- Record: found
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- Article: found
Maternal high-fat diet in mice induces cerebrovascular, microglial and long-term behavioural alterations in offspring
research-article
Author(s):
Maude Bordeleau
1
,
2 ,
Cesar H. Comin
3 ,
Lourdes Fernández de Cossío
1
,
4 ,
Chloé Lacabanne
1 ,
Moises Freitas-Andrade
5
,
6
,
7 ,
Fernando González Ibáñez
2
,
8
,
9 ,
Joanna Raman-Nair
5
,
6
,
7 ,
Michael Wakem
10 ,
Mallar Chakravarty
1
,
11
,
12
,
13 ,
Luciano da F. Costa
14 ,
Baptiste Lacoste
5
,
6
,
7
,
,
Marie-Ève Tremblay
2
,
8
,
9
,
15
,
16
,
Publication date (Electronic):
11 January 2022
Journal:
Communications Biology
Publisher:
Nature Publishing Group UK
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Abstract
Various environmental exposures during pregnancy, like maternal diet, can compromise, at critical periods of development, the neurovascular maturation of the offspring. Foetal exposure to maternal high-fat diet (mHFD), common to Western societies, has been shown to disturb neurovascular development in neonates and long-term permeability of the neurovasculature. Nevertheless, the effects of mHFD on the offspring’s cerebrovascular health remains largely elusive. Here, we sought to address this knowledge gap by using a translational mouse model of mHFD exposure. Three-dimensional and ultrastructure analysis of the neurovascular unit (vasculature and parenchymal cells) in mHFD-exposed offspring revealed major alterations of the neurovascular organization and metabolism. These alterations were accompanied by changes in the expression of genes involved in metabolism and immunity, indicating that neurovascular changes may result from abnormal brain metabolism and immune regulation. In addition, mHFD-exposed offspring showed persisting behavioural alterations reminiscent of neurodevelopmental disorders, specifically an increase in stereotyped and repetitive behaviours into adulthood.
Abstract
In order to advance our understanding of the effects of maternal high-fat diet (mHFD) on the cerebrovascular health of offspring, Bordeleau et al. use a translational mouse model of mHFD exposure. They demonstrate that mHFD induces cerebrovascular and microglial changes in the offspring as well as behavioural alterations that are reminiscent of neurodevelopmental disorders associated with repetitive behaviours at adulthood.
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Most cited references68
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- Abstract: found
- Article: found
Microglial Interactions with Synapses Are Modulated by Visual Experience
Introduction Upon invasion of the central nervous system during embryonic and early postnatal development, bone-marrow-derived microglia become involved in apoptosis and phagocytic elimination of supernumerary neurons [1]–[3]. As they complete their differentiation, microglia change their morphology from amoeboid to ramified and are thought to become quiescent [4]. In the event of pathological insult, microglia rapidly become activated, thicken and retract their processes, migrate to the site of injury, proliferate, and participate in the presentation of antigens, phagocytosis of cellular debris, and secretion of proteases that promote microglial motility, as well as myelin and extracellular matrix degradation [5]–[7]. Additionally, activated microglia can separate presynaptic axon terminals from postsynaptic neuronal perikarya or dendrites in a process known as synaptic stripping [8]. Even though microglia are quiescent under non-pathological conditions, their highly motile processes continually survey the local environment and make transient contacts with astrocytes, neuronal perikarya, axon terminals, and dendritic spines in vivo [9]–[11]. Microglial apposition with astrocytic and neuronal elements has also been observed with electron microscopy (EM) in situ [9], but a detailed analysis of microglial ultrastructural relationships is still lacking. Reports of spontaneous engulfment of cellular debris [10] suggest that resting microglia may exert phagocytic roles in the healthy brain. Because changes in the level of neuronal activity can also modify the volume of neuropil that microglia sample [10], as well as their frequency of contact with axon terminals [9], Wake et al. [9] proposed that resting microglia could monitor the functional state of synapses. However, aside from immune surveillance, the fate of synaptic architecture under the care of microglia remains poorly understood. The dynamic nature of microglial processes and their interaction with synapses suggest that microglia could effect structural changes at synapses, which are crucial to circuit remodeling and brain plasticity. To begin to investigate this possible task of quiescent microglia at synapses, we verified whether microglial interactions with synapses occur at random or coincide with structural synaptic changes and alterations in sensory experience. Specifically, we characterized the ultrastructural and structural/dynamic interactions between microglia and synapse-associated elements during normal sensory experience, sensory deprivation, and subsequent light exposure in the primary visual cortex (V1) of juvenile mice. In addition to revealing the three-dimensional (3-D) geometry of cell–cell contacts between microglia and all synapse-associated elements (dendritic spines, axon terminals, perisynaptic astrocytic processes, and synaptic clefts), our observations uncovered new modes of microglia–synapse interaction under non-pathological conditions, particularly the regulation of the perisynaptic extracellular space and the phagocytosis of synaptic elements. Moreover, we found that microglia specifically localize to the vicinity of a subset of synaptic elements in vivo, in particular the structurally dynamic and transient dendritic spines. Lastly, we demonstrate that several modalities of microglia–synapse interactions are regulated by sensory experience. Thus, our findings indicate that microglia are not activated only during early brain development or pathological conditions; rather, they also subtly change their behavior toward synapses in correspondence with sensory experience. This raises the intriguing possibility that microglia may contribute to fine-tuning the plastic capacities of individual synapses in the healthy brain. Results Ultrastructural Relationships between Microglia and Synapses To provide a detailed view of the modes of interaction between microglia and excitatory synapses, we analyzed their ultrastructural relationships in layer II of mouse V1 on postnatal day (P) 28, around the peak of the critical period for experience-dependent plasticity [12]. Using immunocytochemical EM with an antibody against the microglia-specific marker IBA1 [13] (see Figure S1 for IBA1 immunostaining at the light microscopic level), we found that microglial cell bodies, as well as proximal and distal processes, juxtaposed synapse-associated elements including synaptic clefts (Figures 1A–1C), an area generally thought to be exclusively reserved for astrocytic processes. Quantitative analysis revealed that the vast majority of microglial process profiles directly contacted at least one of the synapse-associated elements (synaptic index: 94%±0.6%; ∼1,000 µm2 of neuropil in each of three animals). Axon terminals, dendritic spines, perisynaptic astrocytic processes, and synaptic clefts were contacted by microglial processes, in decreasing order of frequency (n = 150 IBA1-positive microglial processes; three animals; see Table S1 for detailed analysis), and more than one synapse-associated element was generally contacted by each process (68%±4%; see Table S2). 10.1371/journal.pbio.1000527.g001 Figure 1 Ultrastructural interactions between microglia and synapses during normal sensory experience. (A–C) EM images showing IBA1-immunostained microglial (m+) cell bodies (A), as well as large (B) and small (C) processes, surrounded by extended extracellular space (asterisks) and contacting axon terminals (blue), dendritic spines (pink), perisynaptic astrocytes (green), and synaptic clefts (arrowheads). d, dentrite; N, nucleus; p, perikaryon. Scale bars = 250 nm. (D) EM image showing extended microglia-associated extracellular spaces (asterisks) after glutaraldehyde instead of acrolein fixation. The unlabeled microglial process (m) makes direct contacts with dendritic spines (s) and axon terminals (t), and displays an inclusion (in), as well as a clathrin-coated pit (black arrow) at the site of contact with a spine. Scale bar = 250 nm. (E) Extracellular space areas with or without contact with IBA1-positive microglial process (mean ± SEM). **, p 0.9; n = 24 terminals and 29 putative contacts in three animals; Figure S7B and S7C), nor correlated to initial terminal size or putative microglial contact duration (p>0.5 and p>0.2, respectively; Figure S7D and S7E). In contrast, 62% of dendritic spines grew, 32% shrank, and 6% remained stable during putative contact; additionally, we observed a significant increase in average spine sizes in the presence versus in the absence of putative microglial contact (size differential: 9%±3%; p 0.9; Figure 3C). No correlation between size change and putative contact duration was noted (p>0.9; Figure S8A), but the size change and initial spine size were significantly correlated (r = 0.28; p 0.3 and p>0.1, respectively). Surprisingly, contacts with synaptic clefts were more frequent in DA animals (p 0.2 versus control; p>0.09 versus DA; n = 3 animals per condition; Figure 4F) while microglia-associated extracellular space area was significantly reduced following light reexposure (p 0.2 versus DA; n = 3 animals per condition; Figure 4E). Similarly, contacts with synaptic clefts (p 0.9 versus DA; n = 3 animals per condition; Figure 4H) and perimeters of contact with dendritic spines (p 0.4 versus DA; Figure 4I) and axon terminals (p 0.6 versus DA), but not with perisynaptic astrocytes (p>0.2 versus control and DA), remained extended. Future experiments with longer light exposures after deprivation will be needed to determine whether these phenomena can be reversed with further light reexposure. Taken together, these observations reveal a complex interaction between sensory-driven activity and microglial behavior. While the expansion of microglial processes and associated extracellular spaces reversed after brief reexposure to daylight, microglial ensheathment of dendritic spines and axon terminals, as well as their phagocytic inclusion, were still increased. Dynamic Relationships between Microglia and Synapses during Alterations in Sensory Experience To assess the dynamic changes in microglia–synapse interactions during visual deprivation, we used two-photon imaging of layers I/II of V1 in juvenile mice that were subjected to DA for 8–10 d, from the beginning to the peak of the critical period [12]. Microglial processes appeared thickened and sparse, and more often terminated into crown-like structures resembling phagocytic cups than in control animals [30] (Figure 5A and 5B; Video S2, as well as Figure S12 and Videos S4 and S5 for comparison of microglial morphology in control and DA animals). We also found that the average motility of microglial processes was significantly reduced (Figure 5C and 5D) when assayed in two ways: comparing morphology over a 5-min interval (motility index; control: 8%±0.8%; DA: 6%±0.5%; p 0.5; Figure 5E), in agreement with an observed reduction in synaptic strength during binocular deprivation [31],[32] and the smaller sizes of dendritic spines during synaptic depression [33],[34]. Surprisingly, dendritic spines putatively contacted by microglia in DA animals were significantly bigger than non-contacted spines (p 0.8), suggesting that increased coverage of synaptic elements by microglia is not a result of longer duration of contact. Similarly, the frequency of putative contacts with individual dendritic spines was unchanged by sensory experience (DA: 1±0.08; control: 1±0.07 contacts per 40 min; p>0.6; Figure S11B). Lastly, most dendritic spines shrank during microglial contact (29% grew, 57% shrank, and 14% remained stable; n = 12 spines and 14 putative contacts in three animals), unlike in the control condition, where most spines grew. While average size changes during contact were not significant (size differential: −4%±4%; p>0.3; Figure S11C), interestingly, dendritic spine shrinkage persisted after microglial contact, with a significant difference in size between before and after the contact (p 0.2 versus control; p 0.1 versus control; p 0.1 versus control and DA; n = 14 contacted spines in three DA+light animals; Figure 5E), while non-contacted spines were significantly bigger in animals reexposed to light (p 0.9; Figure 5E), indicating that microglia no longer localize to specific spine types as in control or DA animals. Lastly, microglia–synapse interactions showed similar structural effects on dendritic spines as in control conditions. Most dendritic spines increased in size during putative microglial contact (67% grew, 20% shrank, and 13% remained stable; average size differential: 13%±5%; p 0.8 comparing size differential before/after contact; Figure 5F). These results reveal additional changes in microglial behavior that can be reversed by brief reexposure to daylight, particularly their motility and preference of contact for subsets of dendritic spines. Discussion Beyond immune surveillance, the physiological roles of quiescent microglia at synapses are unknown. Here we show that different modalities of microglial behavior are subtly altered by sensory experience, including regulation of extracellular spaces, apposition and phagocytosis of synaptic elements, dynamic interaction with subsets of dendritic spines, and motility of microglial processes. Microglial Regulation of Perisynaptic Extracellular Spaces One of the most striking findings of our study was the demonstration of distinctive extracellular spaces closely correlated with the presence of microglial processes. Our EM observations revealed large electron-lucent pockets of extracellular space surrounding microglia. To our further surprise, we also observed changes in the distribution of these microglia-associated extracellular spaces during alterations in visual experience: an expansion during light deprivation and shrinkage during light reexposure. Although alteration of these spaces by brain fixation and embedding for EM may warrant further investigation, we observed them under all conditions tested. In future experiments, it will be important to determine whether microglial processes create this extracellular space themselves or move in to fill space that is created by an unknown mechanism. In any case, our findings of microglia-specific extracellular spaces suggest that microglia have an intimate relationship with their extracellular milieu and may even regulate their surrounding environment in a unique and specific way that is determined by physiological conditions. If microglia directly modulate the extracellular space, they may do this through the secretion of various proteases that degrade extracellular matrix proteins, including cathepsins, metalloproteases, and tissue-type plasminogen activator [35]. This proteolytic degradation of specific matrix proteins could, in turn, facilitate microglial motility, as suggested by the finding that the migratory behavior of cathepsin S–deficient microglia is severely impaired in vitro [35]. In line with this, the volume of extracellular space greatly decreases during postnatal cortical development, concomitant with changes in extracellular matrix composition and reductions in cell migration and process elongation [14]–[16],[36],[37]. The appearance of extracellular spaces specifically associated with microglial processes could therefore reflect their highly motile behavior, relative to other structural elements of neuropil in juvenile cortex. Additionally, regulation of the extracellular matrix composition by microglia-derived proteases could contribute to dendritic spine motility and pruning, as well as activity-dependent and experience-dependent plasticity, which are profoundly affected in vitro and in vivo by treatments with proteases that degrade extracellular matrix proteins [38]–[43]. Microglial Apposition and Phagocytosis of Synaptic Elements Immunocytochemical EM and SSEM with 3-D reconstructions enabled us to analyze the morphology of microglial processes and their ultrastructural relationships with the other subcellular compartments of neuropil—astrocytic processes, axon terminals, and dendritic spines—in situ at high spatial resolution. Building on previous EM observations that microglia contact axon terminals and dendritic spines [9],[44],[45], our quantitative analysis revealed that most microglial processes directly appose not only axon terminals and dendritic spines, but also perisynaptic astrocytic processes and synaptic clefts. Our SSEM with 3-D reconstructions also uncovered the 3-D relationships between microglia and synapses, revealing that microglial processes contact multiple synapse-associated elements at multiple synapses simultaneously. Additionally, we found clathrin-coated pits at interfaces between microglia and dendritic spines, axon terminals, or perisynaptic astrocytic processes, suggesting clathrin-mediated endocytosis of membrane-bound receptors and their ligands, a phenomenon known to initiate various cellular signaling events [46]. Since clathrin-coated pits also occur at the tips of most spinules undergoing invagination, as previously observed in small dendritic spines, axon terminals, and perisynaptic astrocytic processes of mouse hippocampus [47], this may suggest trans-endocytosis of membrane-bound receptors and their ligands, in addition to a direct exchange of cytoplasm, between microglia and synapse-associated elements. To better understand the functional significance of these forms of molecular communication between microglia and synapse-associated elements, it will be important to identify the molecules that are being internalized during their dynamic interactions. Following visual deprivation and reexposure to daylight, our EM results revealed that microglial processes change their morphology, appose synaptic clefts more frequently, and envelop synapse-associated elements more extensively. Since glial presence at the synaptic cleft was classically restricted to astrocytic processes regulating synaptic function through modification of the extracellular space geometry and bidirectional communication with synaptic elements [48],[49], this novel finding suggests that microglia may also contribute uniquely to synaptic transmission and plasticity in the healthy brain. Additionally, the ensheathment of synaptic elements and synaptic clefts by microglial processes may imply their participation in activity-dependent synapse elimination [50], through a separation of pre- and postsynaptic elements reminiscent of synaptic stripping, as previously reported between axon terminals and neuronal cell bodies during immune responses [8]. Lastly, our EM and two-photon in vivo imaging observations revealed that a subpopulation of microglial processes displays phagocytic structures, with an increasing prevalence during alterations in visual experience, thus providing evidence for a microglial role in phagocytic engulfment under non-pathological conditions. This is supported by observations that quiescent microglia can spontaneously engulf tissue components in vivo [10] and that activated microglia play an essential role in phagocytosis of cellular debris [1]–[3],[5],[6]. At the ultrastructural level, we also found cellular inclusions that resembled dendritic spines or axon terminals, suggesting that quiescent microglia can phagocytose synaptic elements in juvenile cortex. In line with this, classical complement proteins C1q and C3 were recently shown to be involved in the pruning of inappropriate retino-geniculate connections during early postnatal development [51]. Since C1q and downstream complement protein C3 can trigger a proteolytic cascade leading to microglial phagocytosis, this finding supports a model in which microglia may contribute to synaptic pruning under non-pathological conditions [52]. Taken together, our findings indicate that distinct modes of microglial interactions with synapses, most notably apposition, ensheathment, and phagocytosis, are subtly regulated by sensory experience. Dynamic Microglial Interaction with Subsets of Dendritic Spines Two-photon visualization of microglia and synaptic elements with two different colors in CX3CR1-GFP/Thy1-YFP mice enabled clear distinction of the separate structures, which facilitated identification of their putative contacts. This approach revealed transient localization of microglial processes to the vicinity of dendritic spines and axon terminals, in agreement with observations from a recent study [9]. In our study, clear distinction of the separate structures also allowed, for the first time, measurement of dendritic spine and axon terminal morphological changes during episodes of proximity with microglial processes. Our results revealed a specificity of these putative microglial contacts for a subset of small, transiently growing, and frequently eliminated dendritic spines. This is consistent with previous findings showing that in mouse visual and somatosensory cortical areas in vivo, small dendritic spines are more motile and more frequently eliminated than their larger counterparts [20],[22],[24],[53],[54]. Dendritic spines undergoing long-term potentiation transiently or persistently enlarge in vitro [33],[34],[55],[56], but since dendritic spine structural changes can occur independently of changes in synaptic strength [57], a connection between putative microglial contact and synaptic plasticity remains to be determined. During light deprivation, microglia preferentially localized to larger dendritic spines that persistently shrank, akin to spines undergoing long-term depression in vitro [33],[34] or shrinking before elimination in vivo [58], while during reexposure, microglia reversed to contacting spines that transiently grew, as in control animals. This uncovers a specificity of microglial interaction for subsets of structurally dynamic and transient dendritic spines, a specificity that, surprisingly, changes with sensory experience. In future experiments, correlating the temporal dynamics of microglial contact with dendritic spine activity could be achieved by in vivo labeling of neurons with electroporated calcium indicators [59] or viral delivery of genetic calcium indicators [60]. A challenge with such techniques will be their invasive nature; the mechanical disruption of the brain alone during indicator delivery can result in microglial activation. Additionally, it will be important to determine whether microglial contacts occur in response to structural changes in dendritic spines and whether such contacts instruct subsequent spine elimination. This will, however, require new technical advances enabling interference with microglial contacts while preserving physiological conditions. Importantly, whether direct microglial contacts, such as observed with EM, are required for these structural changes or whether microglia can exert effects on synapses without close apposition will need to be explored. Motility of Microglial Processes Several lines of evidence, including those presented here, suggest that microglial motility may be regulated by neuronal activity. A pioneer study reported an increase in the volume sampled by quiescent microglia in vivo over a period of 1 h after application of the ionotropic GABA receptor blocker bicuculline, whereas the sodium channel blocker tetrodotoxin had no significant effects [10]. More recently, microglia were shown to retract their processes and reduce their frequency of contact with axon terminals in vivo, over a period of 4–6 h after binocular enucleation or intraocular injection of tetrodotoxin [9]. In our study, two-photon analysis revealed a global decrease in microglial motility during light deprivation without correlated changes in the duration or frequency of microglial contact with individual dendritic spines. This may highlight differences between short-term (1–6 h) and long-term (8–10 d) microglial responses to sensory deprivation or distinguish between more invasive manipulations, which approximate nervous system injury, and more physiological paradigms such as DA. Our observations further suggest that subsets of microglial processes may have different behavior: those in contact with dendritic spines may be highly motile, while others (for example bulky microglial processes displaying phagocytic structures) may become less motile. It is also possible that spindly microglial processes ( 0.2 µm) from the axon or bead-like structures along the axon (at least two times the axon diameter), as previously described [54]. While dendritic spine morphology exists as a continuum, different behaviors have been assigned to spines with different structures [71]. Therefore we classified dendritic spines into mushroom, thin, and stubby types based on their length and spine head volume, as previously described [24]. The only three stubby spines contacted by microglia were excluded from the analysis of spine size because of their rarity and the difficulty of measuring their fluorescence intensity. Consequently, we also removed stubby spines from the analysis of size for non-contacted and contacted dendritic spines. Among thin and mushroom spines, we determined a range of normalized dendritic spine sizes. The spines belonging to the first quarter of this range were considered “small,” and those in the other three-quarters were identified as “large.” Filopodia were rarely observed at these ages and were excluded from the analysis. For visualization of microglial contacts with dendritic spines and axon terminals, the green channel was arbitrarily assigned the color red and the yellow channel assigned the color green, enabling the visualization of microglia in yellow and neuronal elements in green (see Figure S6). Microglial contacts were identified manually by stepping through the Z stack without projection. All microglial contacts (colocalization of fluorescence for microglia and synaptic elements) that started and ended during imaging were included in the analysis. For analysis of changes in synaptic structures, the background was subtracted from each of the two channels and the green channel bleedthrough was subtracted from the yellow channel (see Figure S6 for uncorrected and corrected images). In the stacks unadjusted for brightness or contrast, dendritic spines and axon terminals were analyzed for fluorescence at the Z level where they appeared brightest. A line was traced through each element, and a fluorescence plot profile was created, which was then fitted to a Gaussian. Because the majority of dendritic spines are below the resolution of our two-photon microscope [72], the maximal fluorescence (amplitude of the Gaussian fit) was used to assess the relative spine size (see Figure S8C for sizes of large spines assessed with the width of the fluorescence profile [1/e 1/2 radius of the Gaussian fit] to rule out underestimation of their size changes during putative contact). Since axon terminals are generally much larger than dendritic spines, the width of the Gaussian fit to the fluorescence profile was used to determine their relative size. To rule out contamination of neuronal measurements by any unsubtracted bleedthrough of microglial fluorescence, we analyzed thin fluorescent axons (n = 1 axon in each of 14 animals) that were contacted by microglia, using the height of the Gaussian fit to assess relative size. Axon size measured in this manner was relatively unchanged between populations with and without microglial contact (average size differential of −0.1%±1%; see Figure S6D and S6E), as expected for these structurally stable elements. Size differentials of terminals and spines were calculated as the ratio of size difference with and without contact over the size without contact. In order to compare spine size between animals, we normalized spine fluorescence by the maximal fluorescence in the adjacent dendrite. Axon terminals and dendritic spines with size differentials under 1% were considered stable. For presentation purposes, we normalized the size of terminals and spines by dividing each individual structure's average size in the presence of microglial contact (with or during) by its average size in the absence of microglial contact (without, or before or after). We also normalized the size of terminals and spines by dividing each individual structure's average size by the size of the largest terminal or spine. To determine the turnover of dendritic spines that were or were not contacted by microglia in the first imaging session, the position of individual dendritic spines was compared between time points separated by 2 d. The proportion of eliminated dendritic spines was defined as the proportion of spines from the original population not observed on the second day of imaging. Spines located more than 0.8 µm laterally from their previous location were considered to be new spines. For measurement of microglial motility, images centered on the cell body from five consecutive Z levels were projected into two dimensions, for each microglia and each time point analyzed (0, 5, and 25 min). For each microglia, the images were aligned and grouped into a stack. Stacks were adjusted for brightness and contrast, and then binarized. For each microglia, the difference between images taken at the 0- and 5- or 25-min time points was calculated. A motility index was determined, as the proportion of the pixels that differed between the two images. Statistical Analysis Analyses were performed with Prism 5 software (GraphPad Software). All values reported in the text are mean ± standard error of the mean (SEM). For all statistical tests, significance was set to p<0.05. Two-tailed unpaired Student's t tests and linear regressions were used for both EM and two-photon analyses. For two-photon analyses, two-tailed paired Student's t tests were also used to compare the size of the same dendritic spines or axon terminals in the presence versus in the absence of microglial contact. Sample size (n) represents individual animals for EM (except for correlation analysis, where n represents individual extracellular space and microglial process areas) and synaptic elements or microglial contacts for two-photon analyses (except for analysis of dendritic spine turnover, where n represents individual animals). Supporting Information Figure S1 Light microscopic image showing immunoperoxidase staining for IBA1, under the same immunocytochemical conditions as used for EM. The staining is restricted to microglia, which shows its specificity. Scale bar = 50 µm. (0.88 MB TIF) Click here for additional data file. Figure S2 Other views of the 3-D reconstruction that further reveal the geometry of the microglia-associated extracellular spaces. In (A), axon terminals (blue), dendritic spines (red), and perisynaptic astrocytes (green) are made semitransparent. Taupe indicates microglia. In (B), the axon terminals are removed from the display, while in (C) only the microglial process and extracellular space are shown. Scale bars = 250 nm. (1.45 MB TIF) Click here for additional data file. Figure S3 SSEM images showing additional examples of coated pits at the sites of cell–cell contact between microglia and synapse-associated elements. In these examples, the vesicles that appear coated (black arrows) are found inside a microglial process (m), at the sites of contact with an astrocytic process (a) (A), a dendritic spine (s) (B), and an axon terminal (t) (C). d, dendrite; N, nucleus; p, perikaryon. Scale bar = 250 nm. (2.05 MB TIF) Click here for additional data file. Figure S4 SSEM images from an animal undergoing normal visual experience, showing the phagocytic engulfment of cellular debris by a microglial process. Images are separated by 65 nm. *, extracellular space; a, astrocyte; d, dendrite; in, cellular inclusion; m, microglial process; N, nucleus; p, perikaryon; s, dendritic spine; t, axon terminal. Scale bars = 250 nm. (4.02 MB TIF) Click here for additional data file. Figure S5 Z projections showing the morphology of microglia in brain sections of animals perfused after two-photon in vivo imaging. The imaged area is shown in (A), and the corresponding contralateral area is shown in (B). The pial surface is presented at the top of the image in both cases. The similar polarity, thickness, and density of microglial cell bodies and processes (green) confirm that microglia are not activated by transcranial imaging. Scale bar = 15 µm. (1.47 MB TIF) Click here for additional data file. Figure S6 Separation of GFP and YFP fluorescence in CX3CR1-GFP/Thy1-YFP mice. (A) Two-photon images from the yellow channel (YFP+GFP; left) and green channel (GFP; right) in their uncorrected state. (B) Two-photon image from the yellow channel corrected with subtraction of background and GFP fluorescence for analysis of axon terminal and dendritic spine sizes. (C) Merge of the yellow and green channels (assigned the colors green and red, respectively) and adjustment of brightness and contrast for visualization of microglial contacts (yellow) with neuronal elements (green). (D) Two-photon image showing a contact between a microglial process (yellow) and a small axon (green; white arrowhead) during normal sensory experience. (E) Axon size without versus with microglia contact, normalized to the first condition for presentation purposes. au, arbitrary units. (1.42 MB TIF) Click here for additional data file. Figure S7 Structural/dynamic interactions between microglia and axon terminals during normal visual experience in vivo. (A) Time-lapse image showing an axon terminal (green; blue arrowhead) contacted by microglial processes with bulbous endings (yellow; white arrowhead) over 20 min. Scale bar = 5 µm. (B and C) Axon terminal size without versus with microglial contact (B) or before, during, and after contact (C), normalized to the first condition for presentation purposes. (D and E) Lack of correlation between microglial contact duration or initial terminal size (normalized to largest axon terminal) and the change in axon terminal size during microglial contact. au, arbitrary units. (1.17 MB TIF) Click here for additional data file. Figure S8 Additional analysis of structural/dynamic interactions between microglia and dendritic spines during normal visual experience in vivo. (A) Lack of correlation between microglial contact duration and the change in dendritic spine size during contact. (B and C) Change in the size of large dendritic spines assessed with the amplitude of the Gaussian fit to the fluorescent profile (as for other analyses of dendritic spine size presented in the Results section) or with the width of the fluorescent profile (1/e 1/2 radius of the Gaussian fit; see Materials and Methods section), confirming that the size changes of bigger spines were not underestimated with assessments of maximal fluorescence. (0.13 MB TIF) Click here for additional data file. Figure S9 Additional analysis of ultrastructural interactions between microglia and synapse-associated elements during altered visual experience. (A) EM image taken in a DA animal showing a microglial (m+) perikarya that contains vacuole (vac) and cellular inclusions (in). *, extracellular space; ma, myelinated axon; N, nuleus. Scale bar = 250 nm. (B) Correlation between the areas of microglial processes and associated extracellular space in DA animals. (C) Total number of IBA1-immunopositive microglial processes in a surface of 1,000 µm2 of neuropil, in control versus DA animals (n = 3 animals per experimental condition; mean ± SEM). (D) Synaptic index in control versus DA animals (n = 3 animals per experimental condition; mean ± SEM). (E) Proportion of simultaneous microglial contacts with one, two, or three synapse-associated elements (n = 3 control and 3 DA animals; mean ± SEM). (1.17 MB TIF) Click here for additional data file. Figure S10 EM images showing additional examples of microglial processes contacting multiple synapse-associated elements, including synaptic clefts, in the different experimental conditions. a, perisynaptic astrocytic process; d, dendrite; m, microglial process, s, dendritic spine; t, axon terminal. White arrowheads indicate synaptic clefts. Scale bars = 250 nm. In (A–C; control animals), the small microglial processes (m+) are devoid of cellular inclusions and surrounded by narrow extracellular space. In (D–F; DA animals), larger microglial processes (m+) display bulky (D and E) or spindly (F) morphologies. While the bulky processes contain cellular inclusions resembling profiles of dendritic spine (“s”), axon terminal (“t”), or cellular membranes (“cm”), the spindly process is surrounded by extended extracellular space. In (G–I; DA+light animals), the large-to-small microglial processes display vacuole (vac) and cellular inclusions resembling terminals (“t”) as well as a synapse between a dendritic spine (“s”) and a terminal (“t”). Little extracellular space is observed. (7.32 MB TIF) Click here for additional data file. Figure S11 Additional analysis of structural/dynamic interactions between microglia and dendritic spines during altered visual experience in vivo. (A and B) Duration and frequency of microglial contacts with individual dendritic spines during 40-min imaging sessions, in DA and control animals (mean ± SEM). (C) Dendritic spine size without versus with microglial contact in DA animals (left) and DA+light animals (right), normalized to the first condition for presentation purposes. *, p<0.05. (0.14 MB TIF) Click here for additional data file. Figure S12 Time-lapse images showing additional examples of microglial morphology and motility in the different experimental conditions. Images from control (A), DA (B), and DA+light (C) animals are shown. In (B), note the thickening of microglial processes, which also are sparse. Scale bars = 10 µm. See also Videos S4–S6. (6.14 MB TIF) Click here for additional data file. Table S1 Changes in the ultrastructural interactions between microglia and synapse-associated elements with visual experience. (0.21 MB TIF) Click here for additional data file. Table S2 Changes in the ultrastructural interactions between microglia and different combinations of synapse-associated elements with visual experience. (0.14 MB TIF) Click here for additional data file. Table S3 Diversity in microglia-associated extracellular space volumes. (0.16 MB TIF) Click here for additional data file. Video S1 Time-lapse video showing the structural/dynamic interactions between microglia and dendritic spines during normal visual experience in vivo. Multiple contacts between microglial processes displaying bulbous endings (yellow) and dendritic spines (green), as well as contacts with dendritic branches (green), are observed over 1 h. Images were acquired at the same depth, approximately 50 µm below the pial surface. (7.75 MB AVI) Click here for additional data file. Video S2 Z stack showing examples of microglial processes ending in phagocytic cups during DA in vivo. Images of microglia (yellow) were acquired 1 µm apart, starting approximately 50 µm below the pial surface. Two phagocytic cups are shown (arrows), with the smallest one surrounding a YFP-positive neuronal element (green). (0.77 MB AVI) Click here for additional data file. Video S3 Z stack showing examples of microglial processes ending in phagocytic cups in a DA+light animal in vivo. Images were acquired 1 µm apart, starting approximately 50 µm below the pial surface. Three phagocytic cups are shown (arrows), with the small one at the left surrounding a YFP-positive neuronal element (green). (0.97 MB AVI) Click here for additional data file. Video S4 Time-lapse video showing the morphology and motility of microglia during normal visual experience in vivo. Five microglia (yellow) contacting multiple axons (green) and axon terminals (green) are shown. Note the presence of a small structure resembling a phagocytic cup (arrow) near YFP-positive axonal elements (green). Images were acquired at the same depth, approximately 50 µm below the pial surface, every 5 min for 30 min. (0.32 MB AVI) Click here for additional data file. Video S5 Time-lapse video showing the morphology and motility of microglia during DA in vivo. Four microglia (yellow) displaying sparse processes in the neuropil are shown. Images were acquired at the same depth, approximately 50 µm below the pial surface, every 5 min for 30 min. (0.48 MB AVI) Click here for additional data file. Video S6 Time-lapse video showing the morphology and motility of microglia during DA followed by reexposure to light in vivo. Three microglia (yellow) contacting multiple dendrites (green) and dendritic spines (green) are shown. Images were acquired at the same depth, approximately 50 µm below the pial surface, every 5 min for 30 min. (0.76 MB AVI) Click here for additional data file.
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- Abstract: found
- Article: not found
The neurocircuitry of fear, stress, and anxiety disorders.
Lisa Shin, Israel Liberzon (2010)
Anxiety disorders are a significant problem in the community, and recent neuroimaging research has focused on determining the brain circuits that underlie them. Research on the neurocircuitry of anxiety disorders has its roots in the study of fear circuits in animal models and the study of brain responses to emotional stimuli in healthy humans. We review this research, as well as neuroimaging studies of anxiety disorders. In general, these studies have reported relatively heightened amygdala activation in response to disorder-relevant stimuli in post-traumatic stress disorder, social phobia, and specific phobia. Activation in the insular cortex appears to be heightened in many of the anxiety disorders. Unlike other anxiety disorders, post-traumatic stress disorder is associated with diminished responsivity in the rostral anterior cingulate cortex and adjacent ventral medial prefrontal cortex. Additional research will be needed to (1) clarify the exact role of each component of the fear circuitry in the anxiety disorders, (2) determine whether functional abnormalities identified in the anxiety disorders represent acquired signs of the disorders or vulnerability factors that increase the risk of developing them, (3) link the findings of functional neuroimaging studies with those of neurochemistry studies, and (4) use functional neuroimaging to predict treatment response and assess treatment-related changes in brain function.
- Record: found
- Abstract: found
- Article: found
Microglia and Beyond: Innate Immune Cells As Regulators of Brain Development and Behavioral Function
Kathryn Lenz, Lars H Nelson (2018)
Innate immune cells play a well-documented role in the etiology and disease course of many brain-based conditions, including multiple sclerosis, Alzheimer’s disease, traumatic brain and spinal cord injury, and brain cancers. In contrast, it is only recently becoming clear that innate immune cells, primarily brain resident macrophages called microglia, are also key regulators of brain development. This review summarizes the current state of knowledge regarding microglia in brain development, with particular emphasis on how microglia during development are distinct from microglia later in life. We also summarize the effects of early life perturbations on microglia function in the developing brain, the role that biological sex plays in microglia function, and the potential role that microglia may play in developmental brain disorders. Finally, given how new the field of developmental neuroimmunology is, we highlight what has yet to be learned about how innate immune cells shape the development of brain and behavior.
Author and article information
Contributors
Baptiste Lacoste:
ORCID: http://orcid.org/0000-0002-1922-8801
blacoste@uottawa.ca
Marie-Ève Tremblay:
ORCID: http://orcid.org/0000-0003-2863-9626
evetremblay@uvic.ca
Journal
Journal ID (nlm-ta): Commun Biol
Journal ID (iso-abbrev): Commun Biol
Title:
Communications Biology
Publisher:
Nature Publishing Group UK
(London
)
ISSN
(Electronic):
2399-3642
Publication date
(Electronic):
11
January
2022
Publication date PMC-release: 11
January
2022
Publication date Collection: 2022
Volume: 5
Electronic Location Identifier: 26
Affiliations
[1
]GRID grid.14709.3b, ISNI 0000 0004 1936 8649, Integrated Program in Neuroscience, , McGill University, ; Montreal, QC Canada
[2
]GRID grid.23856.3a, ISNI 0000 0004 1936 8390, Neurosciences Axis, CRCHU de Québec-Université Laval, ; Québec, QC Canada
[3
]GRID grid.411247.5, ISNI 0000 0001 2163 588X, Department of Computer Science, , Federal University of São Carlos, ; São Carlos, SP Brazil
[4
]GRID grid.266100.3, ISNI 0000 0001 2107 4242, Department of Neurosciences, , University of California, La Jolla, ; San Diego, CA USA
[5
]GRID grid.28046.38, ISNI 0000 0001 2182 2255, Department of Cellular and Molecular Medicine, Faculty of Medicine, , University of Ottawa, ; Ottawa, ON Canada
[6
]GRID grid.28046.38, ISNI 0000 0001 2182 2255, University of Ottawa Brain and Mind Research Institute, ; Ottawa, ON Canada
[7
]GRID grid.412687.e, ISNI 0000 0000 9606 5108, Ottawa Hospital Research Institute, , Neuroscience Program, ; Ottawa, ON Canada
[8
]GRID grid.23856.3a, ISNI 0000 0004 1936 8390, Département de médecine moléculaire, , Université Laval, ; Québec, QC Canada
[9
]GRID grid.143640.4, ISNI 0000 0004 1936 9465, Division of Medical Sciences, , University of Victoria, ; Victoria, BC Canada
[10
]GRID grid.472783.d, Genetic Sciences Division, , Thermo Fisher Scientific, ; Burlington, ON Canada
[11
]GRID grid.14709.3b, ISNI 0000 0004 1936 8649, Cerebral Imaging Center, Douglas Mental Health University, , McGill University, ; Montréal, QC Canada
[12
]GRID grid.14709.3b, ISNI 0000 0004 1936 8649, Department of Psychiatry, , McGill University, ; Montréal, QC Canada
[13
]GRID grid.14709.3b, ISNI 0000 0004 1936 8649, Department of Biological and Biomedical Engineering, , McGill University, ; Montréal, QC Canada
[14
]GRID grid.11899.38, ISNI 0000 0004 1937 0722, São Carlos Institute of Physics, , University of São Paulo, ; São Carlos, SP Brazil
Author information
Baptiste Lacoste
http://orcid.org/0000-0002-1922-8801
Marie-Ève Tremblay
http://orcid.org/0000-0003-2863-9626
Article
Publisher ID: 2947DOI: 10.1038/s42003-021-02947-9
PMC ID: 8752761
PubMed ID: 35017640
SO-VID: 1d359ff0-c17a-4e13-ac18-f9fa8dff04e7
Copyright © © The Author(s) 2022
License:
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.
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