- Research article
- Open Access
Axotomy-induced neurotrophic withdrawal causes the loss of phenotypic differentiation and downregulation of NGF signalling, but not death of septal cholinergic neurons
© Lazo et al; licensee BioMed Central Ltd. 2010
- Received: 3 August 2009
- Accepted: 19 January 2010
- Published: 19 January 2010
Septal cholinergic neurons account for most of the cholinergic innervations of the hippocampus, playing a key role in the regulation of hippocampal synaptic activity. Disruption of the septo-hippocampal pathway by an experimental transection of the fimbria-fornix drastically reduces the target-derived trophic support received by cholinergic septal neurons, mainly nerve growth factor (NGF) from the hippocampus. Axotomy of cholinergic neurons induces a reduction in the number of neurons positive for cholinergic markers in the medial septum. In several studies, the reduction of cholinergic markers has been interpreted as analogous to the neurodegeneration of cholinergic cells, ruling out the possibility that neurons lose their cholinergic phenotype without dying. Understanding the mechanism of cholinergic neurodegeneration after axotomy is relevant, since this paradigm has been extensively explored as an animal model of the cholinergic impairment observed in neuropathologies such as Alzheimer's disease.
The principal aim of this study was to evaluate, using modern quantitative confocal microscopy, neurodegenerative changes in septal cholinergic neurons after axotomy and to assess their response to delayed infusion of NGF in rats.
We found that there is a slow reduction of cholinergic cells labeled by ChAT and p75 after axotomy. However, this phenomenon is not accompanied by neurodegenerative changes or by a decrease in total neuronal number in the medial septum. Although the remaining axotomized-neurons appear healthy, they are unable to respond to delayed NGF infusion.
Our results demonstrate that at 3 weeks, axotomized cholinergic neurons lose their cholinergic phenotype without dying and down-regulate their NGF-receptors, precluding the possibility of a response to NGF. Therefore, the physiological role of NGF in the adult septal cholinergic system is to support phenotypic differentiation and not survival of neurons. This evidence raises questions about the relationship between transcriptional regulation of the cholinergic phenotype by retrograde-derived trophic signaling and the transcriptional changes experienced when retrograde transport is impaired due to neuropathological conditions.
- Nerve Growth Factor
- Glial Fibrillary Acidic Protein
- Cholinergic Neuron
- Medial Septum
- Nerve Growth Factor Receptor
Basal forebrain cholinergic neurons (BFCN) account for most of the cholinergic innervation of the hippocampus and cortical mantle, and have a key role in the regulation of synaptic activity and modulation of memory and attention in rodents, primates and humans [1–5].
The physiology of septal cholinergic neurons is regulated by the trophic support offered by their target, the hippocampus. Disconnection of septal cholinergic neurons from their target by an experimental transection of the fimbria-fornix, reduces the number of neurons positive for cholinergic markers such as choline acetyl transferase (ChAT) [1, 6–13]. One of the best-studied trophic factors for septal cholinergic neurons is nerve growth factor (NGF). Levels of NGF mRNA and protein are consistently detected in the hippocampal formation and cortex. In addition, it is well established that NGF is retrogradely transported from the hippocampus to the septal area and that intracerebroventricular application of NGF or intracerebral transplant of NGF-releasing cells prevents the reduction in the proportion of neurons positive for cholinergic markers after axotomy [14, 15, 7, 12, 16–18].
The mechanisms of cholinergic neurodegeneration after axotomy are poorly characterized. Most of the studies performed have considered the loss of ChAT, acetyl cholinesterase (AChE) or p75 neurotrophin receptor (p75) immunoreactivity as an analogue of neurodegeneration [19–22], ruling out the possibility that neurons lose their cholinergic phenotype without dying.
Septal cholinergic neurons express two different types of receptors for NGF: the tyrosine kinase receptor TrkA (TrkA), which specifically binds NGF; and the p75 neurotrophin receptor (p75), which binds all neurotrophins. Together, TrkA and p75 activate pro-survival gene expression and influence growth . Conversely, p75 signaling mediates neuronal death by apoptosis in different neuronal systems, including neurodegenerative models such as corticospinal axotomy and seizure-induced apoptosis of septal cholinergic neurons [24–29]. Activation of NGF receptors up-regulates several cholinergic-specific genes, such as the high-affinity choline transporter and the acetylcholine synthesizing enzyme ChAT, which share a common gene locus with the vesicular acetylcholine transporter (VAChT) [30–33].
Axotomy-induced cholinergic decay in the basal forebrain has been explored as an animal model for cognitive decline due to cholinergic impairment, similar to that observed in aging and neuropathologies such as Alzheimer's disease (AD) [34, 35]. Due to its role as a neurotrophic factor for cholinergic neurons, NGF gene therapy is currently in phase 1 clinical trial for AD treatment [12, 36–39].
The principal aim of this research was to re-evaluate, using modern quantitative confocal microscopy, neurodegenerative changes in septal cholinergic neurons after axotomy and to assess their response to delayed infusion of NGF in rats. To pursue this goal, we have performed a stereological analysis of the rat septal area, using quantitative double- and triple-labeling confocal microscopy analysis of axotomized brains with different cell markers and neurodegenerative labels at different time-points after axotomy. Furthermore, we have assessed the response of cholinergic neurons to delayed infusion of NGF three weeks after axotomy.
As shown before, we found that there is a slow reduction in the number of immunoreactive cholinergic cells after axotomy [6, 40]. However, this phenomenon is not accompanied by neurodegenerative changes or a decrease of total neuronal number in the medial septum. Although the remaining axotomized-cells appear healthy, they are unable to respond to delayed NGF infusion. These results demonstrate that axotomized cholinergic neurons down-regulate their NGF receptors, precluding the possibility of a response to NGF.
Our results suggest that the physiological role of NGF in the adult septal cholinergic system is to support phenotypic differentiation and not neuron survival. This evidence raises the question of how the connection with the target regulates the transcription of cholinergic markers in vivo and of which other factors could re-induce the cholinergic phenotype when retrograde transport is impaired due to neuropathological conditions such as AD and Down syndrome [38, 41, 42].
Expression profile of ChAT and p75 immunopositive septal cholinergic neurons after axotomy of the fimbria-fornix: a time course study
Septal cholinergic neurons do not degenerate through apoptotic cell death after axotomy
To standardize the immunostaining of cleaved caspase-3 and p53, we induced cell death by injecting H2O2 to the medial septum. In these conditions, cleaved caspase-3 and p53 clearly labeled damaged neurons as indicated by their co-localization with altered Neurotrace labeling (Additional file 1). Neurotrace is a fluorescent Nissl staining that only labels neurons and does not co-localize with markers of different glial cells such as astro, micro or oligodendroglia (Additional file 2).
There is no loss of neurons in the medial septum three weeks after axotomy
Total number of neurons in the medial septum.
Number of neurons
Left (± SD)
Right (± SD)
Number of brains considered
Number of sections per brain
1090 ± 319 (CL)
1103 ± 233 (IPL)
1085 ± 137
1057 ± 82
Response of axotomized septal neurons to delayed NGF infusion
In the present study, we report the response of basal forebrain cholinergic cells to axotomy based on the use of time-course analyses, multiple immunostainings and quantitative confocal microscopy. Our results establish that cholinergic cells do not respond to disconnection of their target, as induced by axotomy, with apoptotic cell-death, but rather with down-regulation of the neurotransmitter-synthesizing enzyme ChAT and NGF receptors. As opposed to other nervous system injury models, such as corticospinal and olfactory bulb axotomy and kainic acid-induced cytotoxicity [27, 29, 51], p75 receptor does not play a role in axotomy-induced cholinergic cell neurodegeneration. Of note, like septal cholinergic cells, Purkinje cells in the cerebellum are also resistant to axotomy . We are yet far from explaining the differences in the response to axotomy between cholinergic and Purkinje neurons and cortico-spinal and olfactory bulb neurons. One possibility is that cholinergic neurons became resistant to the withdrawal of neurotrophic factor, as postulated by Snider  for sensory neurons. It is well established that during development, or in neonates, axotomy or neurotrophic deprivation induces apoptotic cell death of dorsal root ganglia (DRG) neurons . However, adult DRGs 'as opposed to embryonic DRGs- differentially express DeltaNp73a, a pro-survival protein related to the p53 family . The expression of this protein renders cells resistant to axotomy or neurotrophic factor withdrawal. Another gene involved in protection of neurons after injury is the heat shock protein Hsp27, which directly or indirectly activates the Akt survival pathway. The expression of Hsp27 inhibits JNK-mediated apoptosis in superior cervical ganglion neurons and adult DRGs [56, 57]. It would be interesting to study whether adult cholinergic cells express DeltaNp73 and/or Hsp27. As mentioned above, other adult neurons such as corticospinal neurons are not resistant to axotomy and die in a p75-dependent fashion, probably because they fail to express any of the abovementioned survival programs after injury [29, 58].
One unexpected observation of our study was that only GFAP-positive cells were immunopositive for cleaved caspase-3. GFAP positive cells appear hypertrophic and strongly stained in the whole tissue, thus revealing a generalized astrogliosis. This reaction may account for a global inflammatory response to the lesion which may cause apoptosis of astrocytes, as has been reported in other injury models or in neurodegenerative diseases . It is interesting that apoptotic cell death in the septal nucleus after axotomy was previously reported by electron microscopy studies . However, the quality of the microscopy precludes discrimination of the cell type. Therefore, it is possible that the apoptotic septal cells mentioned in this study are glial cells rather than neurons.
Disruption of the connection between septal cholinergic cells and the hippocampus results in down-regulation of ChAT, but not neuronal death. A consequence of this is that the basal forebrain cholinergic cells affected during aging and in pathological states such as AD may still be alive. Therefore, the possibility of restoring the cholinergic phenotype by identifying new factors that influence cholinergic function would be an interesting point for further investigation. Some candidates are the BMPs and neurosteroids. Studies by Lopez-Coviella and colleagues have shown that BMP-9 is a robust factor for induction and maintenance of the cholinergic phenotype in vitro [60, 61], which also synergizes with NGF to enhance neuronal transcriptional response [30, 62]. In addition, neurosteroids such as estrogen and retinoic acid have also been shown to up-regulate the cholinergic phenotype in the septal basal forebrain [63, 64].
Taking this into account, future studies could search for components that up-regulate the cholinergic phenotype in the absence of NGF receptors. This may open new avenues for therapeutic intervention for the treatment of the cholinergic deficit observed in AD.
Male Sprague-Dawley rats were maintained with free access to water and fed with normal rat chow at the Pontificia Universidad Catolica animal care facilities. Experimental procedures were in accordance with institutional standards for care and use of laboratory animals.
Rats weighting 280-300 g were anesthetized (xylazine 2 mg/ketamine 20 mg i.p. and lidocaine 9% locally applied on the ears) and positioned in a stereotaxic apparatus. Coordinates were calculated based on the Paxinos and Watson atlas of the rat brain . After surgery, the animals were injected i.p with antibiotic (enrofloxacine 7.5 mg) and maintained under observation and temperature control for one hour. For histological preparation of brain tissue, rats were transcardially perfused with 250 ml of 0.9% NaCl, and 250 ml of 4% paraformaldehyde (PFA) in phosphate buffer. After extraction, the brain was post-fixed overnight in 4% PFA, left on 30% sucrose for 24 hours, and coronally sectioned (40 μm) on a cryostat.
Unilateral axotomy of the septo-hippocampal pathway was induced by aspirative lesion of the fimbria-fornix, as has been previously described . In brief, anesthetized rats were positioned in a stereotaxic apparatus and a small piece of skull was removed at the stereotaxic coordinates 1.8 mm caudal to the bregma, and 0.0-4.0 mm lateral to the midline. After excision of the dura, we performed a syringe aspiration of the dorsal fornix-fimbria. We also used a syringe to aspirate part of the cingulate and parietal cortices, 3.5 mm ventral from the brain surface. Rats were sacrificed 3-21 days after axotomy.
H2O2injection in the medial septum
Two μl of 0.1 M H2O2 were injected in the septal areas of two adult rats: 0.35 mm rostral to bregma, 0.5 mm lateral to midline and 7 mm dorso-ventral. After 72 hours, rats were perfused and brain sections were prepared for immunostaining.
Intracerebroventricular infusion of NGF
Artificial cerebrospinal fluid (ACSF) containing 150 mM NaCl, 1.8 mM CaCl2, 1.2 mM MgSO4, 2 mM KH2PO4 and 10 mM glucose, pH 7.4 with or without NGF (Alomone Labs, Jerusalem, Israel) at a concentration of 0.2 μg/ml was infused for 14 days (2.5 μL/hour) by using a brain infusion kit (Alza Corp., Palo Alto, CA), connected to a model 2002 Alzet osmotic pump (Alza Corp., Palo Alto, CA), as described previously . The cannulae and connector tube were filled with ACSF only or with ACSF plus NGF and attached to a loaded pump. Using the arm of the stereotaxic apparatus, the cannula was lowered into the brain at left ventricle coordinates (0.8 mm caudal to bregma, 1.2 mm lateral to midline and 3.5 mm ventral to the brain surface) and finally anchored to the skull with a screw and glued with dental acrylic. Axotomized rats were infused for 14 days immediately after axotomy (simultaneous NGF infusion) or 3 weeks after axotomy (delayed NGF infusion).
Serial coronal cryostat sections (40 μm) were collected in 0.1 M phosphate buffer (pH 7.4), washed in 65 mM sodium maleate (pH 6.0) and incubated for staining, as floating sections, for 1 hour at room temperature in 0.05 mg/mL acetylthiocholine iodide, 0.1 tetra-isopropyl-pyrophosphatamide, 0.05 mM potassium ferricyanide, 0.3 mM CuSO4, 0.5 mM sodium citrate, and 65 mM sodium maleate (pH 6.0), as described previously .
ChAT immunohistochemistry was performed as follows: (i) 15 min incubation in 0.03% H2O2 in 0.1 M Tris-HCl, 150 mM NaCl, pH 7.4 (TBS) to block endogenous peroxidase; (ii) 30 min incubation at 4°C with 0.4% Triton-X100 in TBS; (iii) 1.5 hr incubation at 4°C with 0.2% Triton-X100, 5% rabbit serum, 5% BSA in TBS; (iv) 48 hr incubation at 4°C with goat anti-ChAT antibody (Chemicon, CA, USA) diluted 1:300 in TBS plus 0.2% Triton-X100 and 5% serum; (v) 1.5 hr incubation at room temperature with biotin-conjugated rabbit anti-goat IgG (1:300 in TBS; DakoCytomation, CA, USA); (vi) 1 hr incubation with peroxidase-conjugated avidin ABC (DakoCytomation, CA, USA), followed by visualization of peroxidase activity with diaminobenzidine (DAB, 1 mg/mL) 0.01% H2O2 in TBS.
NGF immunohistochemistry was preceded by 15 min incubation in 50% ethanol, followed by the same protocol already described. Rabbit anti-NGF (Alomone Labs, Jerusalem, Israel) and rabbit anti-p75 (Upstate, NY, USA) were used at 1:300 and 1:500, respectively.
Single or double-immunofluorescence was performed in floating brain sections as follows: (i) 15 min incubation in TBS; (ii) 15 min incubation in NaBH4 10 mg/mL; (iii) 30 min incubation at 4°C with 0.4% Triton-X100 in TBS; (iv) 1.5 hr incubation at 4°C in 0.2% Triton-X100, 5% rabbit serum, 5% BSA in TBS; (v) 48 hr incubation at 4°C with primary antibodies in 0.2% Triton-X100 5% serum in TBS; (v) 1.5 hr incubation at room temperature with fluorochrome-conjugated (Molecular Probes, Oregon, USA) or biotin-conjugated secondary antibodies (directly labeled antibodies were used 1:100 in TBS; antibodies amplified with biotin were diluted 1:300 in TBS), followed by 1.5 hours with fluorochrome-conjugated streptavidin (Molecular Probes, Oregon, USA). Mouse anti-Neu-N, anti-parvalbumin, and rabbit anti-GFAP (labeling astroglia) antibodies were from Chemicon (Temecula, CA, USA). Mouse anti-CD11B (labeling microglia) was from Serotec, Oxford, UK. Rabbit polyclonal anti-OMgp (labeling oligodendroglia) was kindly provided by Dr. Colman (McGill University, Montreal, Canada). Polyclonal rabbit anti-cleaved caspase-3 was purchased from Cell Signaling Technology (Danvers, MA, USA). Monoclonal anti-p53 was from Santa Cruz Biotechnologies, CA, USA; polyclonal rabbit anti-TrkA antibody was kindly provided by Dr. L. Reichardt (University of California, San Francisco, CA, USA).
Neurotrace and Fluorojade C staining
Staining with fluorescent probes such as Neurotrace (fluorescent Nissl stain; Molecular Probes, Oregon, USA) and Fluorojade C (specific marker for degenerating neurons; Chemicon, CA, USA) was performed as indicated by the manufacturers' instructions. Briefly, Neurotrace staining was performed after immunostaining by incubating brain sections for 20 min in a 1:200 dilution of Neurotrace in TBS. Sections were then rinsed, air-dried and mounted in Mowiol. Fluorojade staining was performed after Neurotrace as follows: brain sections were rinsed twice in TBS, re-hydrated for 2 minutes in distilled water and incubated for 10 minutes in 0.06% potassium permanganate. Finally, brain sections were washed for 2 minutes in distilled water and incubated for 10 minutes in 0.0002% Fluorojade C solution in 0.1% acetic acid. Sections were immediately rinsed in distilled water, mounted, air-dehydrated, cleared with xylene and mounted in Entellan.
Septal cholinergic cell counting
The cholinergic cell count was performed essentially as described before  but modified for rat. Septal cholinergic neurons were defined by using anatomical landmarks in accordance with the rat brain atlas . The ventral border of the medial septum was defined dorsal to the anterior commissure, and the rostral beginning was indicated by the meeting of the body of the corpus callosum at the midline. The caudal end of the septal nucleus was defined by the appearance of the fornix and the midline crossing of the anterior commissure. Four 40-μm-thick coronal sections were examined for each rat (n = 4 animals per time point), starting 0.7 mm caudal to bregma and 200 μm apart, to avoid counting the same cell twice. For each section, immunopositive neuronal profiles (labeled with ChAT or p75) were counted on images digitized on an Olympus BX51 (Tokyo, Japan) optic microscope (40× objective), equipped with a CoolSnap-Pro digital camera (Media Cybernetics, Maryland, USA) and connected to an image analysis system based on Image-pro express software, version 126.96.36.199 (Media Cybernetics, Maryland, USA). The pictures were analyzed by using the Sigmascan software (SPSS; Chicago, IL, USA). The criterion for identifying ChAT- or p75- immunopositive cells was the appearance of a clear nucleus or, in cases when staining was too dark, clear neuronal morphology.
Total neuron counts
The anatomical landmark we used to define the medial septum nucleus (MS) was the same previously described for septal cholinergic cell counting. Three 40-μm-thick coronal sections were considered for each rat (n = 5 animals), starting 0.7 mm caudal to the bregma and 200 μm apart, to avoid counting the same cell twice. The sections were double-stained with Neurotrace (fluorescent Nissl stain; Molecular Probes, Oregon, USA; as indicated by the manufacturer instructions) and anti-Neu-N diluted 1:250 and developed with a secondary antibody conjugated to Alexa 488 fluorochrome. The area for counting was defined as a triangle: its base was a horizontal line crossing the middle point of the left and right anterior commissures, and its sides were the anatomical borders of the medial septum, as shown in Figure 5. Eight pictures of this area (four per side) were obtained using a confocal microscope with a 63× objective and scanned with an optical section of 10.3 μm. The criterion used to define healthy neurons was Neu-N and/or Neurotrace staining: cells with cytoplasmic and nucleolar staining, as shown in Figure 3, were counted as healthy neurons. Other patterns of staining corresponding to neurons undergoing degeneration [67, 68], such as perinuclear Nissl (or chromatolytic) staining and eccentric distribution of the nucleus, were scarcely observed after axotomy (and clearly observed after H2O2 injection) and were not considered.
Confocal images for counting double-labeled ChAT/p75 neurons or total neurons labeled with Neurotrace and Neu-N were collected on a Zeiss LSM Pascal 5 (including a triple laser module [Arg 458/488/514 nm, HeNe 543 nm, HeNe 633 nm; Carl Zeiss, Thornwood, NY]) connected to an inverted microscope (Axiovert 2000). A lower objective (20×) was used to have a panoramic view of the septal nucleus of each brain section and a higher magnification objective (63×) was used to scan the total area as described in Figure 5. The images were analyzed using the SigmaScan software (SPSS, Chicago, IL, USA).
Comparisons between the axotomized or unlesioned side of the septum were statistically validated by performing a Student's t-test to determine significance level (p < 0.05). The analyses were performed using the septum contralateral to the lesioned side as a control (100%).
We wish to thank Dr. Ariel Reyes (Universidad Diego Portales, Santiago, Chile) for technical advise about stereotaxic surgery, and Waldo Cerpa and Jeniffer Serrano for their collaboration in the initial steps of this study. This work was supported by DIPUC, FUNDACION ANDES, FONDAP Center for Biomedicine (13980001), CARE (Conicyt PFB12/2007), FONDECYT 1040799, 1085273, and MINREB (Millennium Center for Regenerative Biology).
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