Introduction
The brain develops in a controlled environment secured by the blood–brain barrier located at the endothelium of the cerebral microvessels and by the blood–CSF barrier located at the epithelium of the choroid plexuses [
1,
2]. The development of the vascular network is intimately linked to that of the neuronal network [
3]. The choroid plexus-CSF system is a key element of neuroimmune regulation [
4], and is involved in the neuroinflammatory response following perinatal injuries, in part through the secretion of multiple cytokines an chemokines [
5]. Apart from these immune functions, these molecules fulfill important physiological functions in the context of brain development [
6]. Severe neonatal jaundice results from liver immaturity that leads to increased levels of unconjugated bilirubin in the blood. This condition specifically affects the brain and can induce neurological impairment ranging from mild neurodevelopmental disorders to dramatic kernicterus [
7‐
9]. The free (plasma protein unbound) fraction of unconjugated bilirubin circulating in the blood increases during severe hyperbilirubinemia. Free bilirubin, being lipophilic, easily penetrates into the central nervous system (CNS) by diffusion across brain barrier cells, and reaches intracerebral concentrations that lead to neurologic alterations. The neuropathological mechanisms associated with this condition remain elusive. Experimental evidence points to the occurrence of oxidative stress and inflammation directly or indirectly induced by unconjugated bilirubin in the brain, both events affecting neural cell proliferation, differentiation and apoptosis [
10‐
15]. Cells forming the blood–brain interfaces require active oxidative metabolism to maintain their tightness and fulfill their functions of protection towards the maturing brain. These cells are the first CNS cells exposed to increased plasma levels of unconjugated bilirubin, and accordingly cerebral endothelial cells have been considered as targets for bilirubin toxicity (reviewed in [
16]). In vitro, bilirubin was shown to lead to endothelial apoptosis and/or to the secretion of factors that are pro-angiogenic and induce barrier dysfunctions [
16‐
19].
We therefore questioned whether pathologic neonatal hyperbilirubinemia alters the postnatal neurovascular network development, and blood–brain and blood–CSF barrier integrity, which would in turn impact brain maturation and functions. Because the choroid-plexus–CSF system is central to neuroimmune interactions and to the initiation of neuroinflammation, we further hypothesized that pathological hyperbilirubinemia would elevate CSF cytokine levels, sustaining neuroinflammation and impacting brain development. To test these hypotheses, we used the Gunn rat characterized by a deficiency in bilirubin conjugation induced by a spontaneous mutation in the 4th common exon of the UDP-glucuronosyltransferase UGT1a enzymes [
20]. Homozygous pup rats, born from heterozygous mothers, display normal plasma levels of unconjugated bilirubin at birth, that sharply increase a few hours after birth and up to 17 days [
21]. This model therefore recapitulates nicely the postnatal profile of unconjugated bilirubin concentration in plasma observed in babies with severe jaundice. The hyperbilirubinemic animals are characterized by a cerebellar hypoplasia and altered behavioral capacities [
22].
In this paper we evaluated the complexity of the vascular network, the integrity of blood–brain interfaces using sucrose as a polar, non-metabolized tracer, and the cytokine content in plasma and CSF during postnatal development in normobilirubinemic rats in order to increase our knowledge on these parameters which remain little characterized during development. In particular, developmental concentration profiles of most cytokines in CSF are currently unknown. In parallel, we examined whether these three parameters are modified in hyperbilirubinemic Gunn animals. We also assessed whether the neuroinflammatory response of the choroid plexus-CSF system to a bacterial stimulus was exacerbated in hyperbilirubinemic newborn rats. Altogether, the data point out important early postnatal functional changes in the blood–brain interfaces/CSF system, and show that among brain cells, those forming the blood–brain interfaces do not appear to be primary targets of bilirubin toxicity.
Material and methods
Animals, genotyping, fluid and tissue sampling
Gunn rats were obtained from the SPF animal facility of the University of Trieste and the colony was maintained at the Lyon Neurosciences Research Center animal facility. Animal homozygous for the mutation (jj genotype) were characterized by a yellow skin color clearly visualized shortly after birth, in contrast to wildtype animals (NN genotype) and animals bearing the heterozygous mutation (Nj genotype). They also developed a mild cerebellar hypoplasia (Additional file
1: Fig. S1). Total serum bilirubin concentration strongly increased to 11 mg/dl after birth, and kept increasing up to 15 mg/dl (257 µM) 17 days after birth in jj animals (Additional file
1: Fig. S1). We previously reported the calculated free bilirubin concentration in the blood of jj animals, which was 154 nM in 2-day-old rats, was maintained at a high value (117 nM) in 9-day-old animals, and decreased thereafter with the parallel increase in albumin concentration [
21]. These high bilirubin levels observed in the early postnatal stages correspond to severe hyperbilirubinemia necessitating phototherapy or exchange transfusion in human babies [
23]. In contrast, Total serum bilirubin levels measured in Nj animals were similar or only slightly above control levels in the early postnatal stages (Additional file
1: Fig. S1), and the calculated free bilirubin concentration did not exceed 1 nM [
21]. In jj animals, bilirubin diffused into the central nervous system, as shown by the yellow color of the CSF and brain tissue. The genotyping needed to differentiate wildtype (NN) from heterozygous (Nj) animals was performed by PCR using the forward primer 5′-GGG ATT CTC AGA ATC TAG ACA TTG T-3′ and the reverse primer 5′-TCG TTT GTT CTT TTC TAT TAC TGA CC-3′ (detailed conditions in Additional file
1: Fig. S1). The amplicon was then digested for 3 h at 60 °C with BstN1 (NEB BioLabs), whose consensus sequence is suppressed by the single mutation deletion in the mutated gene. Gel electrophoresis allows to discriminate samples from NN animals (2 bands: 231 and 80 bp) from Nj animals (3 bands: 311, 231 and 80 bp) (Additional file
1: Fig. S1).
For immunohistochemistry, cytokine measurements, and CSF leukocyte counts, fluid and tissue were sampled as follows: animals were injected intraperitoneally with pentobarbital (Euthasol, 0.34 (1 or 2-day-old), 0.2 (9-day-old), 0.1 (17-day-old) mg of pentobarbital/g as an 1/8 dilution in saline), blood was withdrawn on heparin by intracardiac puncture, and the cisterna magna was exposed. CSF was sampled using a glass micropipette, collected in a low binding tube, and its volume measured. Heparinized blood samples were centrifuged at 5000 rpm and plasma were collected. Fluids were frozen at − 80 °C until use. Following fluid sampling, the brains were dissected out of the cranial box, quickly frozen in isopentane at − 45 °C, and stored at − 80 °C until used for immunohistochemistry.
Permeability measurements
The permeability of the blood–brain barrier (at the levels of cortex, cerebellum, pons) and of the blood–CSF barrier was measured towards [
14C]-sucrose used as a polar marker of the barrier cell integrity. The influx constants, apparent brain K
in (App K
in) and true CSF K
in for [
14C]-sucrose were determined as described previously [
24,
25]. Briefly, [
14C]-sucrose was injected intraperitoneally to animals under slight isoflurane anesthesia. Plasma and CSF were sampled after 20 min as described above. Their radioactive content was measured by liquid scintillation counting in a TRICARB 4910 TR (Packard). The brain was collected and frozen on dry ice. Pieces of cortex, pons, and cerebellum, free of meninges, were dissected from the frozen brain in a − 20 °C chest freezer, weighted, and digested in 1 M sodium hydroxide overnight at 4 °C. The digested tissues were further homogenized with a micro-pestle before being analyzed for radioactive content. [
14C]-Sucrose plasma concentration x time area-under-the-curves (AUCs) from 0 to 20 min (AUC
0→20 min) were calculated for each individual [
14C]-sucrose plasma concentration at 20 min, using the shape of a generic time x sucrose plasma concentration curve obtained by the litter-based method on animals sacrificed at various time points between 2 and 30 min. As no differences were found in sucrose plasma concentrations over time between males and females, the values were pooled to build up the generic sucrose plasma concentration x time curves for 1-day-old and 9-day-old animals. The curves were built separately for NN/Nj and for jj animals, as the jj genotype slightly affected the shape of the curves at both ages (Additional file
1: Fig. S2). K
in values were calculated using the following equation:
$${\text{K}}_{{{\text{in}}\,csf}} = \, [ {^{{{14}}} {\text{C}}} ]\text{-} {\text{sucrose C}}_{{{2}0{\text{min}}}} /{\text{AUC}}_{{0 \to {2}0{\text{min}}}} ,$$
where C
20min is [
14C]-sucrose concentration in CSF at 20 min, expressed as µl . ml (of csf)
−1. min
−1, i.e. 10
–3 min
−1.
$${\text{App K}}_{{{\text{in}}\,brain}} = \, [ {^{{{14}}} {\text{C}}} ] \text{-} {\text{sucrose C}}_{{{2}0{\text{min}}}} /{\text{AUC}}_{{0 \to {2}0{\text{min}}}} ,$$
where C
20min is [
14C]-sucrose concentration in discrete brain regions at 20 min, expressed as µl . g
−1 . min
−1.
For 9-day-old animals, App Kin brain values were corrected for the amount of sucrose associated to the brain vascular space to generate true Kin brain, using the following plasma volume determined as described infra: Cerebral cortex: 5.0 ± 1.4, Pons: 5.7 ± 0.7, Cerebellum 12.6 ± 0.6 µl/g, (mean ± SEM, n = 3 to 6 from 3 different litters).
Plasma volume measurement in brain tissue
After pentobarbital anesthesia, a blood sample was collected by intracardiac puncture and plasma was frozen. Half of the animals from one litter underwent a transcardiac perfusion with a physiological solution (Hank’s balanced salt solution supplemented with 10,000 U/l heparin. Perfusion was performed at a rate of 0.75 ml/min for 20 min in newborn animals, 1.4 ml/min for 15 min in 9-day-old animals, and 3 ml/min for 12 min in 18-day-old animals. Brains from perfused and non-perfused animals were collected and frozen on dry ice. Pieces of cerebral tissue, free of meninges, were dissected out of the frozen brain in a − 20 °C chest freezer, weighted, and homogenized in phosphate buffer saline (PBS; in mM: NaCl 150; Na2HPO4 12; KH2PO4 2; pH 7.4) (5 s sonication, amplitude 50%, UP50H Hielscher sonicator). Rat IgG concentrations were measured in plasma and brain homogenate samples by ELISA (Bethyl Laboratories, E110-128 Kit). The vascular volume, expressed as µl/g, was calculated as being the difference of the IgG amount per gram of tissue between non-perfused and perfused animals, divided by the IgG plasma concentration in the non-perfused animals.
Analysis of the neurovascular network
Rat brains were cut at − 20 °C using a NX50 Microm Microtech cryostat. Ten-µm thick sagittal sections were collected on glass slides and stored at − 20 °C until used for immunostaining. Sections were fixed at room temperature in 4% paraformaldehyde in PBS for 10 min, then blocked for 1 h in 1% BSA, 8% normal goat serum, 0.3% Triton in PBS. After overnight incubation at 4 °C with the anti- RECA-1 antibody (Bio-rad, MCA970GA, 1.7 µg/ml) in PBS supplemented with 1% BSA, 1% normal goat serum, and 0.3% Triton, sections were washed 3 times with PBS supplemented with 0.3% Triton, and incubated with secondary Alexa Fluor 488-conjugated goat antirabbit antibodies (Invitrogen, Carlsbad, CA; 2 µg/ml) for 1 h at room temperature. Sections were counterstained with DAPI for 10 min and then mounted in aqueous medium. Negative controls were performed by omitting the first antibody. For each area of analysis, three serial sections were examined at X160 with a fluorescence stereo zoom microscope (Zeiss Axio Zoom.V16) equipped with an AxioCam 503 camera, and image acquired through zen 2.3 Pro software (Zeiss Microscopy). Quantitative analysis was done using ImageJ software to determine the number of vessel segments per field as well as the vessel surface area (µm2). Areas with large vessels were omitted from the study by introducing a filter in the analysis that takes into account only vessel profiles whose surface area were comprised between 10 and 25,000 µm2. For each animal and each area of analysis, data from the 3 serial sections were averaged. The number of microvessel segments per field of observation and the vessel area per field of observation were used as proxies for the quantification of microvascular network development and vascular volume, respectively.
Plasma and CSF cytokine content
Cytokines, including several chemokines were analyzed by use of a MILLIPLEX® MAP Rat Cytokine/Chemokine Magnetic Bead Panel RECYTMAG-65 K (Millipore) according to the manufacturer’s protocol. Sample volume needed for plasma and CSF analyses, in singlicate, at the minimal required dilution recommended by the manufacturer (ie 2), was 15 µl. When CSF sample withdrawn from one animal was smaller than 15 µl, samples from 2 animals were pooled, and plasma of these two animals were pooled in the same proportions. Analyte quantification was performed on the Bioplex-200 platform.
Assessment of leukocyte infiltration in CSF
Eight-day-old rats were injected intraperitoneally with 1 mg/kg PAM3CSK4 (P3C, Merck Millipore France), or 0.9% saline. CSF was sampled 14 h later [
5], and total leukocytes and polymorphonuclear neutrophiles (PMNs) were counted in a Bürker chamber after staining with Türk’s solution (Sigma-Aldrich). The phenotypic analysis of whole blood immune cells was performed by immunocytochemistry following red blood cell lysis as previously described [
26].
Statistical analysis
Statistical tests and significance are described in figure legends when appropriate.
Discussion
This work first brings forwards developmental specificities of the brain vasculature and shows that the latter is not impacted by hyperbilirubinemia. The analysis of the number of vessel segments and of the vessel surface per surface area of thin brain sections provides an index of the microvascular network complexity and volume, respectively. It shows that the parenchymal microvascular network undergoes extensive growing complexity between 9 and 18 days after birth in the cerebral cortex of rodents as previously described in pioneer studies [
27,
28] and more recent works based on refined imaging analysis [
29‐
32]. The data also show that this growing complexity extends to other brain areas such as the cerebellum and midbrain, and reveals a region-to-region heterogeneity in the vascular network development during the early postnatal period, that is levelled by day 18.
The cerebral capillaries have both scaffold and paracrine signaling functions that modulate the development and differentiation of multiple neural cell populations, including those migrating radially from neurogenic niches to cortical areas [
3,
33]. Conversely, a change in neural activity affects postnatal angiogenesis. Reducing sensory input decreases the cortical vascular density and branching, while an enhancement of neural activity leads to different types of vascular alterations according to different studies [
30,
34]. Both the formation of cerebral vessels and the postnatal cerebral blood flow which increases drastically in parallel with the neurovascular network between postnatal day 10 and 20, are sensitive to oxidative stress such as induced by hypoxia [
29,
35]. Given these complex interactions between neuronal activity, vascular network development, and cerebral blood flow, and considering that bilirubin induces neuronal toxicity and generates oxidative stress, we hypothesized that the development of the postnatal vascular network would be altered in animals with pathological hyperbilirubinemia. We did not observe any effect during vasculogenesis on postnatal day 9, nor after vasculogenesis on postnatal day 18 and in adult. This holds true even in the cerebellum whose postnatal growth is strongly impacted by hyperbilirubinemia (this paper, [
22]). These data indicate that the mechanisms linking the vascularization to the overall growth of brain structures are maintained in hyperbilirubinemic animals, and suggest that oxygen supply to the brain during postnatal development is not impacted by chronic exposure to bilirubin. Whether the growth of the cerebellum in jj animals adapts to the development of the vasculature, or the vasculature adapts to the reduced cerebellar growth remains to be understood.
The blood–brain permeability to sucrose measured in different brain regions is not altered by bilirubin either, whether sucrose transfer across the blood–brain barrier is measured a few hours after bilirubin concentration rises in the blood, or following a chronic exposure of endothelial cells to bilirubin, such as in 9-day-old rats. Similarly, no change in the blood–CSF permeability to sucrose is observed following choroidal exposure to high plasma concentrations of bilirubin. Barrier permeabilities were evaluated by measuring sucrose k
in constants which represent the fraction of the blood-borne molecule that crosses barriers to reach the CSF and neuropil. The distribution of polar tracers such as sucrose into the CSF and brain depends on the efficacy of tight junctions to seal the paracellular pathway between blood and brain/CSF compartments, the formation of transcytotic pathways across barriers cells, the surface area available for exchange, the rate of CSF-extracellular fluid exchange, the brain-to-blood backflux, and the CSF turnover rate. The impact of the last two parameters is minimized in our 20-min experimental setting. Endothelial and choroidal junctions are already tight at birth [
1,
36], but may be more fragile when facing a pathophysiologic, or toxicologic stress. Together with the formation of tight junctions, non-specific endothelial transcytosis decreases rapidly during development under the influence of pericytes [
37,
38]. Overall this explains the efficiency of blood–brain interfaces to prevent the movement of polar compounds from blood to the developing brain. Still, higher k
in values towards sucrose have been measured in 1-day-old rats as compared to 9-day-old animals, not only in CSF but also in tissue despite the less developed vascular network in 1-day-old animals. The reasons for this age-related decrease in apparent permeability are not fully elucidated and have been discussed elsewhere [
24,
36,
39]. While in 1-day-old rats, part of the microvessels are not perfused, most (> 90%) of them are perfused in 9-day-old animals [
40]. As the extent of the vascular network does not change in hyperbilirubinemic animals at either stage of development (Fig.
1), the similar sucrose permeability measured in normo- and hyperbilirubinemic animals indicates that the integrity of the blood–brain barrier is not impacted by bilirubin, and suggests that bilirubin does not affect the extent of vessel perfusion in early postnatal life either. The preservation of the choroidal blood–CSF barrier in hyperbilirubinemic animals is corroborated by the maintenance of the barrier integrity in a cellular model formed by a tight monolayer of differentiated choroid plexus epithelial cells in primary culture, chronically exposed to a pathophysiologically relevant concentration of bilirubin [
41].
Alteration of the blood–brain barrier involving tight junction disorganization has been reported at the adult stage in animal models of hepatic encephalopathy in which serum bilirubin levels are abnormally high [
42‐
44]. The data however point to the implication of noxious agents other than bilirubin in this alteration. Such agents could be ammonia, or selected inflammatory cytokines released in the plasma as a result of liver failure. No imaging studies in babies investigated the influence of a pathological postnatal rise in plasma bilirubin concentration on the integrity of blood–brain barriers. One postmortem analysis reported an increased vascularization and signs of tight junction alterations, possibly linked to VEGF signaling, in the brain of a baby born prematurely with kernicterus associated with signs of severe hypoxia [
45]. Our histological and functional data indicate that bilirubin itself is unlikely to be responsible for these alterations, especially as confounding factors such as hypoxia and ischemia are known to activate VEGF signaling. This is in line with an experimental study coupling hypoxia-induced acidosis with injection of bilirubin in 3-day-old mice. An alteration of the blood–brain barrier was observed in these animals, that resulted from acidosis, and not from bilirubin exposure [
46]. Finally, Roger et al.found that bilirubin entered the brain without any sign of blood–brain barrier alteration following a two-hour intravenous infusion of bilirubin at a pathological concentration in 10-day-old rats [
47]. The brain barriers harbor numerous transporters, among which neuroprotective ABC transporters, whose expression is developmentally regulated [
48]. Evidence that unconjugated bilirubin alters the expression of some of these transporters during postnatal development and in adult has been brought forwards [
49], and reviewed [
50,
51]. Collectively, our data collected in the Gunn rat which brain is exposed to endogenous plasma bilirubin at a pathologically relevant concentration, and the literature based on exogenously injected bilirubin, indicate that search for direct effects of bilirubin on brain barriers should be oriented towards changes in transport functions rather than an overt impairment of the barrier integrity.
The choroid plexuses are rapidly activated following systemic inflammation and the CSF supports pro-inflammatory mediator circulation [
4]. We detected a number of immune mediators in the CSF of normobilirubinemic animals. Our data show that in developing animals cytokine profiles in CSF and plasma are not correlated. In plasma the most abundant cytokines include RANTES and MCP-1 as observed in human newborns [
52]. The source of most cytokines found in plasma is likely to be circulating immune cells. In CSF the data also unravel different kinetics of cytokines during development. For instance, CSF levels of CINC-1/CXCL1, IL-6, GM-CSF, MIP-2/CXCL2, IL-4 and IL-1α were low in 1-day-old animals and increased strongly thereafter, while the opposite was observed for IL-18, MCP-1/CCL2, VEGF, IFNγ, IP-10/CXCL10, fractalkine/CX3CL1, IL-1β, IL-17α, eotaxin/CCL11, MIP-1α/CCL3. These data suggest that these immune mediators fulfill brain-specific physiological functions during development, independent of their functions as inflammatory modulators. Indeed, besides VEGF whose function is well understood in neurovascular development [
53], cytokines classically associated with the immune system are also involved in neuronal differentiation/migration, synaptic plasticity, and neuroendocrine organization during development [
6,
54,
55]. As examples CINC-1 induces the proliferation and limits the migration of oligodendrocyte progenitors [
56]. Fractalkine also promotes oligodendrogenesis [
57] and controls microglial functions necessary for early postnatal brain maturation [
58]. Indirect evidence based on intrauterine growth restriction associated with IL-4 overproduction indicates that IL-4 regulates oligodendrogenesis during postnatal development [
59]. IL-6 is involved in cortical white matter development, motor development, and shapes long-term social behavior [
60], hence participating in the development of autistic syndromes [
61]. IFNγ acts as an efficient negative regulator of neural precursor cell activity and differentiation into neurons in adult [
62], a function that remains to be explored in the context of postnatal development.
To our knowledge our study is the first to report specific CSF developmental profiles for these immune cytokines in the healthy rat during the postnatal period. The physiological meaning of the changes in CSF concentration for each individual cytokine needs to be investigated, as is their cellular origin. The choroid plexus epithelial cells, ependymal cells including tanycytes and immune and non-immune cells harbored in the ventricular, subarachnoid, and cisternal spaces are potential sources, in line with the role attributed to the choroid plexus-CSF system in securing brain development [
4,
63]. Astrocytes and microglial cells found in the periventricular neurogenic niches, and even neurons located deeper in the brain parenchyma are also potential sources of cytokines circulating in CSF [
56,
64]. Transporters for selected cytokines have been described at blood–brain interfaces [
65]. However, plasma is unlikely to be the main source of CSF cytokines, because developmental changes observed in plasma did not overlap those observed in CSF.
Considering the proximity of CSF compartments with periventricular and hippocampal stem cell niches which harbor the main cellular targets of cytokines during development, CSF-borne cytokines are likely to fulfill important endocrine signaling. An alteration of the developmental profile of these bioactive molecules in CSF, induced by perinatal injuries such as sepsis, may not only participate in spreading systemic inflammation to the brain, but also directly lead to unbalanced neuronal signaling. This would ultimately alter normal neural network organization and lead to neurodevelopmental diseases [
6,
66,
67]. As bilirubin impacts only marginally CSF cytokine content, it is unlikely that hyperbilirubinemia-induced cerebral dysfunctions involve a similar pathophysiological mechanism. Of note the somewhat large standard errors that can be attributed to both inter- and intra-litter variability, and to the analytical performances of the multiplex method, have limited the statistical power of the study, which would deserve further investigation by analyzing selected cytokines such as those listed in Fig.
5 with more sensitive methods. The cytokine concentrations measured in CSF of NN rats tend to be higher than concentrations found for Nj rats with a mild physiological hyperbilirubinemia, or jj rats presenting high pathological levels of free bilirubin in plasma. This effect could be related to the systemic anti-inflammatory, or immunosuppressive properties attributed to bilirubin [
68‐
70], independently of its toxic effect on neural cells. In line with this we also observed a decrease in innate immune cell infiltration in the CSF of hyperbilirubinemic as compared to normobilirubinemic animals following systemic exposure to a gram negative bacteria lipopeptide. Whether this reflects a direct impact of bilirubin on circulating innate immune cells, especially PMNs, or a change in the choroidal attributes that set the migration of these cells into the CSF remains to be elucidated. Altogether these data suggest that hyperbilirubinemia does not activate choroid plexus and does not induce an important inflammatory response in CSF that could trigger the neurological impairment observed in pathological jaundice.
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