brain research 1564 (2014) 9–21
Available online at www.sciencedirect.com
www.elsevier.com/locate/brainres
Research Report
Nitric oxide/cyclic GMP signaling regulates motility of a microglial cell line and primary microglia in vitro Hannah Scheiblicha, Frank Roloffa, Vikramjeet Singhb,c, Martin Stangelb,c, Michael Sterna, Gerd Bickera,c,n a Division of Cell Biology, University of Veterinary Medicine Hannover, Bischofsholer Damm 15/102, D-30173 Hannover, Germany b Department of Neurology, Hannover Medical School, Germany c Center for Systems Neuroscience, Hannover, Germany
art i cle i nfo
ab st rac t
Article history:
Microglia are the resident immune cells of the brain, which become rapidly activated and
Accepted 30 March 2014
migrate to the site of insult in brain infection and disease. Activated microglia generate large
Available online 5 April 2014
amounts of the highly reactive messenger molecule nitric oxide (NO). NO is able to raise cyclic
Keywords:
GMP levels via binding to soluble guanylyl cyclase. We investigated potential mechanistic
BV-2 cells
links between inflammation, NO signaling, and microglial migration. To monitor cell
Cytoskeleton
migration, we used a scratch wound assay and compared results obtained in the BV-2
Inflammation
microglial line to primary microglia. Incubation with lipopolysaccharide (LPS) as stimulator of
iNOS
acute inflammatory processes enhanced migration of both microglial cell types. LPS activated
Primary microglia
NO production in BV-2 cells and application of an NO donor increased BV-2 cell migration while an NO scavenger reduced motility. Pharmacological inhibition of soluble guanylyl cyclase and the resulting decrease in motility can be rescued by a membrane permeant analog of cGMP. Despite differences in the threshold towards stimulation with the chemical agents, both BV-2 cells and primary microglia react in a similar way. The important role of NO/cGMP as positive regulator of microglial migration, the downstream targets of the signaling cascade, and resulting cytoskeletal changes can be conveniently investigated in a microglial cell line. & 2014 Elsevier B.V. All rights reserved.
1.
Introduction
Local inflammation within the central nervous system (CNS) has been shown to be a common mechanism causing the neuronal loss in a diversity of chronic neurodegenerative
diseases such as Alzheimer's disease, Parkinson's disease, and multiple sclerosis. Inflammatory reactions following acute spinal cord injury induce detrimental, yet also beneficial post-traumatic cellular responses in the CNS parenchyma (Hausmann, 2003). There are several cell types that
n Corresponding author at: Division of Cell Biology, University of Veterinary Medicine Hannover, Bischofsholer Damm 15/102, D-30173 Hannover, Germany. Fax: þ49 511 856 7687. E-mail addresses: [email protected] (H. Scheiblich), [email protected] (F. Roloff), [email protected] (V. Singh), [email protected] (M. Stangel), [email protected] (M. Stern), [email protected] (G. Bicker).
http://dx.doi.org/10.1016/j.brainres.2014.03.048 0006-8993/& 2014 Elsevier B.V. All rights reserved.
10
brain research 1564 (2014) 9–21
have been implicated in the response to inflammatory stimuli, but it has been postulated that excessive activation or unregulated activation of microglia are mediators of inflammation-mediated neuronal loss in most neurodegenerative circumstances. As resident immune effector cells in the CNS microglia constantly survey their microenvironment to respond to homeostatic changes. In the healthy adult brain, the baseline motility of microglia is characterized by the constant movement of their highly branched cellular processes (Ohsawa and Kohsaka, 2011). Upon detecting environmental stimuli like the inflammogen lipopolysaccharide (LPS), a component of the outer membrane of Gramnegative bacteria, microglia become rapidly activated, retract their processes, and migrate to the tissue damage or the site of injury (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Kreutzberg, 1996) to secrete cytotoxic substances (Liu, 2003; Moss and Bates, 2001) and tumor necrosis factor alpha (Lee et al., 2006) or to release neurotrophic compounds for regeneration and anti-inflammatory factors (Batchelor et al., 2002; Liao, 2004; Morgan et al., 2004; Polazzi et al., 2001; Streit et al., 2004). Insights into mechanisms of activation and migration may contribute to pinpoint microglia as a key pharmacological target in the treatment of neurodegeneration and traumatic injury. The messenger molecule nitric oxide (NO) which is generated at the lesion site has been shown to act as chemoattractant for activated microglia toward the site of injury and as a potential stop signal to cause accumulation of microglia at the lesion (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Haynes et al., 2006; Ohsawa et al., 2007). Several other studies have demonstrated that NO signal transduction regulates cellular motility during the development of vertebrate and invertebrate nervous systems (Gutièrrez-Mecinas et al., 2007; Haase and Bicker, 2003; Mandal et al., 2013; Peunova et al., 2007; Tegenge and Bicker, 2009; Tegenge et al., 2011; Trimm and Rehder, 2004) NO is generated by the enzyme nitric oxide synthase (NOS). In microglia the inducible NOS isoform (iNOS) has been identified to synthesize continuously high levels of NO (Minghetti and Levi, 1998; Nathan, 1992; Vincent, 1994) that is responsible for the defense of the CNS. Microglia are particularly resilient to high concentrations of this compound (Minghetti and Levi, 1998). The main target of NO is the cyclic guanosine monophosphate (cGMP) synthesizing enzyme soluble guanylate cyclase (sGC) that generates cGMP from guanosine triphosphate (GTP) (Garthwaite, 2008; Ignarro, 1990; Moncada et al., 1989). Cyclic GMP itself activates downstream effectors such as protein kinases G (PKG) (Hofmann et al., 2006) and cyclic nucleotidegated ion channels (Bender and Beavo, 2006). PKGs are mediators of NO/cGMP effects (Baltrons et al., 2008) like the phosphorylation of effector proteins, for example members of the Rho GTPase family (Garthwaite, 2008) which are known to be involved in the organization of the actin cytoskeleton and are key regulators of cell migration. A variety of methods at different microscopic scales have been applied to investigate cytoskeletal changes and cell motility, ranging from chemotaxis assays across micropore membranes (Boyden, 1962) to sophisticated biophysical measurements of the force generated by actin polymerization during lamellipodial protrusion (Fuhs et al., 2014). Here, we used a scratch wound assay on both the microglial cell line
BV-2 and primary microglia to study the modulation of microglial migration by NO/cGMP signal transduction in vitro. This rather simple assay allows for microscopical imaging of cell morphology. We confirmed that incubation in LPS upregulates the production of NO in BV-2 cells (Henn et al., 2009). Application of small bioactive enzyme activators and inhibitors of NO/cGMP signaling caused significant cytoskeletal changes. The NO/cGMP signaling pathway positively modulates migration of both BV-2 cells and primary microglia in a functional assay.
2.
Results
2.1.
LPS-induced microglial migration
To model the migration of activated microglia, we applied LPS as inflammatory stimulus to adherent cell cultures of BV-2 cells and primary microglia. Even though BV-2 cells needed tenthousandfold higher LPS concentrations than primary microglia (Fig. 2a and b), the migration distance in a scratch wound migration assay for both cell types was significantly enhanced upon LPS stimulation. Concentrations of 10 mg/ml LPS increased migration of BV-2 cells more than twofold up to 210.1726.2% (Fig. 2a) of control levels. This amounts to an enhancement of the cell migration velocity from 16.1272.65 mm/h (control) up to 30.5374.57 mm/h. In primary microglia 1 ng/ml LPS increased the response of the cells up to 130.375.2% (Fig. 2b), corresponding to an increment of the absolute migration velocity of 14.327 1.47 mm/h (control) up to 18.9671.47 mm/h. Higher concentrations of LPS reduced for both cell types the motility in the scratch wound assay. To visualize effects of LPS stimulation on cell morphology, we performed phalloidin staining against the F-actin cytoskeleton after migration experiments. Both microglial cell types migrated with long F-actin-positive processes into the scratch (Fig. 2c and d). High magnifications of LPS-treated BV-2 cells revealed a polar and spindle-shaped phenotype (Fig. 2e). This morphology was characterized by F-actin-rich microspike projections around the soma or at the terminations of elongated thin cell extensions. LPS-treated primary microglial cells reorganized into a flat and stretched shape (Fig. 2f). Most of the F-actin network formed thick filamentous bundles. Untreated BV-2 cell controls displayed a round to ameboid shape with thick but short F-actin processes (Fig. 2g), resembling the untreated primary microglial morphology (Fig. 2h). Inflammation in brain tissue is accompanied by an increased NO synthesis in microglial cells via induction of iNOS. To visualize this enzyme in BV-2 cells, we used NADPHdiaphorase staining as a cytochemical marker for the presence of NOS. Cultures of both BV-2 cells and primary microglia showed strong labeling for NADPH-diaphorase (Fig. 3a and b). Using the Griess-Assay for nitrite production, we measured endogenous NO formation in BV-2 cells (control: 0.270.07 mM after 8 h; 1.4970.16 mM after 24 h). Application of LPS caused a dose- and time-dependent increase in NO production after 8 h (11.870.21 mM) and 24 h (54.1671.21 mM) of incubation (Fig. 3c). A rather sensitive receptor molecule for NO is soluble guanylyl cyclase (sGC), an enzyme that synthesizes cGMP (Fig. 1d). We used an enzyme immunoassay to investigate whether BV-2 cells can produce cGMP (Fig. 3d). Because
brain research 1564 (2014) 9–21
11
Fig. 1 – Schematic drawing of the in vitro scratch wound assay: (a)–(c) the extent of cell motility was evaluated by measuring the migration during the closure of a wound that is scratched into a confluent cell monolayer. Closure of the wound was followed by monitoring the advancement of the cell front over 8 h for BV-2 cells and 6 h for primary microglia. (d) Manipulation of NO/cGMP signaling by chemical agents. In microglia the inducible nitric oxide synthase (iNOS) has been identified to synthesize nitric oxide (NO) in response to immunological stimuli such as LPS. NO binds to the soluble guanylyl cyclase (sGC) and regulates activity of the enzyme. Exogenous NO level can be regulated by the NO donor NOC-18 or the NO scavenger cPTIO. The sGC activity can be blocked by the inhibitor ODQ. The membrane-permeable cGMP analog 8-Br-cGMP raises the intracellular cGMP level. Scale bar: 200 lm.
12
brain research 1564 (2014) 9–21
Fig. 2 – LPS was used as a tool to assess inflammatory-mediated response of microglial migration behavior. Application of LPS significantly increased the movement of BV-2 cells (a) and primary microglia (b) at lower concentrations range. Higher concentrations resulted in a decreased migration distance down to the control level. BV-2 cells (c, e) and primary microglia (d, f) incubated with LPS extended long F-actin-positive processes into the scratch. Cells aquired polarity and a spindleshaped phenotype (arrowheads) compared to a control without treatment (BV-2: g; primary microglia: h). Data are presented as mean7SEM of at least three independent experiments. (*po0.05, **po0.01, ***po0.001 compared to control). Cyan: DAPI; green: Phalloidin. Scale bar: 100 lm (cþd); 20 lm (e–h).
brain research 1564 (2014) 9–21
13
cultures in control medium (0.0370.01 pmol/ml), application of the sGC activator YC-1 (3-(50 -Hydroxymethyl-20 -furyl)-1-benzyl indazole) (100 mM) in combination with IBMX (1 mM) caused a highly significant increase in cGMP levels (0.1770.03 pmol/ml). Application of the fast-releasing NO donor NOC-12 (N-Ethyl-2(1-ethyl-2-hydroxy-2-nitrosohydrazino)ethanamine) (500 mM) in the presence of IBMX did also increase cGMP levels significantly (0.1370.02 pmol/ml). The co-application of YC-1þNOC-12þ IBMX again caused an increase in cGMP (0.1770.04 pmol/ml). However, this increase did not show an additive effect of the YC-1 and NOC-12 compounds. Similarly, the incubation in IBMX alone increased cGMP to comparable levels (0.1570.3 pmol/ml), indicating that BV-2 cells already produce cGMP endogenously.
2.2. Exogenous nitric oxide stimulates microglial cell migration To test for a potential role of NO as modulator of microglial motility, we investigated the effect of exogenous NO on BV-2 cells and primary microglial cultures. Application of NOC-18 (2,20 -(hydroxynitrosohydrazino)bis-ethanamine) significantly enhanced the migration of BV-2 cells into the scratch in a dose-dependent manner up to 164.9721.64% (500 mM; 24.297 3.52 mm/h) compared to the control (12.7571.48 mm/h) (Fig. 4a). Migration distance and cell migration velocity of primary microglia was significantly increased only at 10 mM NOC-18 (27.7572.13 mm/h) while application of a higher concentration of 100 mM NOC-18 did not change motility compared to controls (18.2671.98 mm/h) (Fig. 4b). Similar to application of LPS, NOC-18 caused a cytoskeletal reorganization of BV-2 cells and primary microglia resulting in a spindle-shaped phenotype (Fig. 4c and d). Application of the NO scavenger cPTIO (2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide) caused a dose-dependent reduction of BV-2 cell migration distance and velocity down to 11.9476.12% (1.370.6 mm/h) (500 mM) compared to vehicle control (10.0770.76 mm/h) (Fig. 4e). We could not observe any phenotypical difference in cell morphology compared to untreated vehicle control cells (Fig. 4f and f0 ). These data indicate that exogenous NO modulates migration and causes cytoskeletal reorganization of both BV-2 cells and primary microglia. Fig. 3 – We used NADPH-diaphorase staining in BV-2 cells (a) and primary microglia (b) as a marker for NOS expression. When BV-2 cells were stimulated with the inflammogen LPS, nitrite production was highly increased (c). To measure the effect of the fast-releasing NO donor NOC-12 (500 lM) and the sGC activator YC-1 (100 lM) in combination with the PDE inhibitor IBMX (1 mM) on the intracellular cGMP level, a cGMP EIA was performed (d). Data are presented as mean7SEM of three independent experiments (*po0.05, **po0.01, ***po0.001 compared to control). In all cases the detected intracellular cGMP level was increased compared to a control. Scale bar: 50 lm.
cytosolic cGMP is efficiently metabolized by phosphodiesterases (PDEs), we measured cGMP levels in the presence of the PDE blocker IBMX (3-Isobutyl-1-methylxanthine). Compared to
2.3. NO/cGMP signaling is a positive regulator of microglial motility Next, we asked whether the NO/cGMP signaling pathway is involved in the control of microglial motility. We employed small bioactive enzyme activators and inhibitors in both gain and loss of function experiments. In a loss of function experiment we tested whether the sGC inhibitor ODQ (1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one) affects microglial motility in a dose-dependent manner. Indeed, application of ODQ above a concentration of 50 mM ODQ significantly inhibited migration of BV-2 cells and primary microglia (Fig. 5a and b) compared to the vehicle controls. By quantifying the impact of ODQ on cell migration velocity we found that BV-2 cell migration was slowed down to 5.1271.11 mm/h (control: 14.4271.63 mm/h) similar to migration velocity of primary microglia (5.257 1.54 mm/h vs. control 12.8871.53 mm/h). We could not observe
14
brain research 1564 (2014) 9–21
Fig. 4 – Exogenous NO level regulates microglial migration. Exogenous NO released from NOC-18 enhanced motility of BV-2 cells (a) and primary microglia (b). When cells were incubated with NOC-18 (100 lM) morphological changes could be observed. BV-2 cells (c) and primary microglia (d) acquired a spindle-shaped phenotype (arrowheads). The NO scavenger cPTIO trapped NO, resulting in a significantly decreased migration distance of BV-2 cells compared to the vehicle control (e). We could not detect any morphological changes (f) compared to the vehicle control (f0 ). Data are mean7SEM of at least three independent experiments. (*po0.05, **po0.01, ***po0.001 compared to control). Cyan: DAPI; green: Phalloidin. Scale bar: 20 lm.
any morphological difference when BV-2 cells were incubated with ODQ compared to vehicle control (Fig. 5c and c0 ). In contrast primary microglia exhibited a round and flat shape with many tiny microspikes after incubation with ODQ (Fig. 5d and d0 ). Co-application of the cell membrane permeant analog of cGMP, 8-Br-cGMP (8-Bromoguanosine 30 ,50 -cyclic monophosphate sodium salt), reversed the inhibitory effect of ODQ to control levels for both cell types (Fig. 5e and f).
This rescue experiment provides direct evidence for NO/cGMP signal transduction as a positive regulator of microglial cell migration. To exclude cytotoxic effects of the chemical compounds on motility, we measured the cell viability in an Alamar Blue assay directly after migration experiments. All the chemicals used in our migration assay did not affect cell viability of BV-2 cells and primary microglia (Fig. 6).
brain research 1564 (2014) 9–21
15
Fig. 5 – Blocking of the cGMP synthesizing enzyme sGC with ODQ resulted in a decreased cell movement of BV-2 cells (a) and primary microglia (b) compared to the vehicle controls. While BV-2 cells (c) did not change their morphology compared to the vehicle control (c0 ) primary microglia (d) exhibited a round and flat shape with many tiny microspikes in contrast to the vehicle control (d0 ). Exogenous application of the cGMP analog, 8-Br-cGMP, rescued the blocking of cell migration by sGC inhibitors in both cell types (e, f). Data are mean7SEM of at least three independent experiments (*po0.05, **po0.01, ***po0.001 compared to control). Cyan: DAPI; green: Phalloidin. Scale bar: 20 lm.
3.
Discussion
3.1.
Microglial BV-2 cells as a model for primary microglia
In the present paper, we have used chemical manipulations to demonstrate that the NO/cGMP signaling system enhances motility of the BV-2 microglial line in a scratch wound assay (Fig. 1). These results are in line with cell culture and in vivo
studies showing NO/cGMP as positive regulator for primary microglial motility (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Haynes et al., 2006; Ohsawa et al., 2007). In the scratch assay, we have also analyzed the motility of rat primary microglia in response to an enzyme activator (Fig. 4) and inhibitor of the NO/cGMP cascade (Fig. 5). To test for unspecific side effects of the chemical agent on cell
16
brain research 1564 (2014) 9–21
Fig. 6 – Cell viability assay of chemical compounds used in the experiments. The highest concentrations of all used chemical compounds did not affect viability of BV-2 cells (a) and primary microglia (b). Data are mean7SEM in [%] of at least three independent experiments (*po0.05, **po0.01, ***po0.001 compared to control).
migration behavior, we performed in parallel rescue experiments with BV-2 and primary rat microglia (Fig. 5e and f). The isolation of primary microglia from rat brains is a time consuming procedure yielding rather low cell numbers for culturing. For biochemical experiments it is necessary to expand these cell cultures, which is restricted by the limited proliferation capacity of the primary cells (Henn et al., 2009). Moreover, for the investigation of a specific signal transduction pathway it is desirable to avoid the heterogeneity in cellular phenotypes that are characteristic of brain-derived microglia (Guillemin and Brew, 2004; Hanisch and Kettenmann, 2007; Mahe et al., 2001). To reduce animal experimentation, it has been suggested that the rapidly proliferating microglial cell line BV-2 might serve as a valid substitute for primary microglia (Henn et al., 2009). The BV-2 cell line has been derived from immortalized murine neonatal microglia (Blasi et al., 1990). The validity of BV-2 cells as a microglial model system has been suggested on the basis of similar morphological properties, the expression of microglial cell markers, transcriptome analysis, the secretion of cytokines, and functional properties (Blasi et al., 1990; Henn et al., 2009; Horvath et al., 2008; for review see: Stansley et al., 2012). BV-2 cells exhibit short, thick processes similar to the morphology of activated primary microglia (Horvath et al., 2008). They express specific microglial markers including the microglia activation marker Iba-1 (ionized calcium binding adaptor molecule 1), MAC 1 and MAC 2 (macrophage antigen complex 1 and 2), but do not express MAC 3 (macrophage antigen complex 3), GFAP (glial fibrillary acidic protein, astrocyte marker), or galactocerebroside (oligodendrocyte marker) (Horvath et al., 2008). Moreover, BV-2 cells were tested 90% positive for non-specific esterase activity and all cells lacked peroxidase activity, indicative of microglia (Blasi et al., 1990; Giulian and Baker, 1986). Transcriptome analysis indicated that 90% of the genes induced in BV-2 cells by LPS were also found in primary microglia (Henn et al., 2009) and the secretion of cytokines was increased dose dependently (Blasi et al., 1990). Previous studies have also pointed towards a similar physiological behavior of BV-2 cells and primary microglia in in vitro transwell migration experiments or in assays for iNOS expression and generation of NO (Henn et al., 2009).
There are, however, also indications that BV-2 cells can only partially model primary microglia (Henn et al., 2009; for review see: Stansley et al., 2012). One study compared BV-2 cells and primary microglia with regard to activation markers, cell motility and releasable factors (Horvath et al., 2008). Across the dosage range of ng/ml to mg/ml of LPS stimulation, BV-2 cells generated lesser but still substantial amounts of NO compared to primary microglia. Despite some discrepancies in microglial phenotype, our migration assay revealed a striking modulation of BV-2 cell motility by functional NO/cGMP signal transduction. The robust response in the migration assay, cellular homogeneity, and the unlimited proliferation potential make the BV-2 microglial line an ideal model for the investigation of NO/cGMP mediated motility changes.
3.2.
Inflammatory activation of microglia
We found in the scratch wound assay a link between LPSmediated inflammatory activation and cellular motility for both BV-2 cells and primary microglia (Fig. 2). Similar investigations on murine macrophages, human monocytes, and neutrophils showed an LPS-induced increase of cell migration (Kukulski et al., 2007; Maa et al., 2008). In our functional assay, LPS treatment resulted in a significantly increased migration distance and changes in morphology (Fig. 2). Application of LPS promoted the outgrowth of processes in both BV-2 cells and primary microglia. Cells became polarized and one large and flattened process per cell extended into the scratch. Primary microglia was far more sensitive to LPS stimulation (Fig. 2), whereas the BV-2 cells responded better to NO stimulation than the primary microglia (Fig. 4). However, for a straightforward comparison of both cell types, a caveat should be kept in mind. Primary microglia were isolated from neonatal rodent brains and initially kept in a primary mixed glial culture. Compared to adult microglia, this neonatal origin and the subsequent cell culture conditions may have modified the reaction potential of our primary microglia (Henn et al., 2009). Moreover, primary microglia cells are a heterogeneous population. In a pathological context, microglia from different anatomical regions within the brain can exhibit different reaction
brain research 1564 (2014) 9–21
patterns resulting from regional adaptations (Guillemin and Brew, 2004; Hanisch and Kettenmann, 2007; Mahe et al., 2001), whereas BV-2 cells perform more uniformly (Blasi et al., 1990; Henn et al., 2009). Thus, BV-2 cells may serve as useful model in assays where robust microglial cell behavior is required. In macrophages, microglia, and endothelial cells LPS stimulation is accompanied by enhancing the synthesis level of iNOS, resulting in an increased release of NO (Arias-Salvatierra et al., 2011; Fiebich et al., 1998; Lieb et al., 2003; Maa et al., 2008; Stangel and Compston, 2001). Similar to primary microglia, BV-2 cells labeled for NADPH-diaphorase, a cytochemical marker for the presence of NOS (Fig. 3a and b). Moreover, LPS induced in a doseand time-dependent manner the production of NO (Fig. 3c), confirming results from Henn et al. (2009) and Terazawa et al. (2013) about the induction of iNOS transcripts and protein in BV-2 cells. Low concentrations of NO can enhance the movement of migrating neutrophils while high concentrations inhibited the movement and acted as a stop signal to cause accumulation at the site of effect (VanUffelen et al., 1998, 1996). For similar reasons higher LPS concentrations may have no effect on the migration of BV-2 cells and primary microglia in the scratch assay.
3.3. The effect of NO/cGMP signaling on microglial migration To show that BV-2 cells can synthesize cGMP, we used an enzyme immunoassay (Fig. 3d). Treatment with the PDE blocker IBMX caused a significant increase in cGMP levels, indicating endogenous cGMP synthesis. This increase in cGMP could not be enhanced by the additional application of the fast-releasing NO donor NOC-12 and the NO independent sGC activator YC-1, suggesting that endogenous production of cGMP in BV-2 cells already runs at maximum capacity. During neural development NO signal transduction is a positive regulator of motility in various cell types, such as human neuronal precursor cells (Tegenge and Bicker, 2009; Tegenge et al., 2011), immature mouse neurons of the medial ganglionic eminence (Mandal et al., 2013), and insect enteric neuron migration (Haase and Bicker, 2003). Similar mechanisms might influence microglial motility in inflammated neural tissue. Under pathological conditions, NO released from degenerating neurons seems to act as a chemoattractant for recruiting microglia toward the site of injury (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Haynes et al., 2006; Ohsawa et al., 2007). In the leech NO controls the direction of microglial movement while ATP is required for the activation of microglia or for movement initiation (Duan et al., 2009). In mice NO alone was not sufficient to act as a chemoattractant for microglia when ATP was removed from the extracellular environment (Dibaj et al., 2010). We showed in the scratch assay that exogenous NO can act alone as a positive modulator of microglial migration. The application of the slow-releasing NO donor NOC-18 enhanced migratory behavior significantly while the treatment with the NO scavenger cPTIO attenuated movement (Fig. 4). In our cell culture model the blocking of the NO-induced activation of sGC by ODQ reduced the migration of both BV-2 cells and primary microglia significantly (Fig. 5a and b). This blocking of cGMP signaling was not accompanied by any striking changes in the morphology of BV-2 cells (Fig. 5c)
17
compared to vehicle controls (Fig. 5c0 ). Since we could not observe outgrowth of processes under incubation with ODQ the BV-2 morphology appears of a resting cell phenotype. Our findings are in line with in vivo experiments on mice demonstrating that ODQ blocked the migratory microglial response to laser lesions (Dibaj et al., 2010). The co-application of a cGMP analog completely rescued the decreased cell migration caused by ODQ (Fig. 5e and f). The rescue of the loss of function effect indicates that a certain level of cGMP is required for microglial migration. Application of cGMP to microglia with intact NO/cGMP signaling did not increase motility (data not shown). Presumably, the endogenous cGMP production may keep the motility in an optimal range. Taken together, our data indicate that the NO/cGMP signaling pathway may positively modulate the migration of microglial BV-2 cells in a similar way than in primary microglia in the in vitro scratch assay. In our cell culture system the downstream targets are not known, but cGMP can mobilize the microglial cytoskeleton (Dibaj et al., 2010) by activating the PKG or cyclic nucleotide-gated ion channels (Bender and Beavo, 2006; Hofmann et al., 2006). Further investigations are required to understand the detailed molecular mechanisms by which NO/cGMP signaling regulates cytoskeletal rearrangement and cell motility. Several studies have demonstrated that the regulation of the actin cytoskeleton is caused downstream of the cGMP pathway (Borán and García, 2007; Mandal et al., 2013). PKG regulates the expression and phosphorylation of RhoA, a small GTPase, which in turn activates the Rho kinase (ROCK). ROCK has been shown to regulate various cellular functions, amongst others cytoskeletal reorganization through the phosphorylation of the regulatory subunit of myosin (Pfitzer, 2001; Raftopoulou and Hall, 2004). Inhibition of the Rho/ROCK pathway induces dramatic morphological changes (Borán and García, 2007) and accelerated migration of astrocytes (Hö ltje et al., 2005; Ramakers and Moolenaar, 1998), leukocytes (Alblas et al., 2001), and human neutrophils (Niggli, 1999) suggesting that RhoA/ROCK signaling is critical for the regulation of cellular migration (Sauzeau et al., 2003). Phosphorylation of Enabled/ vasodilator-stimulated phosphoprotein family proteins by PKG has also been found as mechanism for the regulation of actin polymerization (Chen et al., 2007; Lindsay et al., 2007; Sporbert et al., 1999). In summary, this in vitro investigation study showed that the BV-2 microglial line is a useful model for primary microglia in cell migration experiments. Despite some differences in the threshold towards stimulation with chemical agents, both BV-2 cells and primary microglia react in a similar way. We demonstrated in the scratch migration assay that the NO/cGMP signal transduction cascade enhances microglial motility for both cell types. Since this assay allows for optical imaging of the cytoskeletal dynamics, we can now investigate mechanistic links between drug stimulation and resulting microglial migration behavior.
4.
Experimental procedure
4.1.
Materials
All substances were obtained from Sigma (Taufkirchen, Germany) unless otherwise noted.
18
4.2.
brain research 1564 (2014) 9–21
Cell culture
All animals used for cell isolation were treated according to the legal and ethical requirements of the Hannover Medical School (Germany). The procedures complied with the guidelines of animal welfare as laid down by the German Research Council (DFG). Primary microglia cells were isolated by the method of Giulian and Baker (1986) as described previously (Singh et al., 2012). Briefly, brains from neonatal Sprague-Dawley rats were stripped of the meninges and dissociated using mechanical shearing and trypsin. Cells of one to two brains were plated on poly-L-lysine coated T75 culture flasks (Iwaki, BibbySterilin, Stone, UK) and cultivated in DMEM (Gibco, Paisley, UK) supplemented with 10% fetal calf serum (FCS; Bio-Whittaker, Wokingham, UK) and 1% penicillin/streptomycin (Sigma-Aldrich, Steinheim, Germany). While using an orbital shaker-incubator, loosely attached microglia were removed from the astrocytic monolayer and mobilized into the supernatant after an incubation period of 7–10 days. Cells were plated at 5 105 per well in 24-well-plates (Costar, Corning GmbH, Kaiserslautern, Germany) and incubated overnight at 37 1C/5% CO2 to build a confluent monolayer. The murine microglial BV-2 cell line developed by (Blasi et al., 1990) was a gift from Prof. Dr. M. Leist (Department of In vitro Toxicology and Biomedicine, University of Konstanz, Germany). Semi-adherent cells were maintained and cultivated in T175 culture flasks (Nunc, Thermo Fisher Scientific, Langenselbold, Germany) in DMEM (Gibco-Invitrogen, Karlsruhe, Germany) supplemented with 10% FCS and 1% penicillin/streptomycin (Gibco-Invitrogen). Confluent cultures were passaged by using a cell scraper (Greiner Bio-One, Frickenhausen, Germany) for a maximum of 30 passages and plated into poly-D-lysine coated 24-well plates (Costar, Corning GMBH, Kaiserslautern, Germany) in serum-free DMEM to avoid the FCS-mediated activation of the cells (Laurenzi et al., 2001). The cells (3.5 105 cells/well) were allowed to adhere overnight in an incubator at 37 1C/5% CO2 to build a confluent monolayer.
4.3.
Scratch wound assay
To analyze microglial migration we performed a scratch wound assay (Fig. 1a–c) as previously described by Liang et al. (2007). The confluent primary microglia or BV-2 cell monolayer was scraped with a P10 pipette tip to create a wound, followed by a wash with BV-2 ringer (130 mM NaCl, 3 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, 4.5 g D-Glucose, pH 7.4) to remove debris and replaced with 1 ml DMEM-containing chemical compounds interfering with the NO/cGMP pathway (Fig. 1d). cPTIO, ODQ, and YC-1 were dissolved in dimethylsulfoxide (DMSO) as 100 mM stock. NOC-18 (Calbiochem, Darmstadt, Germany) was prepared as a 100 mM stock solution in 10 mM of NaOH solution. All other chemical compounds were directly added into the medium. The cultures were treated with chemicals for at least 6 h (primary microglia) and 8 h (BV-2 cells). Stimulation of primary microglia with LPS occurred with low concentrations (0.1, 1, 10 ng/ml) while BV-2 microglia needed ten-thousand fold higher LPS concentrations (0.1, 1, 10, 100 mg/ml).
To determine the migration of the cells, we acquired images at defined time points. Images were taken at 4.3 magnification in a marked sector as reference point. Scratch widths were measured before the treatment and wound closure was calculated by dividing widths measured after incubation. Wound widths were quantified at 15 distinct locations within the marked sector.
4.4.
Cell viability assay
To monitor cell viability and cytotoxic effects of chemical compounds we used the Alamar Blues assay (Trek Diagnostic Systems, East Grinstead, UK). Directly after the scratch wound assay the medium was completely changed and replaced with 1 ml cell culture medium containing 5% Alamar Blue. Fluorescence intensity of Alamar Blue was measured with a microplate reader (Infinite M200, Tecan, Männedorf, Switzerland) after 2 h.
4.5.
F-actin labeling
Cultures were fixed in 4% paraformaldehyde (PFA) dissolved in phosphate-buffered saline (PBS, 10 mM sodium phosphate, 150 mM NaCl, pH 7.4) for 15 min and permeabilized by washing them tree times for 5 min with PBS containing 0.1% Triton X-100 (PTX). Blocking solution containing PTX and 5% normal goat serum (Vector Laboratories, Burlingame, CA, USA) was applied for 20 min. The F-actin-marker Alexa Fluors 488 phalloidin (1:200, Invitrogen, Karlsruhe, Germany) was applied for 30 min followed by three more washing steps. For nuclear counter-staining we used 40 ,6-diamidino-20 phenylindol-dihydrochloride (DAPI) (Invitrogen) at 0.1 mg/ml for 10 min in PBS.
4.6.
NADPH-diaphorase staining
BV-2 cells and primary microglia were fixed in 4% PFA for 15 min on ice and washed in 0.1 M Tris–HCl (pH 7.8) with 0.1% Triton X-100 three times. Afterwards staining solution (2 mg nitro-blue tetrazolium and 20 mg NADPH in 10 ml PTX) was applied for 60 min at room temperature in the dark. Staining was stopped with three more washing steps with PBS for 5 min. In parallel we used hemocytes from Locusta migratoria as a positive control (Stern et al., 2010) and staining solution without NAPDH on BV-2 cells as a negative control (data not shown). Staining was visualized via light microscopy.
4.7.
Nitric oxide determination
The NO production was determined by measuring the stable breakdown product nitrite using the colorimetric Griess Reagent System (Promega Corporation, Madison, WI, USA) in culture medium. BV-2 cells (8 104 cells/well) in 96-well plates were stimulated for 8 h and 24 h with LPS in the concentration range used for the migration experiments. Supernatant was collected and assay was performed according to the manufacturer's protocol. Absorbance between 520 and 550 nm was read in a microplate reader (Infinite M200, Tecan, Männedorf, Switzerland). Concentration of nitrite was determined by interpolation using a nitrite standard curve.
brain research 1564 (2014) 9–21
4.8.
Measurement of intracellular cGMP
Intracellular cGMP levels were quantified using the Direct cGMP EIA kit (Enzo Life Sciences, Lö rrach, Germany). Briefly, cells (1 106 cells/well) in 6-well plates were allowed to adhere overnight in an incubator and treated 20 min prior to cell lysis with chemical compounds except LPS, LPSþIBMX and IBMX which were applied 24 h before measurements. IBMX was dissolved in DMSO as 100 mM stock. In contrast to migration experiments were NOC-18 was used as a slowreleasing NO donor (half-life of 20 h, 37 1C), NOC-12 (half-life of 327 min, 37 1C) was applied as a fast-releasing NO donor to measure intracellular cGMP. NOC-12 was prepared as a 100 mM stock in 10 mM of NaOH solution. Cell lysis was enhanced by adding 0.1% Triton X-100 to 0.1 M HCl and verified visually via light microscopy. Supernatants were assayed after centrifugation using the optional overnight acetylated version according to the manufacturer's protocol. Optical density at 405 nm was determined photometrically with a microplate reader (Infinite M200, Tecan, Männedorf, Switzerland). Concentrations of intracellular cGMP levels were calculated by interpolation using a cGMP standard curve.
4.9.
Microscopy and data analysis
The scratch wound assay and immunocytochemical stainings were examined with a Zeiss Axiovert 200 fluorescence microscope (Zeiss, Gö ttingen, Germany) equipped with a Cool SNAP digital camera (Visitron Systems GmbH, Puchheim, Germany). Acquired images were processed using ImageJ (Wayne Rusband, National Institute of Health, USA). The data were presented as mean7SEM of at least three independent experiments. Statistical comparisons of controls versus treatment were performed with one-way analysis of variance. Levels of significance are indicated as *po0.05, **po0.01, ***po0.001.
Acknowledgment This study was supported by a grant from the German Research Foundation (DFG, FG1103) to Gerd Bicker and to Martin Stangel. We would like to thank Prof. Dr. M. Leist (University of Konstanz, Germany) for the gift of the BV-2 cell line and Saime Tan for technical support.
r e f e r e n c e s
Alblas, J., Ulfman, L., Hordijk, P., Koenderman, L., 2001. Activation of RhoA and ROCK are essential for detachment of migrating leukocytes. Mol. Biol. Cell 12, 2137–2145. Arias-Salvatierra, D., Silbergeld, E.K., Acosta-Saavedra, L.C., Calderon-Aranda, E.S., 2011. Role of nitric oxide produced by iNOS through NF-κB pathway in migration of cerebellar granule neurons induced by lipopolysaccharide. Cell. Signal. 23, 425–435. Baltrons, M.A., Bora´n, M.S., Pifarre´, P., Garcı´a, A., 2008. Regulation and function of cyclic GMP-mediated pathways in glial cells. Neurochem. Res. 33, 2427–2435.
19
Batchelor, P.E., Porritt, M.J., Martinello, P., Parish, C.L., Liberatore, G.T., Donnan, G.A., Howells, D.W., 2002. Macrophages and microglia produce local trophic gradients that stimulate axonal sprouting toward but not beyond the wound edge. Mol. Cell. Neurosci. 21, 436–453. Bender, A.T., Beavo, J.A., 2006. Cyclic nucleotide phosphodiesterases: molecular regulation to clinical use. Pharmacol. Rev. 58, 488–520. Blasi, E., Barluzzi, R., Bocchini, V., Mazzolla, R., Bistoni, F., 1990. Immortalization of murine microglial cells by a v-raf/v-myc carrying retrovirus. J. Neuroimmunol. 27, 229–237. Bora´n, M.S., Garcı´a, A., 2007. The cyclic GMP-protein kinase G pathway regulates cytoskeleton dynamics and motility in astrocytes. J. Neurochem. 102, 216–230. Boyden, S., 1962. The chemotactic effect of mixtures of antibody and antigen on polymorphonuclear leucocytes. J. Exp. Med. 115, 453–466. Chen, A., Kumar, S.M., Sahley, C.L., Muller, K.J., 2000. Nitric oxide influences injury-induced microglial migration and accumulation in the leech CNS. J. Neurosci. 20, 1036–1043. Chen, H., Levine, Y.C., Golan, D.E., Michel, T., Lin, A.J., 2007. ANPinitiated CcGMP pathways regulate VASP phosphorylation and angiogenesis in vascular endothelium. J. Biol. Chem. 283, 4439–4447. Dibaj, P., Nadrigny, F., Steffens, H., Scheller, A., Hirrlinger, J., Schomburg, E.D., Neusch, C., Kirchhoff, F., 2010. NO mediates microglial response to acute spinal cord injury under ATP control in vivo. Glia 58, 1133–1144. Duan, Y., Sahley, C.L., Muller, K.J., 2009. ATP and NO dually control migration of microglia to nerve lesions. Dev. Neurobiol. 69, 60–72. Fiebich, B.L., Butcher, R.D., Gebicke-Haerter, P.J., 1998. Protein kinase C-mediated regulation of inducible nitric oxide synthase expression in cultured microglial cells. J. Neuroimmunol. 92, 170–178. Fuhs, T., Goegler, M., Brunner, C.A., Wolgemuth, C.W., Kaes, J.A., 2014. Causes of retrograde flow in fish keratocytes. Cytoskeleton 71, 24–35. Garthwaite, J., 2008. Concepts of neural nitric oxide-mediated transmission. Eur. J. Neurosci. 27, 2783–2802. Giulian, D., Baker, T.J., 1986. Characterization of ameboid microglia isolated from developing mammalian brain. J. Neurosci. 6, 2163–2178. Guillemin, G.J., Brew, B.J., 2004. Microglia, macrophages, perivascular macrophages, and pericytes: a review of function and identification. J. Leukoc. Biol. 75, 388–397. Gutie`rrez-Mecinas, M., Crespo, C., Blasco-Iba´n˜ez, J.M., Na´cher, J., Varea, E., Martı´nez-Guijarro, F.J., 2007. Migrating neuroblasts of the rostral migratory stream are putative targets for the action of nitric oxide. Eur. J. Neurosci. 26, 392–402. Haase, A., Bicker, G., 2003. Nitric oxide and cyclic nucleotides are regulators of neuronal migration in an insect embryo. Development 130, 3977–3987. Hanisch, U.-K., Kettenmann, H., 2007. Microglia: active sensor and versatile effector cells in the normal and pathologic brain. Nat. Neurosci. 10, 1387–1394. Hausmann, O.N., 2003. Post-traumatic inflammation following spinal cord injury. Spinal Cord 41, 369–378. Haynes, S.E., Hollopeter, G., Yang, G., Kurpius, D., Dailey, M.E., Gan, W.-B., Julius, D., 2006. The P2Y12 receptor regulates microglial activation by extracellular nucleotides. Nat. Neurosci. 9, 1512–1519. Henn, A., Lund, S., Hedtja¨rn, M., Schrattenholz, A., Po¨rzgen, P., Leist, M., 2009. The suitability of BV2 cells as alternative model system for primary microglia cultures or for animal experiments examining brain inflammation. ALTEX 26, 83–94.
20
brain research 1564 (2014) 9–21
Hofmann, F., Feil, R., Kleppisch, T., Schlossmann, J., 2006. Function of cGMP-dependent protein kinases as revealed by gene deletion. Physiol. Rev. 86, 1–23. Ho¨ltje, M., Hoffmann, A., Hofmann, F., Mucke, C., Große, G., Van Rooijen, N., Kettenmann, H., Just, I., Ahnert-Hilger, G., 2005. Role of Rho GTPase in astrocyte morphology and migratory response during in vitro wound healing. J. Neurochem. 95, 1237–1248. Horvath, R.J., Nutile-McMenemy, N., Alkaitis, M.S., DeLeo, J.A., 2008. Differential migration, LPS-induced cytokine, chemokine, and NO expression in immortalized BV-2 and HAPI cell lines and primary microglial cultures. J. Neurochem. 107, 557–569. Ignarro, L.J., 1990. Nitric oxide. A novel signal transduction mechanism for transcellular communication. Hypertension 16, 477–483. Kreutzberg, G.W., 1996. Microglia: a sensor for pathological events in the CNS. Spec. Issue Glial Signal. 19, 312–318. Kukulski, F., Yebdri, F.B., Lefebvre, J., Warny, M., Tessier, P.A., Se´vigny, J., 2007. Extracellular nucleotides mediate LPSinduced neutrophil migration in vitro and in vivo. J. Leukoc. Biol. 81, 1269–1275. Laurenzi, M.A., Arcuri, C., Rossi, R., Marconi, P., Bocchini, V., 2001. Effects of microenvironment on morphology and function of the microglial cell line BV-2. Neurochem. Res. 26, 1209–1216. Lee, J.Y., Jhun, B.S., Oh, Y.T., Lee, J.H., Choe, W., Baik, H.H., Ha, J., Yoon, K.-S., Kim, S.S., Kang, I., 2006. Activation of adenosine A3 receptor suppresses lipopolysaccharide-induced TNF-α production through inhibition of PI 3-kinase/Akt and NF-κB activation in murine BV2 microglial cells. Neurosci. Lett. 396, 1–6. Liang, C.-C., Park, A.Y., Guan, J.-L., 2007. In vitro scratch assay: a convenient and inexpensive method for analysis of cell migration in vitro. Nat. Protoc. 2, 329–333. Liao, H., 2004. Tenascin-R plays a role in neuroprotection via its distinct domains that coordinate to modulate the microglia function. J. Biol. Chem. 280, 8316–8323. Lieb, K., Engels, S., Fiebich, B.L., 2003. Inhibition of LPS-induced iNOS and NO synthesis in primary rat microglial cells. Neurochem. Int. 42, 131–137. Lindsay, S.L., Ramsey, S., Aitchison, M., Renne´, T., Evans, T.J., 2007. Modulation of lamellipodial structure and dynamics by NO-dependent phosphorylation of VASP Ser239. J. Cell Sci. 120, 3011–3021. Liu, B., 2003. Role of microglia in inflammation-mediated neurodegenerative diseases: mechanisms and strategies for therapeutic intervention. J. Pharmacol. Exp. Ther. 304, 1–7. Maa, M.-C., Chang, M.Y., Chen, Y.-J., Lin, C.-H., Yu, C.J., Yang, Y.L., Li, J., Chen, P.-R., Tang, C.-H., Lei, H.-Y., Leu, T.-H., 2008. Requirement of inducible nitric-oxide synthase in lipopolysaccharide-mediated Src induction and macrophage migration. J. Biol. Chem. 283, 31408–31416. Mahe, D., Fisson, S., Montoni, A., Morel, A., Couez, D., 2001. Identification and IFNγ-regulation of differentially expressed mRNAs in murine microglial and CNS-associated macrophage subpopulations. Mol. Cell. Neurosci. 18, 363–380. Mandal, S., Stanco, A., Buys, E.S., Enikolopov, G., Rubenstein, J.L. R., 2013. Soluble guanylate cyclase generation of cGMP regulates migration of MGE neurons. J. Neurosci. 33, 16897–16914. Minghetti, L., Levi, G., 1998. Microglia as effector cells in brain damage and repair: focus on prostanoids and nitric oxide. Prog. Neurobiol. 54, 99–125. Moncada, S., Palmer, R., Higgs, E., 1989. The biological significance of nitric oxide formation from L-arginine. Biochem. Soc. Trans. 17, 642–644. Morgan, S.C., Taylor, D.L., Pocock, J.M., 2004. Microglia release activators of neuronal proliferation mediated by activation of
mitogen-activated protein kinase, phosphatidylinositol-3kinase/Akt and delta-Notch signalling cascades: microglia enhance neuronal survival and proliferation. J. Neurochem. 90, 89–101. Moss, D.W., Bates, T.E., 2001. Activation of murine microglial cell lines by lipopolysaccharide and interferon-gamma causes NOmediated decreases in mitochondrial and cellular function. Eur. J. Neurosci. 13, 529–538. Nathan, C., 1992. Nitric oxide as a secretory product of mammalian cells. FASEB J. 6, 3051–3064. Niggli, V., 1999. Rho-kinase in human neutrophils: a role in signalling for myosin light chain phosphorylation and cell migration. FEBS Lett. 445, 69–72. Ohsawa, K., Irino, Y., Nakamura, Y., Akazawa, C., Inoue, K., Kohsaka, S., 2007. Involvement of P2 4 and P2Y12 receptors in ATP-induced microglial chemotaxis. Glia 55, 604–616. Ohsawa, K., Kohsaka, S., 2011. Dynamic motility of microglia: purinergic modulation of microglial movement in the normal and pathological brain. Glia 59, 1793–1799. Peunova, N., Scheinker, V., Ravi, K., Enikolopov, G., 2007. Nitric oxide coordinates cell proliferation and cell movements during early development of Xenopus. Cell Cycle 6, 3132–3144. Pfitzer, G., 2001. Invited Review: regulation of myosin phosphorylation in smooth muscle. J. Appl. Physiol. 91, 497–503. Polazzi, E., Gianni, T., Contestabile, A., 2001. Microglial cells protect cerebellar granule neurons from apoptosis: evidence for reciprocal signaling. Glia 36, 271–280. Raftopoulou, M., Hall, A., 2004. Cell migration: Rho GTPases lead the way. Dev. Biol. 265, 23–32. Ramakers, G.J.A., Moolenaar, W.H., 1998. Regulation of astrocyte morphology by RhoA and lysophosphatidic acid. Exp. Cell Res. 245, 252–262. Sauzeau, V., Rolli-Derkinderen, M., Marionneau, C., Loirand, G., Pacaud, P., 2003. RhoA expression is controlled by nitric oxide through cGMP-dependent protein kinase activation. J. Biol. Chem. 278, 9472–9480. Singh, V., Voss, E.V., Be´nardais, K., Stangel, M., 2012. Effects of 2-chlorodeoxyadenosine (Cladribine) on primary rat microglia. J. Neuroimmune Pharmacol. 7, 939–950. Sporbert, A., Mertsch, K., Smolenski, A., Haseloff, R.F., Scho¨nfelder, G., Paul, M., Ruth, P., Walter, U., Blasig, I.E., 1999. Phosphorylation of vasodilator-stimulated phosphoprotein: a consequence of nitric oxide-and cGMP-mediated signal transduction in brain capillary endothelial cells and astrocytes. Mol. Brain Res. 67, 258–266. Stangel, M., Compston, A., 2001. Polyclonal immunoglobulins (IVIg) modulate nitric oxide production and microglial functions in vitro via Fc receptors. J. Neuroimmunol. 112, 63–71. Stansley, B., Post, J., Hensley, K., 2012. A comparative review of cell culture systems for the study of microglial biology in Alzheimer’s disease. J. Neuroinflamm. 9, 115. Stern, M., Bo¨ger, N., Eickhoff, R., Lorbeer, C., Kerssen, U., Ziegler, M., Martinelli, G.P., Holstein, G.R., Bicker, G., 2010. Development of nitrergic neurons in the nervous system of the locust embryo. J. Comp. Neurol., 1157–1175. Streit, W.J., Mrak, R.E., Griffin, W.S.T., 2004. Microglia and neuroinflammation: a pathological perspective. J. Neuroinflamm. 1, 14. Tegenge, M.A., Bicker, G., 2009. Nitric oxide and cGMP signal transduction positively regulates the motility of human neuronal precursor (NT2) cells. J. Neurochem. 110, 1828–1841. Tegenge, M.A., Rockel, T.D., Fritsche, E., Bicker, G., 2011. Nitric oxide stimulates human neural progenitor cell migration via cGMP-mediated signal transduction. Cell. Mol. Life Sci. 68, 2089–2099.
brain research 1564 (2014) 9–21
Terazawa, R., Akimoto, N., Kato, T., Itoh, T., Fujita, Y., Hamada, N., Deguchi, T., Iinuma, M., Noda, M., Nozawa, Y., Ito, M., 2013. A kavalactone derivative inhibits lipopolysaccharide-stimulated iNOS induction and NO production through activation of Nrf2 signaling in BV2 microglial cells. Pharmacol. Res. 71, 34–43. Trimm, K.R., Rehder, V., 2004. Nitric oxide acts as a slow-down and search signal in developing neurites. Eur. J. Neurosci. 19, 809–818. VanUffelen, B.E., Koster, B.M., de, Broek, den, P.J.V., VanSteveninck, J., Elferink, J.G., 1996. Modulation of neutrophil
21
migration by exogenous gaseous nitric oxide. J. Leukoc. Biol. 60, 94–100. VanUffelen, B.E., Van Der Zee, J., De Koster, B.M., Vansteveninck, J., Elferink, J.G., 1998. Intracellular but not extracellular conversion of nitroxyl anion into nitric oxide leads to stimulation of human neutrophil migration. Biochem. J. 330, 719–722. Vincent, S.R., 1994. Nitric oxide: a radical neurotransmitter in the central nervous system. Prog. Neurobiol. 42, 129–160.
Available online at www.sciencedirect.com
www.elsevier.com/locate/brainres
Research Report
Nitric oxide/cyclic GMP signaling regulates motility of a microglial cell line and primary microglia in vitro Hannah Scheiblicha, Frank Roloffa, Vikramjeet Singhb,c, Martin Stangelb,c, Michael Sterna, Gerd Bickera,c,n a Division of Cell Biology, University of Veterinary Medicine Hannover, Bischofsholer Damm 15/102, D-30173 Hannover, Germany b Department of Neurology, Hannover Medical School, Germany c Center for Systems Neuroscience, Hannover, Germany
art i cle i nfo
ab st rac t
Article history:
Microglia are the resident immune cells of the brain, which become rapidly activated and
Accepted 30 March 2014
migrate to the site of insult in brain infection and disease. Activated microglia generate large
Available online 5 April 2014
amounts of the highly reactive messenger molecule nitric oxide (NO). NO is able to raise cyclic
Keywords:
GMP levels via binding to soluble guanylyl cyclase. We investigated potential mechanistic
BV-2 cells
links between inflammation, NO signaling, and microglial migration. To monitor cell
Cytoskeleton
migration, we used a scratch wound assay and compared results obtained in the BV-2
Inflammation
microglial line to primary microglia. Incubation with lipopolysaccharide (LPS) as stimulator of
iNOS
acute inflammatory processes enhanced migration of both microglial cell types. LPS activated
Primary microglia
NO production in BV-2 cells and application of an NO donor increased BV-2 cell migration while an NO scavenger reduced motility. Pharmacological inhibition of soluble guanylyl cyclase and the resulting decrease in motility can be rescued by a membrane permeant analog of cGMP. Despite differences in the threshold towards stimulation with the chemical agents, both BV-2 cells and primary microglia react in a similar way. The important role of NO/cGMP as positive regulator of microglial migration, the downstream targets of the signaling cascade, and resulting cytoskeletal changes can be conveniently investigated in a microglial cell line. & 2014 Elsevier B.V. All rights reserved.
1.
Introduction
Local inflammation within the central nervous system (CNS) has been shown to be a common mechanism causing the neuronal loss in a diversity of chronic neurodegenerative
diseases such as Alzheimer's disease, Parkinson's disease, and multiple sclerosis. Inflammatory reactions following acute spinal cord injury induce detrimental, yet also beneficial post-traumatic cellular responses in the CNS parenchyma (Hausmann, 2003). There are several cell types that
n Corresponding author at: Division of Cell Biology, University of Veterinary Medicine Hannover, Bischofsholer Damm 15/102, D-30173 Hannover, Germany. Fax: þ49 511 856 7687. E-mail addresses: [email protected] (H. Scheiblich), [email protected] (F. Roloff), [email protected] (V. Singh), [email protected] (M. Stangel), [email protected] (M. Stern), [email protected] (G. Bicker).
http://dx.doi.org/10.1016/j.brainres.2014.03.048 0006-8993/& 2014 Elsevier B.V. All rights reserved.
10
brain research 1564 (2014) 9–21
have been implicated in the response to inflammatory stimuli, but it has been postulated that excessive activation or unregulated activation of microglia are mediators of inflammation-mediated neuronal loss in most neurodegenerative circumstances. As resident immune effector cells in the CNS microglia constantly survey their microenvironment to respond to homeostatic changes. In the healthy adult brain, the baseline motility of microglia is characterized by the constant movement of their highly branched cellular processes (Ohsawa and Kohsaka, 2011). Upon detecting environmental stimuli like the inflammogen lipopolysaccharide (LPS), a component of the outer membrane of Gramnegative bacteria, microglia become rapidly activated, retract their processes, and migrate to the tissue damage or the site of injury (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Kreutzberg, 1996) to secrete cytotoxic substances (Liu, 2003; Moss and Bates, 2001) and tumor necrosis factor alpha (Lee et al., 2006) or to release neurotrophic compounds for regeneration and anti-inflammatory factors (Batchelor et al., 2002; Liao, 2004; Morgan et al., 2004; Polazzi et al., 2001; Streit et al., 2004). Insights into mechanisms of activation and migration may contribute to pinpoint microglia as a key pharmacological target in the treatment of neurodegeneration and traumatic injury. The messenger molecule nitric oxide (NO) which is generated at the lesion site has been shown to act as chemoattractant for activated microglia toward the site of injury and as a potential stop signal to cause accumulation of microglia at the lesion (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Haynes et al., 2006; Ohsawa et al., 2007). Several other studies have demonstrated that NO signal transduction regulates cellular motility during the development of vertebrate and invertebrate nervous systems (Gutièrrez-Mecinas et al., 2007; Haase and Bicker, 2003; Mandal et al., 2013; Peunova et al., 2007; Tegenge and Bicker, 2009; Tegenge et al., 2011; Trimm and Rehder, 2004) NO is generated by the enzyme nitric oxide synthase (NOS). In microglia the inducible NOS isoform (iNOS) has been identified to synthesize continuously high levels of NO (Minghetti and Levi, 1998; Nathan, 1992; Vincent, 1994) that is responsible for the defense of the CNS. Microglia are particularly resilient to high concentrations of this compound (Minghetti and Levi, 1998). The main target of NO is the cyclic guanosine monophosphate (cGMP) synthesizing enzyme soluble guanylate cyclase (sGC) that generates cGMP from guanosine triphosphate (GTP) (Garthwaite, 2008; Ignarro, 1990; Moncada et al., 1989). Cyclic GMP itself activates downstream effectors such as protein kinases G (PKG) (Hofmann et al., 2006) and cyclic nucleotidegated ion channels (Bender and Beavo, 2006). PKGs are mediators of NO/cGMP effects (Baltrons et al., 2008) like the phosphorylation of effector proteins, for example members of the Rho GTPase family (Garthwaite, 2008) which are known to be involved in the organization of the actin cytoskeleton and are key regulators of cell migration. A variety of methods at different microscopic scales have been applied to investigate cytoskeletal changes and cell motility, ranging from chemotaxis assays across micropore membranes (Boyden, 1962) to sophisticated biophysical measurements of the force generated by actin polymerization during lamellipodial protrusion (Fuhs et al., 2014). Here, we used a scratch wound assay on both the microglial cell line
BV-2 and primary microglia to study the modulation of microglial migration by NO/cGMP signal transduction in vitro. This rather simple assay allows for microscopical imaging of cell morphology. We confirmed that incubation in LPS upregulates the production of NO in BV-2 cells (Henn et al., 2009). Application of small bioactive enzyme activators and inhibitors of NO/cGMP signaling caused significant cytoskeletal changes. The NO/cGMP signaling pathway positively modulates migration of both BV-2 cells and primary microglia in a functional assay.
2.
Results
2.1.
LPS-induced microglial migration
To model the migration of activated microglia, we applied LPS as inflammatory stimulus to adherent cell cultures of BV-2 cells and primary microglia. Even though BV-2 cells needed tenthousandfold higher LPS concentrations than primary microglia (Fig. 2a and b), the migration distance in a scratch wound migration assay for both cell types was significantly enhanced upon LPS stimulation. Concentrations of 10 mg/ml LPS increased migration of BV-2 cells more than twofold up to 210.1726.2% (Fig. 2a) of control levels. This amounts to an enhancement of the cell migration velocity from 16.1272.65 mm/h (control) up to 30.5374.57 mm/h. In primary microglia 1 ng/ml LPS increased the response of the cells up to 130.375.2% (Fig. 2b), corresponding to an increment of the absolute migration velocity of 14.327 1.47 mm/h (control) up to 18.9671.47 mm/h. Higher concentrations of LPS reduced for both cell types the motility in the scratch wound assay. To visualize effects of LPS stimulation on cell morphology, we performed phalloidin staining against the F-actin cytoskeleton after migration experiments. Both microglial cell types migrated with long F-actin-positive processes into the scratch (Fig. 2c and d). High magnifications of LPS-treated BV-2 cells revealed a polar and spindle-shaped phenotype (Fig. 2e). This morphology was characterized by F-actin-rich microspike projections around the soma or at the terminations of elongated thin cell extensions. LPS-treated primary microglial cells reorganized into a flat and stretched shape (Fig. 2f). Most of the F-actin network formed thick filamentous bundles. Untreated BV-2 cell controls displayed a round to ameboid shape with thick but short F-actin processes (Fig. 2g), resembling the untreated primary microglial morphology (Fig. 2h). Inflammation in brain tissue is accompanied by an increased NO synthesis in microglial cells via induction of iNOS. To visualize this enzyme in BV-2 cells, we used NADPHdiaphorase staining as a cytochemical marker for the presence of NOS. Cultures of both BV-2 cells and primary microglia showed strong labeling for NADPH-diaphorase (Fig. 3a and b). Using the Griess-Assay for nitrite production, we measured endogenous NO formation in BV-2 cells (control: 0.270.07 mM after 8 h; 1.4970.16 mM after 24 h). Application of LPS caused a dose- and time-dependent increase in NO production after 8 h (11.870.21 mM) and 24 h (54.1671.21 mM) of incubation (Fig. 3c). A rather sensitive receptor molecule for NO is soluble guanylyl cyclase (sGC), an enzyme that synthesizes cGMP (Fig. 1d). We used an enzyme immunoassay to investigate whether BV-2 cells can produce cGMP (Fig. 3d). Because
brain research 1564 (2014) 9–21
11
Fig. 1 – Schematic drawing of the in vitro scratch wound assay: (a)–(c) the extent of cell motility was evaluated by measuring the migration during the closure of a wound that is scratched into a confluent cell monolayer. Closure of the wound was followed by monitoring the advancement of the cell front over 8 h for BV-2 cells and 6 h for primary microglia. (d) Manipulation of NO/cGMP signaling by chemical agents. In microglia the inducible nitric oxide synthase (iNOS) has been identified to synthesize nitric oxide (NO) in response to immunological stimuli such as LPS. NO binds to the soluble guanylyl cyclase (sGC) and regulates activity of the enzyme. Exogenous NO level can be regulated by the NO donor NOC-18 or the NO scavenger cPTIO. The sGC activity can be blocked by the inhibitor ODQ. The membrane-permeable cGMP analog 8-Br-cGMP raises the intracellular cGMP level. Scale bar: 200 lm.
12
brain research 1564 (2014) 9–21
Fig. 2 – LPS was used as a tool to assess inflammatory-mediated response of microglial migration behavior. Application of LPS significantly increased the movement of BV-2 cells (a) and primary microglia (b) at lower concentrations range. Higher concentrations resulted in a decreased migration distance down to the control level. BV-2 cells (c, e) and primary microglia (d, f) incubated with LPS extended long F-actin-positive processes into the scratch. Cells aquired polarity and a spindleshaped phenotype (arrowheads) compared to a control without treatment (BV-2: g; primary microglia: h). Data are presented as mean7SEM of at least three independent experiments. (*po0.05, **po0.01, ***po0.001 compared to control). Cyan: DAPI; green: Phalloidin. Scale bar: 100 lm (cþd); 20 lm (e–h).
brain research 1564 (2014) 9–21
13
cultures in control medium (0.0370.01 pmol/ml), application of the sGC activator YC-1 (3-(50 -Hydroxymethyl-20 -furyl)-1-benzyl indazole) (100 mM) in combination with IBMX (1 mM) caused a highly significant increase in cGMP levels (0.1770.03 pmol/ml). Application of the fast-releasing NO donor NOC-12 (N-Ethyl-2(1-ethyl-2-hydroxy-2-nitrosohydrazino)ethanamine) (500 mM) in the presence of IBMX did also increase cGMP levels significantly (0.1370.02 pmol/ml). The co-application of YC-1þNOC-12þ IBMX again caused an increase in cGMP (0.1770.04 pmol/ml). However, this increase did not show an additive effect of the YC-1 and NOC-12 compounds. Similarly, the incubation in IBMX alone increased cGMP to comparable levels (0.1570.3 pmol/ml), indicating that BV-2 cells already produce cGMP endogenously.
2.2. Exogenous nitric oxide stimulates microglial cell migration To test for a potential role of NO as modulator of microglial motility, we investigated the effect of exogenous NO on BV-2 cells and primary microglial cultures. Application of NOC-18 (2,20 -(hydroxynitrosohydrazino)bis-ethanamine) significantly enhanced the migration of BV-2 cells into the scratch in a dose-dependent manner up to 164.9721.64% (500 mM; 24.297 3.52 mm/h) compared to the control (12.7571.48 mm/h) (Fig. 4a). Migration distance and cell migration velocity of primary microglia was significantly increased only at 10 mM NOC-18 (27.7572.13 mm/h) while application of a higher concentration of 100 mM NOC-18 did not change motility compared to controls (18.2671.98 mm/h) (Fig. 4b). Similar to application of LPS, NOC-18 caused a cytoskeletal reorganization of BV-2 cells and primary microglia resulting in a spindle-shaped phenotype (Fig. 4c and d). Application of the NO scavenger cPTIO (2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide) caused a dose-dependent reduction of BV-2 cell migration distance and velocity down to 11.9476.12% (1.370.6 mm/h) (500 mM) compared to vehicle control (10.0770.76 mm/h) (Fig. 4e). We could not observe any phenotypical difference in cell morphology compared to untreated vehicle control cells (Fig. 4f and f0 ). These data indicate that exogenous NO modulates migration and causes cytoskeletal reorganization of both BV-2 cells and primary microglia. Fig. 3 – We used NADPH-diaphorase staining in BV-2 cells (a) and primary microglia (b) as a marker for NOS expression. When BV-2 cells were stimulated with the inflammogen LPS, nitrite production was highly increased (c). To measure the effect of the fast-releasing NO donor NOC-12 (500 lM) and the sGC activator YC-1 (100 lM) in combination with the PDE inhibitor IBMX (1 mM) on the intracellular cGMP level, a cGMP EIA was performed (d). Data are presented as mean7SEM of three independent experiments (*po0.05, **po0.01, ***po0.001 compared to control). In all cases the detected intracellular cGMP level was increased compared to a control. Scale bar: 50 lm.
cytosolic cGMP is efficiently metabolized by phosphodiesterases (PDEs), we measured cGMP levels in the presence of the PDE blocker IBMX (3-Isobutyl-1-methylxanthine). Compared to
2.3. NO/cGMP signaling is a positive regulator of microglial motility Next, we asked whether the NO/cGMP signaling pathway is involved in the control of microglial motility. We employed small bioactive enzyme activators and inhibitors in both gain and loss of function experiments. In a loss of function experiment we tested whether the sGC inhibitor ODQ (1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one) affects microglial motility in a dose-dependent manner. Indeed, application of ODQ above a concentration of 50 mM ODQ significantly inhibited migration of BV-2 cells and primary microglia (Fig. 5a and b) compared to the vehicle controls. By quantifying the impact of ODQ on cell migration velocity we found that BV-2 cell migration was slowed down to 5.1271.11 mm/h (control: 14.4271.63 mm/h) similar to migration velocity of primary microglia (5.257 1.54 mm/h vs. control 12.8871.53 mm/h). We could not observe
14
brain research 1564 (2014) 9–21
Fig. 4 – Exogenous NO level regulates microglial migration. Exogenous NO released from NOC-18 enhanced motility of BV-2 cells (a) and primary microglia (b). When cells were incubated with NOC-18 (100 lM) morphological changes could be observed. BV-2 cells (c) and primary microglia (d) acquired a spindle-shaped phenotype (arrowheads). The NO scavenger cPTIO trapped NO, resulting in a significantly decreased migration distance of BV-2 cells compared to the vehicle control (e). We could not detect any morphological changes (f) compared to the vehicle control (f0 ). Data are mean7SEM of at least three independent experiments. (*po0.05, **po0.01, ***po0.001 compared to control). Cyan: DAPI; green: Phalloidin. Scale bar: 20 lm.
any morphological difference when BV-2 cells were incubated with ODQ compared to vehicle control (Fig. 5c and c0 ). In contrast primary microglia exhibited a round and flat shape with many tiny microspikes after incubation with ODQ (Fig. 5d and d0 ). Co-application of the cell membrane permeant analog of cGMP, 8-Br-cGMP (8-Bromoguanosine 30 ,50 -cyclic monophosphate sodium salt), reversed the inhibitory effect of ODQ to control levels for both cell types (Fig. 5e and f).
This rescue experiment provides direct evidence for NO/cGMP signal transduction as a positive regulator of microglial cell migration. To exclude cytotoxic effects of the chemical compounds on motility, we measured the cell viability in an Alamar Blue assay directly after migration experiments. All the chemicals used in our migration assay did not affect cell viability of BV-2 cells and primary microglia (Fig. 6).
brain research 1564 (2014) 9–21
15
Fig. 5 – Blocking of the cGMP synthesizing enzyme sGC with ODQ resulted in a decreased cell movement of BV-2 cells (a) and primary microglia (b) compared to the vehicle controls. While BV-2 cells (c) did not change their morphology compared to the vehicle control (c0 ) primary microglia (d) exhibited a round and flat shape with many tiny microspikes in contrast to the vehicle control (d0 ). Exogenous application of the cGMP analog, 8-Br-cGMP, rescued the blocking of cell migration by sGC inhibitors in both cell types (e, f). Data are mean7SEM of at least three independent experiments (*po0.05, **po0.01, ***po0.001 compared to control). Cyan: DAPI; green: Phalloidin. Scale bar: 20 lm.
3.
Discussion
3.1.
Microglial BV-2 cells as a model for primary microglia
In the present paper, we have used chemical manipulations to demonstrate that the NO/cGMP signaling system enhances motility of the BV-2 microglial line in a scratch wound assay (Fig. 1). These results are in line with cell culture and in vivo
studies showing NO/cGMP as positive regulator for primary microglial motility (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Haynes et al., 2006; Ohsawa et al., 2007). In the scratch assay, we have also analyzed the motility of rat primary microglia in response to an enzyme activator (Fig. 4) and inhibitor of the NO/cGMP cascade (Fig. 5). To test for unspecific side effects of the chemical agent on cell
16
brain research 1564 (2014) 9–21
Fig. 6 – Cell viability assay of chemical compounds used in the experiments. The highest concentrations of all used chemical compounds did not affect viability of BV-2 cells (a) and primary microglia (b). Data are mean7SEM in [%] of at least three independent experiments (*po0.05, **po0.01, ***po0.001 compared to control).
migration behavior, we performed in parallel rescue experiments with BV-2 and primary rat microglia (Fig. 5e and f). The isolation of primary microglia from rat brains is a time consuming procedure yielding rather low cell numbers for culturing. For biochemical experiments it is necessary to expand these cell cultures, which is restricted by the limited proliferation capacity of the primary cells (Henn et al., 2009). Moreover, for the investigation of a specific signal transduction pathway it is desirable to avoid the heterogeneity in cellular phenotypes that are characteristic of brain-derived microglia (Guillemin and Brew, 2004; Hanisch and Kettenmann, 2007; Mahe et al., 2001). To reduce animal experimentation, it has been suggested that the rapidly proliferating microglial cell line BV-2 might serve as a valid substitute for primary microglia (Henn et al., 2009). The BV-2 cell line has been derived from immortalized murine neonatal microglia (Blasi et al., 1990). The validity of BV-2 cells as a microglial model system has been suggested on the basis of similar morphological properties, the expression of microglial cell markers, transcriptome analysis, the secretion of cytokines, and functional properties (Blasi et al., 1990; Henn et al., 2009; Horvath et al., 2008; for review see: Stansley et al., 2012). BV-2 cells exhibit short, thick processes similar to the morphology of activated primary microglia (Horvath et al., 2008). They express specific microglial markers including the microglia activation marker Iba-1 (ionized calcium binding adaptor molecule 1), MAC 1 and MAC 2 (macrophage antigen complex 1 and 2), but do not express MAC 3 (macrophage antigen complex 3), GFAP (glial fibrillary acidic protein, astrocyte marker), or galactocerebroside (oligodendrocyte marker) (Horvath et al., 2008). Moreover, BV-2 cells were tested 90% positive for non-specific esterase activity and all cells lacked peroxidase activity, indicative of microglia (Blasi et al., 1990; Giulian and Baker, 1986). Transcriptome analysis indicated that 90% of the genes induced in BV-2 cells by LPS were also found in primary microglia (Henn et al., 2009) and the secretion of cytokines was increased dose dependently (Blasi et al., 1990). Previous studies have also pointed towards a similar physiological behavior of BV-2 cells and primary microglia in in vitro transwell migration experiments or in assays for iNOS expression and generation of NO (Henn et al., 2009).
There are, however, also indications that BV-2 cells can only partially model primary microglia (Henn et al., 2009; for review see: Stansley et al., 2012). One study compared BV-2 cells and primary microglia with regard to activation markers, cell motility and releasable factors (Horvath et al., 2008). Across the dosage range of ng/ml to mg/ml of LPS stimulation, BV-2 cells generated lesser but still substantial amounts of NO compared to primary microglia. Despite some discrepancies in microglial phenotype, our migration assay revealed a striking modulation of BV-2 cell motility by functional NO/cGMP signal transduction. The robust response in the migration assay, cellular homogeneity, and the unlimited proliferation potential make the BV-2 microglial line an ideal model for the investigation of NO/cGMP mediated motility changes.
3.2.
Inflammatory activation of microglia
We found in the scratch wound assay a link between LPSmediated inflammatory activation and cellular motility for both BV-2 cells and primary microglia (Fig. 2). Similar investigations on murine macrophages, human monocytes, and neutrophils showed an LPS-induced increase of cell migration (Kukulski et al., 2007; Maa et al., 2008). In our functional assay, LPS treatment resulted in a significantly increased migration distance and changes in morphology (Fig. 2). Application of LPS promoted the outgrowth of processes in both BV-2 cells and primary microglia. Cells became polarized and one large and flattened process per cell extended into the scratch. Primary microglia was far more sensitive to LPS stimulation (Fig. 2), whereas the BV-2 cells responded better to NO stimulation than the primary microglia (Fig. 4). However, for a straightforward comparison of both cell types, a caveat should be kept in mind. Primary microglia were isolated from neonatal rodent brains and initially kept in a primary mixed glial culture. Compared to adult microglia, this neonatal origin and the subsequent cell culture conditions may have modified the reaction potential of our primary microglia (Henn et al., 2009). Moreover, primary microglia cells are a heterogeneous population. In a pathological context, microglia from different anatomical regions within the brain can exhibit different reaction
brain research 1564 (2014) 9–21
patterns resulting from regional adaptations (Guillemin and Brew, 2004; Hanisch and Kettenmann, 2007; Mahe et al., 2001), whereas BV-2 cells perform more uniformly (Blasi et al., 1990; Henn et al., 2009). Thus, BV-2 cells may serve as useful model in assays where robust microglial cell behavior is required. In macrophages, microglia, and endothelial cells LPS stimulation is accompanied by enhancing the synthesis level of iNOS, resulting in an increased release of NO (Arias-Salvatierra et al., 2011; Fiebich et al., 1998; Lieb et al., 2003; Maa et al., 2008; Stangel and Compston, 2001). Similar to primary microglia, BV-2 cells labeled for NADPH-diaphorase, a cytochemical marker for the presence of NOS (Fig. 3a and b). Moreover, LPS induced in a doseand time-dependent manner the production of NO (Fig. 3c), confirming results from Henn et al. (2009) and Terazawa et al. (2013) about the induction of iNOS transcripts and protein in BV-2 cells. Low concentrations of NO can enhance the movement of migrating neutrophils while high concentrations inhibited the movement and acted as a stop signal to cause accumulation at the site of effect (VanUffelen et al., 1998, 1996). For similar reasons higher LPS concentrations may have no effect on the migration of BV-2 cells and primary microglia in the scratch assay.
3.3. The effect of NO/cGMP signaling on microglial migration To show that BV-2 cells can synthesize cGMP, we used an enzyme immunoassay (Fig. 3d). Treatment with the PDE blocker IBMX caused a significant increase in cGMP levels, indicating endogenous cGMP synthesis. This increase in cGMP could not be enhanced by the additional application of the fast-releasing NO donor NOC-12 and the NO independent sGC activator YC-1, suggesting that endogenous production of cGMP in BV-2 cells already runs at maximum capacity. During neural development NO signal transduction is a positive regulator of motility in various cell types, such as human neuronal precursor cells (Tegenge and Bicker, 2009; Tegenge et al., 2011), immature mouse neurons of the medial ganglionic eminence (Mandal et al., 2013), and insect enteric neuron migration (Haase and Bicker, 2003). Similar mechanisms might influence microglial motility in inflammated neural tissue. Under pathological conditions, NO released from degenerating neurons seems to act as a chemoattractant for recruiting microglia toward the site of injury (Chen et al., 2000; Dibaj et al., 2010; Duan et al., 2009; Haynes et al., 2006; Ohsawa et al., 2007). In the leech NO controls the direction of microglial movement while ATP is required for the activation of microglia or for movement initiation (Duan et al., 2009). In mice NO alone was not sufficient to act as a chemoattractant for microglia when ATP was removed from the extracellular environment (Dibaj et al., 2010). We showed in the scratch assay that exogenous NO can act alone as a positive modulator of microglial migration. The application of the slow-releasing NO donor NOC-18 enhanced migratory behavior significantly while the treatment with the NO scavenger cPTIO attenuated movement (Fig. 4). In our cell culture model the blocking of the NO-induced activation of sGC by ODQ reduced the migration of both BV-2 cells and primary microglia significantly (Fig. 5a and b). This blocking of cGMP signaling was not accompanied by any striking changes in the morphology of BV-2 cells (Fig. 5c)
17
compared to vehicle controls (Fig. 5c0 ). Since we could not observe outgrowth of processes under incubation with ODQ the BV-2 morphology appears of a resting cell phenotype. Our findings are in line with in vivo experiments on mice demonstrating that ODQ blocked the migratory microglial response to laser lesions (Dibaj et al., 2010). The co-application of a cGMP analog completely rescued the decreased cell migration caused by ODQ (Fig. 5e and f). The rescue of the loss of function effect indicates that a certain level of cGMP is required for microglial migration. Application of cGMP to microglia with intact NO/cGMP signaling did not increase motility (data not shown). Presumably, the endogenous cGMP production may keep the motility in an optimal range. Taken together, our data indicate that the NO/cGMP signaling pathway may positively modulate the migration of microglial BV-2 cells in a similar way than in primary microglia in the in vitro scratch assay. In our cell culture system the downstream targets are not known, but cGMP can mobilize the microglial cytoskeleton (Dibaj et al., 2010) by activating the PKG or cyclic nucleotide-gated ion channels (Bender and Beavo, 2006; Hofmann et al., 2006). Further investigations are required to understand the detailed molecular mechanisms by which NO/cGMP signaling regulates cytoskeletal rearrangement and cell motility. Several studies have demonstrated that the regulation of the actin cytoskeleton is caused downstream of the cGMP pathway (Borán and García, 2007; Mandal et al., 2013). PKG regulates the expression and phosphorylation of RhoA, a small GTPase, which in turn activates the Rho kinase (ROCK). ROCK has been shown to regulate various cellular functions, amongst others cytoskeletal reorganization through the phosphorylation of the regulatory subunit of myosin (Pfitzer, 2001; Raftopoulou and Hall, 2004). Inhibition of the Rho/ROCK pathway induces dramatic morphological changes (Borán and García, 2007) and accelerated migration of astrocytes (Hö ltje et al., 2005; Ramakers and Moolenaar, 1998), leukocytes (Alblas et al., 2001), and human neutrophils (Niggli, 1999) suggesting that RhoA/ROCK signaling is critical for the regulation of cellular migration (Sauzeau et al., 2003). Phosphorylation of Enabled/ vasodilator-stimulated phosphoprotein family proteins by PKG has also been found as mechanism for the regulation of actin polymerization (Chen et al., 2007; Lindsay et al., 2007; Sporbert et al., 1999). In summary, this in vitro investigation study showed that the BV-2 microglial line is a useful model for primary microglia in cell migration experiments. Despite some differences in the threshold towards stimulation with chemical agents, both BV-2 cells and primary microglia react in a similar way. We demonstrated in the scratch migration assay that the NO/cGMP signal transduction cascade enhances microglial motility for both cell types. Since this assay allows for optical imaging of the cytoskeletal dynamics, we can now investigate mechanistic links between drug stimulation and resulting microglial migration behavior.
4.
Experimental procedure
4.1.
Materials
All substances were obtained from Sigma (Taufkirchen, Germany) unless otherwise noted.
18
4.2.
brain research 1564 (2014) 9–21
Cell culture
All animals used for cell isolation were treated according to the legal and ethical requirements of the Hannover Medical School (Germany). The procedures complied with the guidelines of animal welfare as laid down by the German Research Council (DFG). Primary microglia cells were isolated by the method of Giulian and Baker (1986) as described previously (Singh et al., 2012). Briefly, brains from neonatal Sprague-Dawley rats were stripped of the meninges and dissociated using mechanical shearing and trypsin. Cells of one to two brains were plated on poly-L-lysine coated T75 culture flasks (Iwaki, BibbySterilin, Stone, UK) and cultivated in DMEM (Gibco, Paisley, UK) supplemented with 10% fetal calf serum (FCS; Bio-Whittaker, Wokingham, UK) and 1% penicillin/streptomycin (Sigma-Aldrich, Steinheim, Germany). While using an orbital shaker-incubator, loosely attached microglia were removed from the astrocytic monolayer and mobilized into the supernatant after an incubation period of 7–10 days. Cells were plated at 5 105 per well in 24-well-plates (Costar, Corning GmbH, Kaiserslautern, Germany) and incubated overnight at 37 1C/5% CO2 to build a confluent monolayer. The murine microglial BV-2 cell line developed by (Blasi et al., 1990) was a gift from Prof. Dr. M. Leist (Department of In vitro Toxicology and Biomedicine, University of Konstanz, Germany). Semi-adherent cells were maintained and cultivated in T175 culture flasks (Nunc, Thermo Fisher Scientific, Langenselbold, Germany) in DMEM (Gibco-Invitrogen, Karlsruhe, Germany) supplemented with 10% FCS and 1% penicillin/streptomycin (Gibco-Invitrogen). Confluent cultures were passaged by using a cell scraper (Greiner Bio-One, Frickenhausen, Germany) for a maximum of 30 passages and plated into poly-D-lysine coated 24-well plates (Costar, Corning GMBH, Kaiserslautern, Germany) in serum-free DMEM to avoid the FCS-mediated activation of the cells (Laurenzi et al., 2001). The cells (3.5 105 cells/well) were allowed to adhere overnight in an incubator at 37 1C/5% CO2 to build a confluent monolayer.
4.3.
Scratch wound assay
To analyze microglial migration we performed a scratch wound assay (Fig. 1a–c) as previously described by Liang et al. (2007). The confluent primary microglia or BV-2 cell monolayer was scraped with a P10 pipette tip to create a wound, followed by a wash with BV-2 ringer (130 mM NaCl, 3 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, 4.5 g D-Glucose, pH 7.4) to remove debris and replaced with 1 ml DMEM-containing chemical compounds interfering with the NO/cGMP pathway (Fig. 1d). cPTIO, ODQ, and YC-1 were dissolved in dimethylsulfoxide (DMSO) as 100 mM stock. NOC-18 (Calbiochem, Darmstadt, Germany) was prepared as a 100 mM stock solution in 10 mM of NaOH solution. All other chemical compounds were directly added into the medium. The cultures were treated with chemicals for at least 6 h (primary microglia) and 8 h (BV-2 cells). Stimulation of primary microglia with LPS occurred with low concentrations (0.1, 1, 10 ng/ml) while BV-2 microglia needed ten-thousand fold higher LPS concentrations (0.1, 1, 10, 100 mg/ml).
To determine the migration of the cells, we acquired images at defined time points. Images were taken at 4.3 magnification in a marked sector as reference point. Scratch widths were measured before the treatment and wound closure was calculated by dividing widths measured after incubation. Wound widths were quantified at 15 distinct locations within the marked sector.
4.4.
Cell viability assay
To monitor cell viability and cytotoxic effects of chemical compounds we used the Alamar Blues assay (Trek Diagnostic Systems, East Grinstead, UK). Directly after the scratch wound assay the medium was completely changed and replaced with 1 ml cell culture medium containing 5% Alamar Blue. Fluorescence intensity of Alamar Blue was measured with a microplate reader (Infinite M200, Tecan, Männedorf, Switzerland) after 2 h.
4.5.
F-actin labeling
Cultures were fixed in 4% paraformaldehyde (PFA) dissolved in phosphate-buffered saline (PBS, 10 mM sodium phosphate, 150 mM NaCl, pH 7.4) for 15 min and permeabilized by washing them tree times for 5 min with PBS containing 0.1% Triton X-100 (PTX). Blocking solution containing PTX and 5% normal goat serum (Vector Laboratories, Burlingame, CA, USA) was applied for 20 min. The F-actin-marker Alexa Fluors 488 phalloidin (1:200, Invitrogen, Karlsruhe, Germany) was applied for 30 min followed by three more washing steps. For nuclear counter-staining we used 40 ,6-diamidino-20 phenylindol-dihydrochloride (DAPI) (Invitrogen) at 0.1 mg/ml for 10 min in PBS.
4.6.
NADPH-diaphorase staining
BV-2 cells and primary microglia were fixed in 4% PFA for 15 min on ice and washed in 0.1 M Tris–HCl (pH 7.8) with 0.1% Triton X-100 three times. Afterwards staining solution (2 mg nitro-blue tetrazolium and 20 mg NADPH in 10 ml PTX) was applied for 60 min at room temperature in the dark. Staining was stopped with three more washing steps with PBS for 5 min. In parallel we used hemocytes from Locusta migratoria as a positive control (Stern et al., 2010) and staining solution without NAPDH on BV-2 cells as a negative control (data not shown). Staining was visualized via light microscopy.
4.7.
Nitric oxide determination
The NO production was determined by measuring the stable breakdown product nitrite using the colorimetric Griess Reagent System (Promega Corporation, Madison, WI, USA) in culture medium. BV-2 cells (8 104 cells/well) in 96-well plates were stimulated for 8 h and 24 h with LPS in the concentration range used for the migration experiments. Supernatant was collected and assay was performed according to the manufacturer's protocol. Absorbance between 520 and 550 nm was read in a microplate reader (Infinite M200, Tecan, Männedorf, Switzerland). Concentration of nitrite was determined by interpolation using a nitrite standard curve.
brain research 1564 (2014) 9–21
4.8.
Measurement of intracellular cGMP
Intracellular cGMP levels were quantified using the Direct cGMP EIA kit (Enzo Life Sciences, Lö rrach, Germany). Briefly, cells (1 106 cells/well) in 6-well plates were allowed to adhere overnight in an incubator and treated 20 min prior to cell lysis with chemical compounds except LPS, LPSþIBMX and IBMX which were applied 24 h before measurements. IBMX was dissolved in DMSO as 100 mM stock. In contrast to migration experiments were NOC-18 was used as a slowreleasing NO donor (half-life of 20 h, 37 1C), NOC-12 (half-life of 327 min, 37 1C) was applied as a fast-releasing NO donor to measure intracellular cGMP. NOC-12 was prepared as a 100 mM stock in 10 mM of NaOH solution. Cell lysis was enhanced by adding 0.1% Triton X-100 to 0.1 M HCl and verified visually via light microscopy. Supernatants were assayed after centrifugation using the optional overnight acetylated version according to the manufacturer's protocol. Optical density at 405 nm was determined photometrically with a microplate reader (Infinite M200, Tecan, Männedorf, Switzerland). Concentrations of intracellular cGMP levels were calculated by interpolation using a cGMP standard curve.
4.9.
Microscopy and data analysis
The scratch wound assay and immunocytochemical stainings were examined with a Zeiss Axiovert 200 fluorescence microscope (Zeiss, Gö ttingen, Germany) equipped with a Cool SNAP digital camera (Visitron Systems GmbH, Puchheim, Germany). Acquired images were processed using ImageJ (Wayne Rusband, National Institute of Health, USA). The data were presented as mean7SEM of at least three independent experiments. Statistical comparisons of controls versus treatment were performed with one-way analysis of variance. Levels of significance are indicated as *po0.05, **po0.01, ***po0.001.
Acknowledgment This study was supported by a grant from the German Research Foundation (DFG, FG1103) to Gerd Bicker and to Martin Stangel. We would like to thank Prof. Dr. M. Leist (University of Konstanz, Germany) for the gift of the BV-2 cell line and Saime Tan for technical support.
r e f e r e n c e s
Alblas, J., Ulfman, L., Hordijk, P., Koenderman, L., 2001. Activation of RhoA and ROCK are essential for detachment of migrating leukocytes. Mol. Biol. Cell 12, 2137–2145. Arias-Salvatierra, D., Silbergeld, E.K., Acosta-Saavedra, L.C., Calderon-Aranda, E.S., 2011. Role of nitric oxide produced by iNOS through NF-κB pathway in migration of cerebellar granule neurons induced by lipopolysaccharide. Cell. Signal. 23, 425–435. Baltrons, M.A., Bora´n, M.S., Pifarre´, P., Garcı´a, A., 2008. Regulation and function of cyclic GMP-mediated pathways in glial cells. Neurochem. Res. 33, 2427–2435.
19
Batchelor, P.E., Porritt, M.J., Martinello, P., Parish, C.L., Liberatore, G.T., Donnan, G.A., Howells, D.W., 2002. Macrophages and microglia produce local trophic gradients that stimulate axonal sprouting toward but not beyond the wound edge. Mol. Cell. Neurosci. 21, 436–453. Bender, A.T., Beavo, J.A., 2006. Cyclic nucleotide phosphodiesterases: molecular regulation to clinical use. Pharmacol. Rev. 58, 488–520. Blasi, E., Barluzzi, R., Bocchini, V., Mazzolla, R., Bistoni, F., 1990. Immortalization of murine microglial cells by a v-raf/v-myc carrying retrovirus. J. Neuroimmunol. 27, 229–237. Bora´n, M.S., Garcı´a, A., 2007. The cyclic GMP-protein kinase G pathway regulates cytoskeleton dynamics and motility in astrocytes. J. Neurochem. 102, 216–230. Boyden, S., 1962. The chemotactic effect of mixtures of antibody and antigen on polymorphonuclear leucocytes. J. Exp. Med. 115, 453–466. Chen, A., Kumar, S.M., Sahley, C.L., Muller, K.J., 2000. Nitric oxide influences injury-induced microglial migration and accumulation in the leech CNS. J. Neurosci. 20, 1036–1043. Chen, H., Levine, Y.C., Golan, D.E., Michel, T., Lin, A.J., 2007. ANPinitiated CcGMP pathways regulate VASP phosphorylation and angiogenesis in vascular endothelium. J. Biol. Chem. 283, 4439–4447. Dibaj, P., Nadrigny, F., Steffens, H., Scheller, A., Hirrlinger, J., Schomburg, E.D., Neusch, C., Kirchhoff, F., 2010. NO mediates microglial response to acute spinal cord injury under ATP control in vivo. Glia 58, 1133–1144. Duan, Y., Sahley, C.L., Muller, K.J., 2009. ATP and NO dually control migration of microglia to nerve lesions. Dev. Neurobiol. 69, 60–72. Fiebich, B.L., Butcher, R.D., Gebicke-Haerter, P.J., 1998. Protein kinase C-mediated regulation of inducible nitric oxide synthase expression in cultured microglial cells. J. Neuroimmunol. 92, 170–178. Fuhs, T., Goegler, M., Brunner, C.A., Wolgemuth, C.W., Kaes, J.A., 2014. Causes of retrograde flow in fish keratocytes. Cytoskeleton 71, 24–35. Garthwaite, J., 2008. Concepts of neural nitric oxide-mediated transmission. Eur. J. Neurosci. 27, 2783–2802. Giulian, D., Baker, T.J., 1986. Characterization of ameboid microglia isolated from developing mammalian brain. J. Neurosci. 6, 2163–2178. Guillemin, G.J., Brew, B.J., 2004. Microglia, macrophages, perivascular macrophages, and pericytes: a review of function and identification. J. Leukoc. Biol. 75, 388–397. Gutie`rrez-Mecinas, M., Crespo, C., Blasco-Iba´n˜ez, J.M., Na´cher, J., Varea, E., Martı´nez-Guijarro, F.J., 2007. Migrating neuroblasts of the rostral migratory stream are putative targets for the action of nitric oxide. Eur. J. Neurosci. 26, 392–402. Haase, A., Bicker, G., 2003. Nitric oxide and cyclic nucleotides are regulators of neuronal migration in an insect embryo. Development 130, 3977–3987. Hanisch, U.-K., Kettenmann, H., 2007. Microglia: active sensor and versatile effector cells in the normal and pathologic brain. Nat. Neurosci. 10, 1387–1394. Hausmann, O.N., 2003. Post-traumatic inflammation following spinal cord injury. Spinal Cord 41, 369–378. Haynes, S.E., Hollopeter, G., Yang, G., Kurpius, D., Dailey, M.E., Gan, W.-B., Julius, D., 2006. The P2Y12 receptor regulates microglial activation by extracellular nucleotides. Nat. Neurosci. 9, 1512–1519. Henn, A., Lund, S., Hedtja¨rn, M., Schrattenholz, A., Po¨rzgen, P., Leist, M., 2009. The suitability of BV2 cells as alternative model system for primary microglia cultures or for animal experiments examining brain inflammation. ALTEX 26, 83–94.
20
brain research 1564 (2014) 9–21
Hofmann, F., Feil, R., Kleppisch, T., Schlossmann, J., 2006. Function of cGMP-dependent protein kinases as revealed by gene deletion. Physiol. Rev. 86, 1–23. Ho¨ltje, M., Hoffmann, A., Hofmann, F., Mucke, C., Große, G., Van Rooijen, N., Kettenmann, H., Just, I., Ahnert-Hilger, G., 2005. Role of Rho GTPase in astrocyte morphology and migratory response during in vitro wound healing. J. Neurochem. 95, 1237–1248. Horvath, R.J., Nutile-McMenemy, N., Alkaitis, M.S., DeLeo, J.A., 2008. Differential migration, LPS-induced cytokine, chemokine, and NO expression in immortalized BV-2 and HAPI cell lines and primary microglial cultures. J. Neurochem. 107, 557–569. Ignarro, L.J., 1990. Nitric oxide. A novel signal transduction mechanism for transcellular communication. Hypertension 16, 477–483. Kreutzberg, G.W., 1996. Microglia: a sensor for pathological events in the CNS. Spec. Issue Glial Signal. 19, 312–318. Kukulski, F., Yebdri, F.B., Lefebvre, J., Warny, M., Tessier, P.A., Se´vigny, J., 2007. Extracellular nucleotides mediate LPSinduced neutrophil migration in vitro and in vivo. J. Leukoc. Biol. 81, 1269–1275. Laurenzi, M.A., Arcuri, C., Rossi, R., Marconi, P., Bocchini, V., 2001. Effects of microenvironment on morphology and function of the microglial cell line BV-2. Neurochem. Res. 26, 1209–1216. Lee, J.Y., Jhun, B.S., Oh, Y.T., Lee, J.H., Choe, W., Baik, H.H., Ha, J., Yoon, K.-S., Kim, S.S., Kang, I., 2006. Activation of adenosine A3 receptor suppresses lipopolysaccharide-induced TNF-α production through inhibition of PI 3-kinase/Akt and NF-κB activation in murine BV2 microglial cells. Neurosci. Lett. 396, 1–6. Liang, C.-C., Park, A.Y., Guan, J.-L., 2007. In vitro scratch assay: a convenient and inexpensive method for analysis of cell migration in vitro. Nat. Protoc. 2, 329–333. Liao, H., 2004. Tenascin-R plays a role in neuroprotection via its distinct domains that coordinate to modulate the microglia function. J. Biol. Chem. 280, 8316–8323. Lieb, K., Engels, S., Fiebich, B.L., 2003. Inhibition of LPS-induced iNOS and NO synthesis in primary rat microglial cells. Neurochem. Int. 42, 131–137. Lindsay, S.L., Ramsey, S., Aitchison, M., Renne´, T., Evans, T.J., 2007. Modulation of lamellipodial structure and dynamics by NO-dependent phosphorylation of VASP Ser239. J. Cell Sci. 120, 3011–3021. Liu, B., 2003. Role of microglia in inflammation-mediated neurodegenerative diseases: mechanisms and strategies for therapeutic intervention. J. Pharmacol. Exp. Ther. 304, 1–7. Maa, M.-C., Chang, M.Y., Chen, Y.-J., Lin, C.-H., Yu, C.J., Yang, Y.L., Li, J., Chen, P.-R., Tang, C.-H., Lei, H.-Y., Leu, T.-H., 2008. Requirement of inducible nitric-oxide synthase in lipopolysaccharide-mediated Src induction and macrophage migration. J. Biol. Chem. 283, 31408–31416. Mahe, D., Fisson, S., Montoni, A., Morel, A., Couez, D., 2001. Identification and IFNγ-regulation of differentially expressed mRNAs in murine microglial and CNS-associated macrophage subpopulations. Mol. Cell. Neurosci. 18, 363–380. Mandal, S., Stanco, A., Buys, E.S., Enikolopov, G., Rubenstein, J.L. R., 2013. Soluble guanylate cyclase generation of cGMP regulates migration of MGE neurons. J. Neurosci. 33, 16897–16914. Minghetti, L., Levi, G., 1998. Microglia as effector cells in brain damage and repair: focus on prostanoids and nitric oxide. Prog. Neurobiol. 54, 99–125. Moncada, S., Palmer, R., Higgs, E., 1989. The biological significance of nitric oxide formation from L-arginine. Biochem. Soc. Trans. 17, 642–644. Morgan, S.C., Taylor, D.L., Pocock, J.M., 2004. Microglia release activators of neuronal proliferation mediated by activation of
mitogen-activated protein kinase, phosphatidylinositol-3kinase/Akt and delta-Notch signalling cascades: microglia enhance neuronal survival and proliferation. J. Neurochem. 90, 89–101. Moss, D.W., Bates, T.E., 2001. Activation of murine microglial cell lines by lipopolysaccharide and interferon-gamma causes NOmediated decreases in mitochondrial and cellular function. Eur. J. Neurosci. 13, 529–538. Nathan, C., 1992. Nitric oxide as a secretory product of mammalian cells. FASEB J. 6, 3051–3064. Niggli, V., 1999. Rho-kinase in human neutrophils: a role in signalling for myosin light chain phosphorylation and cell migration. FEBS Lett. 445, 69–72. Ohsawa, K., Irino, Y., Nakamura, Y., Akazawa, C., Inoue, K., Kohsaka, S., 2007. Involvement of P2 4 and P2Y12 receptors in ATP-induced microglial chemotaxis. Glia 55, 604–616. Ohsawa, K., Kohsaka, S., 2011. Dynamic motility of microglia: purinergic modulation of microglial movement in the normal and pathological brain. Glia 59, 1793–1799. Peunova, N., Scheinker, V., Ravi, K., Enikolopov, G., 2007. Nitric oxide coordinates cell proliferation and cell movements during early development of Xenopus. Cell Cycle 6, 3132–3144. Pfitzer, G., 2001. Invited Review: regulation of myosin phosphorylation in smooth muscle. J. Appl. Physiol. 91, 497–503. Polazzi, E., Gianni, T., Contestabile, A., 2001. Microglial cells protect cerebellar granule neurons from apoptosis: evidence for reciprocal signaling. Glia 36, 271–280. Raftopoulou, M., Hall, A., 2004. Cell migration: Rho GTPases lead the way. Dev. Biol. 265, 23–32. Ramakers, G.J.A., Moolenaar, W.H., 1998. Regulation of astrocyte morphology by RhoA and lysophosphatidic acid. Exp. Cell Res. 245, 252–262. Sauzeau, V., Rolli-Derkinderen, M., Marionneau, C., Loirand, G., Pacaud, P., 2003. RhoA expression is controlled by nitric oxide through cGMP-dependent protein kinase activation. J. Biol. Chem. 278, 9472–9480. Singh, V., Voss, E.V., Be´nardais, K., Stangel, M., 2012. Effects of 2-chlorodeoxyadenosine (Cladribine) on primary rat microglia. J. Neuroimmune Pharmacol. 7, 939–950. Sporbert, A., Mertsch, K., Smolenski, A., Haseloff, R.F., Scho¨nfelder, G., Paul, M., Ruth, P., Walter, U., Blasig, I.E., 1999. Phosphorylation of vasodilator-stimulated phosphoprotein: a consequence of nitric oxide-and cGMP-mediated signal transduction in brain capillary endothelial cells and astrocytes. Mol. Brain Res. 67, 258–266. Stangel, M., Compston, A., 2001. Polyclonal immunoglobulins (IVIg) modulate nitric oxide production and microglial functions in vitro via Fc receptors. J. Neuroimmunol. 112, 63–71. Stansley, B., Post, J., Hensley, K., 2012. A comparative review of cell culture systems for the study of microglial biology in Alzheimer’s disease. J. Neuroinflamm. 9, 115. Stern, M., Bo¨ger, N., Eickhoff, R., Lorbeer, C., Kerssen, U., Ziegler, M., Martinelli, G.P., Holstein, G.R., Bicker, G., 2010. Development of nitrergic neurons in the nervous system of the locust embryo. J. Comp. Neurol., 1157–1175. Streit, W.J., Mrak, R.E., Griffin, W.S.T., 2004. Microglia and neuroinflammation: a pathological perspective. J. Neuroinflamm. 1, 14. Tegenge, M.A., Bicker, G., 2009. Nitric oxide and cGMP signal transduction positively regulates the motility of human neuronal precursor (NT2) cells. J. Neurochem. 110, 1828–1841. Tegenge, M.A., Rockel, T.D., Fritsche, E., Bicker, G., 2011. Nitric oxide stimulates human neural progenitor cell migration via cGMP-mediated signal transduction. Cell. Mol. Life Sci. 68, 2089–2099.
brain research 1564 (2014) 9–21
Terazawa, R., Akimoto, N., Kato, T., Itoh, T., Fujita, Y., Hamada, N., Deguchi, T., Iinuma, M., Noda, M., Nozawa, Y., Ito, M., 2013. A kavalactone derivative inhibits lipopolysaccharide-stimulated iNOS induction and NO production through activation of Nrf2 signaling in BV2 microglial cells. Pharmacol. Res. 71, 34–43. Trimm, K.R., Rehder, V., 2004. Nitric oxide acts as a slow-down and search signal in developing neurites. Eur. J. Neurosci. 19, 809–818. VanUffelen, B.E., Koster, B.M., de, Broek, den, P.J.V., VanSteveninck, J., Elferink, J.G., 1996. Modulation of neutrophil
21
migration by exogenous gaseous nitric oxide. J. Leukoc. Biol. 60, 94–100. VanUffelen, B.E., Van Der Zee, J., De Koster, B.M., Vansteveninck, J., Elferink, J.G., 1998. Intracellular but not extracellular conversion of nitroxyl anion into nitric oxide leads to stimulation of human neutrophil migration. Biochem. J. 330, 719–722. Vincent, S.R., 1994. Nitric oxide: a radical neurotransmitter in the central nervous system. Prog. Neurobiol. 42, 129–160.
