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Keywords:

  • actin;
  • astrocyte;
  • cytoskeleton;
  • GFAP;
  • intermediate filaments;
  • microtubules;
  • mobility;
  • vesicle;
  • vimentin

Abstract

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References

Exocytotic vesicles in astrocytes are increasingly viewed as essential in astrocyte-to-neuron communication in the brain. In neurons and excitable secretory cells, delivery of vesicles to the plasma membrane for exocytosis involves an interaction with the cytoskeleton, in particular microtubules and actin filaments. Whether cytoskeletal elements affect vesicle mobility in astrocytes is unknown. We labeled single vesicles with fluorescent atrial natriuretic peptide and monitored their mobility in rat astrocytes with depolymerized microtubules, actin, and intermediate filaments and in mouse astrocytes deficient in the intermediate filament proteins glial fibrillary acidic protein and vimentin. In astrocytes, as in neurons, microtubules participated in directional vesicle mobility, and actin filaments played an important role in this process. Depolymerization of intermediate filaments strongly affected vesicle trafficking and in their absence the fraction of vesicles with directional mobility was reduced.

Astrocytes are the most abundant glial cells in the central nervous system and have important roles in neuronal function (1). They maintain local ion concentrations and pH homeostasis in the extracellular space, supply neurons with nutrients and remove byproducts of metabolism and neurotransmitters released into the synaptic cleft (1,2). Astrocytes also regulate synapse development, modulate synaptic strength and probably help regulate brain microcirculation (1,3,4). Understanding how astrocytes communicate with neighboring cells is therefore of fundamental importance.

Astrocytes store and release many neuroactive substances, such as chemical transmitters, eicosanoids, steroids, neuropeptides and growth factors (2,5). Some are lipophylic and are released from cells by diffusion through the plasma membrane. Others are transported and released by membrane-bound vesicles (2). Some neuroactive substances in membrane-bound vesicles may be released by Ca2+-dependent exocytosis (5–11), but the mechanism of vesicle transport through the cytoplasm to exocytotic sites remains poorly explained. Exocytotic vesicles in astrocytes show two types of mobility (12): nondirectional mobility, which probably involves free diffusion, and directional mobility, which may involve the cytoskeleton, as in secretory cells (13).

The cytoskeleton and its regulatory and signaling molecules and molecular motors are important in organelle transport within eukaryotic cells (14). Transport along microtubules is relatively rapid in neurons (15,16) and is reported to be comparable in astrocytes (12), but whether this process involves cytoskeleton has not been addressed directly in astrocytes.

Intracellular vesicles can be rapidly transported by motor proteins along a network of microtubules and actin filaments (17). In addition to these components, the cytoskeleton in astrocytes consists of intermediate filaments, which are composed of glial fibrillary acidic protein (GFAP), vimentin and nestin in reactive astrocytes and of GFAP and vimentin in nonreactive astrocytes in the adult (18). In mature astrocytes, GFAP and vimentin are the main cytoskeletal components (19). Unlike actin filaments and microtubules, intermediate filaments lack enzymatic activity and are the least understood part of the cytoskeleton (14,18,19). They form extensive networks in the cytoplasm of most vertebrate cells (14), including astrocytes (19). Their known roles include maintaining the mechanical stability and shape of the cell and providing a scaffold for the organization of the cytoplasm and organelles (20). Intermediate filaments also contribute to astrocyte motility and activation and to the early and late stages of reactive gliosis after brain or spinal cord injury (21–23). In mice deficient in GFAP and vimentin (GFAP−/−Vim−/−), reactive gliosis is attenuated, and various aspects of central nervous system regeneration are improved (19,21,23–28).

To assess the role of the cytoskeleton in astrocytic vesicle mobility, we used fluorescently tagged atrial natriuretic peptide (ANP), proANP-Emd (29), to label single exocytotic vesicles containing native ANP (7,10,12) in rat astrocytes in which individual cytoskeletal components had been pharmacologically depolymerized. These studies confirmed the role of microtubules in directional vesicle mobility and showed that actin filaments are important in this process. Studies in rat astrocytes and astrocytes from GFAP−/−Vim−/− mice also showed that intermediate filaments determine the fraction of vesicles with directional mobility.

Results

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References

Increased [Ca2+]i reduces vesicle numbers but does not influence vesicle mobility

To confirm that an increase in intracellular calcium concentration ([Ca2+]i) triggers the release of proANP-Emd-labeled vesicles from astrocytes (7), we stimulated cells with ionomycin, which evoked a 1- to 2-μm increase in [Ca2+]i. Within 1 min, the percentage of fluorescently labeled vesicles was significantly less than in unstimulated cells (data not shown). However, the mean velocity of vesicles was similar before and after stimulation (0.29 ± 0.01 and 0.27 ± 0.01 μm/second, respectively; n = 167).

Microtubule disassembly predominantly affects directional mobility

To address the role of microtubules in vesicle transport, we began by treating rat astrocytes with nocodazole to depolymerize microtubules (30). Incubation with 200 μm nocodazole for 1 h at room temperature efficiently depolymerized microtubules (Figure 1D) apparently without noticeably altering the shape of the cells (Figure 1A,B).

image

Figure 1. Depolymerizing microtubules with nocodazole reduces vesicle mobility in astrocytes. Transmission light microscopic images of a control astrocyte (−Noc.) (A) and a nocodazole-treated rat astrocyte (+Noc.) (B). Nocodazole (200 μm, 1 h, room temperature) did not alter cell shape. (C and D) The same cells, labeled with monoclonal antibody against mouse α-tubulin and secondary antibody against mouse IgG. Bars: 10 μm. (E and F) Circles denote vesicles whose mobility was plotted for 15 seconds. Bars: 2.5 μm. In control cells (E), vesicles showed both directional mobility (track appears almost rectilinear) and nondirectional mobility (track appears as a dot). In cells with depolymerized microtubule filaments (F), vesicles showed only nondirectional mobility. Relationship between maximal displacement and length of vesicle tracks (directionality index) in control cells (G) and in nocodazole-treated cells (H). The populations of directional and nondirectional vesicles are indicated by lines fitted to the data. The slopes of the lines fitted were 0.4765 ± 0.0641 for directional vesicles (above dashed line) and 0.0700 ± 0.0083 for nondirectional vesicles (below dashed line) (G). n denotes the number of analyzed vesicles. Dashed line represents 1 μm.

In rat astrocytes transfected with the proANP-Emd plasmid (7,12,29) and incubated 24 h later with nocodazole, vesicles showed only nondirectional mobility (Figure 1F). Vesicles in untreated control cells, however, showed both directional and nondirectional mobility (Figure 1E). Vesicles were characterized as directional and nondirectional as reported previously (12). Further analysis showed that microtubule depolymerization significantly reduced the mobility of vesicles (Figure 1G,H), including their average velocity, track length (the length of the pathway that vesicle traveled in 15 seconds) and maximal displacement (a measure for the net translocation of vesicles in 15 seconds) (Figure 4).

image

Figure 4. Depolymerization of microtubules, actin filaments and intermediate filaments significantly reduces vesicle mobility in rat astrocytes. A) Mean velocity of vesicles was 0.30 ± 0.0003 μm/second in cells with depolymerized microtubules (+Noc.), 0.08 ± 0.01 μm/second in cells with depolymerized actin filaments (+CST), 0.21 ± 0.01 μm/second in cells with depolymerized intermediate filaments (+Cal A) and 0.39 ± 0.01 μm/second in controls. B) Mean track lengths of vesicles were 4.42 ± 0.05 μm in cells with depolymerized microtubules, 1.12 ± 0.12 μm in cells with depolymerized actin filaments, 3.10 ± 0.13 μm in cells with depolymerized intermediate filaments and 5.84 ± 0.102 μm in controls. C) Maximal displacement of vesicles was 0.22 ± 0.002 μm in cells with depolymerized microtubules, 0.22 ± 0.01 μm in cells with depolymerized actin filaments, 0.47 ± 0.03 μm in cells with depolymerized intermediate filaments and 1.10 ± 0.08 μm in controls. (A–C) *p < 0.001, **p < 0.0001, ***p < 0.00001 versus control. Numbers in the columns represent the number of analyzed vesicles.

Actin filaments contribute to vesicle mobility

To examine the role of actin filaments, we treated astrocytes with Clostridium spiroforme toxin (CST). Actin filaments were efficiently depolymerized by incubation with 15 nm concentrations of each CST subunit (Sa and Sb) for 1 h at room temperature (Figure 2D). This treatment altered the shape of the cells (Figure 2A,B). In astrocytes transfected with proANP-Emd and treated 24 h later with CST, depolymerization of actin filaments significantly reduced directional mobility (Figure 2E,F) and the average velocity, track length and maximal displacement of the vesicles (Figure 4).

image

Figure 2. Depolymerizing actin filaments with CST reduces vesicle mobility in rat astrocytes. Transmission light microscopic images of a control rat astrocyte (−CST) (A) and a CST-treated rat astrocyte (+CST) (B). Treatment with CST (Sa and Sb: 15 nm, 1 h, room temperature) altered the shape of the cell. Fluorescence micrographs showing actin filaments labeled with rhodamine-phalloidin (red) in the control rat astrocyte (C) and in the CST-treated astrocyte (D). Bars: 10 μm. Relationship between maximal displacement and track length (directionality index) in control vesicles (E) and in CST-treated cells (F). The slopes of the lines fitted were 0.4762 ± 0.0755 for directional vesicles (above dashed line) and 0.0701 ± 0.010 for nondirectional vesicles (below dashed line) (E). n denotes the number of analyzed vesicles. Dashed line represents 1 μm.

Intermediate filaments affect the directional mobility of vesicles

Although they are not thought to participate in vesicle transport, intermediate filaments interact with microtubules (14) and play an important role in astrocyte motility (22). To determine if they contribute to vesicle mobility, we depolymerized intermediate filaments in rat astrocytes with the phosphatase inhibitor calyculin A (Cal A), as described by Chang and Goldman (14). Efficient depolymerization was achieved by incubation with 20 nm Cal A for 20 min at room temperature (Figure 3D). In these conditions, the structure of microtubules and actin filaments appeared nonaffected (data not shown), similarly as observed previously (31). This treatment altered the shape of the cells (Figure 3A,B) and significantly reduced vesicle mobility (Figure 3E,F). The average velocity, track length and maximal displacement of vesicles were all significantly lower in cells with depolymerized intermediate filaments (Figure 4).

image

Figure 3. Depolymerization of intermediate filaments with Cal A reduces vesicle mobility in rat astrocytes. (A and B) Transmission light microscopic images of a control rat astrocyte (−Cal A) and a Cal-A-treated astrocyte (+Cal A). Incubation with Cal A (20 nm, 15 min, room temperature) affected cell shape. The treated astrocyte appears to be smaller. (C and D) The same cells, labeled with a polyclonal antibody against vimentin and secondary antibody against rabbit IgG to show intermediate filaments. Bars: 10 μm. Relationship between maximal displacement and vesicle track length (directionality index) in control cells (E) and in Cal-A-treated cells (F). The slopes of the lines fitted were 0.4762 ± 0.0755 for directional vesicles (above dashed line) and 0.0701 ± 0.010 for nondirectional vesicles (below dashed line) (E). n denotes the number of analyzed vesicles. Dashed line represents 1 μm.

Because the reduced mobility could reflect a nonspecific action of Cal A that affected the phosphorylation of other substrates (14), we analyzed proANP-Emd-labeled vesicles in astrocytes from GFAP−/−Vim−/−mice, which are devoid of intermediate filaments (18) and from matched wild-type mice (Figure 5). In wild-type and GFAP−/−Vim−/− astrocytes, the mean track lengths of directional vesicles were similar [5.97 ± 0.11 μm (n = 313) versus 5.86 ± 0.11 μm (n = 229), respectively] to those of nondirectional vesicles [4.31 ± 0.05 μm (n = 635) versus 4.33 ± 0.04 (n = 688), respectively]. In both groups, the mean maximal displacements for directional vesicles were almost identical in wild-type (2.22 ± 0.07 μm) and in GFAP−/−Vim−/− astrocytes (2.20 ± 0.08 μm) and similarly for nondirectional vesicles, 0.37 ± 0.01 μm in wild-type and 0.35 ± 0.01 μm in GFAP−/−Vim−/− astrocytes. However, the fraction of vesicles showing directional mobility was lower in GFAP−/−Vim−/− astrocytes [21 versus 34%, as in rat astrocytes (12)]. The fraction of directional vesicles calculated per cell was 34 ± 4% (n = 15) in wild-type astrocytes and 21 ± 4% (n = 15, p = 0.03) in GFAP−/−Vim−/− astrocytes.

image

Figure 5. The absence of intermediate filaments reduces the percentage of vesicles showing directional mobility in mouse astrocytes. Relationship between maximal displacement and track length of vesicles in wild-type astrocytes (A) and GFAP−/−Vim−/− astrocytes (B). In 34% of wild-type (WT) astrocytes, but only 21% of GFAP−/−Vim−/− astrocytes, maximal displacement was >1 μm (dashed line). Frequency histograms of maximal displacement of vesicles in astrocytes from wild-type mice (C) and GFAP−/−Vim−/− mice (D). Horizontal lines represent the intersection of the cumulative plot (curves) with the maximal displacement threshold (1 μm, dashed line). The fraction of vesicles with maximal displacement longer than 1 μm is smaller in GFAP−/−Vim−/− cells. Frequency histograms of the directionality index of vesicles in astrocytes from wild-type mice (E) and in cells from GFAP−/−Vim−/− mice (F). The fraction of vesicles with a higher directionality index is smaller in GFAP−/−Vim−/− cells. (C–F) Ordinates on the right show cumulative plots of histograms that allow the determination of the fraction of vesicle mobilities with maximal displacements >1 μm or directionality indexes >0.2. n denotes the number of analyzed vesicles; n d, directional vesicles; n n, nondirectional vesicles.

Next, we explored the mechanism for the influence of intermediate filaments on vesicle mobility. Because intermediate filaments and microtubules interact (14), we reasoned that depolymerization of microtubules with nocodazole would have a greater effect on vesicle mobility in wild-type astrocytes than in GFAP−/−Vim−/− astrocytes. Analysis of proANP-Emd-labeled vesicles showed that the average track lengths were similar in GFAP−/−Vim−/− and wild-type astrocytes (Figure 6A). After treatment with nocodazole, the mean track length was significantly reduced to a similar level in both groups (Figure 6A). The maximal displacement of vesicles was also reduced to a similar extent in both groups (Figure 6A). These results showed no difference in vesicle mobility following the depolymerization of microtubules in both types of cells.

image

Figure 6. Depolymerization of microtubules with nocodazole reduces vesicle mobility in astrocytes from wild-type and GFAP−/−Vim−/− mice. A) Average track lengths were similar in untreated wild-type (4.64 ± 0.06 μm) and GFAP−/−Vim−/− (4.51 ± 0.05 μm) astrocytes. After depolymerization of microtubules, mean track lengths were reduced to similar levels in wild-type (3.22 ± 0.06 μm) and GFAP−/−Vim−/− (3.10 ± 0.02 μm) astrocytes. **p < 0.00001. B) Mean maximal displacement of vesicles in 15 seconds was 1.02 ± 0.04 μm in wild-type (WT) cells and 0.87 ± 0.03 μm in GFAP−/−Vim−/− cells. *p < 0.01. After depolymerization of microtubules with nocodazole (+Noc.), mean maximal displacement was significantly reduced in both groups of cells (to 0.26 ± 0.01 μm in wild-type and to 0.33 ± 0.01 μm in GFAP−/−Vim−/− cells (**p < 0.00001). (A and B) Numbers in the columns represent the number of analyzed vesicles.

Next, we addressed whether genetic ablation of astrocyte intermediate filaments triggered a compensatory response from the microtubular system. We used quantitative real-time polymerase chain reaction (rtPCR) to assess the expression of microtubule-associated protein-2 (MAP-2), which was previously implicated in reactive gliosis (32). MAP-2 expression was 52% higher at postnatal day 1 (P1) GFAP−/−Vim−/− brains (p < 0.005) and 45% higher in primary astrocyte-enriched cultures prepared from P1 GFAP−/−Vim−/− brains (p < 0.01) compared with wild-type brains and astrocyte cultures, respectively. To establish whether the elevated MAP-2 expression in GFAP−/−Vim−/− persists in adult animals in a pathological context, we used the electrically induced cortical lesion as a neurotrauma model. Four days after injury in the tissue surrounding the lesion, MAP-2 expression was 64% higher in GFAP−/−Vim−/− mice compared with wild-type mice (p < 0.05). Thus, genetic ablation of astrocyte intermediate filaments seems to have a specific effect on the microtubular system.

Discussion

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References

This study shows that cytoskeletal components fulfill important functions in vesicle transport within astrocytes. As in neurons, microtubules were important in directional mobility, but actin filaments also contributed significantly. A novel finding was the influence of intermediate filaments in vesicle transport. The mechanism did not appear to be mediated through an interaction between intermediate filaments and microtubules. More likely, the dense network of intermediate filaments provides a steric hindrance/channeling system that helps direct vesicle movements.

Previously, we found that vesicles in resting astrocytes exhibit directional, almost rectilinear mobility and nondirectional mobility (12), and we hypothesized that rapid directional mobility required interaction with the cytoskeleton. Our findings in this study support that hypothesis. They also support the notion that proANP-Emd-labeled vesicles are released from astrocytes by calcium-dependent exocytosis (7) because increasing [Ca2+]i by treatment with ionomycin reduced the number of labeled vesicles. However, vesicle mobility in the cytoplasm was not affected, consistent with a report for secretory vesicles in differentiated PC12 cells (33).

Microtubules and directional mobility of vesicles

The maximal velocity of vesicles in control cells was up to 3.4 μm/second. Such rapid trafficking is made possible by the molecular motors kinesin and dynein, which transport cargo along microtubules (34–36). Depolymerizing microtubules with nocodazole significantly reduced vesicle mobility, as indicated by the marked reduction in maximal displacement, but the effects on mean velocity and vesicle length were less pronounced than in cells with depolymerized actin or intermediate filaments (Figure 4). Because microtubules were effectively depolymerized (Figure 1D), the moderate reduction in vesicle track length in nocodazole-treated cells may indicate that the mobility of proANP-Emd-labeled vesicles is dependent on elements other than microtubules. However, the strong effect of nocodazole on the maximal displacement of vesicles confirms the role of microtubules in directional mobility (37).

Actin filaments and vesicle mobility

In astrocytes with depolymerized actin filaments, all measures of vesicle mobility were significantly reduced (Figures 2 and 4). Despite apparent changes in cell shape (Figure 2), microtubules were still able to transport vesicles with a maximal velocity of 1.9 μm/second, which is faster than the measured maximal velocity of myosin molecules (38). In control astrocytes, vesicles were mobile in opposite directions. Besides kinesin and dynein, this kind of transport may involve myosins, several isotypes of which (I, II, V and VI) are found in mammalian astrocytes (39). After depolymerization of actin filaments, however, vesicle mobility was severely impaired (Figure 2F), indicating that actin depolymerization prevented an interaction between vesicles and microtubules. Similarly, in ARPE-19 human retina cells, vesicles remained trapped in a mesh of depolymerized actin filaments and were mobile by free diffusion (40).

Actin filaments may play a role in retaining organelles in a particular part of the cell (41). In PC12 neuroendocrine cells, the mean velocity of vesicles near the plasma membrane decreased in control cells but increased slightly in cells with depolymerized actin filaments. This finding suggested that vesicles were liberated from the peripheral area and regained access to the microtubule tracks (42). In contrast, we found that depolymerization of actin decreased vesicle mobility. Thus, rather than constraining cytoplasmic vesicle transport, actin filaments likely increase vesicle mobility in astrocytes.

We speculate that vesicles are shuttled along actin filaments until mechanisms of microtubular transport are engaged. This possibility is consistent with the notion that actin filaments are used for short-range transport (43) and with the finding that the same cargo can move on both microtubules and actin filaments, switching between motors during transport (37). Moreover, the actin filament network is randomly oriented and has sufficient density to make a good local transport system (37). In Xenopus melanophores, cells regulate transport by controlling how often granules switch from one filament type to another rather than by altering individual motor activity at the single-molecule level or by relying on structural changes in the network (44).

Intermediate filaments affect directional mobility

In cells treated with Cal A to depolymerize intermediate filaments, all measures of vesicle mobility were significantly reduced (Figure 4). However, a loss of interactions between different cytoskeleton elements after depolymerization of intermediate filaments might have had indirect effects on vesicle mobility (14,31). To rule out this possibility, we analyzed GFAP−/−Vim−/− astrocytes, which are devoid of intermediate filaments. The percentage of vesicles exhibiting directional mobility was lower in the cells without intermediate filaments than in wild-type astrocytes (21 versus 34%) (Figure 5), confirming a direct role for intermediate filaments in vesicle mobility. In GFAP−/−Vim−/− cells, we found five times less vesicles with track length higher than 9 μm compared with wild-type cells (5 versus 27 vesicles, respectively; Figure 5). It is likely that intermediate filaments act as a scaffold representing a conduit for highly mobile vesicles. This is also supported by the finding that the fraction of vesicles exhibiting directional mobility was lower in the GFAP−/−Vim−/− compared with wild-type astrocytes.

Although the mobility of intermediate filaments can be inhibited with substances that depolymerize microtubules (14), we found no difference in vesicle mobility between wild-type and GFAP−/−Vim−/− mouse astrocytes after treatment with nocodazole (Figure 4). Therefore, interactions between intermediate filaments and microtubules do not seem to be a major determinant of vesicle mobility in astrocytes.

Genetic ablation of astrocyte intermediate filaments seems to trigger a partial compensatory response of the microtubular system. We found elevated expression levels of MAP-2 both in GFAP−/−Vim−/− P1 brains and in astrocyte cultures prepared from these brains. Moreover, MAP-2 expression was also increased in the tissue surrounding the cortical lesions in the electrically induced neurotrauma model. MAP-2 was proposed to have a direct effect on stabilization of microtubules and their cross-linking as well as cross-linking of microtubules with intermediate filaments (45). This may explain the relatively minor difference between the mobility of vesicles in GFAP−/−Vim−/− and wild-type astrocytes (Figure 6).

Possible role of intermediate filaments in vesicle transport

Intermediate filament proteins exist in different forms, such as free monomers, short stretches of filaments and long filaments, and can be connected to actin filaments and microtubules (14). Short segments of intermediate filaments are highly mobile structures transported along microtubules by kinesin and dynein (46–49). Interestingly, their mobility parameters (velocity, maximal displacement, two modes of mobility) are similar to those of proANP-Emd-labeled vesicles and vimentin particles (12,47).

Two hypotheses have been proposed to explain the role of intermediate filaments in particle mobility (50). One hypothesis predicts that intermediate filaments (such as neurofilaments) act as an anchor or ligand between molecular motors and a cargo or organelle (50). Consistent with this hypothesis, intermediate filaments have been proposed to participate in recruiting the adaptor protein AP-3 for vesicle formation, in the uncoating of vesicles and in recruiting motor proteins to AP-3 vesicles (51). The second hypothesis holds that the three-dimensional lattice of intermediate filaments acts as a physical barrier that extends radially from the nucleus to the cell surface, conferring advantages in coordinating cytoskeletal activities and relaying information between the cell surface and the innermost compartments of the cell (14).

We found no differences in measures of vesicle mobility in wild-type and GFAP−/−Vim−/− astrocytes after treatment with nocodazole. Therefore, it is unlikely that interactions between intermediate filaments and microtubules significantly modulate vesicle mobility in astrocytes. Our results support the hypothesis that intermediate filaments are required for long-range directional vesicle mobility, by acting as a three-dimensional lattice.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References

Cell cultures and GFAP/Vim/ mice

Astrocytes from the cortex of 3- day-old rats (Wistar) and 1-day-old GFAP−/−Vim−/− and wild-type mice (a mixed 129Sv–129Ola genetic background) were isolated and maintained as described previously (12,18,52–56). GFAP−/−Vim−/− mice (18,21,23,24) were obtained by cross-breeding of mice lacking vimentin (57) and mice lacking GFAP (54).

Cytoskeleton

Actin filaments were depolymerized with 15 nm CST (a gift from Dr M. R. Popoff, Institut Pasteur, Paris, France), microtubules with 200 μm nocodazole (Sigma, Diesenhofen, Germany) and intermediate filaments with 20 nm Cal A (Calbiochem, San Diego, CA, USA). For immunolabeling, primary antibodies against α-tubulin (1:200; Sigma) and vimentin (1:200; Abcam, Cambridge, UK) and secondary antibodies against mouse or rabbit IgG (Alexa Fluor 546, 1:500; Molecular Probes, Eugene, OR, USA) were used. Actin filaments were labeled with rhodamine-conjugated phalloidin (Molecular Probes). Cells were immunostained as described by Kreft et al. (10).

Measurements of [Ca2+]i

To increase [Ca2+]i, cells were perfused with 10 μm ionomycin in extracellular solution: (in mm) 130 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 d-glucose and 10 HEPES, pH 7.2. To measure [Ca2+]I, cells were preloaded with 4 μm Fura-2/AM (Molecular Probes) and sequentially excited at 340 and 380 nm with a monochromator (Polychrome IV; Till Photonics, Gräfelfing, Germany). Emission was monitored with a 440-nm filter (Omega Optical Inc., Brattleboro, VT, USA).

Transfection

Cells were transfected with DNA proANP-Emd (a gift from Dr Ed Levitan, Department of Pharmacology, University of Pittsburgh, Pittsburgh, PA, USA), using FuGene transfecting reagent (Roche, Mannheim, Germany) as recommended by the manufacturer.

Microscopy

Cells were examined with an inverted Zeiss LSM 510 confocal microscope (oil-immersion objective × 63/NA 1.4, He/Ne laser, long-pass filter, with a cutoff below 560 nm). Tracking of vesicles in cells was recorded every 300 mseconds by an inverted microscope (Zeiss Axiovert 135; Carl Zeiss Inc., Jena, Germany) equipped with a charge-coupled device camera and a water-immersion objective (C-Apochromat × 63/NA 1.2). Cells were excited at 470 nm with a monochromator (Polychrome IV, Till Photonics), and emission light was collected through a 510- to 533-nm filter and a 500-nm dichroic mirror (Omega Optical Inc.).

Vesicle tracking

Vesicle tracking was analyzed with custom software (ParticleTR, Celica, Slovenia) as described by Potokar et al. (12). We estimated current time (time from the beginning of tracking for a single vesicle), step length (displacement of a vesicle in the time interval 300 mseconds), track length (the total length of the analyzed vesicle pathway), velocity, maximal displacement and the directionality index (maximal displacement/total track length) of vesicles as described previously (12,56). Vesicle mobility was analyzed from three independent astrocyte cultures, rat and mouse, respectively. The analysis of vesicle mobility was performed for epochs of 15 seconds. Statistical significance was determined with the two-tailed t-test for equal variances. Values are expressed as mean ± SEM.

Surgical procedures

Electrically induced injury of the cerebral cortex was performed as described (58) in four wild-type and four GFAP−/−Vim−/− 6-month-old mice. All mice were on a mixed genetic background (129Sv–129Ola) and were maintained in a barrier animal facility. Anesthetized mice were placed in a stereotactic frame, and a hole was drilled through the skull. A fine-needle electrode was inserted through the skull 2.25 mm laterally at the level of bregma and lowered 1.0 mm (measured from the meningeal level) into the cortex of the right hemisphere. A second electrode was attached to the root of the tail. Using Lesion maker (Ugo Basile, Comerio, Italy), a direct current of 5 mA was applied for 10 seconds. The mice were kept in heated cages until they recovered from anesthesia. Four days later, the mice were killed by cervical dislocation. Part of the right frontotemporal cortex containing the lesion with the surrounding tissue (approximately 5 × 5 × 5 mm) was dissected out, freezed in liquid nitrogen and stored at −70°C.

RNA preparation from astrocyte cultures, brains and cortical lesions

When confluent, the astrocyte cultures were harvested by scraping in ribonuclease (RNAse)-free PBS and centrifuged at 500 × g for 5 min at 4°C. The supernatant was discarded and the cell pellet stored at −70°C. TRIZOL Reagent (Invitrogen, Carlsbad, CA, USA) was added to each sample – frozen cell pellets, whole brains or lesioned frontotemporal cortex. The cell pellets were homogenized by vortexing, the tissue was homogenized by Polytron PT 2100 (Kinematica AG, Littau, Switzerland) at 4°C and extracted four times by phenol/chloroform. Glycogen (coprecipitant) and isopropanol were added to precipitate the RNA; the pellet was washed with 80% ethanol, dried, resuspended in RNAse-free water and stored at −70°C. The purity of the RNA was assessed by gel electrophoresis on 1% agarose gel containing MOPS buffer (CPG Inc., Lincoln Park, NJ, USA) and 1 m formaldehyde.

Reverse transcription and quantitative rtPCR

Complementary DNA was generated using the iScript cDNA Synthesis Kit (Bio-Rad Laboratories, Hercules, CA, USA) with a mixture of random hexamers and oligo(dT) primers, according to the manufacturer's instructions; the incubation time at 42°C was increased from 30 to 60 min. The reverse transcription was run in duplicates in 10-μL reactions (59) using 1.5 μg of total RNA extracted from primary astrocyte cultures, brains or tissue from cortical lesions.

Gene-specific SYBR-Green-based PCR assays were designed for MAP-2 (GenBank accession number M21041). Formation of expected PCR products was confirmed with agarose gel electrophoresis and melting curve analysis. Primer sequences for MAP-2 were 5′-TCA GGA GAC AGG GAG GAG AA-3′ (forward) and 5′-GGG GTA GTA GGT GTG GAG GTG-3′ (reverse). The rtPCR experiments were run on a Rotor-Gene 3000 (Corbett Research, Sydney, Australia) and analyzed as described elsewhere (59,60). All gene expression data were normalized against total RNA concentration (61). Statistical significance between GFAP−/−Vim−/− and wild-type mice was tested with Student's t-test. PCR run conditions and thermocycler programs are available upon request.

Acknowledgments

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References

We thank Dr Ulrika Wilhelmsson for performing the microsurgery, Dr Mikael Kubista, Dr Michael Nilsson and TATAA Biocenter in Göteborg, Sweden, for their contribution to MAP-2 rtPCR experiments. This work was supported by the Ministry of Education, Sciences and Sports of The Republic of Slovenia grant P3 310 0381, the EC grant #QLG3 2001-2004, EC support DECG, CLG3-CT-2001-02004 and the Swedish Research Council 11548, ALF Göteborg, Hjärnfonden, the Swedish Stroke Association and Torsten och Ragnar Söderbergs stiftelser, Rune och Ulla Amlövs stiftelse and Trygghansa to MP, and the Swedish Medical Society 16850, Wilhelm och Martina Lundgrens stiftelse, Hjärnfonden and Åhlén-stiftelsen to UW.

References

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References