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Genes to Cells (2008) 13, 1171-1183. doi:10.1111/j.1365-2443.2008.01235.x
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N-acetyltransferase ARD1-NAT1 regulates neuronal dendritic development

 Noriaki Ohkawa1,2, Shunichiro Sugisaki1,3, Eri Tokunaga1,2, Kazuko Fujitani1, Takahiro Hayasaka1,4,a, Mitsutoshi Setou1,4,a and Kaoru Inokuchi1,2,3,*

1 Mitsubishi Kagaku Institute of Life Sciences, MITILS, 11 Minamiooya, Machida, Tokyo 194-8511, Japan
2 JST, CREST, Honcho 4-1-8, Kawaguchi 332-0012, Japan
3 Graduate School of Environment and Information Sciences, Yokohama National University, Yokohama 240-8501, Japan
4 National Institute for Physiological Science, 5-1 Higashiyama, Myodaiji-cho, Okazaki 444-8787, Japan


   Abstract
Top
 Abstract
Introduction
 Results
 Discussion
 Experimental procedures
 References
 
ARD1 and NAT1 constitute an N-acetyltransferase complex where ARD1 holds the enzymatic activity of the complex. The ARD1–NAT1 complex mediates N-terminal acetylation of nascent polypeptides that emerge from ribosomes after translation. ARD1 may also acetylate the internal lysine residues of proteins. Although ARD1 and NAT1 have been found in the brain, the physiological role and substrates of the ARD1–NAT1 complex in neurons remain unclear. Here we investigated role of N-acetyltransferase activity in the process of neuronal development. Expression of ARD1 and NAT1 increased during dendritic development, and both proteins colocalized with microtubules in dendrites. The ARD1–NAT1 complex displayed acetyltransferase activity against a purified microtubule fraction in vitro. Inhibition of the complex limited the dendritic extension of cultured neurons. These findings suggest that the ARD1–NAT1 complex has acetyltransferase activity against microtubules in dendrites. Regulation by acetyltransferase activity is a novel mechanism that is required for dendritic arborization during neuronal development.


   Introduction
Top
 Abstract
 Introduction
Results
 Discussion
 Experimental procedures
 References
 
The morphological development of neuronal dendrites is a critically important process in the formation of neuronal circuits in the brain (Jan & Jan 2003). The cerebellar Purkinje cell (PC) exhibits one of the most highly complicated dendritic branching patterns seen among the various neuronal cell types. The PC dendritic tree extends directly towards the pial surface, with dendritic branching beginning to develop during the second and thirdrd postnatal weeks in rodents (Altman 1972; Weiss & Pysh 1978). The total area of the dendritic field and the length of the dendritic branches both peak around the end of this period (Weiss & Pysh 1978). The specific molecular mechanisms modulating the dendritic architecture of PCs remain unclear.

Acetylation, a post-translational protein modification that modulates the function of many proteins, is one of the most common protein modifications in eukaryotes. N{alpha}- and N{varepsilon}-acetylation are the two acetylation pattern types. N{alpha}-acetylation, also known as N-terminal acetylation, is observed on approximately 80%–90% of cytosolic mammalian proteins, and a presumed cotranslational enzymatic reaction that acts on the {alpha}-amino group of the N-terminal amino acid (Polevoda & Sherman 2003). The physiological significance of N-terminal acetylation varies with the particular protein, with some proteins requiring this acetylation pattern for the regulation of protein–protein interactions, catalytic activity and stability (Polevoda & Sherman 2003). Many proteins are also known to be acetylated on the {varepsilon}-amino group of the lysine residue, a pattern known as N{varepsilon}-acetylation. This acetylation pattern mediates many diverse functions of proteins, including DNA recognition, protein–protein interactions, and protein stability (Kouzarides 2000). Moreover, {alpha}-tubulin acetylation is primarily associated with stabilized microtubules (MTs) (MacRae 1997; Umeshima et al. 2007). Thus, evidence suggests that protein acetylation is one of the most important reactions to maintain biological processes, including neuronal development (Shalizi et al. 2006).

ARD1 and NAT1 form a stable N-terminal acetyltransferase complex and are conserved from yeast to human (Park & Szostak 1992; Sugiura et al. 2003; Arnesen et al. 2005). It is assumed that ARD1 functions as a catalytic subunit, whereas NAT1 acts as a cofactor in the complex (Asaumi et al. 2005). In addition, ARD1 functions as an N{varepsilon}-acetyltransferase that directs acetylation at Lys532 of HIF1{alpha} (Jeong et al. 2002; Kim et al. 2006). ARD1 and NAT1 are expressed in various subregions of the developing brain (Sugiura et al. 2003), but the physiological functions of the N-acetyltransferase complex in neurons have not yet been characterized.

In this study, we investigated the neuronal functions of the ARD1-NAT1 N-acetyltransferase complex in PCs. ARD1 and NAT1 expressions increased during the dendritic development of PCs. We found that ARD1 and NAT1 colocalized with MTs in cultured hippocampal neurons and fibroblasts, and that ARD1 activity post-translationally mediated acetylation in the purified tubulin fraction in vitro. Over-expression of the dominant-negative forms of ARD1 dramatically limited dendritic development in cultured PCs. Similarly, dendrogenesis was inhibited by ARD1 knockdown. These results indicate a novel mechanism by which the N-acetyltransferase activity of the ARD1–NAT1 complex regulates dendritic arborization in neuronal cells.


   Results
Top
 Abstract
 Introduction
 Results
Discussion
 Experimental procedures
 References
 
N-acetyltransferase is expressed in developing PCs

Between postnatal day 9 (P9) and P20, PC dendrites undergo dramatic extension and branching (Weiss & Pysh 1978). Immunohistochemistry using an antibody against the neuron-specific subtype of β-tubulin, type III β-tubulin, clearly showed the dendritic development of PCs (Fig. 1A). We isolated PCs from frozen brain sections of P12 and P15 mice by a laser capture microdissection method (Fig. 1B). To identify molecules that regulate the dendritic development of PCs, we carried out PC-specific RT-PCR differential display using PC mRNA prepared at several time points (Ohkawa et al. 2007). The results of this screen showed that expression of the NAT1 gene increased during PC development (Fig. 1C). It has been previously reported that NAT1 is a cofactor of ARD1, and that these two proteins form an N-acetyltransferase complex (Park & Szostak 1992; Sugiura et al. 2003; Arnesen et al. 2005). Expression of the genes for these two proteins was examined using in situ hybridization in developing PCs. Expression of NAT1 mRNA increased between P12 and P15 (Fig. 1D, upper panels), and the ARD1 transcript gradually increased in PCs between P9 and P18 (Fig. 1D, lower panels). Immunoblotting showed that the expression of the NAT1 and ARD1 proteins also increased in the cerebellum during the period of PC development (Fig. 1E–H). These data support the possibility that the ARD1–NAT1 complex is involved in PC growth during dendritic development.


Figure 1
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  		Figure 1  Expression of ARD1 and NAT1 in the developing cerebellum. (A) Immunohistochemistry of neuron-specific β-tubulin III. Brains were dissected at P9, P12, P15, or P18. Lobules 4–5 (L4–5) are shown. (B) PC collection from frozen cerebellar sections (30-µm thick, stained with toluidine blue) using the Laser Capture Microdissection System LM2000. The photograph shows a section after laser treatment, with the PCs removed. (C) PC-specific differential display (PC-DD) shows up-regulation of NAT1 gene expression between P12 and P15. Total cellular RNA was prepared independently from the PCs of four animals (two at P12 and two at P15) and subjected to differential display. Arrowhead indicates NAT1 cDNA. (D) In situ hybridization was carried out with ARD1 and NAT1 antisense probes and cerebellar frozen sections (10-µm thick). Brains were dissected at P9, P12, P15 and P18. L4-5 are shown. Scale bars: A, 50 µm; D, 0.1 mm. IGL, internal granule cell layer; ML, molecular layer; PCL, PC layer (indicated by arrowheads). (E and G) Immunoblotting of NAT1, ARD1, and acetylated {alpha}-tubulin using a whole cell lysate from the developing cerebellum. (F and H) Signal level, normalized by {alpha}-tubulin level, was quantified from three independent samples of different animals at each time point. Error bars indicate mean ± SEM.

 
ARD1 and NAT1 colocalize with MTs in dendrites

FLAG epitope-tagged ARD1 (ARD1-FLAG) and Myc epitope–tagged NAT1 (NAT1-Myc) were over-expressed in PtK2 cells to investigate the subcellular distributions of ARD1 and NAT1. Both the tagged proteins displayed a fibrous distribution and colocalized with EGFP-tagged {alpha}-tubulins (Fig. 2). We observed the distribution of endogenous ARD1 and NAT1 using primary cultured hippocampal neurons under low-density culture conditions (Fig. 3), and found that both proteins colocalized with {alpha}-tubulins in the dendritic shafts (Fig. 3A). Both ARD1 and NAT1 also colocalized with acetylated {alpha}-tubulin in neuronal dendrites (Fig. 3B); acetylated {alpha}-tubulin accumulates in stable MT configurations (MacRae 1997; Umeshima et al. 2007), suggesting an interaction of ARD1-NAT1 with MTs. In contrast, ARD1 and NAT1 did not colocalize with F-actin, another cytoskeletal structure (Fig. 3C).


Figure 2
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  		Figure 2  Over-expressed ARD1 and NAT1 colocalized with MTs in cultured epithelial cells. ARD1-FLAG– or NAT1-Myc–pcDNA3 and pEGFP–{alpha}-tubulin were cotransfected in PtK2. Cells were fixed for immnocytochemistry. An anti-FLAG or anti-Myc antibody was used as the primary antibody, and rhodamine-conjugated secondary antibody was used to visualize antibody binding. Scale bar, 20 µm.

 

Figure 3
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  		Figure 3  Intracellular localization of ARD1 and NAT1 in hippocampal neurons under low-density culture conditions (DIV-21–DIV23). Immunocytochemical micrographs of {alpha}-tubulin (A) or acetylated {alpha}-tubulin (B) and ARD1 (left panels) or NAT1 (right panels). The low-density culture condition allowed precise observation of the distribution of ARD1 and NAT1 within individual neurons. (C) Immunocytochemistry of ARD1 and NAT1 was performed with a rhodamine–phalloidin reaction for F-actin labeling. Scale bars, 10 µm.

 
ARD1 activity regulates acetylation levels of the tubulin fraction

ARD1 and NAT1 form a stable complex in mammalian cells (Sugiura et al. 2003). We hypothesized that the ARD1–NAT1 complex regulates the acetylation state of an MT-associated substrate candidate. A number of conserved amino acid residues in ARD1 have been found to be functionally sufficient for N-acetyltransferase enzymatic activity (Table 1) (Tercero et al. 1992; Lu et al. 1996; Neuwald & Landsman 1997). We produced functional mutants of ARD1, 2 and 4 pm, by introducing point mutations at the consensus amino acids of the N-acetyltransferase motif (Table 1). A point mutation of ARD1 similar in nature to that of the 4 pm mutant was previously found to reduce the N-terminal acetyltransferase activity of the ARD1–NAT1 complex (Asaumi et al. 2005).


Figure 1
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  		Table 1  Design of mouse ARD1 point mutants

 
We used an in vitro acetyltransferase (AT) assay using the purified tubulin fraction from porcine brain as a substrate. Immunoprecipitates of the ARD1–NAT1 complex were used as it was difficult to prepare a recombinant protein over 100 kDa, such as NAT1. Enzyme immunoprecipitation has previously been used in tubulin deacetylase (Matsuyama et al. 2002) and polyglutamylase (Janke et al. 2005) assays in vitro. ARD1-FLAG or the FLAG tagged point mutant of ARD1 were over-expressed with NAT1-Myc in COS7 cells (Fig. 4A). Immunoprecipitation of ARD1-FLAG using an anti-FLAG antibody showed that 2 pm ARD1 had an NAT1-binding potential similar to wild-type ARD1 (Fig. 4A, lanes 6 and 7). In contrast, 4 pm ARD1 showed a lower binding affinity to NAT1 than either wild-type or 2 pm ARD1 (Fig. 4A, lane 8). The immunoprecipitated complexes displayed acetyltransferase activity against the purified tubulin fraction in vitro (Fig. 4B). The complex containing wild-type ARD1 significantly increased the acetylation level of the substrates compared with the immunoprecipitate from mock-transfected cells. In contrast, the immunoprecipitate containing 2 or 4 pm ARD1 did not enhance the acetylation level of the tubulin fraction (fold index: vector control, 1; ARD1, 3.08 ± 0.41; 2 pm, 1.33 ± 0.07, 4 pm, 1.08 ± 0.15). These results suggest that acetylation of the purified tubulin fraction is regulated by the ARD1–NAT1 complex in an ARD1 enzymatic activity-dependent manner.


Figure 4
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  		Figure 4  The ARD1–NAT1 complex positively regulates acetylation of tubulins in an ARD1 acetyltransferase activity–dependent manner. (A) ARD1 and NAT1 form a stable complex. ARD1-FLAG and NAT1-Myc were co-expressed in COS7 cells and then immunoprecipitation (IP) was performed with anti-FLAG antibody (M2)-conjugated agarose beads. 2 and 4 pm are the point mutants of ARD1-FLAG shown in Table 1. Lanes 1 and 5 show the mock-transfected control. IP, immunoprecipitation; WT, wild-type. (B) In vitro acetyltransferase assay with the tubulin fraction purified from porcine brain. The IP samples shown in (A) were incubated with [3H] acetyl-CoA and the tubulin fraction. Radioactivity was determined by scintillation counting after washout of free [3H] acetyl-CoA. Averaged data were obtained from three independent experiments. Error bars indicate mean ± SEM. Asterisk (*) indicates where P < 0.01 vs. a condition including WT ARD1-FLAG, NAT1-Myc, and tubulin fraction, using Student's t-test.

 
Over-expression of mutant ARD1 limits dendritic development in cultured PCs

The ARD1 mutants 2 and 4 pm exhibited reduced acetyltransferase activity against the purified tubulin fraction, as described above (Fig. 4). These mutants were over-expressed in primary cultured PCs (Fig. 5A,B), and transfected PCs were identified at 14 days in vitro (DIV-14) by staining with anti-calbindin and anti-FLAG antibodies (to visualize wild-type or ARD1 point mutants) or by staining with an anti-calbindin antibody and observation of GFP fluorescence. We observed the localization of endogenous ARD1 in dendritic shaft of cultured PCs (data not shown). Over-expression of wild-type ARD1 produced no obvious change, compared with GFP over-expression, in the total dendritic length in cultured PCs (GFP, 557 ± 53 µm; ARD1, 508 ± 62 µm). In contrast, dendritic development was dramatically limited by over-expression of 2 pm ARD1 (2 pm, 21 ± 9 µm). Over-expression of 4 pm ARD1 also reduced the total dendritic length, but with a milder effect (4 pm, 264 ± 39 µm). The 2 pm mutant interacted more strongly with NAT1 than did the 4 pm mutant (Fig. 4A), and over-expressed ARD1 did not localize to the PC nuclei (Fig. 5C). These results suggest that these mutants function in a dominant-negative manner against the endogenous ARD1–NAT1 complex by sequestering NAT1 in the cytoplasm. This in turn indicates that blocking the function of the endogenous ARD1–NAT1 complex limits dendritic development in cultured PCs.


Figure 5
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  		Figure 5  ARD1 mutants function as dominant-negative forms that limit dendritic extension in cultured PCs. (A) Fluorescence micrographs of GFP-, ARD1-, and HDAC6-over-expressing PCs in primary culture. Cells were transfected at DIV-0 and fixed at DIV-14. Calbindin (red) was used as a PC marker. Expression of GFP, FLAG, or Myc-tagged proteins is shown in green. Scale bar, 50 µm. (B) Quantification of total dendritic branch length of GFP-, ARD1-, and HDAC6-expressing PCs in primary culture. At least three independent experiments were carried out for each constructs. Number of PCs analyzed: GFP, n = 27; ARD1, n = 18; 2 pm, n = 27; 4 pm, n = 22; HDAC6, n = 67; HDAC6 ARD1, n = 27. Error bars indicate mean ± SEM. P-values from Student's t-test are shown in the graph. (C) Micrographs display ARD1-FLAG (green) and nuclei (blue) signals in cultured PCs. Note that for (A) the maximum-intensity projection images were prepared to detect dendritic extension in transfectants; thus, there seem to be ARD1 signals in the nucleus. However, the low intensity image showed that ARD1-FLAG did not distribute to the nuclei of PCs. Arrowheads indicate the location of nucleus. Scale bar, 20 µm. (D) Co-expression of exogenous ARD1 and HDAC6 was observed in all cultured PCs introduced with CMV-L7-ARD1-FLAG and L7-HDAC6-Myc (n = 19). Both expression vectors were co-transfected at DIV-0, and the cells were subjected to immunocytochemistry with anti-FLAG and anti-Myc antibodies at DIV-14. Three independent experiments were performed. Scale bar, 20 µm.

 
Histone deacetylase 6 (HDAC6), a class II HDAC, predominantly localizes in cytoplasm (Verdel et al. 2000), interacts with MTs in vitro, and colocalizes with MTs in cells (Matsuyama et al. 2002; Zhang et al. 2003). One substrate of HDAC6 activity is {alpha}-tubulin (Hubbert et al. 2002; Matsuyama et al. 2002). We next investigated whether over-expression of MT-associated deacetylation activity has a similar effect as ARD1 mutants on the dendritic development of cultured PCs. The total dendritic length of HDAC6 transfectants was significantly reduced compared with that of GFP transfectants (HDAC6, 317 ± 24 µm) (Fig. 5A,B). This effect was rescued by ARD1 co-expression (HDAC6 + ARD1, 434 ± 53 µm), whereby all PCs expressing exogenous ARD1 co-expressed exogenous HDAC6 (n = 19) (Fig. 5D). These results indicate that ARD1 and HDAC6 have opposite functions in the same pathway. Taken together, we suggest that acetyltransferase activity on MTs is a novel cellular mechanism of dendritic development.

ARD1 knockdown significantly reduces dendritic extension in cultured PCs

Finally, we performed an RNA interference experiment to investigate the role played by endogenous ARD1 in dendritic extension (Fig. 6). Anti-ARD1 siRNA decreased the expression of rat (Fig. 6C), human, and mouse (data not shown) ARD1 in HEK293T cells. We introduced Alexa Fluor-546-conjugated non-silencing siRNA and anti-ARD1 siRNA into cultured PCs at DIV-0, and observed the dendritic extension at DIV-14. The RNA interference activity of synthetic siRNAs persists for at least 3 weeks in cultured mammalian neurons (Omi et al. 2004). PCs transfected with anti-ARD1 siRNA showed a significant 42% reduction in the total dendritic length (non-silencing siRNA, 461 ± 30 µm; anti-ARD1 siRNA, 269 ± 49 µm) (Fig. 6A,B). This clearly indicates that ARD1 is an essential factor for the normal dendritic development of cultured PCs.


Figure 6
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  		Figure 6  Knockdown of endogenous ARD1 reduces dendritic growth in PCs. (A) Typical images of PCs in primary culture transfected with non-silencing or anti-ARD1 siRNAs. The siRNAs were introduced into cells at DIV-0 and fixed at DIV-14. Alexa Fluor546-conjugated siRNAs were used in this experiment. Scale bar indicates 50 µm. Arrowheads indicate the same location of each cell. (B) Quantification of total dendritic branch length of PCs transfected with non-silencing siRNA or anti-ARD1 siRNA. At least six independent experiments were carried out. Number of PCs analyzed: non-silencing siRNA, n = 21; anti-ARD1 siRNA, n = 21. Error bars indicate mean ± SEM. Asterisk (*): P < 0.005 according to Student's t-test. (C) Efficacy of rat ARD1 knockdown by siRNAs. Immunoblotting of whole cell lysate from HEK293T cells transfected with the indicated siRNAs and rat ARD1-FLAG expression vector. {alpha}-tubulin provided a loading control.

 

   Discussion
Top
 Abstract
 Introduction
 Results
 Discussion
Experimental procedures
 References
 
Expression of ARD1 and NAT1 are related to neuronal development

The ARD1–NAT1 complex is a highly conserved N-acetyltransferase, and the two factors are present in almost all tissues (Sugiura et al. 2001; Jeong et al. 2002; Sugiura et al. 2003). We identified NAT1 as a gene increased during PC development by screening of a PC-specific differential display. ARD1 expression was also increased during the same period of PC development (Fig. 1). While the expression of both genes is down-regulated from embryonic day to neonatal day in developing brain subregions, the expression returns during postnatal development in the hippocampus (Sugiura et al. 2003). This suggests that the regulation of ARD1 and NAT1 expression is related to neuronal development, especially in the hippocampus and in the PCs of the cerebellum. Neurons in these two brain subregions are known to mediate neuronal plasticity. Thus, the expression of NAT1 and ARD1 may be regulated by and correlated with the development of specific neurons which participate in plasticity after maturation.

The ARD1–NAT1 complex is an MT-localizing N-acetyltransferase in neurons

We found that ARD1 and NAT1 displayed a fibrous distribution and had colocalized with MTs in cultured neuronal cells (Figs 2 and 3). We confirmed that mouse ARD1 and NAT1 formed an N-acetyltransferase complex (Fig. 4). An in vitro AT assay indicated that the acetylation of proteins included in the brain tubulin fraction was mediated by ARD1 acetyltransferase activity (Fig. 4B). The neuronal phenotype caused by over-expression of the MT-interacting deacetylase HDAC6 was altered by co-expression of ARD1. These results suggest that the ARD1–NAT1 complex is an MT-localizing N-acetyltransferase in neurons, and that the candidate substrates are also closely associated with MTs.

N-acetyltransferase positively regulates dendritic arborization

There are no reports describing the physiological function of N-acetyltransferase in neuronal cells. Over-expression of the catalytic mutants of ARD1, 2 and 4 pm, limited the dendritic extension of cultured PCs (Fig. 5). These mutants appeared to function as dominant-negative forms whose inhibitory effect correlated with their ability to interact with NAT1. As over-expression of ARD1 alone elicited no obvious change in dendritic development, a balance between the expression levels of ARD1 and NAT1 may be required for the N-acetyltransferase to function in dendritic growth. Moreover, ARD1 knockdown reduced the total branch length of dendrites in developing cultured PCs (Fig. 6). Over the course of PC development, levels of ARD1 and NAT1 mRNAs gradually and coincidently increased (Fig. 1C,D). These data indicate that the ARD1–NAT1 complex positively regulates dendritic development in PCs.

A previous report stated that ARD1 appears in the cytoplasm and is excluded from the nucleus of rat fibroblast cells (Sugiura et al. 2003). We also observed that over-expressed ARD1 predominantly localized to the cytoplasm of cultured PCs (Fig. 5) and HEK293T cells (data not shown). Therefore, it is predicted that the inhibition of dendritic development by the over-expression of ARD1 mutants (Fig. 5) is mediated by a dominant negative effect of the mutants on the cytoplasmic function of the N-acetyltransferase complex. We suggest that the primary function of cytoplasmic ARD1-NAT1 activity is the regulation of dendritic development. That is, the nuclear activity of the N-acetyltransferase complex, for example histone acetylation, is unlikely to be involved in the phenomena described in this study.

Substrates of the ARD1–NAT1 complex for dendritic development and neuronal functions

The identity of substrates of the ARD1–NAT1 complex that participate in dendritic development remain unclear, because we have not identified the substrate from tubulin fraction that contains {alpha}/β-tubulin and so on. Although over-expression of HDAC6, one of the major deacetylases of {alpha}-tubulin (Hubbert et al. 2002; Matsuyama et al. 2002; Zhang et al. 2003), significantly reduced the dendritic arborization of cultured PCs, the phenotype was rescued by ARD1 co-expression (Fig. 5). These results suggest that the activities of ARD1 and HDAC6 oppositely target same pathway. In addition, the two factors may compete with each other for interaction with the same targets, as the effect of HDAC6 on dendritic arborization was affected by ARD1 alone (Fig. 5). The common substrate may be {alpha}-tubulin. Acetylation of {alpha}-tubulin is closely correlated with MT stability (MacRae 1997), which is required for proper dendritic growth (Vaillant et al. 2002; Ohkawa et al. 2007). A recent report indicated that SIRT2 also deacetylases {alpha}-tubulin (North et al. 2003) and negatively regulates oligodendroglial arborization (Li et al. 2007). Therefore, the ARD1–NAT1 complex may mediate acetylation of {alpha}-tubulin and MT stability for dendritic development. To argue this hypothesis, it should be examined in future works.

Kinesins are motor proteins which facilitate trafficking along MTs, and participate in dendritic transport (Setou et al. 2000, 2004), assisting in the regulation of neuronal function (Ikegami et al. 2007). A recent report indicates that MT acetylation promotes binding of kinesin-1 to MT, and influences the recruitment of kinesin-1 to specific MT tracks (Reed et al. 2006). Kinesin-1/KIF5 binds to and transports vesicles containing AMPA receptors and RNA granules containing {alpha}CaMKII and Arc mRNAs in dendrites (Setou et al. 2002; Matsumoto et al. 2007). Thus, it is suggested that {alpha}-tubulin acetylation influences the dendritic transport of components important for synaptic functions. If the ARD1–NAT1 complex mediates tubulin acetylation, the N-acetyltransferase activity may also regulate neural functions. Indeed, the ARD1 and NAT1 expression are maintained in adult hippocampus and cerebellum, both of which are major regions of synaptic plasticity (Sugiura et al. 2003).


   Experimental procedures
Top
 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
References
 
Reagents and antibodies

Fluorescein (FITC)- and rhodamine-conjugated donkey anti-mouse secondary antibodies and anti-rabbit secondary antibodies were purchased from Chemicon (Temecula, CA). The FITC-conjugated donkey anti-chick secondary antibody was purchased from Jackson ImmunoResearch (West Grove, PA). The primary antibodies were from the following sources: rabbit anti-neuron-specific class III β-tubulin, Dr Arimatsu, MITILS, Japan; rabbit and mouse anti-FLAG antibodies, mouse anti-{alpha}-tubulin antibody B-5-1-2, and mouse anti-acetylated {alpha}-tubulin antibody 6-11B-1, Sigma-Aldrich (St. Louis, MO); rabbit (A-14) and mouse (9E10) anti-Myc antibodies, Santa Cruz Biotechnology, Inc (Santa Cruz, CA); and rabbit and mouse anti-calbindin antibodies, Chemicon and Swant Swiss Antibodies (Bellinzona, Switzerland), respectively. DRAQ5 (Biostatus Ltd, Leicestershire, UK), which fluoresces in the far red, was used for nuclear staining.

Laser capture microdissection and PCR-differential display

Laser capture microdissection was performed as described in our previous reports (Fukazawa et al. 2003; Ohkawa et al. 2007). Briefly, mice were sacrificed at P12 or P15, and the brains were immediately frozen on dry ice powder. Sagittal cryosections cut at 30 µm thickness were fixed with 70% ethanol, dehydrated with ethanol, immersed in xylene, and then air-dried. The PC layers were collected using the Laser Capture Microdissection System (LM2000, Arcturus Eng. Inc., Mountain View, CA). Total cellular RNA was extracted from the collected PCs using Sepasol-RNA I (Nacalai Tesque, Kyoto, Japan). PCR-differential display was performed as described in our previous reports (Inokuchi et al. 1996; Hegde et al. 1997; Matsuo et al. 2000). One of the candidate genes was amplified with an arbitrary primer (GGGGTGACGA) and a T12GC anchor primer. DNA sequence analysis and a database search revealed that it included nucleotides 2657–2757 of mouse NMDA receptor-regulated gene 1 (NARG1) (GENBANK accession number: AK078042 [GenBank] ), which is same as the mouse NAT1.

Plasmid construction

A mouse ARD1 cDNA that was FLAG-epitope-tagged at the C-terminus (ARD1-FLAG) was obtained by PCR using the 5' primer 5'-CGGGATCCGCCACCATGAACATCCGCAAT GCTAGGCCCGAAGACC-3', the FLAG-tag–encoding 3' primer 5'-GGAATTCCTACTTGTCATCATCGTCCTTGTA GTCGGAGGCAGAGTCAGAGGCCTCTGAGCTG-3', and the dT-primed cDNAs prepared from mouse brain as templates. The underlined sequences in the 5' and 3' primers correspond to mouse ARD1 cDNA. To obtain human and rat ARD1-FLAG cDNA, dT-primed cDNAs were prepared from placental and brain mRNA, respectively. The same 5' and 3' primers mentioned above were used for PCR. A mouse NAT1-Myc cDNA was similarly obtained using the 5' primer 5'-GGGGTACCGCCACCA TGCCGGCCGTGAGCCTCCCGCCCAA-3' and the Myc-tag-encoding 3' primer 5'-GGGATATCTCACAGGTCTTCCTC ACTGATCAGCTTCTGTTCCTCGATTTCATTGGCCAG TTCTTCCGTTTCTGCAG-3'. An EGFP-{alpha}-tubulin was purchased from Clontech (Mountain View, CA).

PCR mutagenesis to generate mouse ARD1 mutants (2 and 4 pm, see Table 1) was carried out using an overlap-extension reaction (Higuchi et al. 1988) with ARD1-FLAG as a template. PCR products were ligated into the cloning vector pCRII-TOPO (Invitrogen, Carlsbad, CA) according to the manufacturer's instructions. An HDAC6-Myc cDNA was produced. An KpnI and ApaI fragment of 5' region of HDAC6-cHA donated from Dr Yoshida (RIKEN, Japan) was subcloned into pcDNA3 (Invitrogen). A C-terminally Myc-tagged 3' region of HDAC6 cDNA was obtained by PCR using the 5' primer 5'-ATGGGCCCCGCAT GGGTGATGCTGATTACC-3' and the Myc-tag–encoding 3' primer 5'-TGGGCCCGGTACCTTACAGGTCTTCCTCAC TGATCAGCTTCTGTTCCTCGTGTGAGTGGGGCATGT CCTCCCCAAAC-3'. The PCR product was digested with ApaI, and then subcloned into pcDNA3 containing the 5' region of HDAC6 in the appropriate direction.

To generate a mouse ARD1-FLAG or a mouse NAT1-Myc expression vector, ARD1-FLAG or NAT1-Myc cDNA was introduced into pcDNA3 (Invitrogen). To generate a PC-specific mouse ARD1-FLAG or HDAC6-Myc expression plasmid (CMV-L7ARD1-FLAG or L7-HDAC6-Myc), ARD1-FLAG or HDAC6-Myc was introduced into exon 4 of the L7 gene cassette (Oberdick et al. 1990; Ichise et al. 2000), and then, except for L7-HDAC6-Myc, each L7-cDNA fragment was subcloned into the NotI site of pcDNA3. A PC-specific GFP expression vector was described previously (Ohkawa et al. 2007). The L7 promoter in this CMV promoter cassette directed cDNA expression specifically and efficiently to PCs under in vitro culture conditions.

Animals and brain sections

All procedures involving the use of animals complied with the guidelines of the National Institute of Health and were approved by the Animal Care and Use Committee of Mitsubishi Kagaku Institute of Life Sciences. C57BL/6 mice were purchased from CLEA Japan Inc. (Tokyo, Japan). For preparation of frozen brain sections, mice were killed and their brains were dissected and immediately frozen on dry ice. Cryosections (10-µm thick) were air-dried and stored at –80 °C until use for in situ hybridization and immunohistochemistry.

In situ hybridization

The following mouse cDNA fragments were used to generate cRNA probes: NAT1, nucleotides 19–291 of NARG1 EST (GENBANK accession number: AA474587 [GenBank] ); and ARD1, nucleotides 630–830 of ARD1 cDNA (GENBANK accession number: BC027219 [GenBank] ). The NAT1 and ARD1 cDNAs obtained by PCR were subcloned into the vector pCRII-TOPO. The vector was digested with SpeI or EcoRV to generate a template for in vitro transcription to produce an antisense or sense cRNA probe, respectively. Digoxigenin-labeled cRNA probes were produced by transcription with T7 or Sp6 RNA polymerase. In situ hybridization was performed as described previously (Kato et al. 1998; Matsuo et al. 2000).

Preparation of whole cell extracts from cerebellum

Mice were sacrificed at P12, P15, or P18, and dissected cerebellums were immediately frozen with liquid nitrogen. The cerebellums were homogenized in TNE buffer (50 mM Tris–HCl pH 7.5, 150 mM NaCl, and 2 mM EDTA) containing 1x Protease inhibitor cocktail (Sigma) and 1% NP-40. After measurement of the protein concentration using the BCA protein assay kit (Pierce, Rockford, IL), the whole cell lysates were subjected to polyacrylamide gel electrophoresis and immunoblotting.

Immunohistochemistry

Brain sections were fixed with 10% formaldehyde neutral buffer (pH 7.0; Nacalai Tesque) at room temperature (RT) for 30 min. After being washed with phosphate-buffered saline (PBS), the sections were treated with PBST (PBS supplemented with 0.2% Triton X-100) three times at RT for 3 min each, followed by three washes with PBS for 5 min each. The sections were then treated with blocking buffer (2% bovine serum albumin (BSA) in PBS) at RT for 1 h. Reactions with primary antibodies were performed in blocking buffer containing rabbit anti-neuron-specific β-tubulin III antibody (1 : 500) at 4 °C overnight. After three washes with PBS for 10 min each, the sections were incubated with rhodamine-conjugated secondary antibodies at 4 °C overnight. Sections were washed with PBS three times for 10 min each, and then mounted with ProLong Gold antifade reagents (Molecular Probes, Eugene, OR). The fluorescent signals were examined with a laser-scanning confocal microscope (LSM 5 PASCAL, Carl Zeiss, Jena, Germany).

RNA interference

Alexa Fluor-546-labeled non-silencing (designed by Qiagen) and anti-ARD1 siRNA duplexes were synthesized by Qiagen (Valencia, CA). The ARD1 target sequence used was CCAUGGACAUAU CACCUCA, as previously reported (Bilton et al. 2005), which is conserved in mouse, rat, and human ARD1 genes. HEK293T cells were plated in a 35-mm dish, and then transfected with ARD1-FLAG–pcDNA3 (4 µg) and siRNA (final concentration 100 nM) 24 h later using Lipofectamine 2000 (Gibco, Rockville, MD). Two days after the transfection, cells were collected and lysed in SDS sample buffer for SDS polyacrylamide gel electrophoresis and immunoblotting.

Cell culture

COS7 and HEK293T cells were cultured with Dulbecco's Modified Eagle Medium (DMEM) (Invitrogen) supplemented with 10% fetal bovine serum at 37 °C in 10% CO2.

Primary culture and transfection

Cultivation of dissociated primary hippocampal neurons was carried out as described (Goslin et al. 1998; Shoji-Kasai et al. 2007). Briefly, rat hippocampi were dissected from embryonic day 18 embryos and dissociated with papain. The neurons were then plated at a density of approximately 1.0 x 104 cells/cm2 on cover slips coated with poly-L-lysine (1 mg/mL) in DMEM supplemented with 5% horse serum and 5% fetal calf serum. After the neurons had attached to the substrate, the cover slips were inverted onto a dish containing a monolayer of astrocytes and maintained for 17 days in serum-free DMEM with N2 supplement (Gibco), 1 mM sodium pyruvate, and 0.1% ovalbumin.

Mixed cerebellar cell cultures were prepared from rats at embryonic day 18 according to the previously published methods (Furuya et al. 1998; Tabata et al. 2000), except for the culture medium. Briefly, prepared cells were plated at a final density of 4 x 106 cells/mL on poly-L-ornithine-coated coverslips in 24-well plates filled with 40 µL of seeding solution (DMEM/F12 containing 10% fetal calf serum). After incubation for 3 h at 37 °C, 400 µL of culture medium (DMEM/F12 containing B27 (final dilution, 1 : 100; Gibco), N2 supplement (1 : 100; Gibco), and 0.5 ng/mL tri-iodothyronine (Sigma-Aldrich)) was added to each well. The GFP, ARD1-FLAG, and/or HDAC6-Myc expression plasmids (1 µg each) or siRNA (final concentration 100 nM) were transfected using Lipofectamine 2000 (Gibco) immediately after the addition of the culture medium at DIV-0. In the siRNA experiment, the PCs selected and observed were those in which the Alexa Fluor546 signal was detected within the PC soma (revealed by calbindin signal) in Z-axis-stacked confocal micrographs.

Immunocytochemistry

Primary cultured cerebellar and hippocampal neurons or PtK2 cells were fixed in 10% formaldehyde neutral buffer (pH 7.0) at 4 °C for 45 min or at 37 °C for 15 min, respectively. After washing with PBS, the cells were treated with PBST at RT for 3 min, followed by two washes with PBS for 10 min each. The cells were then treated with blocking buffer (2% BSA in PBS) at RT for 1 h. To detect ARD1-FLAG, NAT1-Myc, and HDAC6-Myc transfectants, a mouse or rabbit anti-calbindin antibody (1 : 500–1000) and rabbit anti-FLAG antibody (1 : 1000), or a mouse anti-Myc antibody (1 : 100) diluted in blocking buffer were incubated with the cells at 4 °C overnight. Rabbit anti-ARD1 (from Dr Lillehaug, University of Bergen, 1 : 100), chick anti-NAT1/Tbdn (Ab1272 from Dr Gendron, Memorial University of Newfoundland, 1 : 100) (Gendron et al. 2000), mouse anti-{alpha}-tubulin (1 : 100), and anti-acetylated {alpha}-tubulin (1 : 500) primary antibodies were used to detect endogenous proteins. After three washes with PBS for 10 min each, the cells were incubated with FITC- or rhodamine-conjugated secondary antibodies (1 : 100) and/or Rhodamine–phalloidin (Sigma-Aldrich, 1 : 200) at 4 °C overnight. Cells were washed with PBS (containing DRAQ5 for nuclear staining) three times for 10 min, and then mounted with the ProLong Gold antifade reagent (Molecular Probes). The fluorescent signals were examined using an LSM 5 PASCAL laser-scanning confocal microscope.

Immunoprecipitation and acetyltransferase (AT) assay

Immunoprecipitation was performed essentially as described previously (Yao et al. 2007). COS7 cells were plated on a 100-mm plate, and then transfected with ARD1-FLAG– and NAT1-Myc–pcDNA3 (30 µg each) 24 h later using Lipofectamine 2000 (Gibco). Forty-eight hours after transfection, the cells were collected and lysed in lysis buffer (TNE buffer containing 1% Triton X-100, 1 mM PMSF, 10 µg/mL leupeptin, and 10 µg/mL aprotinin) at 4 °C for 30 min, followed by centrifugation at 14 000 g for 10 min at 4 °C. The supernatant was collected, the protein concentration was measured using a BCA protein assay kit (Pierce), and the supernatant was then used for immunoprecipitation. FLAG-tagged proteins were immunoprecipitated from a total 1.5 mg input cell lysate with anti-FLAG M2 agarose affinity gel (Sigma) overnight at 4 °C. For immunoblotting analysis, the immunoprecipitate was denatured in the SDS sample buffer for SDS polyacrylamide gel electrophoresis. For the AT assay, the immunoprecipitate was lysed in 80 µL of reaction buffer (50 mM Tris–HCl pH 8.0, 10% glycerol, 1 mM DTT, 100 µM EDTA) after a final wash with TBS (20 mM Tris–HCl pH 7.5, 150 mM NaCl). The acetyltransferase reaction was performed as follows: in a 20 µL total reaction volume, 14.3 µL of immunoprecipitate, 2 µL of tubulin fraction purified from porcine brain (Peloquin et al. 2005), 10 µM [3H] acetyl-CoA (TRK688, Amersham Biosciences, Piscataway, NJ), and 2 mM GTP (Sigma) were combined and incubated at 30 °C for 12 h. After the reaction, the sample was applied to a GF/F glass filter (Whatman, Maidstone, UK), and TCA precipitation was performed. Radioactivity was determined by scintillation counting.

Immunoblotting

Chick anti-ARD1 (GenWay Biotech, San Diego, CA, 1 : 100), rabbit anti-NAT1 (from Dr Lillehaug, University of Bergen, 1 : 200), rabbit anti-FLAG (Sigma, 1 : 1000), rabbit anti-Myc (Santa Cruze, 1 : 1000), mouse anti-{alpha}-tubulin (B-5-1-2, Sigma, 1 : 1000), and mouse anti-acetylated {alpha}-tubulin (6-11B-1, Sigma, 1 : 1–5000) antibodies were used as primary antibodies, and peroxidase-conjugated goat anti-rabbit (Jackson ImmunoResearch), rabbit anti-chick (Chemicon), and donkey anti-mouse (Chemicon) antibodies were used as secondary antibodies. SuperSignal West Femto Maximum Sensitivity Substrate (Pierce) was used as the peroxidase substrate for signal detection. The chemiluminescent signals were detected using an LAS1000 image analyzer (Fuji Film, Tokyo, Japan) and measured with Image Gauge software (Fuji Film).

Data analysis

All micrograph analyses were performed using Metamorph Software (Molecular Devices, Downingtown, PA). To measure the signal length alongside the PC dendrites, maximum-intensity projection images were prepared. Signals in each image were traced and calculated using the "Distance" function in the software. All statistical analyses were performed using StatView Software (Abacus Concepts, Berkeley, CA).


   Acknowledgements
 
Authors thank Dr A. Aiba for the L7 cassette; Dr M. Yoshida for the HDAC6 cDNA; Dr Y. Arimatsu for the rabbit anti-neuron-specific class III β-tubulin (Mβ6) antibody; Drs J. R. Lillehaug and T. Arnesen for rabbit anti-ARD1 and NAT1 antibodies; Dr R. L. Gendron for the chick anti-NAT1/Tbdn antibody; and F. Ozawa, R. Okubo-Suzuki, M. Sekiguchi, and Y. Shoji-Kasai for technical assistance. This work was supported by Special Coordinate Funds for Promoting Science and Technology, and in part by a Grant-in-Aid for Scientific Research on the Priority Areas "Neural Circuit Project", "Advanced Brain Science Project", and "Molecular Brain Science" from the Ministry of Education, Culture, Sports, Science and Technology of the Japanese Government to K.I.


   Footnotes
 
Communicated by: Noriko Osumi

aPresent address: Department of Molecular Anatomy, Molecular Imaging Advanced Research Center, Hamamatsu Medical School, Handayama 1-20-1, Hamamatsu, Shizuoka 431-3192, Japan Back

* Correspondence: kaoru{at}mitils.jp


   References
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 Abstract
 Introduction
 Results
 Discussion
 Experimental procedures
 References
 
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Accepted: 21 August 2008




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