Hormonal and morphological study of the pituitaries in reeler mice

Abstract

Reelin is a neuronal glycoprotein that plays a crucial role in brain layer formation during prenatal development. The reeler mutant mouse lacks Reelin, leading to abnormalities in the neuronal layering of cerebral cortex and cerebellum, producing ataxia, tremor and abnormal locomotion. Reeler mice are reported to have growth retardation and most of them are sterile or unable to bring up their newborns. Since the brain is one of the main regulator of pituitary hormone secretion and no information was reported regarding pituitary function and structure in these mutant mice, we studied pituitary endocrine activity and morphology in reeler mice. Mice were classified in three groups as reeler homozygote (RHM), reeler heterozygote (RHT) or control (CO). Pituitary hormone blood levels were assessed by enzyme immunoassay (EIA) and immunoradiometric assay (IRMA). Animals and their pituitaries were weighted and pituitaries were studied by histology, immunohistochemistry and electron microscopy. Results showed statistically significant differences in body weight and in adrenocorticotropic hormone (ACTH) and luteinizing hormone (LH) blood levels between the three groups. In contrast, growth hormone (GH) blood levels showed a high individual variation and no decrease in reeler groups compared with CO. Morphological studies revealed no differences in pituitary cell types except that somatotrophs appeared to be slightly smaller in RHM and RHT. Although it seems that pituitary hypofunction is not responsible for growth retardation, more studies are needed to obtain a deeper insight into the endocrine status of these mutant mice to elucidate the cause of their low body weight and reproductive behaviour.

The reeler mouse, first recognized by Falconer in 1951, is an autosomal, recessive, spontaneous mutant exhibiting ataxia, tremor, imbalance, a characteristic reeling gait and behavioural disturbances. Falconer (1951) described them pointing out that the most striking abnormality was the mouse inability to keep its handquarters upright and when walking or running they frequently fall over on their sides. He also suggested that reeler mice must be mentally deficient because they showed no signs of the normal aggressive behaviour toward strange mice.

The gene which is mutated in reeler mice has been identified and is called reelin (D'Arcangelo et al. 1995; Goffinet 1995). It encodes an extra-cellular matrix protein of 385–420 kDa (D'Arcangelo et al. 1995; Smalheiser et al. 2000). The Reelin protein is synthesized mainly in the Cajal-Retzius neurons of the brain (Ogawa et al. 1995; Schiffmann et al. 1997) but it is also expressed in the spinal cord and retina (Schiffmann et al. 1997) and outside the nervous system, mainly in the liver but also in spleen, thymus, kidney, testis and ovary as well as adrenal medulla and can also be detected in the blood (Ikeda & Terashima 1997; Smalheiser et al. 2000; Samama & Boehm 2005). The Reelin protein binds to several receptors [very-low-density-lipoprotein receptor (VLDLR) and apolipoprotein E receptor 2 (ApoE2), both lipoprotein receptors (D'Arcangelo et al. 1999; Trommsdorff et al. 1999] and inhibits neuronal adhesion, influences neuronal migration and regulates lamination (Curran & D'Arcangelo 1998). Reelin plays a vital role during development but it seems that it performs an important function during the adulthood too. Reelin is highly expressed during the period of development when brain cells are migrating and it guides newly formed neurons to their proper final destination. It is only then that normal synaptic connections may be established between neurons. The lack of this protein guide leads to the neuronal patterning in disarray (Tissir et al. 2002). After birth, Reelin regulates synaptic plasticity and neurotransmitter synthesis (Pesold et al. 1998) and plays a role in stabilizing the cyto-architecture and in remoulding in adult organs (Ikeda & Terashima 1997). In the reeler homozygote mouse (rl/rl) the Reelin protein is not produced leading to disruption of cellular organization, severe hypoplasia of the cerebellum which lacks foliation, neuronal ectopia in laminated brain structures such as the cerebral and cerebellar cortices and the hippocampus (Hamburgh 1960, 1963; D'Arcangelo & Curran 1998). Several other neurons are also abnormally positioned (Yip et al. 2000; Nishikawa et al. 2003). Reeler heterozygote mice (rl/+) expressing Reelin at 50% of wild type levels, apparently present normal behaviour (Podhorna & Didriksen 2004), have normal brains but exhibit a progressive loss of Purkinje cells, a neuronal target of Reelin action, during ageing (Hadj-Sahraoui et al. 1997; Tueting et al. 1999).

The brain plays a crucial role in the regulation of pituitary hormone secretion (Horvath & Kovacs 1991). Since brain development is markedly altered in the reeler mice, we decided to investigate their pituitary function and structure. To our knowledge, information is limited regarding endocrine status in the reeler mice. It is known that the majority of male and female reeler mice show growth retardation and appear to be sterile, and the females that breed are not able to rear their young (Falconer 1951). It was shown that the Reelin protein is expressed in the pituitary pars intermedia of rats and is colocalized with alpha-melanocyte-stimulating hormone (α-MSH) (Smalheiser et al. 2000). It was also reported that Reelin blood levels do not change after hypophysectomy (Smalheiser et al. 2000).

Materials and methods

All mice were handled in accordance with the procedures of the Guiding Principles for the Care and Use of Animal Research (86/609/EU) at the University of Santiago de Compostela, Campus of Lugo, Lugo, Spain. Fifteen adult non-pregnant female mice from our in-house breeding colony were used in this experiment. The animals were divided in three groups: five animals were classified as reeler homozygote (RHM), five mice were Reeler heterozygotes (RHT) and five animals were normal and considered as the control group (CO). They were housed at room temperature with controlled photoperiod, and fed a diet of mouse pellets and water ad libitum. All animals were weighted and then killed with deep pentothal anaesthesia.

Hormone assays

Blood was collected with a syringe directly from the heart and stored in vials with heparin at 4 °C. Then, blood was centrifuged at 3600 g for 20 min at 4 °C, and plasma was collected and frozen till used for hormonal assay. Plasma growth hormone (GH), follicle-stimulating hormone (FSH), and luteinizing hormone (LH) were determined by means of enzyme immunoassay (EIA). Plasma adrenocorticotropic hormone (ACTH) was measured using a two-site immunoradiometric assay (IRMA). All the results are expressed in ng/ml.

In brief, the GH assay was performed in 96-well-microtitre plates. The determination was based on the sandwich principle. Duplicates of mouse GH standards [0, 1, 2.5, 5, 10, 50 and 100 ng/ml (DSL-10-72101 to DSL-10-72106)], controls (DSL 10-72151 and DSL-10-72152), and plasma samples were incubated in precoated wells with anti-mouse GH (DSL-10-72110). Present GH was bound to the wells and unbound material was removed by washing. Anti-mouse GH conjugated with peroxidase (DSL-10-72120) was added, and incubated for 90 min at room temperature while shaking. The unbound material was removed by washing. Next, a chromogene substrate solution (TMB Neogen, Lexington, KY, USA) was added in each well and incubated for 15–20 min at room temperature. This reaction was stopped with 10% H₂SO₄. Then, the intensity of colour development was read using a spectrophotometer set at 600–450 nm. The standard curve was obtained by blotting GH concentration vs. absorbance using specific software (elisa aids, Eurogenetics, Tessenderlo, Belgium). The GH plasma concentration of the specimens were determined from the curve. The coefficients of intra-assay and inter-assay variations were <5% and <14% respectively. The minimum detectable concentration of GH was 0.13 ng/ml and the recovery value was 98.5%.

The FSH and LH concentrations in plasma were measured by EIA based on the sandwich principle in two steps. Briefly, the assay was performed in 96-well-microtitre plates precoated with anti-mouse FSH or LH (mFSH: NIDDK-anti-rFSH-S-10; mLH: NIDDK-anti-rLH-S-11). Duplicates of mouse FSH or LH standards [0, 1, 2.5, 5, 10, 50 and 100 ng/ml (FSH: AFP-5318 and LH: AFP-5306A)] controls, and plasma samples were added and incubated for 120 min at room temperature. Next, the second antibody (anti-mouse IgG conjugated with peroxidase, A3673, Sigma, St Louis, MO, USA) was added and incubated for 60 min at room temperature. The liquid was removed and wells were washed five times. Then, a chromogene substrate solution (TMB Neogen) was added in each well and incubated for 15–20 min at room temperature. This reaction was stopped with 10% H₂SO₄. The intensity of colour development was read using a spectrophotometer set at 600–450 nm. The data were analysed using specific software (elisa aids; Eurogenetics). The coefficients of intra-assay and inter-assay variations were <5% and <14% respectively. The minimum detectable concentration of FSH and LH were 0.20 and 0.15 ng/ml, respectively. The recovery value was 98.0% for FSH and 96.5% for LH.

Plasma ACTH was measured using a two-site IRMA. In short, for assay in duplicate, 100 μl of first specific antibody anti-ACTH (DSL-5110) were added to plastic tubes except tubes 1 and 2. Tubes 3–16 were used for ACTH standards (0, 6, 29, 55, 225, 750 and 2000 pg/ml DSL-5101 to DSL-7107) and the other tubes were for plasma samples. The second antibody was added (anti-ACTH radiolabelled with ₁₂₅I, DSL 5210) to all tubes and they were incubated for 18–20 h at room temperature. Then, all the tubes, except tubes 1 and 2, were washed three times with wash solution and the bound radioactivity was measured by a scintillation gamma counter for 1 min. The two uncoated tubes filled with ₁₂₅I were used as the ‘total count’. The results were processed by specific software (elisa aids, Eurogenetics) to assess the ACTH plasma concentration. Intra- and inter-assay coefficients of variation were <5% and <9% respectively. The minimum detectable concentration of ACTH was 1.3 pg/ml and the recovery value was 99%.

Morphological methods

Simultaneously, the pituitaries were quickly removed, weighted and immersed in 10% buffered formalin. The pituitaries of three animals per group were divided in two halves. One half was fixed in 10% buffered formalin as well as the endocrine organs (thyroid, thymus, pancreas, adrenals and ovaries) and embedded in paraffin using an automatic processor. Paraffin sections were cut of 5 μm in thickness, mounted on poly-l-lysine coated slides and dried at 37 °C. Slides were then dewaxed in xylene, rehydrated, and transferred to distilled water. Sections were pretreated for antigen retrieval by microwaving with 0.01 M citric buffer, pH 6.0. After heating, the slides were removed from the microwave and allowed to cool for 15 min at room temperature. Slides were then rinsed in distilled water and in TRIS-buffered saline (TBS) for 5 min. Subsequently treated slides were immnunostained to identify the hormone producing cells in the anterior pituitary by the avidin–biotin–peroxidase complex method using various specific polyclonal antibodies. The sections were pretreated with 1.5% H₂O₂ in absolute methanol for 30 min to inactivate endogenous peroxidase, 5% normal goat serum for 30 min to block non-specific binding of the secondary antibodies. The primary antisera were diluted in 1% of normal goat serum and incubated overnight at 4 °C [rabbit anti-rat GH 1:3000, anti-rat PRL 1:2500, anti-rat ACTH 1:4000, anti-rat β-thyroid-stimulating hormone (TSH) 1:3000, anti-rat β-FSH 1:4000 and anti-rat β-LH 1:3000]. Slides were washed three times in TBS for 5 min each and incubated for 1 h with biotinylated secondary antibody. The buffer rinses were repeated and the slides were incubated for 1 h with Avidin-Peroxidase Complex (Vectastain ABC kit; Vector Laboratories, Inc., Burlingame, CA, USA). The final reaction was achieved by incubating the sections for 5 min in a solution of 5 mg 3,3′-diaminobenzidine and 20 μl H₂O₂ in 100 ml TRIS buffer, pH 7.4. The reaction was terminated with TBS. The slides were then counterstained with haematoxylin, coverslipped and examined. To ensure specificity, control testing included both (1) replacement of the primary antibody with TBS, and (2) pre-absorption of the primary antibody with homologous antigen. Immunolabelling was completely abolished in both control tests.

The other half pituitaries were fixed in 4% paraformaldehyde-1% glutaraldehyde for 7 h at 4 °C, embedded in Epon, cut and studied using a Philips 410 LS transmission electron microscope (Philips, Santa, Clara, CA, USA).

Statistical analysis

All numerical results are expressed as mean ± SEM. An analysis of variance (anova) was conducted to estimate the differences between RHM, RHT and CO in the observed parameters, followed by a Student–Newman–Keuls multiple range test. Analyses were performed using spss software, version 12.0S for Windows. The statistical level of significance was set at P < 0.05.

Results

Body weights and pituitary weights

Means and standard errors of the body and pituitary weights from each group are presented in Table 1. Note the very significant differences in the parameter of body weight between the three groups, emphasizing the low body weight of the RHM compared with RHT and CO.

Table 1.

Average and F-values for the data obtained after comparing some parameters from CO, RHM and RHT mice used in this study

	CO		RHM		RHT
	Mean	SEM	Mean	SEM	Mean	SEM	F-values
Body weight (g)	42.4 c	3.01	19.3 a	0.73	25.7 b	0.96	54.29****†
Pituitary weight (mg)	2.9	0.58	1.3	0.21	1.4	0.23	4.70 ns
Blood levels (ng/ml)
GH	47.9	14.96	93.2	58.05	208.3	77.74	2.66 ns
ACTH	1.8 b	0.23	1.0 a	0.19	1.1 a	0.05	8.34**
FSH	2.1	0.07	2.1	0.06	2.0	0.07	0.55 ns
LH	4.9 a	0.10	5.9 b	0.23	5.3 a	0.24	7.06**

^**Significant at P ≤ 0.01

^****significant at P ≤ 0.0001

^†For each row means with different letters are significantly different (P = 0.05) to Student–Newman–Keuls multiple range test.

ns, non-significant.

Pituitary weights showed a strong correlation with body weights as expected. The correlation was significant at P ≤ 0.01 level. However, due to a lower number of pituitaries weighted, the parameter of pituitary weights did not show significant differences between the three groups studied.

Blood hormone levels

Blood levels of GH, ACTH, FSH and LH are displayed in Table 1. Results showed a high individual variation in the GH blood levels expressed by the high SEM but no statistically significant differences were seen between the three groups. ACTH blood levels were significantly decreased (P ≤ 0.01) in reeler groups compared with the CO group, whereas LH levels were significantly higher (P ≤ 0.01) in RHM compared with CO and RTH groups. No differences were found in FSH values between the three groups. We could not measure PRL and TSH blood levels because it was not possible to collect enough blood in the RHM animals.

Histology and immunohistochemistry

By light microscopy all the pituitaries from RHM, RHT as well as CO showed no histologic changes with normal proportion of acidophilic and basophilic cells.

Immunohistochemistry for GH showed somatotrophs with spheroidal or ovoid shape. Somatotrophs made up a large proportion of the adenohypophysis and were distributed diffusely in the gland (Figure 1). The unique difference we could observe was that the size of GH cells from RHM and RHT seemed to be slightly smaller than in CO mice.

Figure 1.

Photomicrographs showing the pituitary cells of the groups studied: CO (first column), RHM (second column) and RHT (third column) immunostained for GH (first row), PRL (second row) and ACTH (third row). All the micrographs have the same magnification. Scale bar, 15 μm.

Lactotrophs were more irregular, polyhedral and their cytoplasmic processes extended between neighbouring cells towards capillaries. Lactotrophs constituted a high percentage of the adenohypophysial cells and were distributed randomly in the gland (Figure 1). No differences were detected between RHM, RHT and CO.

Corticotrophs were small, stellate and characterized by cytoplasmic processes that projected between neighbouring cells. The number of corticotrophs depended on the level of cut. As compared with GH and PRL cells, there were few and exhibited a much different pattern of distribution. Corticotrophs of CO and RHT mice showed a strongly positive cytoplasmic immunostaining. Corticotrophs from RHM mice displayed a stronger immunopositivity mainly close to the cell membrane (Figure 1).

Thyroid-stimulating hormone cells were large, polyhedral cells and their borders were usually indented by adjacent cells. They were limited to the ventral half of the adenohypophysis and usually appeared isolated although sometimes they formed clusters (Figure 2). Apparently, no differences were seen in the thyrotrophs between the three studied groups.

Figure 2.

Photomicrographs showing the adenohypophysis of the three studied groups immunostained for TSH (first row), FSH (second row) and LH (third row). All the micrographs have the same magnification. Scale bar, 15 μm.

The gonadotrophs were polyhedral and irregular in size and shape. They were numerous and distributed all around the adenohypophysis. Several gonadotrophs were isolated, whereas others formed densely packed groups displaying a moderate to strong immunopositivity (Figure 2). No major differences were detected in the gonadotrophs between the three groups.

Transmission electron microscopy

The somatotrophs were spherical with a centrally located nucleus. The cytoplasm contained a large number of organelles, with well developed rough endoplasmic reticulum (RER), showing several parallel cisternae with ribosomes attached. Some somatotrophs had a dilated RER in a paranuclear position, independently of the group studied. The Golgi complexes were located adjacent to the nucleus and were composed of flat sacculi containing immature secretory granules. The mitochondria were ovoid or rod shaped with lamellar cristae. The secretory granules were spherical, with different electron density but homogeneous in size, and were randomly distributed throughout the cytoplasm. The GH cells of RHM (Figure 3) and RHT (Figure 4) seemed to be slightly smaller in size and they were less rich in RER than the somatotrophs of the control group (Figure 2).

Figure 3.

Electron micrographs showing the parenchyma of the pituitaries of mice from the CO group. Dilated RER is apparent in GH cytoplasm. Note the follicle-stellate cells arranged around a pituitary follicle (3A). GH, somatotroph cell; PRL, lactotroph cell; ACTH, corticotroph cell; TSH, thyrotroph cell; FS, follicle-stellate cell. Scale bars, 2.5 μm.

Figure 4.

Electron micrographs showing the pituitaries from the RHM group. GHT, gonadotroph cell; the others are the same as figure 3. Scale bars, 2.5 μm.

Lactotrophs were elongated possessing an ovoid eccentric nucleus. The cytoplasm exhibited several well developed RER cisternae displayed in parallel arrays, with ribosomes attached and mainly in one pole of the cell. The Golgi complexes were well developed and ring shaped. The mitochondria were ovoid or rod shaped. The secretory granules were abundant, spherical or pleomorphic and had different electron density (Figures 3–5).

Figure 5.

Electron micrograph showing the pituitaries from RHT mice. Scale bars, 2.5 μm.

The corticotrophs were angular or irregular possessing a pleomorphic, eccentric nucleus. The cytoplasm was relatively electron lucent with well developed RER and Golgi complexes. Mitochondria were spherical with lamellar cristae. Secretory granules were numerous, spherical and different in size (Figures 3 and 4).

The thyrotrophs were angular or polyhedral with an usually eccentric nucleus irregular in shape. The RER was moderately developed, the Golgi complexes were prominent with dilated, smooth walled sacculi. The mitochondria were ovoid or rod shaped with lamellar cristae and moderately dense matrix. The secretory granules were uniform in size but very small and localized close to the cell membrane. Apparently no differences were seen in the ultrastructure of thyrotrophs between the three groups (Figures 3 and 4).

Gonadotrophs were usually spherical with a generally eccentric nucleus. The RER and Golgi complexes were well developed. Mitochondria were numerous. The secretory granules presented different sizes and electron density and some of them showed a darker core (Figures 4 and 5). No differences were detected between the three groups.

The endocrine organs in the three groups showed no histological abnormalities and no tumours were observed in any of the samples.

Discussion

The present findings confirm previous results which showed that reeler mice with missing Reelin protein have a low body weight, growth retardation and are small during their life time (Falconer 1951). The small stature is accompanied by low pituitary weight. The levels of ACTH, FSH and LH in the blood are in the normal range and blood GH concentrations do not differ significantly from the control values. However, due to extensive fluctuations and the unusually high standard errors these results are difficult to interpret. The marked variability in the GH blood levels between the animals from the same group and between groups could be explained by several factors, such as age, the time of blood collection (although these parameters were the same for the three groups) and hypothalamic activity that could be different in each mouse. No major histologic alterations are apparent in the adenohypophysis and immunohistochemistry demonstrates normal distribution of hormone producing cells. Transmission electron microscopy reveals normal ultrastructure of adenohypophysial cells with the exception of GH cells which seem to be slightly smaller compared with those of the controls. The hormone producing organelles, including the RER, Golgi complexes and mitochondria are well preserved and no change is obvious in their size, appearance and number of secretory granules.

Short stature and dwarfism, a more advanced variant, are intriguing conditions with different aetiology. Dwarfism occurs in children when GH is not produced in sufficient amounts. GH deficiency may be due to tumours such as craniopharyngiomas, other destructive lesions in the sella region, prepubertal hypophysectomy or various genetic abnormalities which suppress the proper development of the pituitary (Lopes et al. 2006). In children with Laron dwarfism, GH secretion is not decreased but the GH receptor gene is mutated and GH can not exert its growth producing effect (Giordano 1992; Hull & Harvey 1999). Other conditions unrelated to the pituitary include prepubertal oestrogen/androgen/glucocorticoid excess, Turner syndrome, various metabolic diseases and skeletal disorders (Martin et al. 2003; Dehner et al. 2004). Several studies provided conclusing evidence that genetic errors may lead to short stature and dwarfism in rodents as well (Efstratiadis 1998; Sellier 2000). The cause of short stature, growth retardation and pituitary hypoplasia in the reeler mice remains obscure. The question can be asked whether these alterations are due to hypopituitarism affecting primarily GH secretion. However, blood GH levels were not reduced significantly suggesting that the small pituitary secreted hormones in sufficient amounts to maintain endocrine homeostasis.

The study of the pituitary hormonal blood levels in mutant mice with nervous symptoms has not been carried out extensively. To our knowledge, information regarding pituitary function in reeler mice was not reported to date. Consequently, little information is available about the hypothalamic–pituitary–adrenal (HPA) axis or hypothalamic–pituitary–gonadal (HPG) axis in mutant mice used as animal models of nervous pathologies.

The hypothalamus and pituitary tightly controls glucocorticoid (GC) synthesis and release. Elevated GC concentrations cause suppression of corticotropin releasing hormone (CRH) and ACTH secretion, leading to homeostasis of GC levels. In animal models chronic pain is associated with activation of the HPA axis, chronic pain acts as an inescapable stressor which is associated with ‘depressive-like’ symptoms in experimental animals (Blackburn-Munro & Blackburn-Munro 2001). Abnormalities in the HPA axis are, so far, the most consistently demonstrated biological markers in depressive illness. Animal models of depression show behavioural abnormalities and upregulation of the HPA axis, thus combine behavioural and neuroendocrine features of human depression (Gass et al. 2001). During stress, there is also a number of key changes in the function of the HPA axis like dysfunction of HPA axis regulation, whether this is due to reduction in negative feedback control of the axis, or altered central drive on the pituitary (Blackburn-Munro & Blackburn-Munro 2001). It may be that in our study, the lower ACTH levels in the reeler groups compared with the CO group is due to high blood levels of GC that would exert a strong negative feedback causing repression of CRH and ACTH secretion.

Although not traditionally associated as regulators of neuronal function, the HPG hormones possess receptors throughout the brain. The effects of the HPG hormones have been reported in the brain related to development, maintenance and cognitive functions. LH is known to cross the blood-brain barrier and receptors of LH are more concentrated in the hippocampus, the region of the brain more vulnerable to Alzheimer's disease (AD) (Meethal et al. 2005). LH also promotes the amyloidogenic processing of the amyloid-beta precursor in vitro (Bowen et al. 2004; Casadesus et al. 2005). Thus evidence has accumulated for a role of LH in promoting neurodegenerative changes. The elevation of LH with dysregulation of the HPG axis at senescence is a physiologically relevant signal that could promote neurodegeneration and be implicated in the pathogenesis of AD (Bowen et al. 2004; Meethal et al. 2005). Although the most studied of HPG hormones were the sex steroids. Because of the feedback loop within the HPG axis, it is difficult to attribute the structural and functional changes during development, adulthood and senescence to a single HPG hormone. Therefore, loss in cognitive function during senescence, typically ascribed to sex steroids, may also result from increased signalling via GnRH, LH or activating receptors (Vadakkadath et al. 2005). In our study, FSH levels were similar in the three groups; however, LH levels showed higher levels in the RHM group compared with the RHT and CO groups. On the basis of our results, it is uncertain whether the higher levels of LH were related to any alterations caused by the reeler gene mutation.

Obviously more studies are needed with larger numbers of reeler mice. Further research should include the investigation of synthesis and release of hypothalamic stimulatory and inhibitory hormones. It may well be that the cerebral changes involve the hypothalamus as well and secretion of hypothalamic hormones may be abnormal. Alternatively, it may well be that the GH receptor is altered, no IGF-1 is formed and GH can not exert its growth promoting effect. It is also possible that low body weight is unrelated to the pituitary and other extrapituitary factors are responsible for the growth retardation.