All animal experiments were approved by the Wayne State University Institutional Review Board and were conducted following the National Institute of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 80-23, revised 1996). The H-Tx rats (H-Tx/hcj strain) originated from Dr. Hazel Jones, University of Florida. Animals were maintained on 12 h dark-light cycles in a controlled environment with free to access food and water at all times.

Brains from hydrocephalic H-Tx rats were examined utilizing immunohistochemistry and immunoblotting at the following postnatal ages: 5d, 12d, 21d and 36d, and compared to non-hydrocephalic age-matched control H-Tx rats (n = 5 for each group). The effect of CSF drainage was examined by inserting a shunt into severely hydrocephalic animals at 15d, and examining them after 6d (21d of age) or after 21d (36d of age). These shunted animals were compared to age-matched un-treated hydrocephalic H-Tx rats and their respective control littermates (n = 5 for each group).

In this study, the H-Tx rat model of congenital hydrocephalus was utilized. Although this is not an exact replica of the human condition, we believe this model appropriately mimics congenital human hydrocephalus. H-Tx animals have a slowly progressive form of hydrocephalus which is primarily caused by an alteration of the cerebral aqueduct occurring between embryonic day 18 and post-natal day 5 2930313233343536. Because the cranial sutures of these young animals are not yet fully fused, the skull is expandable and accommodates the rising ventricular volume, thus allowing for overt visual identification of the hydrocephalic individuals.

To examine the effects of shunting, Teflon-coated catheters coupled to Heyer-Schulte low-pressure neonatal valves (Heyer-Schulte- Integra, New Jersey, USA) were inserted into the lateral ventricle of hydrocephalic H-Tx rats at 15d. This age approximates the stage when hydrocephalus in these animals advances from a moderate to a severe state, and the developing rat cerebral cortex is close to that of a newborn human 37. Animals were given a pre-operative oral dose of Cephalexin antibiotic (50 mg/kg) and the same dose was given twice daily for 5d postoperatively to help prevent infection. Animals were anesthetized with 2% halothane and prepared for sterile surgery, and all procedures were performed under aseptic conditions. A small incision was made over the skull and over the lower back of the animal. A small burr hole was created in the skull 1 mm lateral to the midline and 2 mm posterior to Bregma. After piercing the dura mater, the tip of the shunt catheter was advanced without use of a stylet into the lateral ventricle, and fixed to the skull using one to two drops of ethyl cyanoacrylate (Krazy^©) glue. The distal end of the catheter was left lying in the subcutaneous tissue above the distal lumbar vertebrae of the spine near the caudal vertebrae. This allowed for movement of the distal end of the catheter during growth, and prevented the shunt catheter from being pulled out of the burr hole. The patency of the shunt was tested by withdrawing a few drops of CSF through the distal end of the catheter. Skin incisions were closed using tissue staples, and an ear tag was placed for identification purposes. The animal was removed from anesthesia, allowed to recover on a heated surface, and returned to the cage with its mother and littermates. In the days following shunt insertion, any animals displaying neurologic dysfunction or lethargy, suggesting a shunt failure or infection, were sacrificed and not utilized for this study.

Animals used for histology and immunohistochemistry were deeply anesthetized with (1–4 ml depending on age) 4% chloral hydrate i.p. and perfused transcardially with saline followed by 4% paraformaldehyde fixative in 0.1 M PBS pH 7.4, for optimal preservation of brain morphology. Following perfusion, the brain was removed from the skull, sectioned coronally into thirds (frontal, parietal and occipital sections) and placed into paraformaldehyde. The hydrocephalic animals were classified as having severe hydrocephalus through observation of the dilation of the ventricles and the thickness of the cortex, and only those animals with clearly defined hydrocephalus were included in this study. The control littermates were also verified visually by inspection of the ventricles. Following 2–4 h of post-fixation, the brain was removed from the fixative, rinsed and stored in 0.1 M PBS until paraffin embedding.

Separate groups of rats were used for Western blotting analyses and the tissue was prepared using a fresh frozen technique. Animals were deeply anesthetized by inhalation of a halothane from a soaked cotton swab, decapitated, and the brain rapidly removed and dissected on ice. Samples from the cerebral cortex were snap-frozen in liquid nitrogen and stored at -80°C. Brain tissue was homogenized in buffer containing: 50 mM NaF, 50 mM HEPES (4-(2-Hydroxyethyl) piperazine-1-ethanesulfonic acid) (pH 7.5), 2.5 mM 4-dithiotreitol-d₁₀ (DTT), 120 mM KCl, 4 mM MgOAc, 1 mM ethylenediaminetetraacetic acid (EDTA), 20 mM glycerophosphate, 10 μg/ml pepstatin A, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 1 mM orthovanidate, 250 nM okadaic acid and 1 mM phenylmethanesulfonyl fluoride (PMSF). Protein concentration was determined by the Lowry method. Samples were added to 2X loading buffer containing: 0.125 M Tris-HCl pH 6.8, 4% sodium dodecyl sulfate (SDS), 20% glycerol, and 10% β-mercaptoethanol and boiled for 90 sec. Equal protein amounts were loaded into lanes for electrophoresis procedures.

Aliquots containing 50 μg protein from brain homogenates were electrophoresed in 10% SDS-polyacrylamide gels. To allow for comparison between membranes, a sample from one individual animal was loaded onto every gel, and all samples on the membrane resulting from that gel were standardized using the consistently loaded sample. Beta actin expression was also used frequently to ensure standard loading of gels.

Proteins were then electrophoretically transferred onto a nitrocellulose membrane, which was fixed in 10% acetic acid and 25% isopropanol for 15 min to ensure protein immobilization. The membrane was then placed at 4°C for 15 min, and rinsed 10 times in dH₂O followed by one rinse in 50 mM Tris, pH 7.4, 200 mM NaCl. All subsequent procedures were performed at room temperature.

For detection of GFAP, non-specific antibody binding was pre-blocked in 5% low-fat dried milk dissolved in TTBS (100 mM Tris, pH 7.4, 0.9% NaCl, 0.1% v/v Tween-20) for 45 min with gentle agitation. After rinsing with TTBS, the membranes were incubated with anti-GFAP antibody 1:1,000 in TTBS (DAKO, USA) for 60 min and gently agitated at room temperature.

After incubation in primary antibody, membranes were washed 3X in TTBS, and incubated for 30 min in TTBS containing anti-rabbit IgG-horseradish peroxidase conjugate at a 1:10,000 dilution. The membranes were then washed 3X in TTBS, incubated in Enhanced Chemiluminsence (ECL – Amersham, USA) kit detection reagents for 90 sec, drained, covered with plastic wrap and contact-exposed to film. After film development, the bands were quantified by densitometry (Intelligent Quantifier, Bio Image Inc. version 3.0.0, Michigan, USA).

All layers of the occipital cortex and the parietal cortex were examined from perfusion-fixed coronal sections with immunohistochemical techniques using antibodies specific for the astrocytic protein, GFAP, and microglial protein, ILB4. Additionally, the area surrounding the cerebral aqueduct was examined by the same immunohistochemical techniques. Labeled cells were identified using brightfield or fluorescent techniques using a Leica DMRE microscope (Leica Microsystem Products, New Jersey, USA).

Brains were embedded in paraffin and sectioned at 10 μm using standard histology procedures. Before staining, the slides were de-paraffinized and rehydrated, and subjected to antigen retrieval by placing mounted slides into 10 mM citrate buffer (pH 6.0) preheated to 90–100°C for 20 min, cooled for 20 min in buffer after removal from heat, and then washed in 0.1 M PBS.

For GFAP immunostaining, hydrated sections were incubated in 3% H₂O₂ for 10 min. The sections were then washed in dH₂O and incubated in 5% goat serum diluted in 0.1 M PBS for 20 min and then in anti-GFAP antibody, 1:300 (DAKO, USA) diluted in an antibody diluent reagent solution (Zymed, California, USA). Primary antibody was then removed with a modified phosphate buffered saline (MPBS) containing 50 mM K₂HPO₄, 10 mM NaH₂PO₄, and 10 mM NaCl, and an anti-rabbit secondary antibody (1:200, Vector Laboratories, USA) was applied for 10 min. The secondary antibody was rinsed off with MPBS and an avidin-biotin complex (ABC kit, Vector Laboratories, USA) was applied for 30 min. Color was developed for 2–10 min using an un-enhanced DAB kit (Vector Laboratories, USA). Slides were dehydrated by passing them through increasing concentrations of alcohol and three changes (5 min each) of xylene, cover-slipped using Permount (Fisher Scientific, USA), a non-aqueous mounting media, and allowed to dry overnight. For fluorescence microscopy, the same primary GFAP antibody was used as described above, however, following primary incubation, tissue was incubated in a TRITC-labeled anti-rabbit secondary antibody (1:400, Sigma Aldrich, USA) used for 2 h at room temperature. Excess antibody was rinsed off and cover slips applied using Aquamount (Fisher Scientific, USA) or a DAPI fluorescent stain-impregnated hard-set mounting medium (Vector Laboratories, USA) to label nuclei.

ILB4 from Griffonia simplicifolia is a marker for brain microglia and peri-vascular cells 38. Following rehydration, sections underwent antigen retrieval as described above, and were incubated in anti-ILB4 antibody 0.2 mg/ml (HRP labeled, Sigma Aldrich, USA) for 2 h at room temperature. The slides were washed with deionized water, and color developed with a nickel enhanced 3,3'-diaminobenzidine (DAB) kit for up to 10 min (Vector Laboratories, USA). The DAB solution was rinsed off, and the slides were dehydrated and cover slipped as above. For fluorescent staining, FITC-conjugated ILB4 (Sigma Aldrich, USA) primary antibody was used at 0.2 mg/ml and incubated at room temperature for 2 h. The slides were then rinsed and prepared for examination as described above.

Western blots were quantified by scanning densitometry and the data was analyzed using a Mann-Whitney U test for two groups, or a Kruskal-Wallis test for 3 groups, followed by individual Mann-Whitney U tests with a Bonferroni correction for between group comparison. For the quantitative assessment of astrocyte staining, the number of positively stained GFAP cells was graded on a four-point scale. A score of 1 indicated GFAP-labeled cells at a relatively low quantity, 2 and 3 indicated medium and moderate amounts, and 4 indicated that GFAP labeled cells were present in high abundance. The data was then analyzed using the Kruskal-Wallis and Mann-Whitney U tests followed by a Bonferroni correction as above.

ILB4 positive cells were analyzed from their morphologic appearance, which is a commonly accepted method to judge relative reactivity 213940, rather than on the overall numeric density. Developmentally, microglia alter their morphology dramatically during the early postnatal period 41. Microglia change from amoeboid-like cells at postnatal day 0 to completely ramified microglia over the first three weeks of post-natal development as seen in the figure adapted from Orlowski et al (Fig. 1) 39414243. Due to the dramatic changes in normal cellular morphology observed during the first few weeks of life, each age group was treated individually, and was not directly compared to the other age groups. Therefore, for morphologic analysis of microglia in this study, hydrocephalic animals were compared only to their age-matched control and age-matched shunted counterparts.

At sacrifice, all untreated hydrocephalic animals were easily identified by the presence of large, domed heads. Upon removal of the brain, these animals had a noticeable expansion of the lateral ventricles and thinning of the cortex and were classified as having severe hydrocephalus by visual inspection. This dramatic increase in ventricular volume and the thinning of the cortex is evident in low-power images (Figs. 2 and 3). All animals included in this study had similarly thinned cortices.

At the time of sacrifice, both the short-term and long-term shunted animals had decreases in apparent ventricular size when compared to untreated hydrocephalic rats of the same age. Additionally, the cortex of the shunted groups was thicker when compared to untreated hydrocephalic animals, although the thickness did not appear to return to that of control levels. This reduction in ventricular volume and the increase in cortical thickness can be seen in low power images, which depict the brain of a shunted animal at 36d (Figs. 2 and 3).

In the parietal cortex, GFAP levels in 5d hydrocephalic animals as measured by scanning densitometry of Western blots, were significantly increased by 3.68X compared to control animals (p < 0.01) (Fig. 4). In 12d hydrocephalic animals, the GFAP levels were also increased over controls by 1.69X (p < 0.05). The same trend in GFAP continued in the 21d hydrocephalic animals (2.77X; p < 0.01), and also in the 36d animals (2.69X; p < 0.01).

In the occipital cortex, there was a similar trend toward raised levels of GFAP in hydrocephalic animals compared to age-matched controls, although at 12d with a 1.81X increase the difference was not significant (p > 0.05). At the other ages, GFAP expression was increased over age-matched controls by 2.46X at 5d (p < 0.05,) by 5.26X at 21d (p < 0.01), and by 5.23X at 36d (p < 0.01) (Fig. 5).

GFAP levels in the parietal cortex exhibited significant alterations between control, hydrocephalic and shunted rats at 21d (p < 0.01). Shunt-treated animals had an 18% decrease in GFAP when compared to the untreated hydrocephalic animals, but this failed to reach significance (Fig. 4). At 36d, the shunted animals had a 23% decrease in GFAP levels when compared to the untreated 36d hydrocephalic animals (p < 0.05). An example of a representative Western blot is shown in Fig. 4.

The effect of shunting in the occipital cortex was more dramatic in reducing the levels of GFAP. Shunting hydrocephalic animals at 15d and allowing them to recover until 21d significantly reduced the amount of GFAP expression by 77% when compared to the untreated hydrocephalic animals (p < 0.01) (Fig. 5). This was similar to the expression level in control rats at 21d. Shunting at 15d and allowing a three-week post-shunt survival period until 36d, significantly reduced GFAP expression by 48.2% (p < 0.05).

Qualitative histologic examination of tissue labeled with GFAP revealed astrocytes present throughout all layers of the cortex with relative increases in the number of positively stained cells found in untreated hydrocephalic animals, regardless of age (Fig. 2). Following quantitative grading based on the relative abundance of astrocytes in sections, the data was then compared and graphed (Fig. 6).

At 5d, upon visual examination, the control and hydrocephalic animals showed small numbers of GFAP labeled astrocytes, and these were found throughout all cortical layers (Fig. 2). Upon assigning a grade, the 5d hydrocephalic animals had a significant increase in the number of astrocytes present when compared to 5d non-hydrocephalic animals (2.3x, p < 0.05) (Fig. 6). When examining the periventricular white matter (Fig. 7A,B) or in the area surrounding the cerebral aqueduct (Fig. 8A,B), there were no dramatic differences in the appearance of astrocytes at 5 days of age.

Qualitative histologic examination of the 12d animals revealed astrocytes throughout all cortical layers (Fig. 2). When sections were graded for the relative abundance of GFAP positively stained cells, there was a 1.9 × (p < 0.05) increase in cellular amount in the 12d hydrocephalic animals when compared to the 12d control animals. Examination of the tissue sections, in the periventricular white matter (Fig. 7C,D) and the cerebral aqueduct (Fig. 8C,D) showed no obvious differences between hydrocephalic and control rats.

In the 21d hydrocephalic animals, the group with the most severe form of hydrocephalus, GFAP stained cells were distributed evenly throughout all cortical layers whereas in the 21d control animals, they were within cortical layers 1 and 2 (Fig. 2). When tissue sections were graded, there was a significant 1.97x increase in the relative number of astrocytes in the hydrocephalic animals when compared to age matched controls (p < 0.05) (Fig. 6). Additionally, a large concentration of GFAP positive cells was found to be present in the peri-ventricular white matter (Fig. 7E,F), and in the cerebral aqueduct (Fig. 8E,F).

At 36d, the hydrocephalic animals also had an increase in GFAP labeled astrocytes throughout all cortical layers when compared to the 36d control animals (Fig. 2). After grading, a 1.46 × increase in astrocytes was found in the hydrocephalic animals when compared to their control counter parts (p < 0.05). Additionally, GFAP labeled cells were more abundant in both the periventricular white matter (Fig. 7H,I) and the area surrounding the cerebral aqueduct (Fig. 8H,I).

Following shunting, the distribution of astrocytes was altered in both the 21d and 36d animals. In the 21d shunted animals, a marginal decrease in GFAP labeled astrocytes was observed throughout all layers of the brain tissue (Fig. 2). Furthermore, a non-significant decrease of 18.6% was measured after grading (Fig. 6). In the periventricular white matter (Fig. 7G) and the area surrounding the cerebral aqueduct (Fig. 8G), astrocyte density was slightly reduced in the shunted animals when compared to their hydrocephalic counterparts.

In the 36d shunted animals, there was a noticeable decrease in relative abundance of GFAP labeled astrocytes in the cortical layers (Fig. 2). After grading, there was a significant 31.5% decrease (p < 0.05) in the relative number of astrocytes when compared to age-matched untreated hydrocephalic animals, and the grade was close to that of the control rats. Examination of the periventricular white matter (Fig. 7J) and the area surrounding the cerebral aqueduct (Fig. 8J), showed a more dramatic reduction in the numbers of labeled astrocytes in the shunted group when compared to the untreated 36d hydrocephalic littermates.

Although the microglial cells were not quantified, the number of ILB4 positive cells in the cortical sections did not increase or decrease dramatically with developmental age or the severity of hydrocephalus. However, the cellular morphology of the microglia was noticeably different (Fig. 3). This change toward activated microglia has been well documented as a response to injury 394044. In activated microglia, processes become shorter and thicker, while cell bodies and cellular processes stain more intensely. Hydrocephalic animals in all age groups appeared to have at least some activated microglia with a thicker cell body giving off shorter and thicker branches and cellular processes.

Normal differentiating microglia also change morphology. The typical maturation process of a microglial cell is demonstrated beautifully by Orlowski et al 42, and was used for maturation comparison in the current study (Fig. 1). In his paper, Orlowski described immature microglia as having an amoeboid shape with little cellular differentiation. As the microglial cell matured, cellular processes became more pronounced, and by postnatal day 30, the cell was fully ramified with extensively branched and long, fine processes. ILB4-immunostained cells in both control and hydrocephalic animals at the different time-points clearly exhibited the changes in microglial morphology with maturation (Fig. 3 left column). In 5d control animals, the majority of microglia had an amoeboid shape, with very few cells having the beginning stages of cellular processes (Fig. 3, top row). The hydrocephalic 5d animals had microglial cells that looked similar to the postnatal day 0 animals from Orlowski's figure (Fig. 1 top) and there was a small increase in the relative number of these amoeboid-like cells especially in the peri-ventricular white matter, although cell counting was not performed. In 12d control animals, the normal microglial population was more differentiated with established processes, although the processes were not fully ramified as in mature microglial cells (Figs. 1, 3, 2^nd row), while microglia in the hydrocephalic 12d animals had shorter processes, and their cell bodies appeared to be thicker than those in the control animals (Fig. 3, 2^nd row). By 21d, the microglia in control animals had reached their fully ramified state, with branched processes that were long and slender (Fig. 1, 3, 3^rd row). Microglia in the 21d hydrocephalic animals had altered and activated morphology with a wider cell body and shorter, thicker processes (Fig. 3, 3^rd row). These changes in microglia morphology toward that of an activated state continued in the 36d hydrocephalic animals (Fig. 3, 4^th row). There were no dramatic alterations in the microglial populations of the periventricular white matter (Fig. 7) or the cerebral aqueduct (Fig. 8) in any of these untreated animal groups.

Histologic examination of ILB4 stained sections shows that shunting altered the microglial morphology from the appearance of microglia in the untreated brain. Following shunting (both long and short term), the shape of the microglia began to return towards that of a resting microglia cell. Microglia in shunted animals had thicker cell processes than those of resting microglia, but these processes were not as broad and aggravated as the activated microglia in the untreated hydrocephalic animals (Fig. 3, 3^rd and 4^th rows). The microglial cell activation in the 36d shunted animals was almost completely reversed, with less intense staining and a return of fine cellular processes. Additionally, there was a small increase in number of microglia cells in the periventricular area (Fig. 7). Shunting was also found to increase the overall number of microglial cells present in the peri-aqueductal area for both shunt durations, and appeared as small, amoeboid shaped cells on the ventricular surface (Fig. 8G,J).