홈

find - awk example

Wed, 26 Nov 2008 11:25:52 +0900

Calcurate number and total size of specified files

Save it as fs and chmod +x fs

#!/bin/bash
find ~ -name $1 -exec ls -l {} \; | awk '{total+=$5; count+=1;} END {print total,count;}'

Example

fs *.wmv

-> return the total size of wmv file in all directories and number of files

Quick Look Plugin

Mon, 21 Jan 2008 02:53:35 +0900

레오파드에서 새로 추가된 기능 중 하나인 퀵룩 (Quick Look)

원하는 문서를 단지 훑어보기 위해서 무거운 응용 프로그램을 열지 않고도 파인더 내에서 간단히 파일 내용을 스페이스키 하나 치는 것만으로 바로 볼 수 있게 하여 시간을 절약하게 해 준다. (pdf 를 보는 것 정도야 퀵 룩으로 보나 프리뷰나 큰 차이가 없겠지만 만약 파워포인트나 MS Word라면 천지차이일 것이다)

그러나 한 가지 문제라면, 지원하지 않는 문서 포맷이나 폴더 아이콘에서 퀵 룩을 실행하면 이렇게 썰렁한 아이콘만 내밷고 만다. 이거 폴더인지 퀵룩 안해도 다 안다. -.-;;

Zip 파일의 경우도 역시 썰렁하다,

아이콘 확대사진보다는 폴더나 압축파일에 어떤 내용물이 들어있는지 볼 수 있다면 좀 더 유용하지 않을까?

Zip 파일의 경우에도 마찬가지.

즉, 퀵 룩 기능은 플러그인 방식으로 확장이 가능하다는 것이다. 위에서 예를 보인 두 가지 플러그인을 설치하는 방법을 알아보도록 하자.

- 폴더 리스트 보기및 Zip 파일 내용 보기 플러그인을 아래 링크에서 다운로드받는다.

폴더 리스트 보기 다운로드 세부 정보

Zip 파일 내용 보기 다운로드 세부 정보

압축을 풀고, 압축파일에서 .qlgenerator 라는 확장자를 가진 파일 (Folder.qlgenerator) 만을 Macintosh HD->Library->QuickLook에 복사해 넣는다.

* 로그아웃 - 재 로그인을 하거나, 파인더를 재시동한다. 파인더를 재시동하는 가장 간단한 방법은

독의 파인더 아이콘 위에서 "Option-Right Click" (혹은 Option-Ctrl-Click)

이제 파인더에서 폴더나 Zip 파일을 퀵 룩으로 보면 위의 그림처럼 내용을 살펴볼 수 있을 것이다.

현재까지 나온 퀵 룩의 플러그인들을 정리해 둔 사이트로 이곳이 있는데, 대부분의 플러그인 설치법은 위에 설명한 것과 대동소이하므로 (*.qlgenerator 파일을 /Library/QuickLook에 복사 -> 파인더 재시동) 그리 어렵지 않게 설치할 수 있을 것이다.

위의 사이트에서 소개된 플러그인 중 프로그래머들에게 유용할 만한 것은

QLColorCode 라는 플러그인으로써 퀵룩으로 보는 소스코드에 syntax highlighting 을 적용해 준다.

Vim Cheat Sheet

Thu, 10 May 2007 14:35:03 +0900

MBC

Sat, 28 Apr 2007 06:58:43 +0900

Actin-depolymerizing Factor and Cofilin-1 Play Overlapping Roles in Promoting Rapid F-Actin Depolymerization in Mammalian Nonmuscle Cells

Pirta Hotulainen^*, Eija Paunola, Maria K. Vartiainen, and Pekka Lappalainen

Program in Cellular Biotechnology, Institute of Biotechnology, University of Helsinki, 00014 Helsinki, Finland

Actin-depolymerizing factor (ADF)/cofilins are small actin-binding proteins found in all eukaryotes. In vitro, ADF/cofilins promote actin dynamics by depolymerizing and severing actin filaments. However, whether ADF/cofilins contribute to actin dynamics in cells by disassembling "old" actin filaments or by promoting actin filament assembly through their severing activity is a matter of controversy. Analysis of mammalian ADF/cofilins is further complicated by the presence of multiple isoforms, which may contribute to actin dynamics by different mechanisms. We show that two isoforms, ADF and cofilin-1, are expressed in mouse NIH 3T3, B16F1, and Neuro 2A cells. Depleting cofilin-1 and/or ADF by siRNA leads to an accumulation of F-actin and to an increase in cell size. Cofilin-1 and ADF seem to play overlapping roles in cells, because the knockdown phenotype of either protein could be rescued by overexpression of the other one. Cofilin-1 and ADF knockdown cells also had defects in cell motility and cytokinesis, and these defects were most pronounced when both ADF and cofilin-1 were depleted. Fluorescence recovery after photobleaching analysis and studies with an actin monomer-sequestering drug, latrunculin-A, demonstrated that these phenotypes arose from diminished actin filament depolymerization rates. These data suggest that mammalian ADF and cofilin-1 promote cytoskeletal dynamics by depolymerizing actin filaments and that this activity is critical for several processes such as cytokinesis and cell motility.

Introduction

Actin filaments in nonmuscle cells are highly dynamic and play a critical role in numerous cellular processes, including cell migration, cytokinesis, and polarized growth. These processes rely on the correct spatial and temporal organization of actin filaments that is regulated by numerous actin-binding proteins.

The actin-depolymerizing factor (ADF)/cofilins are a family of small (M_r = 15-20) proteins that bind monomeric and filamentous actin.

Unicellular organisms such as yeasts typically have only one ADF/cofilin, whereas multicellular organisms can have several isoforms (reviewed by Bamburg et al., 1999).
In mammals, there are three different ADF/cofilins: cofilin-1, cofilin-2, and ADF.

cofilin-1 is expressed in most embryonic and adult mouse cells,

cofilin-2 is expressed in muscle

ADF is mainly found in epithelial and neuronal cells (Vartiainen et al., 2002).

Based on in vitro studies, ADF/cofilins enhance the rate of actin filament turnover by depolymerizing filaments at their pointed ends, thereby providing a pool of actin monomers for filament assembly.

ADF/cofilins also sever actin filaments and consequently increase the number of filaments ends (reviewed by Bamburg et al., 1999; Carlier et al., 1999). The mammalian ADF/cofilins are quantitatively different in their activities.

ADF is the most efficient at turning over actin filaments and promotes a stronger pH-dependent actin filament disassembly than cofilin-1 or cofilin-2. The muscle cell-specific cofilin-2 has a weaker actin filament depolymerization activity than the other two and promotes filament assembly rather than disassembly in steady-state assays (Vartiainen et al., 2002; Yeoh et al., 2002).

Genetic studies on Saccharomyces cerevisiae, Drosophila melanogaster, and Caenorhabditis elegans demonstrated that ADF/cofilins are essential for viability (Moon et al., 1993; McKim et al., 1994; Gunsalus et al., 1995). However, whether ADF/cofilins contribute to cytoskeletal dynamics by depolymerizing actin filaments at their pointed ends, or by creating new filament barbed ends for F-actin assembly through their severing activity has remained unclear.

Studies on the motility of Listeria and analysis of loss-of-function cofilin mutants in yeast indicated that ADF/cofilins enhance actin dynamics by depolymerizing actin filaments and provide actin monomers to the cytoplasmic pool (Carlier et al., 1997; Rosenblatt et al., 1997; Lappalainen and Drubin, 1997). Furthermore, cytoplasmic actin filaments accumulate when ADF/cofilins are mutated in Drosophila or Caenorhabditis elegans or when ADF/cofilins are inactivated by overexpressing LIM kinase (Gunsalus et al., 1995; Arber et al., 1998; Yang et al., 1998; Ono et al., 1999; Chen et al., 2001). In contrast, studies on epidermal growth factor (EGF)-stimulated rat mammary adenocarcinoma cells suggested that ADF/cofilin's biological role is to increase actin filament nucleation by severing actin filaments and thus create new filament barbed ends for actin assembly (Chan et al., 2000; Zebda et al., 2000; Ichetovkin et al., 2002; Ghosh et al., 2004).

It is important to note that the biological role(s) of ADF/cofilins in mammalian cells has been mainly examined by inactivating these proteins by LIM kinase. Recent studies revealed that LIM kinase also has other targets than cofilin (Roovers et al., 2003), and to accurately understand the role of ADF/cofilins in actin dynamics in mammalian cells, more direct and specific methods are required. Furthermore, ADF and cofilin-1 are coexpressed in many mammalian cells (Vartiainen et al., 2002), but whether these proteins are involved in same or different biological processes has not been examined. To elucidate the biological roles of mammalian ADF/cofilins, we depleted ADF and cofilin-1, either individually or in combination with each other, from various mouse cell-lines by small interfering RNA (siRNA)-induced gene silencing (Elbashir et al., 2001). Analyses of the ADF and cofilin-1 knockdown cells showed that these proteins promote rapid F-actin depolymerization and provide new monomers to the cytoplasmic actin pool. Our studies also demonstrated that the actin dynamics induced by ADF and cofilin-1 are important for normal actin organization, as well as for morphogenesis, motility, and cytokinesis in cultured mammalian cells.

JMB 2002, 315,4 Weed et al.

Sat, 28 Apr 2007 04:43:16 +0900

Determining the differences in actin binding by human ADF and cofilin

Sharon Yeoh¹, Brian Pope¹, Hans G. Mannherz² and Alan Weeds¹^,

Abstract

The actin-depolymerizing factor (ADF)/cofilin family of proteins play an essential role in actin dynamics and cytoskeletal re-organization. Human tissues express two isoforms in the same cells, ADF and cofilin, and these two proteins are more than 70 % identical in amino acid sequence. We show that ADF is a much more potent actin-depolymerizing agent than cofilin: the maximum level of depolymerization at pH 8 by ADF is about 20 μM compared to 5 μM for cofilin, but little depolymerization occurs at pH 6.5 with either protein. However, we find little difference between the two proteins in their binding to filaments, their severing activities or their activation of subunit release from the pointed ends of filaments. Likewise, they show no significant differences in their affinities for monomeric actin: both bind 15-fold more tightly to actin.ADP than to actin.ATP. Complexes between actin.ADP and ADF or cofilin associate with both barbed and pointed ends of filaments at similar rates (close to those of actin.ATP and much higher than those of actin.ADP). This explains why high concentrations of both proteins reverse the activation of subunit release at pointed ends. The major difference between the two proteins is that the nucleating activity of cofilin-actin.ADP complexes is twice that of ADF-actin.ADP complexes and this, in turn, is twice that of actin.ATP alone. It is this weaker nucleating potential of ADF-actin.ADP that accounts for the much higher steady-state depolymerizing activity. The pH-sensitivity is due to the nucleating activity of complexes being greater at pH 6.5 than at pH 8. Sequence analysis of mammalian and avian isoforms shows a consistent pattern of charge differences in regions of the protein associated with F-actin-binding that may account for the differences in activity between ADF and cofilin.

ADF is a more potent depolymerizing agent than cofilin at pH 8.0

Sedimentation assays showed that ADF is a much more potent depolymerizing agent than cofilin at pH 8.0 (Figure 1). Using 30 μM F-actin, the maximum extent of depolymerization with ADF is 21(±6) μM (five assays) and occurs at a molar ratio to actin of about 1:1. At higher ADF concentrations, the extent of depolymerization decreased (Figure 1(b). There was much less depolymerization with cofilin; the apparent “critical concentration” was 4.8(±1.9) μM, also at a 1:1 molar ratio. Similar experiments at pH 6.5 confirmed that neither ADF nor cofilin induced significant depolymerization: not more than 2 μM actin was released into the supernatant (not shown). Although the extent of depolymerization is much less at pH 6.5 than at pH 8, the depolymerizing activity, expressed as a percentage of the maximum activity, showed an identical profile for both proteins (Figure 1(c). There is a sharp transition in depolymerizing activity around pH 7.3.

ADP-actin

Rabbit muscle actin was prepared[59] and labelled with N-pyrenyl iodoacetamide. To prepare “magnesium” F-actin, G-actin was pre-treated with 0.2 mM MgCl₂ and 0.2 mM EGTA for at least six minutes prior to polymerization in F-buffer (10 mM Tris-HCl (pH 8), 1 mM MgCl₂, 100 mM NaCl, 0.1 mM ATP, 0.2 mM EGTA, 1 mM dithiothreitol, 1 mM NaN₃). Actin.ADP was prepared essentially as described.[60] G-actin in G-buffer (2 mM Tris-HCl (pH 8.0), 0.1 mM CaCl₂, 0.1 mM ATP, 1 mM NaN₃) was stripped of free nucleotide by addition of a 20 % volume of a 50 % (w/v) slurry of Dowex 1×8 for one minute, then 0.2 mM ADP was added together with 0.1 mM glucose and two units/ml of hexokinase coupled to agarose beads (Sigma Aldrich), and the mixture incubated for at least two hours on ice. The hexokinase resin was removed by centrifugation and actin.ADP used within a few hours of preparation. ADP was obtained from Boehringer Mannheim (this contains less than 0.1 % residual ATP as measured by luciferin-luciferase assay). This assay was also used to confirm the complete removal of ATP from the actin.ADP.

Journal Club

Sat, 28 Apr 2007 04:33:44 +0900

Actin Related Papers

Twinfilin : PNAS (2007), Paavilainen et al.

Twinfilin : Embo J. (2006)

Cofilin: JMB (2007), Weeds et al.

WAVE1:Science (2006), Kim et al.

Structure of the N-terminal half of gelsolin bound to actin: roles in severing, apoptosis and FAF : Embo J. (2004)

ADF/Cofilin Related Paper

Mol Biol Cell. 2006 May; 17(5): 2190–2199

JMB 2002, 315,4 Weed et al.

Dog Size Paper

Dog Size

ADF/Cofilin Review summary

Wed, 25 Apr 2007 07:48:15 +0900

Annual Review of Cell and Developmental Biology
Vol. 15: 185-230 (Volume publication date November 1999)
(doi:10.1146/annurev.cellbio.15.1.185)

PROTEINS OF THE ADF/COFILIN FAMILY: Essential Regulators of Actin Dynamics

Ubiquitous among eukaryotes, the ADF/cofilins are essential proteins responsible for the high turnover rates of actin filaments in vivo. In vertebrates, ADF and cofilin are products of different genes. Both bind to F-actin cooperatively and induce a twist in the actin filament that results in the loss of the phalloidin-binding site. This conformational change may be responsible for the enhancement of the off rate of subunits at the minus end of ADF/cofilin-decorated filaments and for the weak filament-severing activity. Binding of ADF/cofilin is competitive with tropomyosin. Other regulatory mechanisms in animal cells include binding of phosphoinositides, phosphorylation by LIM kinases on a single serine, and changes in pH. Although vertebrate ADF/cofilins contain a nuclear localization sequence, they are usually concentrated in regions containing dynamic actin pools, such as the leading edge of migrating cells and neuronal growth cones. ADF/cofilins are essential for cytokinesis, phagocytosis, fluid phase endocytosis, and other cellular processes dependent upon actin dynamics.

Introduction

The correct spatial and temporal organization of actin filaments is essential for the development and function of eukaryotic organisms.

actin filament dynamics play an essential role in,,

- Cell motility

- neuronal pathfinding

- membrane dynamics

- establishment of cell polarity cytokinesis

- generation of tension

two actin filament (F-actin) pools occur in most vertebrate cells,

- relatively stable, and often associated with tropomyosin

- more dynamic, found in the cortex, especially at the leading edge of motile cells and ruffling membrane regions.

Both F-actin-stabilizing drugs and drugs that inhibit actin assembly halt cellular processes dependent on this dynamic pool of actin.

* Fluorescence photobleaching and photoactivation experiments using derivatized actins have shown that actin filaments turn over 100–200-fold faster in cells than has been measured for the purified filaments in vitro (reviewed in Zigmond 1993).

- This enhanced turnover was initially ascribed to the actions of many actin-binding proteins, some of which capped filament ends and some of which severed actin filaments and/or sequestered actin monomers.

- the enhanced turnover of filaments observed in vivo is primarily due to the action of proteins in the ADF/cofilin (AC) family (Rosenblatt et al 1997, Carlier et al 1997, Lappalainen et al 1997).

This review..

focuses on the cellular function and regulation of ACs

their important roles in development and cellular degeneration.

Brief History

The first member of the AC family was identified in extracts of embryonic chick brain that had been passed through a DEAE-cellulose column to remove actin (Bamburg et al 1980).

The identifying assay made use of the ability of ADF to increase the actin monomer pool and the previously recognized strong inhibition of DNase I activity by globular actin (G-actin) in the presence of excess F-actin (Blikstad et al 1978).

This assay was also used to identify actin-depolymerizing proteins in porcine serum (Harris et al 1980), which were initially referred to as high (serum) and low (brain) molecular weight ADFs.

However, the serum ADF proved to be a secreted form of gelsolin (Yin et al 1984). The name ADF is now applied to only the low molecular weight species.

Another protein with actin-depolymerizing activity, depactin, was isolated from starfish eggs (Mabuchi 1981), and a similar protein was identified in extracts of sea urchin eggs (Hosoya et al 1982). Depactin also caused rapid reduction in the viscosity of F-actin solutions (Mabuchi 1983). Although its sequence is only 20% identical to human ADF, it shows the conservation in sequence within the important domains shared by all members of this family (see below). Depactin was the first member of this family to have its sequence determined directly (Takagi et al 1988).

The first mammalian ADF to be characterized was isolated from bovine brain extracts (Berl et al 1983). The assay used to fractionate the protein was based on the disappearance of the actin-activated ATPase activity of myosin. Berl and colleagues showed that an approximately 20-kDa protein in brain caused the loss of F-actin and suggested that this protein might be similar to chick brain ADF. Porcine ADF, termed destrin (destroys F-actin), was isolated from extracts of brain (Maekawa et al 1984, Nishida et al 1984a) and kidney (Nishida et al 1985) with a DNase I–agarose column to bind actin and its associated proteins. In 1986, Cooper et al reported the isolation and characterization of an ADF, called actophorin, from Acanthamoeba castellanii. In 1989, chick ADF was isolated from embryonic skeletal muscle (Abe & Obinata 1989). All these proteins were 15–20 kDa and rapidly reduced the viscosity of an F-actin solution.

Cofilin was purified from porcine brain (Maekawa et al 1984) as a 21-kDa protein that bound through actin to a DNase I affinity column. Although it slightly increased the monomeric pool of actin, cofilin was characterized primarily as an F-actin-binding protein (1:1 ratio with actin subunits) and hence was named for its ability to form cofilamentous structures with actin (Nishida et al 1984b). Cofilin was the first member of this family to be cloned and to have its amino acid sequence deduced from the cDNA sequence (Matsuzaki et al 1988)

Structure Primary Sequence

All members of the AC family contain between 118 and 168 amino acids (13–19 kDa), and sequences for about 30 are known. The secondary structural characteristics they share are shown in Figure 1a, along with the positions of the most highly conserved amino acids, using the numbering system for human ADF. Among the most highly conserved regions are the known actin-binding domains, including the single regulatory phosphorylation site. A second region of high homology for surface residues lies outside the actin-binding domain, suggesting that binding to another protein could be important (Lappalainen et al 1997, Fedorov et al 1997). The regions of nonhomology between the most divergent members occur in loops between the conserved secondary structures and do not have a profound impact on their three-dimensional structure. ADF from T. gondii, containing 118 amino acids, is the smallest family member and lacks any amino acid insertions between the conserved secondary structures (Allen et al 1997).

Figure 1 The structure of human ADF. (a) The primary sequence, showing the residues that are conserved in >90% of the different AC family members across phylogeny (shaded) and the conserved secondary structural features (above the sequence: α, α helices; β, β sheet regions). (b) Three-dimensional structure of the human ADF. Ribbon diagram in left panel shows phosphorylatable serine (pS); PIP₂-binding domains (PI) (shaded regions); nuclear localization sequence (NLS) (black); N terminus (N); C terminus (C). Modified from Lappalainen et al 1998, Hatanaka et al 1996. Middle and right panels show front (same as left panel) and back views, respectively, of space filling models with conserved residues darkened.

Nuclear Localization Sequence

The ACs from vertebrates contain the nuclear localization sequence (NLS) PE(E/D) (I/V)KKRKK immediately prior to the second strand of the β-sheet. This sequence resembles the NLS of the SV40 large T antigen (Matsuzaki et al 1988) and can function as an NLS when linked to nonnuclear proteins that are injected into cells (Abe et al 1993). In many vertebrate cells in culture, both ADF and cofilin are predominantly cytoplasmic, but they often can be induced to collect in the nucleus with actin in the form of actin rods (Nishida et al 1987). Nuclear uptake of AC under stress conditions is accompanied by uptake of cytoplasmic actin (Sanger et al 1980), which has no NLS, and the process requires ATP (Figure 2). Mutagenesis within the basic region of the NLS prevented nuclear accumulation of cofilin in response to stress (Iida et al 1992). The NLS is opposite to the actin-binding site on AC, so it is not difficult to understand how the complex can be recognized by elements of the importin pathway, the primary route for nuclear uptake of proteins with this basic type of NLS (reviewed in Gorlich 1997). What is less obvious is why unbound ADF or ADF-actin complex is not concentrated in the nucleus under most conditions. It is likely that the NLS of ACs becomes much less accessible to the importins when ACs bind to F-actin because the protein contacts two actin subunits (McGough et al 1997). In some cell types it was reported that dephosphorylation of phospho-ADF (pADF) or phospho-cofilin (p-cofilin) accompanies nuclear uptake (Ohta et al 1989, Samstag et al 1994). However, dephosphorylation without nuclear uptake occurs in thyroid cells (Saito et al 1994), and nuclear accumulation (but not necessarily of the phosphorylated form) without dephosphorylation occurs in myotubes (Abe et al 1993). Thus it has yet to be demonstrated whether dephosphorylation is required for nuclear uptake or is correlative only under some conditions.

ADF-Cofilin Related Paper

Wed, 25 Apr 2007 04:49:05 +0900

Destrin cDNA Cloning (Moriyama, Yahara et al., 1990, JBC)

5768.pdf

Pelleting assay

The abilities of the purified cofilin to bind to and to depolymerize F-actin were assessed by the pelleting assay ( Yonezawa et al. 1988; Moriyama et al. 1996) with minor modifications. G-actin and cofilin (2.5 μm each) were mixed in 40 μL of polymerization buffer (50 m m Hepes-NaOH, pH 6.8 or 8.3, 50 m m KCl, 2 m m MgCl₂, 0.02 m m ATP, 0.5 mg/mL BSA), left for 1 h at 25 °C, and centrifuged for 40 min at 100 000 g at 4 °C. Equivalent amounts of the pellet and the supernatant fractions were analysed by SDS-polyacrylamide gel (10–20%) electrophoresis.

Buffer for lysis

20mM Tris-HCl

10mM EGTA
protease inhibitors
1mM DTT

Mol Biol Cell. 2006 May; 17(5): 2190–2199

Wed, 25 Apr 2007 00:18:18 +0900

Enhancement of Actin-depolymerizing Factor/Cofilin-dependent Actin

MATERIALS AND METHODS

Pelleting Assay for F-Actin Binding and Depolymerization

Rabbit muscle actin (Pardee and Spudich, 1982 ), C. elegans actin (Ono, 1999 ), and UNC-60B (Ono and Benian, 1998 ) were purified as described previously. Glutathione S-transferase (GST)-UNC-78 with or without mutation was bacterially expressed and purified as described previously (Mohri et al., 2004 ). An F-actin copelleting assay was performed as described previously (Mohri et al., 2004 ) with modifications. Briefly, 10 μM F-actin was incubated with or without 10 or 20 μM UNC-60B and 0.1–2.0 μM GST-UNC-78 in F-buffer (0.1 M KCl, 2 mM MgCl₂, 1 mM dithiothreitol, and 20 mM HEPES-NaOH, pH 7.5) for 30 min at room temperature and centrifuged at 80,000 rpm (285,000 × g) for 20 min in a Beckman TLA-100 rotor. The supernatants and pellets were adjusted to the same volumes and analyzed by SDS-PAGE. Gels were stained with Coomassie brilliant blue R-250 (National Diagnostics, Atlanta, GA) and scanned by a UMAX PowerLook III scanner at 300 dpi, and the band intensity was quantified by Scion Image Beta 4.02 (Scion, Frederick, MD). In some experiments, the conditions for centrifugation were changed to 56,000 rpm (140,000 × g) for 20 min in TLA-100 or 14,000 rpm (18,000 × g) for 10 min in a Beckman Microfuge (Beckman Coulter, Fullerton, CA).

Assay for Barbed End Elongation

A spectroscopic assay to examine actin elongation from filament ends was performed as described previously using rabbit muscle actin as seeds (Yamashiro et al., 2005 ). Briefly, 10 μM F-actin from rabbit muscle was mixed for 30 s with or without 10 μM UNC-60B and with or without 1 μM UNC-78 variants in F-buffer and used as seeds for polymerization of pyrene-labeled rabbit muscle G-actin. The kinetics of actin polymerization was measured by the increase in the fluorescence of pyrene and the initial rate in the presence of UNC-60 proteins was compared with that of actin alone.

Light Scattering Assay

F-actin (5 μM) was mixed with GST-UNC-78 and/or UNC-60B in F-buffer, and light scattering at an angle of 90° and a wavelength of 500 nm was measured with an LS50B fluorescence spectrophotometer (PerkinElmer Life and Analytical Sciences, Boston, MA). Slit width was set at 2.5 nm for both excitation and emission.

Fluorescence Microscopy

Immunofluorescent staining of adult worms was performed as described previously (Finney and Ruvkun, 1990 ) with anti-UNC-60B antibody (Ono et al., 1999 ) or anti-UNC-78 antibody (Mohri and Ono, 2003 ) and anti-actin antibody (C4; MP Biomedicals) as primary antibodies, and Cy3-labeled goat anti-rabbit IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) and Alexa 647-labeled goat anti-mouse antibody (Invitrogen) as secondary antibodies. Staining of worms with tetramethylrhodamine-phalloidin was performed as described previously (Ono, 2001 ). Live worms expressing GFP-fusion proteins were anesthetized by 0.1% tricaine and 0.01% tetramisole and mounted on 2% agarose pads as described previously (Ono and Ono, 2004 ).

Samples were viewed by epifluorescence using a Nikon Eclipse TE2000 inverted microscope with a CFI Plan Fluor ELWD 40× objective (dry; numerical aperture [N.A.] 0.60) or a CFI Plan Apo 60× objective (oil; N.A. 1.4). Images were captured by a SPOT RT monochrome charge-coupled device camera (Diagnostic Instruments, Sterling Heights, MI) and processed by the IPLab imaging software (Scanalytics, Rockville, MD) and Adobe Photoshop 6.0 (Adobe Systems, San Jose, CA).

Vector Maps

Mon, 23 Apr 2007 16:05:02 +0900

Bacterial Expression Vectors

pRSFDuet-1.pdf

Structure of the N-terminal half of gelsolin bound to actin: roles in severing, apoptosis and FAF : Embo J. (2004)

Mon, 23 Apr 2007 16:01:53 +0900

Structure of the N-terminal half of gelsolin bound to actin: roles in severing, apoptosis and FAF

Leslie D Burtnick¹, Dunja Urosev¹, Edward Irobi², Kartik Narayan² and Robert C Robinson²

The actin filament-severing functionality of gelsolin resides in its N-terminal three domains (G1–G3). We have determined the structure of this fragment in complex with an actin monomer. The structure reveals the dramatic domain rearrangements that activate G1–G3, which include the replacement of interdomain interactions observed in the inactive, calcium-free protein by new contacts to actin, and by a novel G2–G3 interface. Together, these conformational changes are critical for actin filament severing, and we suggest that their absence leads to the disease Finnish-type familial amyloidosis. Furthermore, we propose that association with actin drives the calcium-independent activation of isolated G1–G3 during apoptosis, and that a similar mechanism operates to activate native gelsolin at micromolar levels of calcium. This is the first structure of a filament-binding protein bound to actin and it sets stringent, high-resolution limitations on the arrangement of actin protomers within the filament.

Gelsolin is an actin filament capping and severing protein that enhances the rate of cell migration

The control of gelsolin activity

gelsolin is sequestered by phosphatidylinositol 4,5-bisphosphate (PIP₂)

- hydrolysis of PIP₂ releases gelsolin into the cytoplasm;

- calcium activates free gelsolin to allow it to cap and sever actin filaments

- selected filaments are uncapped by a PIP₂-rich membrane to allow actin polymerization to proceed and to render gelsolin inactive (Janmey and Stossel, 1987; Allen, 2003).

Branching of newly uncapped filaments through a mechanism that involves arp2/3 amplifies propulsive actin polymerization (Falet et al, 2002),
le gelsolin-capped filaments are marked for rapid depolymerization by cofilin (Ressad et al, 1998).

In a calcium-free environment, gelsolin exists as a compact, inert arrangement of six related domains, G1–G6 (Burtnick et al, 1997).

On binding calcium, metamorphosis of gelsolin to an activated structure entails the opening at least of three identifiable latches (tail latch, G1–G3 latch and G4–G6 latch) to expose actin-binding surfaces on G2, G1 and G4, respectively (Robinson et al, 1999; Choe et al, 2002).

Tryptophan fluorescence experiments show that gelsolin undergoes two or, possibly, three conformational changes characterized by calcium binding with K_d values of 0.1, 7 and, controversially, 0.3 M (Kinosian et al, 1998; Lin et al, 2000).

New evidence from synchrotron footprinting experiments confirms the tail latch to be released at micromolar calcium concentrations, and identifies 3–6 further calcium ions that bind at around 100 M and are thought necessary to open the G1–G3 and G4–G6 latches (Kiselar et al, 2003). In vitro, gelsolin capping and severing activities occur at a low level under conditions similar to those in a resting cell, where the cytoplasmic Ca²⁺ concentration is approximately 0.2 M (Kinosian et al, 1998). The rate of gelsolin severing accelerates beyond that at resting calcium concentrations in response to calcium transients.

Protein crystallography has identified a conserved Ca²⁺-binding site in each gelsolin domain studied to date (McLaughlin et al, 1993; Burtnick et al, 1997; Choe et al, 2002; Kazmirski et al, 2002). These six variable affinity sites are termed type-2 Ca²⁺-binding sites and are responsible for the structural changes that activate gelsolin. Each type-2 calcium ion acts both to disrupt the structure of calcium-free gelsolin and to stabilize the activated form. Of particular note are the sites within G2 and G6, which are well situated to cooperate in opening the tail latch. Two other Ca²⁺-binding sites, termed type-1 sites, are sandwiched at the actin:G1 and the actin:G4 interfaces, respectively, and likely moderate the affinity of activated gelsolin for actin (Choe et al, 2002). In these sites, coordination of the Ca²⁺ ion involves residue Glu167 from two different actin protomers along with analogous contacts from G1 and G4

Secreted gelsolin prevents elevation of the viscosity of extracellular fluids by severing actin filaments that are released as a result of cell death or injury (Haddad et al, 1990). A hereditary mutation in gelsolin of Asp187 to Asn or Tyr in sufferers of Finnish-type familial amyloidosis (FAF) exposes the Arg172 to Ala173 peptide bond to proteolysis by furin during transport through the trans-Golgi network (Chen et al, 2001). A subsequent extracellular proteolytic event generates a gelsolin fragment, residues 173–243, which self-assembles into amyloid fibrils. Amyloid accumulation effects a range of neuropathies, ophthalmic disorders and dermatological abnormalities.

Appreciation of the mechanism of disintegration of FAF gelsolin has evolved in parallel with knowledge of the conformation and function of normal gelsolin. The structure of calcium-free gelsolin revealed that both the mutation site and primary cleavage site reside in G2, and demonstrated that Asp187 contributes to the stability of G2 through a series of electrostatic interactions (Burtnick et al, 1997). Later, it was realized that proteolysis requires an activated form of FAF gelsolin; the binding of Ca²⁺ to Asp187 protects normal gelsolin from cleavage (Zapun et al, 2000; Chen et al, 2001; Robinson et al, 2001; Kazmirski et al, 2002). Intriguingly, however, overexpression of furin can lead to detectable attack even on normal gelsolin (Huff et al, 2003).

G1–G3 as an independent fragment of gelsolin is biologically relevant, being generated from the intact protein during apoptosis via the actions of caspase-3, -7 and -9 (Kothakota et al, 1997; Azuma et al, 2000). Free of the three C-terminal domains, G1–G3 is relieved of regulation by Ca²⁺ and proceeds to dismantle the actin cytoskeleton unchecked. This outcome is countered by full-length gelsolin and its C-terminal half, G4–G6, which display antiapoptotic activity by blocking voltage-dependant anion channels, inhibiting mitochondrial membrane potential loss and cytochrome c release (Koya et al, 2000; Kusano et al, 2000). In addition, complexes that involve caspases, gelsolin and PIP₂ are able to block cleavage of gelsolin and inhibit apoptosis (Azuma et al, 2000). Gelsolin-deficient cells display a retarded onset of apoptosis, while transient overexpression of G1–G3 induces apoptosis (Kothakota et al, 1997). Clearly, gelsolin plays a multifaceted role in the control and execution of apoptosis, many details of which remain to be discovered.

Atomic coordinates currently are available for intact gelsolin in the calcium-free state (Burtnick et al, 1997) and for the actin complexes with gelsolin fragments G1 (McLaughlin et al, 1993; Irobi et al, 2003) and G4–G6 (Choe et al, 2002). Absent from this set is the structure of the activated F-actin recognition unit, G2–G3. The present work amends this deficiency by detailing the architecture of the apoptotic fragment of gelsolin, G1–G3, bound to an actin monomer. A novel compact arrangement of G2–G3 is revealed. Calcium ions occupy type-2 sites in G1 and G3, but the corresponding site in G2 is vacant. A complete set of conformations for the resting and activated domains of gelsolin, together with published calcium-binding data, allows proposal of schemes for the overall activation of gelsolin and for its actin-modulating activities.

Figure 1
The structure of the G1–G3:actin complex. (A) A schematic representation of the G1–G3:actin complex. The gelsolin domains are colored: G1, red; G2, green; G3, yellow. This scheme is preserved in subsequent figures unless explicitly stated. Actin, with subdomains 1, 3 and 4 indicated, is colored gray. ATP is shown as a ball-and-stick representation with its associated Ca²⁺ in purple. Type-1 and type-2 Ca²⁺ ions are depicted as gold and black spheres, respectively. (B) The structure of the G4–G6:actin complex (PDB 1H1V; Choe et al, 2002) for comparison. Gelsolin domains are colored: G4, pink; G5, dark green; G6, orange. (C) Model of Ca²⁺-free G1–G3 interacting with actin: obtained by taking the structure of G1–G3 excised from Ca²⁺-free, inactive gelsolin (PDB 1D0N; Burtnick et al, 1997) and positioned on actin, in accord with the overlaying of G2 onto the structure presented in (A). (D) Stereo view of a representative portion of the 2F_o–F_c electron density map contoured at 1.

Figure 2
Calcium-induced activation of gelsolin and the severing of F-actin. (A) Levels of calcium activation. Ca²⁺-free gelsolin domains are shown as hexagons and calcium-bound domains are depicted as ovals. Calcium ion concentrations are indicated for each step. The scissors represent the stage at which FAF gelsolin is cleaved. (B) The sequence of events during severing of actin by fully activated gelsolin. Actin protomers are shown in blue.

Figure 3
Regions of gelsolin involved in binding PIP₂. (A) Activated G1 and G2 (pink) bound to actin (pale blue) as observed in the G1–G3:actin structure. PIP₂-binding regions 132–140 and 161–172 are highlighted in red and green, respectively. Lysine residues from the KxKK motifs, within these regions, are drawn and labeled KxKK1 and KxKK2, respectively. The hydrophobic-rich G1–G2 linker, residues 141–160, is colored orange. The G1 type-2 calcium ion is drawn as a gray sphere. (B) G1–G2 excised from the Ca²⁺-free inactive form of gelsolin (PDB 1D0N; Burtnick et al, 1997). Colors are as in (A).

Figure 4
Model of a gelsolin-capped filament. (A) The ADP model of the actin filament (four protomers are drawn in blue and gray), with G1–G3:actin (1RGI) and G4–G6:actin (G4, pink; G5, dark green; G6, orange; PDB 1H1V; Choe et al, 2002) overlaid onto the barbed end. Purple spheres mark Asp371 of G3 and Met412 of G4, a gap of 63.1 Å to be bridged by the G3–G4 linker, which is modeled in purple. (B) A second view of the gelsolin-capped model filament, looking directly at the capped, barbed end. Three actin protomers are depicted. (C) The G1:actin structure (red:blue) with subdomain 2 of a second actin protomer from the ADP model drawn in black. The arrow indicates a steric clash between the long helix of G1 and the ADP helix of actin subdomain 2. (D) The tail latch. G2 (green) and the C-terminal tail (residues 736–755; orange) from inactive gelsolin positioned on the ADP model by overlaying G2 in (A). Only one actin from the filament is shown (gray). In this position, the C-terminal tail obscures the actin-binding site on G2.