive. Western blot analysis revealed that postsynaptic density protein 95 and the presynaptic proteins synapsin I and synaptotagmin increased significantly at 8 wk of diabetes, suggesting increased hypothalamic synaptic density. Thus, in poorly controlled diabetic rats, there is a decrease in the number of hypothalamic astrocytes that is correlated with modifications in synaptic proteins and possibly synaptic inputs. These morphological changes in the arcuate nucleus could be involved in neurosecretory and metabolic changes seen in diabetic animals.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
ASTROCYTES, THE MOST abundant cell type in the mature brain, have a broad spectrum of functions. In addition to providing metabolic and trophic support to neurons, including a fundamental role in brain glucose metabolism (1), astrocytes are involved in maintenance of the extracellular environment and brain-blood barrier (2, 3, 4), neuronal homeostasis and transmission (5, 6), and regulation of cerebral microcirculation (7). Furthermore, astrocytes enable and regulate neurogenesis and gliogenesis from resident precursor cells (8) and perform neuroprotective functions by mediating estrogen’s protective effects (9, 10, 11) and clearing reactive oxygen species (12) and excess glutamate (13) from the local environment.

Synaptic number and activity can also be regulated by glial cells (5, 6, 9, 14, 15, 16, 17). One mechanism is through modifications in their surface density, allowing astrocytes to ensheath neurons to varying extents and influence synaptic communication (9, 14, 15, 16). In the arcuate nucleus, which coordinates growth, reproductive, adrenal, and metabolic functions, inverse changes occur in the number of synapses and glial ensheathing of neurons during development, throughout the estrous cycle, and with sex steroid treatments (9). Not only are astrocytes involved in synaptic reorganization necessary for pubertal onset, but they also produce trophic factors fundamental for this process (18), as well as transmitters that modulate postsynaptic efficacy in the hypothalamus (17). Furthermore, recent studies demonstrated that leptin and ghrelin induce changes in synaptic number in the arcuate nucleus (19), suggesting that synaptic remodeling may be a physiological mechanism in metabolic signaling. It is possible that hypothalamic astrocytes, which express leptin receptors (20, 21), also play a role in this process.

Although it is clear that hypothalamic astrocytes are active in the physiological control of neuroendocrine systems (1, 9, 15, 17, 18, 19), their role in pathological processes remains poorly understood. Type 1 diabetes mellitus is the most common chronic disease in childhood, with an increasing incidence worldwide (22, 23). Long-term poor glycemic control results in serious metabolic and hormonal disturbances in both diabetic patients (24) and experimental animal models (25, 26, 27). Because these functions are under hypothalamic control, we hypothesized that modifications in astrocytes of this brain area could be involved in the endocrine changes in poorly controlled diabetic animals. In this study our aims were to: 1) determine whether there are changes in the levels of glial fibrillary acidic protein (GFAP) and vimentin in the hypothalamus of diabetic rats; 2) investigate whether the number and/or morphology of astrocytes in the arcuate nucleus is modulated; 3) analyze changes in cell proliferation and death in the hypothalamus and determine whether the turnover of astrocytes is affected; and 4) determine whether the levels of specific synaptic proteins are modified in the hypothalamus of diabetic rats.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
All chemicals and reagents were purchased from Sigma (St. Louis, MO) or Merck (Barcelona, Spain) unless otherwise indicated.

Animals
Young adult male Wistar rats (~250 g) were housed under constant light (12 h) and dark (12 h) conditions and given free access to rat chow and tap water. Diabetes was induced by injection with streptozotocin. This toxin enters pancreatic ß-cells causing DNA alkylation and induction of poly-ADP-ribosylation. Poly-ADP-ribosylation leads to depletion of cellular nicotinamide adenine dinucleotide (oxidized form) and ATP, resulting in the formation of superoxide radicals. Streptozotocin also liberates toxic amounts of nitric oxide that participates in DNA damage. As a result of these processes, ß-cells undergo destruction (28).

Experiment 1
Rats were injected with streptozotocin (70 mg/kg, ip, Sigma) and killed by asphyxiation with CO2 and decapitation 8 wk later between 1000 and 1200 h. Blood samples were taken by tail puncture at the time of the injection and at autopsy to verify the diabetic state (glycemia > 300 mg/dl) by measurement of blood glucose (Glucocard Memory 2; Menarini Diagnostic, Florence, Italy) and insulin. Control rats were injected with vehicle [0.1 M citrate buffer (pH 4.5)]. The brains were removed and rapidly frozen on dry ice and stored at –70 C until processed. These diabetic animals are referred to as diabetic for 8 wk (DB8W).

Experiment 2
Rats were injected with streptozotocin (70 mg/kg, ip) and killed as described above 1, 4, 6, or 8 wk later. Blood samples were collected at the time of injection and at autopsy to verify the diabetic state by measurement of blood glucose and insulin. As in experiment 1, control rats received vehicle only.

The following groups of rats were established: diabetic for 1 wk (DB1W), diabetic for 4 wk (DB4W), diabetic for 6 wk (DB6W), and DB8W. Control rats were killed at each time point, but because no differences were found between control rats at the different time points in any of the studied variables (data not shown), control data were pooled for statistical analysis.

For all studies, rats were treated according to the European Community laws for animal care, and the study was approved by the appropriate institutional ethical committee.

Insulin ELISA
This assay was performed according to the manufacturer’s instructions (Linco Research, Inc., St. Charles, MO). Briefly, a microtiter plate coated with pretitered monoclonal mouse antirat insulin antibody was washed three times with wash buffer [50 mM Tris-buffered saline (TBS) containing Tween 20]. Ten microliters of prediluted standards, quality control samples, and serum samples were added to wells in duplicate. Detection antibody (biotinylated antiinsulin, 80 µl) was then added and the plate sealed and incubated for 2 h while shaking. The wells were then washed and 100 µl of enzyme solution (streptavidin-horseradish peroxidase conjugate) added and incubated for 30 min. After washing, 100 µl of substrate solution (3,3',5,5'-tetramethylbenzidine) were added. After the color developed sufficiently (~15 min), 100 µl of stop solution (0.3 M HCl) were added and the plates read at 450 and 590 nm on an automatic plate reader (Tecan infinite M200; Grödig, Austria). The intra- and interassay coefficient of variation were 1.9 and 7.6, respectively. The limit of sensitivity of this assay is 0.2 ng/ml. For those diabetic samples below the limit of detectability, 0.2 ng/ml was used for statistical analysis.

Protein extraction
Tissue derived from six rats from each group was processed for Western blotting and ELISA. The hypothalami were isolated on ice using the following boundaries: an anterior cut was made at the level of the optic chiasm, a posterior coronal section anterior to the mammillary bodies, two sagittal sections parallel to the lateral ventricles, and a dorsal horizontal section at the level of the anterior commissure. Each hypothalamus was then divided in two symmetric halves. One hemihypothalamus was used for cell death detection ELISA and the other half for Western blotting. Tissue for ELISA was homogenized in lysis buffer provided by the manufacturer of the commercial kit (Roche Diagnostics, Mannheim, Germany). Tissue for Western blotting was homogenized on ice in 300 µl radioimmunoprecipitation assay lysis buffer (50 mM NaH2PO4, 100 mM Na2H2PO4, 0.1% sodium dodecyl sulfate, 0.5% NaCl, 1% Triton X-100) with 5 mg/ml sodium deoxycholate, phenylmethanesulfonylfluoride (1 mM), and a cocktail of EDTA-free protease inhibitors (Roche). The lysates were incubated for 1 h on ice and then centrifuged at 12,000 x g for 5 min at 4 C. The clear supernatant was transferred to a new tube and placed overnight at –70 C. It was centrifuged again at 12,000 x g for 5 min at 4 C and the supernatant transferred to a new tube and stored at –70 C until assayed. Total protein concentration was determined by the method of Bradford (protein assay; Bio-Rad Laboratories, Hercules, CA).

Western blotting
Depending on the specific protein to be detected, 2.5, 30, or 60 µg of protein were resolved on a 12% sodium dodecyl sulfate-polyacrylamide gel under denaturing conditions. The proteins were then electrotransferred to polyvinyl difluoride membranes (Bio-Rad). Membranes were blocked in TBS (20 mM) containing 5% nonfat dried milk or 5% BSA and 0.1% Tween 20 for 2 h. Primary antibodies were added at the following concentrations: antiproliferating cell nuclear antigen (PCNA; 1:1000; Signet Laboratories, Dedham, MA), anti-GFAP (1:3000; Sigma), antivimentin (1:1000; Dako, Glostrup, Denmark), antisynapsin I (1:1000; Calbiochem, San Diego, CA), antisynaptotagmin (1:1000; Calbiochem), or antipostsynaptic density (PSD) 95 (1:1000; Calbiochem) and incubated for 2 h at room temperature under agitation. The membranes were subsequently washed and incubated with the corresponding secondary antibody conjugated with peroxidase (Pierce, Rockford, IL). Bound peroxidase activity was visualized by chemiluminescence (PerkinElmer Life Science, Boston, MA) and quantified by densitometry using the BIO-1D system (Vilber Lournat, Marne la Vallee, France). All results were first normalized to actin levels in each lane (antiactin 1:1000; Santa Cruz Biotechnology, Santa Cruz, CA) and then to control values on each blot. All experiments were performed a minimum of two times.

Cell death detection ELISA
This photometric enzyme immunoassay for the quantitative in vitro determination of cytoplasmic histone-associated DNA fragments (mono- and oligonucleosomes) after induced cell death, was carried out according to the manufacturer’s instructions (cell death detection ELISA; Roche) and with the following modifications. Tissue was homogenized in 300 µl of incubation buffer, placed on ice for 1 h, centrifuged at 1200 x g for 5 min at 4 C and the supernatant collected. The microtiter plates were prepared by adding 100 µl of the coating solution (antihistone antibody) to each well and incubating for 1 h at room temperature (RT). The coating solution was removed and 200 µl of incubation buffer added to each well, covered, and incubated for 30 min at RT. The wells were then rinsed three times and the samples (25 µl sample + 75 µl incubation buffer) added and incubated for 90 min at RT. This dilution was chosen after preliminary assays showed it to be the most adequate for detecting changes. After washing, 100 µl of the conjugate solution (anti-DNA-peroxidase) was added. The wells were covered and incubated at RT for 90 min. After washing, 100 µl of substrate solution was added, mixed, and incubated for 15 min. The resulting color was then measured at 405 nm on an automatic microplate analyzer (Biotek Instruments, Inc., Winooski, VT). Each sample was measured in duplicate in each assay. Background measurements were made and this value subtracted from the mean value of each sample This assay has a detection limit of approximately 50 dead cells/well, and results were normalized to protein levels in each sample and are reported as relative levels of cell death, compared with controls. The inter- and intraassay coefficients of variation were 8.5 and 4.3%, respectively.

Tissue sectioning
The brains were allowed to equilibrate in the cryostat chamber (–17 C), trimmed, and embedded in OCT (Tissue-Tek, Elkhart, IN). Coronal sections were cut at 20 µm throughout the entire arcuate nucleus and thaw mounted onto positively charged slides. Tissue slides were stored at –70 C until immunohistochemistry was performed.

Immunohistochemistry
After fixation for 20 min at RT in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4), sections were washed in phosphate buffer and incubated with 1% hydrogen peroxide in 100% methanol for 20 min to inhibit endogenous peroxidase staining. After this treatment, the sections were washed twice for 20 min in 0.1 M phosphate buffer with 0.1% BSA and 0.1% Triton X-100. This buffer was also used in the subsequent washes and incubations. Sections were then incubated overnight at 4 C in a humid chamber with anti-GFAP antiserum at a dilution of 1:400, antivimentin antiserum at 1:500, or anti-OX42 (a marker of activated microglia) antiserum (Serotec, Oxford, UK) at 1:150. Sections were then washed twice in buffer and incubated for 2 h with the secondary antibody at room temperature (horseradish peroxidase conjugated goat antimouse IgG, diluted 1:250; Pierce Biotechnology) and washed three times with buffer. Peroxidase activity was revealed with 0.01% hydrogen peroxide, using 3,3'-diaminobenzidine as the chromogen. Immunostaining was absent when the primary antibody was omitted. For every experiment, sections from both groups were incubated in parallel. Immunostained sections were observed through an optical microscopy (Zeiss Axioplan, Göttingen, Germany). Images were captured using a digital camera (model CV-S3200; JAI Corp., Yokohama, Japan) and were processed using Image-Pro Plus software (version 5.0 for Windows; Media Cybernetics Inc., Silver Spring, MD). Sections analyzed were cut at ca. –2.3 to –3.3 mm from the Bregma, according to the atlas of Paxinos and Watson (29).

Quantitative evaluation
Seven tissue sections throughout the hypothalamic arcuate nucleus per animal (n = 6 from each group) were analyzed; 14 fields in each section were captured using a x50 objective for evaluation (14 fields/section, seven tissue sections, or 98 fields/arcuate nucleus of each rat). Morphometric techniques unbiased for possible variability in cell size were used to quantify the surface density of GFAP-immunoreactive profiles, the number of GFAP-immunoreactive cells, and the extension of GFAP-immunoreactive processes. The surface density quantification of GFAP-immunoreactive profiles was performed on the upper focal plane of each section by using a stereologic grid, according to the point-counting method of Weibel (30) and as previously described (31), in which the ratio of the surface of immunoreactive profiles to the volume of a given structure (surface density) is calculated by the following formula: surface density = 2I/L, where I is the number of points at which the immunoreactive profiles (cell bodies and cell processes) cross the test grid lines, and L is the test line length in the tissue. The test grid used is based on the c16 grid of Wiebel and has 10 x 10 lines of a total length of 2000 µm (L = 2 mm). The number of GFAP-immunopositive cells was counted according to the optical dissector technique as described by West and Gundersen (32). Sholl’s analysis (33) was performed to assess differences in the extension of glial processes as described by Del Cerro et al. (34). Briefly, an overlay of five concentric rings centered on the soma was positioned over 50 astrocytes per section, and all intersections of the GFAP-immunoreactive processes with the lines of the graticule were counted. The separation between the annuli of the graticule was equivalent to 10 µm. All GFAP-immunoreactive processes were considered for quantification. All morphometric analyses were performed without previous knowledge of the experimental group from which the sections were obtained. The error of repetition of the operator was less than 5%.

Double-fluorescent immunohistochemistry
After fixation in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4), sections were washed in phosphate buffer and then equilibrated in TBS [0.1 M (pH 7.4)] with 0.1% Triton X-100 and 0.1% BSA for 20 min. This same buffer was used in the subsequent washes. After this treatment, the sections were blocked for 90 min at RT in TBS with 1% Triton X-100 and 3% BSA. Sections were then incubated for 48 h at 4 C in a humid chamber with anti-GFAP monoclonal antiserum at a dilution of 1:400, and antiserum for phospho-histone H3 (ser10) (1:100; Upstate, Lake Placid, NY), a specific marker of mitosis, was used to detect proliferating cells. Sections were then washed three times in buffer and incubated for 2 h under dark conditions with the secondary antibodies (Alexa Fluor 633 goat antimouse IgG diluted 1:1000; Molecular Probes, Leiden, The Netherlands; and goat antirabbit IgG biotin conjugated diluted 1:500, Pierce Biotechnology) and washed three times with buffer. Sections were then incubated in the dark during 90 min with streptavidin Alexa Fluor 488 conjugate (Molecular Probes) at a dilution of 1:2000. Immunofluorescence was absent when the primary antibodies were omitted. For every experiment, sections for both groups were incubated in parallel. Immunofluorescence was visualized by using a confocal microscope (model DMIRB; Leica, Madrid, Spain).

Terminal deoxyuridine 5-triphosphate nick-end labeling (TUNEL) plus immunohistochemistry
Cell death detection assays were performed following the manufacturer’s instructions (Roche). Briefly, after fixation in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4), sections were washed three times in buffer and incubated for 30 min with a 0.1% sodium citrate and 0.1% Triton X-100 solution to increase tissue permeability. Slides were again washed three times with buffer and incubated with TUNEL solution for 90 min at 37 C in a humid chamber in the dark. After washing, the slides were incubated with GFAP antibody (1:400) in TBS containing 3% BSA and 1% Triton X-100 and left 48 h at 4 C. The slides were incubated with Alexa Fluor anti-fluorescein-488 and -633 conjugated goat antimouse IgG (Molecular Probes) in blocking buffer both at a dilution of 1:1000. Finally, the slides were again washed three times before mounting in glycerol. Results were visualized with a confocal microscope.

Fluoro-Jade B
After fixation in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4), sections were washed three times in buffer and equilibrated in distilled water. Slides were then transferred to a solution of 0.06% KmNO4 for 17 min to ensure consistent background suppression between sections and washed three times with distilled water. Finally, the slides were incubated with 0.001% Fluoro-Jade B (Histo-Chem Inc., Jefferson, AR) diluted in 0.01% acetic acid for 15 min and then washed with distilled water before mounting with glycerol. Immunofluorescence was visualized using a confocal microscope.

Statistical analysis
All experiments were performed a minimum of two times. When a sample was analyzed more than once in separate assays (Western blots) or repeated measures in the same assay (ELISA), the mean value was used for statistical analysis; hence, n represents the total number of animals used in each group. Testing for normality was performed by the Lilliefors test. In experiments with more than two groups, Bartlett’s test was used to determine that the groups have equal variances. Student’s t test was used for comparison when only two groups were analyzed and a one-way ANOVA followed by a Scheffé F test when more than two groups were analyzed. Statistical significance was chosen as P < 0.05. All results are reported as mean ± SEM.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Blood glucose levels
At the time the animals were killed, morning blood glucose levels were as follows: control, 70.0 ± 1.9 mg/dl; DB1W, 520.5 ± 30.9 mg/dl; DB4W, 524.6 ± 37.2 mg/dl; DB6W, 541.1 ± 21.4 mg/dl; and DB8W, 478.6 ± 41.6 mg/dl. Glucose levels remained elevated throughout the study (ANOVA, P < 0.0001), and no significant differences were found between the diabetic groups.

Serum insulin levels
At the time the animals were killed, serum insulin levels were: control, 1.75 ± 0.18 ng/ml; DB1W, 0.23 ± 0.03 ng/ml; DB4W, 0.35 ± 0.13 ng/ml; DB6W, 0.28 ± 0.06 ng/ml; and DB8W, 0.33 ± 0.05 ng/ml. Insulin levels remained decreased throughout the study (ANOVA, P < 0.0001), and no significant differences were found between the diabetic groups.

Effect of 8 wk of diabetes on glial cell markers
There was a marked reduction in GFAP protein in the hypothalamus of diabetic rats after 8 wk of evolution, as shown by the significant 40% reduction in Western blot band density, compared with controls (Fig 1AGo; P < 0.01).


Figure 1
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  		FIG. 1. A, Relative mean signal levels of GFAP in the hypothalamus of diabetic and control rats as determined by Western blot analysis. Data are represented as mean ± SE and are normalized to control results for each experiment (n = 6 in each group). A representative Western blot is shown. B, Representative photographs of sections through the arcuate nucleus of a control (left) and a diabetic rat (right), immunostained for GFAP. Scale bar, 20 µm. C, Relative mean levels of GFAP-immunoreactive surface area in the arcuate nucleus of control and diabetic rats. Data are represented as mean ± SE (n = 6 in each group). D, The number of GFAP-positive cells per cubic millimeter in the arcuate nucleus of control and diabetic rats. Data are represented as mean ± SE (n = 6 in each group). E, Results from the Sholl analysis expressed as the number of intersections (normalized to control) of GFAP-immunoreactive processes with the lines of the test grid. Data are represented as mean ± SE (n = 6 in each group).

 
The inspection of histological sections of the hypothalamus immunostained for GFAP revealed an obvious decrease in immunostaining in DB8W animals: GFAP-immunoreactive cells were more abundant and showed longer processes in the arcuate nucleus of control animals (Fig. 1BGo). This qualitative impression was confirmed by the morphometric analysis. After 8 wk of diabetes, rats had a significant decrease in GFAP-immunoreactive surface density in the arcuate nucleus, compared with controls (Fig. 1CGo; P < 0.01) and a 31.5% decrease in the number of GFAP-labeled cells in the arcuate nucleus (Fig. 1DGo; P < 0.01). In addition, the number of intersections of GFAP-immunoreactive processes with the lines of the grid in the Sholl’s analysis was significantly reduced in diabetic animals (Fig. 1EGo; P < 0.01), indicating a reduction in the extension, branching, and/or number of GFAP-immunoreactive processes per cell.

No significant variation in vimentin protein levels was found by Western blotting in the hypothalamus of diabetic animals with respect to controls (control: 100 ± 32.9 vs. diabetes: 110.4 ± 24.4). In addition, no difference in immunolabeling for anti-OX42, a macrophage marker, was found between the arcuate nucleus of control and diabetic rats (data not shown).

Cell turnover in the hypothalamus of diabetic rats after 8 wk of evolution
A 60% increase (P < 0.05) in cell death was observed in DB8W rats, compared with controls, as assessed by ELISA (Fig. 2AGo; control: 100 ± 8 vs. DB8W: 160 ± 16).


Figure 2
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  		FIG. 2. A, Relative mean cell death levels in the hypothalamus of diabetic and control animals, as measured by ELISA to detect histone-associated DNA fragments and expressed as arbitrary levels of OD. Data are represented as mean ± SE and are normalized to control results for each experiment (n = 6 in each group). B, TUNEL-positive cells that coexpressed GFAP were found in the arcuate nucleus (ArcN) and median eminence (ME) of diabetic rats. White arrows indicate TUNEL and GFAP-positive cells.

 
TUNEL-labeled cells were sparse in the arcuate nucleus of diabetic rats, with approximately one to five labeled cells found per section. The majority of these cells could be clearly identified as GFAP positive (Fig. 2BGo). No TUNEL-labeled cells were observed in the arcuate nucleus of control rats.

No specific Fluoro-Jade B labeling was found in the hypothalamus of either control or diabetic rats, although specific labeling was found scattered in other brain areas, such as the cerebellum, in the diabetic group (data not shown).

In the diabetic rat hypothalamus, the Western blot signal for PCNA levels was reduced approximately 55% with respect to controls (Fig. 3AGo; P < 0.001).


Figure 3
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  		FIG. 3. A, Relative mean signal levels of PCNA in the hypothalamus of diabetic rats, compared with controls, as determined by Western blotting. Data are represented as mean ± SE and are normalized to control results for each experiment (n = 6 in each group). A representative Western blot is shown. B, Double-fluorescent immunohistochemistry for phosphorylated histone H3 (green) and GFAP (red) in the hypothalamic arcuate nucleus of a control rat. White arrows indicate cells immunoreactive for phosphorylated-histone H3 and GFAP.

 
Fewer H3-immunoreactive cells were found in the arcuate nucleus of diabetic rats than controls. These cells were not grouped and did not show any specific distribution pattern but rather were disperse throughout the parenchyma. Double immunohistochemistry revealed that all H3-immunoreactive cells identified, in both diabetic and control rats, were also GFAP positive (Fig. 3BGo).

Synaptic proteins in the hypothalamus of diabetic rats after 8 wk of evolution
Significantly higher levels of all three synaptic proteins measured, synapsin I (Fig. 4AGo; P < 0.05), synaptotagmin (Fig. 4BGo; P < 0.001), and PSD-95 (Fig. 4CGo; P < 0.05), were observed in the hypothalamus of diabetic animals, compared with controls.


Figure 4
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  		FIG. 4. Relative mean signal levels of synapsin I (A), synaptotagmin (B), and PSD-95 (C) in the hypothalamus of control and diabetic rats. Data are represented as mean ± SE and are normalized to control results for each experiment (n = 5). Representative Western blots are shown above each bar graph.

 
Time-dependent changes in GFAP levels and cell turnover
To determine whether the time of evolution of diabetes had an effect on hypothalamic cell turnover, rats were killed 1, 4, 6, and 8 wk after diabetes onset. An increase in cell death was not detected until 4 wk after the onset of diabetes (Fig. 5AGo; P < 0.01). Mean levels of PCNA decreased gradually throughout the study, becoming significantly lower than controls at 8 wk of diabetes (Fig. 5BGo; P < 0.05). Mean GFAP protein levels were significantly decreased only after 6 wk of diabetes (Fig. 5CGo; P < 0.01). Although mean levels of vimentin tended to increase during the early stages of diabetes, this did not reach statistical significance at any time point (control: 100 ± 21.4; DB1W: 149.2 ± 36.2; DB4W: 176.4 ± 20.8; DB6W: 162.3 ± 15.6; DB8W: 129.1 ± 26.7).


Figure 5
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  		FIG. 5. Relative mean signal levels of cell death (A), measured by ELISA, and PCNA (B) and GFAP (C), measured by Western blot, in the hypothalamus of diabetic and control rats. *, ANOVA P < 0.05, compared with control; **, ANOVA P < 0.01, compared with control. C, Control (n = 4 for each diabetic group and n = 8 for controls). Representative Western blots are shown above each bar graph.

 
No change in any of the synaptic proteins measured were found previous to 8 wk of diabetes (data not shown).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In the adult brain, glial cells widely outnumber neurons (35) and commonly react to physiological and pathological states by cellular swelling, hypertrophy-hyperplasia (astrogliosis), proliferation (astrocytosis) (36), and expression of markers of immature glia, such as vimentin (37). These changes in astrocyte reactivity are observed in a variety of pathophysiological situations including ischemia (38), acute injury (39), kainic acid treatment (40), and neurodegenerative diseases as well as in normal aging (41); whereas neurons tend to degenerate and show a relative inability to grow and proliferate (42). However, GFAP protein and immunoreactive density decrease in the cerebellum (43, 44), cerebral cortex, and hippocampus (43) of diabetic animals. Likewise, here we found no evidence of astrocyte activation in the hypothalamus but instead a decrease in GFAP levels by 6 wk of diabetes evolution, with GFAP immunoreactive density, projections, and cell number all being significantly decreased at 8 wk of diabetes. In contrast to the study of Coleman et al. (43), others have reported increased astrocyte reactivity in the hippocampus and cortex of diabetic rats (45) as well as the hippocampus of diabetic mice (46). These discrepancies could be due to differences in the time of diabetes evolution because we found GFAP levels in the cerebellum to increase at 1 wk and decrease significantly thereafter (44). However, here we show that GFAP levels in the hypothalamus were not increased at any time. It is well known that astrocytes behave in a regionally specific fashion; thus, this differential response could be explained by the specific functions that astrocytes play in the hypothalamus (9).

Although GFAP is a hallmark of astrocyte maturation and reactivity, it is also expressed by tanycytes, a specialized glial cell lining the third ventricle (47). However, the observed decrease in GFAP protein levels is most likely due to astrocytes rather than tanycytes because the number of GFAP-immunoreactive astrocytes and extension of GFAP-immunoreactive astrocytic processes were reduced. Moreover, vimentin, expressed by tanycytes or immature astrocytes, did not decrease in the hypothalamus of diabetic animals but tended to increase, although this change was not significant.

The decrease in the number of hypothalamic astrocytes in diabetic rats is due to both increased death and decreased proliferation. Hypothalamic PCNA levels, a marker of cell proliferation, decreased throughout the study in parallel to the decrease in GFAP levels, suggesting the involvement of astrocytes. Furthermore, the number of cells that coexpressed phosphorylated histone H3, a marker of mitosis, and GFAP in the arcuate nucleus was decreased in diabetic rats. Increased cell death was detected by both ELISA and TUNEL. In the arcuate nucleus, death was selective for astrocytes because TUNEL labeling corresponds to GFAP-positive cells. Regional differences in diabetes-induced cell death may exist because vasopressin neurons in the supraoptic nucleus undergo increased apoptosis (48). However, it is also possible that neuronal death is a delayed phenomenon because apoptosis of vasopressin neurons was reported to occur after 6 months of diabetes but not after 6 wk (48). In support of this, we found no Fluoro-Jade B labeling, a marker of neuronal degeneration (49), in the hypothalamus after 8 wk of diabetes.

The change in astrocyte number and morphology could be the consequence of unviable extracellular conditions such as hyperosmolarity, low nutrient availability, or increased oxidative stress. The lack of insulin could be involved because insulin influences astrocyte morphology, differentiation, and GFAP expression (50). IGF-I is also important for astrocyte differentiation and morphological changes (51), and both its circulating levels (52) and local production in the hypothalamus (Ref. 53 and our unpublished observations) are decreased in poorly controlled diabetic animals. Although streptozotocin is reported to be specifically toxic for pancreatic ß-cells, it could conceivably induce cell death in other tissues. However, streptozotocin has a short half-life in the circulation, and pancreatic ß-cells begin to die within hours (54, 55), suggesting that the 4-wk delay in hypothalamic cell death is most likely not a direct effect of streptozotocin. It is also conceivable that these astrocytic changes are not only a secondary complication, but may actively participate in the neuroendocrine response to the diabetic state.

Modifications in some hypothalamic neuropeptides, such as GHRH and somatostatin, are reported to occur within days after streptozotocin injection (56), indicating that the delayed changes in astrocytes and synaptic proteins reported here are not involved in these processes. However, hypothalamic {alpha}MSH levels do not change until after 4 wk of diabetes (57). Likewise, there are both rapid and delayed modifications in systems controlled by the hypothalamus, such as the pituitary and circulating hormones (58, 59, 60). Thus, the observed modifications in hypothalamic astrocytes and synaptic inputs could be related to the delayed endocrine changes seen in diabetic animals, although whether they are causative or are a result of these processes remains to be determined.

Glial coverage of neuronal membranes decreases when astrocytic processes retract, resulting in synaptic contact remodeling (9, 16). This process occurs in physiological phenomenon such as during the estrous cycle, parturition, lactation and in water balance (9, 16). Here the decrease in hypothalamic GFAP levels, due to reduced astrocyte projections per astrocyte and the number of astrocytes, is followed by increased synapsin I levels. Synapsin I is a vesicle-related protein present at presynaptic terminals, and changes in its expression are directly related to changes in synapse number (61). Likewise, there was a significant increase in synaptotagmin, a calcium sensor controlling synaptic vesicle exocytosis (62) that is also present at the presynaptic terminal and in PSD-95, a postsynaptic membrane-associated guanylate kinase (63). Hence, this inverse relationship between the extension of glial processes and synaptic protein levels suggests that poorly controlled diabetes mellitus may induce synaptic remodeling in the arcuate nucleus. These morphological changes could be involved in adapting hypothalamic functions to the present pathophysiological situation, as occurs under physiological conditions (9, 16). However, synaptic remodeling in diabetic animals is not restricted to the arcuate nucleus or hypothalamus because this phenomenon also occurs in the hippocampus (64, 65).

Synaptic inputs in the arcuate nucleus are modified by leptin (19), and astrocytes in this brain area express leptin receptors (20, 21). Furthermore, leptin, synthesized by adipocytes and involved in metabolic control, modulates GFAP levels and is important for normal development of neurons and glia (66). As circulating leptin levels are decreased in streptozotocin-induced diabetic rats (67), it could be involved in the changes observed here. Ghrelin, which has opposite effects to leptin on synaptic inputs in the arcuate nucleus (19), may also be involved as circulating levels of this hormone are increased in streptozotocin-induced diabetic rats (68). However, changes in circulating leptin and ghrelin levels occur relatively rapidly (67, 68), as do the reported synaptic changes in response to these hormones (19). Likewise, glucose levels were elevated and insulin levels reduced at the onset and remained so throughout the study, whereas the changes in astrocytes and synaptic proteins were delayed. Other delayed changes in the diabetic brain have been reported (57, 69, 70, 71), and these modifications could be involved in a gradual resetting of the brain to the prolonged situation of diabetes (72). The effect of these metabolic factors, as well as others, on hypothalamic glial and synaptic changes, and how this correlates with both long and short-term metabolic control, deserves further investigation.

A decrease in the number of astrocytes, as well as the extension of astrocytic processes, could alter glucose sensing in the arcuate nucleus (1, 73, 74, 75), contributing to the impaired biochemical sensing of fuel availability that has been proposed as a basic underpinning for defects in the regulation of food intake, ß-cell function, and liver glucose homeostasis (76). Indeed, hypothalamic glucose sensing is fundamental to systemic metabolic control, and astrocytes are reported to play an active role in this phenomenon (1, 73, 74, 75). Furthermore, brain glucose use is reported to be decreased in poorly controlled diabetes (75), and this could also be related to the decrease in astrocytic function because these glial cells are involved in glucose transport and metabolism (1, 75). Whether these glial changes are beneficial or harmful in situations of prolonged malnutrition remains to be determined.

Although it is well accepted that astrocytes are required for the maintenance of normal neuroendocrine function through a variety of mechanisms, there is little information regarding their role in adaptation to chronic metabolic changes such as in diabetes mellitus. Here we demonstrate that poorly controlled diabetes induces a decrease in the number of astrocytes and changes in their morphology in hypothalamic areas involved in neurosecretory control. Levels of synaptic proteins are also modulated, suggesting that synaptic remodeling occurs. Because astrocytes play an important role in neuronal metabolism, function, and survival, modifications in this cell type may have important implications in the pathophysiology of diabetes mellitus. Whether these alterations in hypothalamic astrocytes and synaptic protein densities are related and result in an adaptive response to the diabetes-induced malnutrition requires further investigation. Furthermore, the underlying cause of these changes and whether they are inhibited by correct glycemic control remains to be determined.


   Acknowledgments
 
The authors thank Dr. Luis Miguel García-Segura for the critical review of this manuscript.


   Footnotes
 
This work was funded by grants from Fondo de Investigación Sanitaria (PI04/0817 and PI051268), Ministerio de Educación (SAF2002-03324), Fundación de Investigación Médica Mutua Madrileña, and Fundación de Endocrinología y Nutrición. A.M.L.-S. is supported by a postdoctoral fellowship from Fondo de Investigación Sanitaria. J.A.C. is supported by the biomedical investigation program of the Consejería de Sanidad y Consumo de la Comunidad de Madrid.

Disclosure statement: A.M.L.-S., A.I.A., L.M.F., C.G.-C., A.D.-R.d.C., and J.A.C. have nothing to declare. J.A. has previously received lecture fees from Serrono Spain and Ferring Spain.

First Published Online July 27, 2006

Abbreviations: DB1W, Diabetic for 1 wk; DB4W, diabetic for 4 wk; DB6W, diabetic for 6 wk; DB8W, diabetic for 8 wk; GFAP, glial fibrillary acidic protein; PCNA, proliferating cell nuclear antigen; PSD, postsynaptic density; RT, room temperature; TBS, Tris-buffered saline; TUNEL, terminal deoxyuridine 5-triphosphate nick-end labeling.

Received June 7, 2006.

Accepted for publication July 18, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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