Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M608234200 on December 18, 2006

J. Biol. Chem., Vol. 282, Issue 7, 4494-4503, February 16, 2007
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
282/7/4494    most recent
M608234200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Gong, H.
Right arrow Articles by Byers, D. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Gong, H.
Right arrow Articles by Byers, D. M.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Neutralization of Acidic Residues in Helix II Stabilizes the Folded Conformation of Acyl Carrier Protein and Variably Alters Its Function with Different Enzymes*

Huansheng Gong, Anne Murphy, Christopher R. McMaster, and David M. Byers1

From the Atlantic Research Centre, Departments of Pediatrics and Biochemistry and Molecular Biology, Dalhousie University, Halifax, Nova Scotia B3H 4H7, Canada

Received for publication, August 28, 2006 , and in revised form, December 5, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Acyl carrier protein (ACP), a small protein essential for bacterial growth and pathogenesis, interacts with diverse enzymes during the biosynthesis of fatty acids, phospholipids, and other specialized products such as lipid A. NMR and hydrodynamic studies have previously shown that divalent cations stabilize native helical ACP conformation by binding to conserved acidic residues at two sites (A and B) at either end of the "recognition" helix II. To examine the roles of these amino acids in ACP structure and function, site-directed mutagenesis was used to replace individual site A (Asp-30, Asp-35, Asp-38) and site B (Glu-47, Glu-53, Asp-56) residues in recombinant Vibrio harveyi ACP with the corresponding amides, along with combined mutations at each site (SA, SB) or both sites (SA/SB). Like native V. harveyi ACP, all individual mutants were unfolded at neutral pH but adopted a helical conformation in the presence of millimolar Mg2+ or upon fatty acylation. Mg2+ binding to sites A or B independently stabilized native ACP conformation, whereas mutant SA/SB was folded in the absence of Mg2+, suggesting that charge neutralization is largely responsible for ACP stabilization by divalent cations. Asp-35 in site A was critical for holo-ACP synthase activity, while acyl-ACP synthetase and UDP-N-acetylglucosamine acyltransferase (LpxA) activities were more affected by mutations in site B. Both sites were required for fatty acid synthase activity. Overall, our results indicate that divalent cation binding site mutations have predicted effects on ACP conformation but unpredicted and variable consequences on ACP function with different enzymes.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Acyl carrier protein (ACP)2 is an acidic and highly conserved protein typically consisting of 70–100 residues and is essential for bacterial growth, communication, and pathogenesis. ACP is responsible for supplying acyl groups for the biosynthesis of a plethora of bacterial molecules, including fatty acids (1), phospholipids (2), lipid A (3), lipoic acid (4), hemolysin (5), acyl homoserine lactones involved in quorum-sensing (6), and the aldehyde substrate of luciferase in bioluminescent bacteria such as Vibrio harveyi (7). Other functions of ACP and its homologues include the production of membrane-derived oligosaccharides (8), rhizobial nodulation signaling factors (9), polyketide (10) and non-ribosomal peptide antibiotics (11), and lipoteichoic acid (12). The list of ACP-binding partners continues to expand based on proteomic efforts (13). The requirement for ACP in these diverse processes suggests that interactions between ACP and its partner enzymes must be specific. Information about how individual amino acid residues contribute to ACP conformation and interactions with functionally diverse enzymes will provide insight into the design of novel antimicrobial agents against ACP-dependent targets that are essential for bacterial growth and pathogenesis.

Structural analyses of type II ACPs from Escherichia coli and several other bacterial species reveal a common three-helix bundle with a long loop connecting helices I and II (14–18). The three helices enclose a hydrophobic binding pocket for the fatty acyl chain (18, 19), which is attached through a thioester linkage to the 4'-phosphopantetheine prosthetic group at Ser-36 of holo-ACP, near the N terminus of helix II. E. coli ACP has an inherently mobile structure that maintains a dynamic equilibrium of at least two conformers, with the loop region and helix II being particularly flexible (20, 21). Electrostatic repulsion undoubtedly contributes to conformational flexibility of ACP in this very acidic central region (with 14 acidic and no positive residues between positions 30 and 60); this flexibility in turn is thought to be important in facilitating rapid association and dissociation of ACP with its various partner enzymes. Indeed, computational docking analyses and mutagenic studies with E. coli fatty acid synthase components and other enzymes (22–24) have implicated the conserved acidic helix II of ACP as a "recognition helix" for universal enzyme interaction (25). The co-crystal structure of Bacillus subtilis ACP and holo-ACP synthase also supports the importance of helix II of ACP in this regard (26).

NMR (27), circular dichroism (28, 29), and proteomic (30) approaches have revealed specific binding of divalent cations to ACP and stabilization of ACP conformation by cation binding. Two divalent cation binding sites exist on E. coli ACP at neutral pH with an average Kd/site of about 80 µM (27), and relaxation perturbed two-dimensional NMR studies have identified seven residues involved in these relatively low affinity interactions (31). Site A (consisting of Glu-30, Asp-35, and Asp-38) resides at the N-terminal end of helix II and site B (Glu-47, Asp-51, Glu-53, and Asp-56) at the other. V. harveyi ACP shares 86% amino acid sequence identity with its E. coli counterpart, and six of the seven putative cation binding residues are identical with Asp replacing Glu at position 30 (Fig. 1). Unlike E. coli ACP, however, V. harveyi ACP is largely unfolded at neutral pH in the absence of millimolar concentrations of Mg2+ or attached acyl chains (32).

The role of acidic helix II residues in maintaining the stability or flexibility of ACP conformation, as well as their contributions to ACP function with specific enzyme partners, remains largely uncharacterized. Moreover, whether divalent cation binding influences ACP function is unknown. In the present study, putative divalent cation-binding residues on recombinant V. harveyi ACP (rACP) were mutagenized (alone or in combination) to corresponding neutral amide forms and the consequences on ACP conformation were examined. Four enzyme systems that employ ACP in different reactions were also investigated to identify ACP regions or residues important for each interaction.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—[9,10-3H]Myristic acid (49 Ci/mmol), [3H]uridine diphosphate N-acetyl-D-glucosamine (39.7 Ci/mmol), and [2-14C]malonyl-CoA (51 mCi/mmol) were from NEN/PerkinElmer Life Sciences. [3H]Acetyl-CoA (4.7 Ci/mmol), Factor Xa restriction protease, pGEX-5X-3 vector, glutathione-Sepharose 4B, and SOURCE 15Q anion exchange resin were from Amersham Biosciences (GE Healthcare). Mini-PROTEAN 3 Electrophoresis System and reagents were from Bio-Rad. GelCode Blue Stain and Micro BCA Protein Assay reagents were from Pierce. Uniplate Silica Gel G 250 micron TLC plates were from Analtech. All other chemicals were of the highest quality available.

Site-directed Mutagenesis and Purification of Mutant V. harveyi ACPs—ACP mutants were constructed on a pGEX-5X-3 vector containing the V. harveyi acpP gene (32) using the Gene-Editor Mutagenesis System (Promega) and confirmed by DNA sequencing. Plasmids encoding GST-ACP fusion proteins were maintained in E. coli JM109 (Promega) and transformed into E. coli BL21 (Stratagene) for protein expression. All mutant ACPs exhibited roughly comparable levels of expression with rACP, and none were lethal to E. coli cells. ACPs were purified essentially as described (32). Crude cell extract of E. coli holo-ACP synthase (ACPS) was prepared (33) and added prior to glutathione-Sepharose chromatography to maximize the conversion of GST-apo-ACPs to the holo form. GST-ACPs were purified using glutathione-Sepharose 4B and cleaved by Factor Xa to produce a recombinant V. harveyi ACP (rACP) with an N-terminal extension of four amino acids (GIPL). ACPs were further purified to homogeneity on a SOURCE 15Q anion exchange column (Waters 650 Advanced Protein Purification System) and quantified using the Micro BCA protein assay. Apo-ACPs were obtained using the same method from preparations without added exogenous ACPS. Each apo-ACP is less negatively charged than its holo counterpart because of the absence of the 4'-phosphopantetheine prosthetic group, and thus was readily separated from holo-ACP by SOURCE 15Q anion exchange chromatography (AktaFPLC). The identities of selected ACPs were confirmed by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry.

Circular Dichroism—Spectra from a Jasco J-810 spectropolarimeter were recorded at 25 °C using a 0.1 cm water-jacketed cell. Holo-ACPs were diluted to 3–10 µM in 10 mM sodium phosphate, 0.1 mM EDTA (pH 7.0), and spectra were measured from 260 to 190 nm in continuous mode at a scanning speed of 10 nm/min directly or within 1 min after addition of MgCl2, CaCl2, or MnCl2.

Native PAGE—ACPs and acyl-ACPs were resolved by conformationally sensitive native gel electrophoresis (34) on a 20% polyacrylamide gel at room temperature. Protein bands were visualized by staining with GelCode Blue Stain reagent.

Holo-ACP Synthase Assay—A modification of the assay described by Lambalot and Walsh (33) was performed at 25 or 37 °C in a final volume of 10 µl containing 50 mM sodium phosphate, pH 7.0, 10 mM MgCl2, 5 mM dithiothreitol, 50 µM apo-ACPs, and either partially purified E. coli ACPS from an over-expressing strain (33) or purified C-terminally His6-tagged E. coli ACPS. [3H]Acetyl-CoA (57 µM, 440 dpm/pmol) was added to start the reaction and 4 µl was removed at the appropriate time and quenched with 800 µl of cold 10% trichloroacetic acid (w/v) and 20 µl of bovine serum albumin (25 mg/ml). After incubation on ice for 15 min, the precipitate was obtained by centrifugation at 12,000 x g for 5 min, washed twice with 800 µl of cold 10% trichloroacetic acid (w/v), and solubilized in 50 µl of formic acid. Product formation was measured by liquid scintillation analysis (Beckman LS6500 Multi-Purpose Scintillation Counter) and corrected for blank values obtained in the absence of added apo-ACP (<2% of control values).

Acyl-ACP Synthetase (AAS) Assay—V. harveyi AAS was partially purified by DEAE-Sepharose and Sephacryl S-300 chromatography (35). In the standard assay, holo-ACPs (50 µM) were incubated at 25 or 37 °C for 30 min with AAS (0.5 µgof total protein corresponding to 2 milliunits of activity), 10 mM MgSO4, 10 mM ATP, 0.1 mM [9,10-3H]myristic acid (888 dpm/pmol) in 100 mM Tris-HCl, pH 7.8, and 5 mM dithiothreitol (25 µl total volume). Samples (10 µl) were spotted on filter paper, washed three times with methanol/chloroform/acetic acid (6:3: 1), and product formation was measured by liquid scintillation counting (35); values were corrected for blanks obtained in the absence of added holo-ACP (<5% of rACP control values).

V. harveyi AAS was also used to prepare myristoyl-ACPs for native PAGE analysis and beta-hydroxymyristoyl-ACP substrates for LpxA assay, except that unlabeled myristic acid and DL-beta-hydroxymyristic acid (Sigma) were used, respectively. Reactions were allowed to proceed up to 24 h at room temperature to maximize conversion to acyl-ACPs. beta-Hydroxymyristoyl-ACPs were further purified by application to a Vivapure Q Mini M Spin Column (Vivascience) equilibrated in 25 mM Tris-HCl, pH 7.8. Following elution with 0.2 M NaCl, ACPs were eluted with 0.5 M NaCl and quantified using the Micro BCA protein assay.

FAS Assay—V. harveyi FAS was partially purified from V. harveyi cultures as described (36) and separated from endogenous ACP using ammonium sulfate fractionation (between 40 and 75% saturation). The standard assay was performed at 25 or 37°C for 1 h in a final volume of 40 µl containing 10 mM Na+/K+-phosphate, pH 7.0, 5 mM dithiothreitol, 0.5 mM NADPH, 0.5 mM NADH, 10 µM acetyl-CoA, 23 µM [2-14C]malonyl-CoA (48 dpm/pmol), FAS preparation (40 µg of total protein), and 10 µM ACPs. The reaction was stopped with 40 µl of 1 M KOH and incubated at 60 °C for 1 h. After acidification with 40 µlof 2 M HCl, fatty acids were extracted into 1 ml of hexane and quantified by liquid scintillation counting. Blank values (minus holo-ACP or NAD(P)H) were <5% of [14C]malonyl-CoA incorporation into acyl-rACP control.

LpxA Assay—Assays were performed at 25 °C as previously described (37). Each 10-µl reaction mixture contained 40 mM Na+-HEPES, pH 8.0, 10 mg/ml bovine serum albumin, 10 µM beta-hydroxymyristoyl-ACP, purified His6-tagged LpxA (~1.2 µM), and 2.5 µM [3H]UDP-GlcNAc (10,000 dpm/pmol). Samples (2 µl) were removed at various times up to 10 min (linear portion of assay) and spotted on a silica gel G TLC plate which was developed in chloroform/methanol/acetic acid/water (25: 25:2:4). Formation of the acylated product (Rf ~ 0.6) was quantified using a Bioscan System 200 Imaging Scanner with Win-Scan software and corrected for blank values (<5% of beta-hydroxymyristoyl-rACP control of 0.5–1.0 pmol/min/µg enzyme).

Fluorescence Assay for ACP-LpxA Interaction—Wild-type and mutant holo-ACPs (0.3 mM) were reduced by incubation with 2.5 mM Tris(2-carboxyethyl)phosphine-HCl for 30 min, followed by reaction with IAEDANS (5 mM final concentration) for 2 h at room temperature. Complete modification of the single phosphopantetheine sulfhydryl group to form AEDANS-ACP was indicated by its decreased mobility on native-PAGE, and the product was purified by SOURCE 15Q anion-exchange chromatography.

AEDANS-ACPs (1 µM) in 0.1 M NaCl, 10 mM sodium phosphate (pH 7.0), 0.1 mM EDTA were titrated sequentially with increasing amounts of LpxA in the same buffer. Fluorescence emission spectra were measured on a PerkinElmer LS50B spectrofluorimeter (10-mm cell) with excitation wavelength of 336 nm and excitation and emission slit widths of 5 and 10 nm, respectively. After correction of fluorescence intensities for dilution (<10% total), the total area under the fluorescence emission curve (400–600 nm) was used to calculate apparent LpxA-ACP dissociation constants using GraphPad Prism software, assuming a 1:1 binding stoichiometry.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Preparation and Conformational Analysis of Mutant ACPs—Of the seven acidic residues implicated in divalent cation binding by E. coli ACP (Fig. 1), positions 35, 38, 47, 56, and to a lesser extent positions 30 and 53, are invariant or highly conserved among ACPs from plants and bacteria. Position 51 in site B is the least conserved and tolerates various non-conservative replacements. The conserved residues are likely to not only play a role in cation binding, but may also be important in the interaction of ACP with enzymes. To examine the individual and combined contribution of these acidic residues, site-directed mutagenesis was employed for substitution with the corresponding neutral amino acids. Nine mutants were constructed, i.e. six individual mutants D30N, D35N, D38N, E47Q, E53Q, and D56N, and three global site-elimination mutants SA (D30N/D35N/D38N), SB (E47Q/D51N/E53Q/D56N), and SA/SB. ACPs were expressed in E. coli BL21 cells as GST fusion proteins and purified to homogeneity to eliminate contamination of the mutant proteins with host cell ACP.


Figure 1
View larger version (42K):
[in this window]
[in a new window]

  		FIGURE 1.
Top, UCSF Chimera (48) ribbon representation of V. harveyi ACP showing the three {alpha}-helices I-III (left) and helix II with putative divalent cation binding residues in sites A and B (right). Bottom, amino acid sequences of V. harveyi (Vh) and E. coli (Ec) ACPs, highlighting site A and B residues (underlined), helices I-III, and the site of phosphopantetheine attachment at Ser-36 (bold). Residues conserved between the two species are indicated by a dot in the E. coli sequence.

 
Circular dichroism (CD) was used to examine the effects of divalent cation binding site mutations on the secondary structure of ACP and dependence of ACP folding on MgCl2 (Fig. 2). This takes advantage of the fact that V. harveyi ACP is largely unfolded at physiological pH and adopts a native-like conformation containing ~50% {alpha}-helix in the presence of millimolar levels of Mg2+ (32). Addition of 10 mM Mg2+ to holo-rACP increased the magnitude of ellipticity at 220 nm by 2.5-fold and shifted the negative trough at 201–208 nm indicative of increased {alpha}-helical content (Fig. 2A); similar Mg2+-dependent conformational transitions were observed for the six individual mutants (data not shown). However, the two site-elimination mutants SA and SB exhibited more residual helical content in the absence of divalent cations, and became folded to a similar extent as rACP upon addition of Mg2+ (Fig. 2A). The effect of Mg2+ on [{theta}]220 was significantly (p < 0.05) greater for the SA mutant than for SB (Fig. 2B and Table 1), whereas the double site-elimination mutant SA/SB contained substantial helical content even in the absence of Mg2+, with only a modest further effect of Mg2+ on [{theta}]220 (1.2-fold increase). The midpoint of the Mg2+-induced conformational transition ([{theta}]220) was comparable for holo forms of SA, SB, and rACP (Table 1), suggesting that sites A and B can bind Mg2+ independently and with similar affinity, and that there is little cooperativity between these sites in divalent cation-induced ACP folding. Similar results were observed with the corresponding apo-ACPs (Fig. 2C), although ~2-fold higher levels of Mg2+ were required to elicit conformational change (Table 1).


View this table:
[in this window]
[in a new window]

  		TABLE 1
CD parameters of ACP titration with Mg2+

Holo- and apo-ACPs (10 µM) were titrated with MgCl2 and mean residue ellipticity at 220 nm ([{theta}]220) was measured as described in the legend to Fig. 2. Each value represents the mean ± S.D. determined from six (holo-ACPs) or three (apo-ACPs) independent titrations.

 


Figure 2
View larger version (24K):
[in this window]
[in a new window]

  		FIGURE 2.
Effect of Mg2+ on the circular dichroism of rACP and mutant ACPs. A, CD spectra of the indicated holo-ACPs (10µM) were obtained at 25 °C at pH 7.0 in the absence (open symbols) and presence (closed symbols) of 3 mM MgCl2 as described in the text. B and C, holo- and apo-ACPs (10 µM) were titrated with increasing concentrations of MgCl2 and mean residue ellipticity at 220 nm ([{theta}]220) was measured. Each value represents the mean ± S.D. of three independent samples.

 


Figure 3
View larger version (24K):
[in this window]
[in a new window]

  		FIGURE 3.
Effects of Mg2+, Ca2+, and Mn2+ on the circular dichroism of rACP. A, CD spectra of holo-rACP (3 µM) were obtained in the absence (open symbols) and presence of 5 mM (gray symbols) or 50 mM (black symbols) of CaCl2 or MnCl2. B, CD signal at 220 nm of holo-rACP was recorded at the indicated levels of Mg2+, Ca2+, or Mn2+. The ellipticity relative to holo-rACP in the presence of each cation at 5 mM is shown.

 


Figure 4
View larger version (89K):
[in this window]
[in a new window]

  		FIGURE 4.
Conformationally sensitive native PAGE analysis of wild type and mutant ACPs. ACPs were prepared with (+) or without (–) modification by exogenous E. coli ACPS during purification as described in the text and separated on a 20% polyacrylamide gel at pH 9.2, either before (–) or after (+) ATP-dependent conversion to myristoyl-ACPs using AAS. An equal volume of the extract was loaded and rACP was included on each gel for comparison. Protein bands were visualized by staining with GelCode Blue Stain reagent. The migration of the apo (A), holo (H), and myristoyl (M) forms of each ACP is indicated for clarity.

 
Conformational changes of rACP were also monitored in response to Ca2+ and Mn2+ which, like Mg2+ increased {alpha}-helical content in the 0.1–1.0 mM range (Fig. 3). However, although all three divalent cations elicited a maximum conformational change at around 5 mM, higher levels of Ca2+ and Mn2+ (but not Mg2+) decreased the secondary structure content of rACP. Results from the CD deconvolution program k2d (38) indicated that 50 mM Ca2+ or Mn2+ caused a 40% decrease in {alpha}-helical content of rACP, compared with 5 mM Ca2+ or Mn2+.

Native PAGE at pH 9.2 is very sensitive to both the intrinsic charge and conformation of ACP, and can readily distinguish apo-, holo-, and fatty acylated forms in which a more compact and thus mobile conformation of this acidic protein is progressively stabilized (34). Overexpression and preparation of V. harveyi rACP and mutant ACPs without treatment with exogenous E. coli holo-ACPS during purification resulted in a mixture of apo- and holo-ACPs in most cases (Fig. 4). Post-lysis incubation with ACPS and coenzyme A increased conversion to holo-ACP, whereas a faster migrating acyl-ACP band was observed upon further incubation with myristic acid and ATP in the presence of V. harveyi AAS. Most apo and holo individual mutants had electrophoretic mobilities that were comparable to the corresponding forms of rACP, with the exception of apo- and holo-D35N, which migrated considerably faster than its rACP counterparts (Fig. 4). In contrast to individual ACP mutants, the three site-elimination mutants SA, SB, and SA/SB exhibited increased mobility relative to their apo- and holo-rACP counterparts. These mutants would be less negatively charged than rACP at pH 9.2, indicating that they have a significantly decreased hydrodynamic radius (i.e. a more compact conformation) under these conditions, consistent with CD results. Upon enzymatic myristoylation, all ACP mutants were able to interact with the covalently attached acyl chain as exhibited by further increased mobility relative to holo-ACPs (Fig. 4).

Mutant ACPs as Enzyme Substrates—Closer inspection of Fig. 4 reveals differences in the relative amounts of apo-, holo-, and acyl-forms of various mutant ACPs, suggesting that divalent cation binding site mutations may affect their substrate properties with specific ACP-dependent enzymes. Whereas rACP was typically ~50% converted to the holo form in vivo by endogenous host cell ACPS, some ACP mutants such as D35N, D56N, SA, and SA/SB appeared mainly as apo-ACPs (Fig. 4). Surprisingly, D38N and E53Q were completely modified to the holo form in vivo, and no apo-ACP was recovered for these mutants.

A more quantitative assay measuring holo-ACP formation from [3H]acetyl-CoA confirmed that E. coli ACPS activity at 37 °C is ≥95% impaired with apo-ACP substrates containing individual site A mutations D35N and D56N, as well as mutants SA and SA/SB (Table 2). In contrast, mutants D30N, E47Q, and SB supported partial ACPS activity. The dependence of ACPS activity on the concentrations of rACP and mutant SB was further examined (Fig. 5A). Similar to previous reports (39, 40), ACPS exhibited substrate inhibition at low rACP concentrations, precluding accurate kinetic evaluation, although this was not observed for the SB mutant for which activity was actually greater than for rACP at higher concentrations (up to 50 µM, not shown). These results suggest that combined site B mutations may decrease affinity of ACP for E. coli ACPS, but alleviate substrate inhibition at higher ACP concentrations.


View this table:
[in this window]
[in a new window]

  		TABLE 2
Enzyme activities with individual ACPs

Enzyme activities with the indicated apo- or holo-ACP substrate were measured at 37 °C as described in the text, and the mean ± S.D. of at least three independent assays are shown.

 
We next examined other ACP-dependent enzymes with different functions and catalytic mechanisms. V. harveyi AAS directly activates medium and long chain fatty acids to form acyl-ACP with hydrolysis of ATP to AMP (35). Partially purified AAS exhibited an apparent Km of 3.2 ± 0.9 µM for rACP based on data at low ACP concentrations (≤ 10 µM), while some substrate inhibition was noted at higher concentrations (Fig. 5B). In marked contrast to ACPS, impairment of AAS was greater with the SB than the SA mutant. This was due primarily to increased Km values (12 ± 5 µM and 49 ± 29 µM for SA and SB, respectively), while apparent Vm was more similar (1.00 ± 0.06, 0.74 ± 0.12, and 0.70 ± 0.30 pmol/min/µg for rACP, SA, and SB). Thus, catalytic efficiency relative to rACP was decreased to 20% for SA and 5% for SB. A similar trend was observed when the assay was performed at 37 °C. None of the individual mutations impaired AAS activity by more than 65% relative to rACP, suggesting a cumulative effect of site B mutations on this enzyme (Table 2). The activity of AAS with the SA/SB double mutant was too low to allow for accurate kinetic analysis, which is also evident from the electrophoresis data shown in Fig. 4, as it is the only mutant not quantitatively acylated by AAS under these conditions.


Figure 5
View larger version (18K):
[in this window]
[in a new window]

  		FIGURE 5.
Selected enzyme activities with rACP and site-elimination mutants. Activities were determined at 25 °C as described in the text with the indicated concentrations of rACP, SB, and/or SA. A, His6-tagged E. coli ACPS (7 ng of enzyme, 2–5 min assays). B, V. harveyi AAS (0.5 µg of protein, 10–30 min assays). C, V. harveyi FAS fraction (13 µg of protein, 10–60 min assays). Each value represents the mean ± S.D. of at least four independent samples.

 


Figure 6
View larger version (30K):
[in this window]
[in a new window]

  		FIGURE 6.
Fluorescence titration of AEDANS-labeled ACPs with E. coli LpxA. Panels A–C, fluorescence emission spectra of the indicated AEDANS-ACP (1 µM) were measured in the absence (dotted line) and presence of 2.4, 7, 13, and 22 µM LpxA, as indicated by lines of increasing density. The results are representative of at least three similar experiments for each ACP. D, increased area under the fluorescence emission curve is plotted as a function of ACP concentration for each ACP; each value is the mean ± S.D. of 3–6 independent titrations.

 
The type II FAS system found in plants and bacteria consists of discrete enzymes that carry out the de novo biosynthesis of fatty acids, with all intermediates covalently bound to ACP (1). As shown in Fig. 5C, incorporation of [14C]malonyl-CoA into hexane extractable products using a partially purified V. harveyi FAS fraction at 25 °C was dependent on rACP concentration with an apparent Km of 2.6 ± 0.1 µM. However, very little FAS activity was observed with site elimination mutants SA or SB, even at the highest ACP concentrations tested (40 µM). FAS activity was decreased by 50% or more with all individual mutants except D56N, for which activity was at least as high as rACP (Table 2). These results suggest that acidic residues in both divalent cation binding sites are important for interaction with one or more of the FAS component enzymes, and their mutation may have a cumulative effect on FAS activity.

ACP is also a central cofactor in the biosynthesis of Gram-negative bacterial lipid A (endotoxin). The first step of lipid A biosynthesis involves the transfer of the acyl chain from beta-hydroxymyristoyl-ACP to the 3-OH position of UDP-N-acetyl-glucosamine (UDP-GlcNAc) and is catalyzed by UDP-GlcNAc acyltransferase (LpxA) (3). As with AAS, LpxA activity was substantially more impaired by site B elimination (<5% of rACP activity with SB mutant at 10 µM), whereas 60% of control activity was observed with the SA mutant (data not shown). LpxA activity for the double site SA/SB mutant could not be determined because of difficulty in preparation of the corresponding acyl-ACP substrate (which requires both ACPS and AAS reactions), although we would anticipate little or no LpxA activity.

Kinetic analysis of LpxA is complicated by the thermodynamically unfavorable reaction in the forward direction (41). To further characterize the ACP site elimination mutants in this system, the phosphopantetheine sulfhydryl of ACP was modified with AEDANS to monitor interaction with LpxA using steady-state fluorescence spectroscopy. As shown in Fig. 6A, titration of AEDANS-rACP with LpxA caused a saturable 2-fold increase in fluorescence emission intensity, with a modest blue shift of the emission peak from 490 to 470 nm. The calculated dissociation constant of LpxA-ACP interaction based on these data (1.8 ± 0.1 µM, Fig. 6D) is very similar to the reported Km of LpxA for its beta-hydroxymyristoyl-ACP substrate and the IC50 for myristoyl-ACP (41). Thus, despite the obvious differences between the attached AEDANS moiety and an acyl group, AEDANS-ACP appears to be a useful conformational probe for LpxA interaction. AEDANS-modified SA mutant also exhibited a fluorescence increase upon titration with LpxA (apparent Kd of 4.1 ± 1.3 µM), although the magnitude of this increase at saturation was only 30% that of AEDANS-rACP (Fig. 6B). No effect of LpxA on the fluorescence emission of AEDANS-SB was observed, indicating either the absence of interaction or no change in the probe environment upon binding (Fig. 6C). These results are consistent with the greater inability of the SB versus SA mutant to support LpxA activity, and suggest that both decreased binding affinity and resulting changes in the probe environment upon LpxA interaction may be impaired by mutation of acidic residues surrounding helix II.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
ACP can be considered among a group of so-called "natively unfolded" proteins (42) because of its extraordinary acidic nature and relatively low composition of nonpolar amino acids. The calculated pI of V. harveyi ACP is 3.8 and 22 of its 76 residues are acidic, resulting in a flexible and mobile structure that may facilitate interaction with multiple enzymes (25). Most ACP-dependent enzymes of known three-dimensional structure, e.g. ACPS (26), LpxA (43), and FAS component enzymes FabA, FabB, FabD, FabF, FabG, FabH, and FabI (1, 22, 23), contain a basic/hydrophobic patch on the surface that attracts and participates in the binding of ACP substrates. Previous studies have highlighted the importance of acidic residues in ACP function. Amidation of 50% of the carboxylates on E. coli ACP induced no change in secondary structure but completely destroyed FAS activity (44). The central portion of ACP, including helix II and the flanking site A and site B cation binding regions are particularly anionic with 15 acidic and no basic residues between positions 25 and 60. This region has been implicated in recognition of several ACP-dependent enzymes (22–25). The present results clearly indicate that while mutations that neutralize acidic residues at site A or site B have similar effects on ACP conformation, they have more diverse consequences on the activity of selected ACP-dependent enzymes.

Any effects of ACP mutations on enzyme activity must be interpreted in the context of their influence on conformation. Unlike previously examined V. harveyi ACP mutants A75H (29) or E41K (36), in which strategic placement of a +1 or +2 charge difference abolished the Mg2+ requirement for conformational stability, all individual site A and B mutants were similar to the rACP template, undergoing transition from largely unfolded to helical conformation in the presence of Mg2+. Divalent cations are believed to stabilize folded ACP conformation at these sites through reduction of electrostatic repulsion between carboxylate groups (28). Note that these sites would be expected to be occupied by Mg2+ under physiological conditions.

As our experiments did not directly measure Mg2+ binding (i.e. in the absence of an observable conformational change), we do not know whether Mg2+-induced folding of individual mutants was the result of cation binding to both sites or to the single unaffected site. However, as one might expect mutation of even a single acidic residue to decrease Mg2+ affinity at that site, the simplest explanation is that cation binding to either site is sufficient to elicit the complete conformation change. Indeed, combined neutralizing mutations at each site did result in partial stabilization of folded ACP in the absence of Mg2+. Moreover, the progressively greater helical content of SA (+3 charge difference), SB (+4), and SA/SB (+7) mutants suggests additive stabilization in response to decreasing electrostatic repulsion in this region of the protein. In any case, the unaffected site in both the SA and SB mutants exhibited a Mg2+ affinity similar to that of rACP, indicating that each site can bind this cation and undergo conformational change independently of the other. The modest increase in Mg2+ concentration required for conformational transition of apo-versus holo-ACP is consistent with recent NMR evidence for transient interaction of the phosphopantetheine group with the polypeptide chain of E. coli ACP (45).

In contrast to the similar consequences of abolishing cation binding site A versus site B on ACP conformation and response to Mg2+, mutation of these sites had markedly different effects on individual ACP-dependent enzyme activities. Among these enzymes, ACPS is quite distinct in that it modifies the ACP polypeptide chain directly: the N terminus of helix II must be positioned at the ACPS active site for the Mg2+-dependent transfer of 4'-phosphopantetheine from CoA. The x-ray structure of the B. subtilis ACP-ACPS complex has identified several acidic residues in the vicinity of helix II, including Asp-35, Asp-38, Glu-41, Asp-48, and Asp-56 that participate in electrostatic interactions with basic residues of ACPS (26). Of these, the most important contact is between Arg-14 of ACPS and both Asp-35 (salt bridge) and Asp-38 (H-bond) of ACP, and helix II is displaced toward the enzyme in the complex relative to the corresponding free ACP in solution (16). Our results are quite consistent with this model in that mutation of Asp-35 in site A or Asp-56 in site B had a profound impact on ACPS activity. We can also speculate that the D38N mutation would not prevent its role as a H-bond acceptor in interaction with ACPS, although why there is greater conversion to the holo form for this mutant is not clear at this time.

Whereas the lack of ACPS activity with SA or SA/SB can likely be attributed to a dominant requirement for Asp-35, interpretation of our results with site elimination mutant SB are more complicated. This mutant appears to exhibit decreased affinity for the enzyme (Fig. 5A) while removing substrate inhibition (39, 40) at higher ACP concentrations. This might involve compensatory effects of individual site B mutations, e.g. D56N (which is a poor substrate on its own) and E53Q (which was completely converted to the holo form in vivo). It is important to note that ACPS appears to require a native ACP substrate conformation in this region, as this enzyme is inactive with an ACP mutant (I54A) that is incapable of folding even in the presence of Mg2+ (32).

In contrast to ACPS, activities of both AAS and LpxA were much more sensitive to neutralization of cation binding site B in the ACP substrate. These enzymes are respectively involved in the attachment of acyl groups to, and their transfer from, the phosphopantetheine sulfhydryl moiety. Although the structure of V. harveyi AAS is still unknown, previous work from our laboratory has implicated this region of ACP, in particular Phe-50 and Ile-54 (32), in interaction with this enzyme. Combined elimination of site A or B acidic residues appeared to have a much more significant effect on Km than on Vm, suggesting that these regions (site B in particular) are important for enzyme binding, although none of the individual acidic residues in site B appeared to be critical for AAS activity. Mutation of Asp-35 in E. coli ACP to cysteine was found to poorly support E. coli acyl-ACP synthetase activity, while similar alterations at Glu-30, Asp-38, and Asp-56 had less effect (24). However, V. harveyi acyl-ACP synthetase has been cloned recently and appears to be more closely related to medium chain acyl-CoA synthetases than to the membrane-bound E. coli AAS enzyme (46).

Our results are a little more surprising in the case of LpxA. TROSY NMR of E. coli ACP has recently revealed that residues 35–41 (i.e. site A region) undergo the greatest chemical shift perturbation in the presence of LpxA, and molecular docking analysis further suggested specific roles for Asp-35, Asp-38, and Glu-41 in LpxA binding (43). Our data also support a role for site A residues in interaction with LpxA, based on the partial decrease in LpxA activity and reduced affinity with the SA mutant (in which Glu-41 would still be present). However, while chemical shift perturbation of site B residues by LpxA was minimal in the NMR study (43), our results clearly indicate that these residues are critical for LpxA activity. Fluorescence studies also provided evidence consistent with limited SB interaction with LpxA. It seems unlikely that the inability of SB to bind Mg2+ at site B would explain its lack of function, as Mg2+ is not required for LpxA activity. This apparent paradox could reflect differences between LpxA interaction with the holo-ACP used in the NMR study (43), versus acyl- or AEDANS-ACP in our experiments. Alternatively, elimination of site B might impair another step in the LpxA reaction, such as a conformational change required to release the acyl group to the enzyme active site. Although further experiments will be required to resolve these issues, it is noteworthy that the site B region is important for maintaining ACP conformation (32) and forms part of the fatty acid-binding pocket.

Although early studies pointed to the involvement of ACP carboxylates in FAS activity (44), the current investigation has further shown that the acidic residues around helix II are collectively important for overall activity of the bacterial type II FAS complex. Recent experimental and modeling analyses have supported a role for residues in this region in interaction with specific FAS component enzymes: Glu-41 and Ala-45 in the case of the condensing enzyme FabH (22), Asp-35, Asp-38, and Ile-54 for ketoacyl-ACP reductase (FabG) (23), and Asp-35 for malonyl-CoA:ACP transacylase (FabD) (24). Clearly, interpretation of our results is limited in that we do not know which component(s) of the FAS complex are affected by specific site A or site B mutations. However, unlike E41K (36), none of the individual mutations analyzed in the current study caused complete loss of FAS activity, while elimination of either site A or site B had a much greater impact. Not surprisingly, FAS displayed normal activity with the D56N mutant, as this residue is exposed on the opposite face of ACP relative to most of the helix II residues implicated in enzyme interaction. Some loss of FAS activity with our mutant ACPs could be caused by impaired Mg2+ binding, as optimal E. coli FAS activity in the presence of 5–10 mM divalent cations has been attributed to interaction of cations with the negatively charged ACP substrate (47). We have observed a similar trend with the V. harveyi FAS system.3

In summary, we have investigated the role of acidic ACP residues in four separate enzyme systems that catalyze distinct ACP-dependent reactions. Overall, our results support the emerging consensus that helix II is a principal recognition motif for many enzymes that utilize ACP or its acylated derivatives as substrates. However, our data have also revealed that the relative importance of acidic residues at either end of this helix varies among these different enzymes, thus providing additional insight into the structure-function relationships of this small but essential bacterial protein.


   FOOTNOTES
 
* This work was supported by the Natural Sciences and Engineering Research Council of Canada. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed: Atlantic Research Centre, Dalhousie University, Rm C-305 Clinical Research Centre, 5849 University Ave., Halifax, Nova Scotia B3H 4H7, Canada. Tel.: 902-494-7084; Fax: 902-494-1394; E-mail: david.byers{at}dal.ca.

2 The abbreviations used are: ACP, acyl carrier protein; AAS, acyl-ACP synthetase; ACPS, holo-ACP synthase; CD, circular dichroism; FAS, fatty acid synthase; GST, glutathione S-transferase; IAEDANS, 5-(2-iodoacetylaminoethyl)aminonaphthalene-1-sulfonic acid; LpxA, acyl-ACP:UDP-GlcNAc acyltransferase; rACP, recombinant V. harveyi ACP; TLC, thin-layer chromatography; UDP-GlcNAc, uridine diphosphate N-acetylglucosamine. Back

3 H. Gong, unpublished results. Back


   ACKNOWLEDGMENTS
 
We thank members of the Atlantic Research Centre for helpful discussions, Drs. Amy Gehring and Christopher Walsh for providing the plasmid encoding E. coli ACPS, and Dr. Annette Henneberry for preparing the E. coli ACPS-His6 construct.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. White, S. W., Zheng, J., Zhang, Y. M., and Rock, C. O. (2005) Annu. Rev. Biochem. 74, 791–831[CrossRef][Medline] [Order article via Infotrieve]
  2. Rock, C. O., and Jackowski, S. (1982) J. Biol. Chem. 257, 10759–10765[Abstract/Free Full Text]
  3. Anderson, M. S., and Raetz, C. R. (1987) J. Biol. Chem. 262, 5159–5169[Abstract/Free Full Text]
  4. Jordan, S. W., and Cronan, J. E., Jr. (1997) J. Biol. Chem. 272, 17903–17906[Abstract/Free Full Text]
  5. Issartel, J. P., Koronakis, V., and Hughes, C. (1991) Nature 351, 759–761[CrossRef][Medline] [Order article via Infotrieve]
  6. Schaefer, A. L., Val, D. L., Hanzelka, B. L., Cronan, J. E., Jr., and Greenberg, E. P. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 9505–9509[Abstract/Free Full Text]
  7. Byers, D. M., and Meighen, E. A. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 6085–6089[Abstract/Free Full Text]
  8. Rumley, M. K., Therisod, H., Weissborn, A. C., and Kennedy, E. P. (1992) J. Biol. Chem. 267, 11806–11810[Abstract/Free Full Text]
  9. Geiger, O., Spaink, H. P., and Kennedy, E. P. (1991) J. Bacteriol. 173, 2872–2878[Abstract/Free Full Text]
  10. Shen, B., Summers, R. G., Gramajo, H., Bibb, M. J., and Hutchinson, C. R. (1992) J. Bacteriol. 174, 3818–3821[Abstract/Free Full Text]
  11. Stein, T., Vater, J., Kruft, V., Otto, A., Wittmann-Liebold, B., Franke, P., Panico, M., McDowell, R., and Morris, H. R. (1996) J. Biol. Chem. 271, 15428–15435[Abstract/Free Full Text]
  12. Heaton, M. P., and Neuhaus, F. C. (1994) J. Bacteriol. 176, 681–690[Abstract/Free Full Text]
  13. Butland, G., Peregrin-Alvarez, J. M., Li, J., Yang, W., Yang, X., Canadien, V., Starostine, A., Richards, D., Beattie, B., Krogan, N., Davey, M., Parkinson, J., Greenblatt, J., and Emili, A. (2005) Nature 433, 531–537[CrossRef][Medline] [Order article via Infotrieve]
  14. Kim, Y., and Prestegard, J. H. (1990) J. Am. Chem. Soc. 112, 3707–3709[CrossRef]
  15. Crump, M. P., Crosby, J., Dempsey, C. E., Parkinson, J. A., Murray, M., Hopwood, D. A., and Simpson, T. J. (1997) Biochemistry 36, 6000–6008[CrossRef][Medline] [Order article via Infotrieve]
  16. Xu, G. Y., Tam, A., Lin, L., Hixon, J., Fritz, C. C., and Powers, R. (2001) Structure 9, 277–287[Medline] [Order article via Infotrieve]
  17. Wong, H. C., Liu, G., Zhang, Y. M., Rock, C. O., and Zheng, J. (2002) J. Biol. Chem. 277, 15874–15880[Abstract/Free Full Text]
  18. Roujeinikova, A., Baldock, C., Simon, W. J., Gilroy, J., Baker, P. J., Stuitje, A. R., Rice, D. W., Slabas, A. R., and Rafferty, J. B. (2002) Structure 10, 825–835[Medline] [Order article via Infotrieve]
  19. Zornetzer, G. A., Fox, B. G., and Markely, J. L. (2006) Biochemistry 45, 5217–5227[CrossRef][Medline] [Order article via Infotrieve]
  20. Kim, Y., and Prestegard, J. H. (1989) Biochemistry 28, 8792–8797[CrossRef][Medline] [Order article via Infotrieve]
  21. Andrec, M., Hill, R. B., and Prestegard, J. H. (1995) Protein Sci. 4, 983–993[Medline] [Order article via Infotrieve]
  22. Zhang, Y. M., Rao, M. S., Heath, R. J., Price, A. C., Olson, A. J., Rock, C. O., and White, S. W. (2001) J. Biol. Chem. 276, 8231–8238[Abstract/Free Full Text]
  23. Zhang, Y. M., Wu, B., Zheng, J., and Rock, C. O. (2003) J. Biol. Chem. 278, 52935–52943[Abstract/Free Full Text]
  24. Worsham, L. M., Earls, L., Jolly, C., Langston, K. G., Trent, M. S., and Ernst-Fonberg, M. L. (2003) Biochemistry 42, 167–176[CrossRef][Medline] [Order article via Infotrieve]
  25. Zhang, Y. M., Marrakchi, H., White, S. W., and Rock, C. O. (2003) J. Lipid Res. 44, 1–10[Abstract/Free Full Text]
  26. Parris, K. D., Lin, L., Tam, A., Mathew, R., Hixon, J., Stahl, M., Fritz, C. C., Seehra, J., and Somers, W. S. (2000) Structure 8, 883–895[Medline] [Order article via Infotrieve]
  27. Tener, D. M., and Mayo, K. H. (1990) Eur. J. Biochem. 189, 559–565[Medline] [Order article via Infotrieve]
  28. Schulz, H. (1975) J. Biol. Chem. 250, 2299–2304[Abstract/Free Full Text]
  29. Keating, M. M., Gong, H., and Byers, D. M. (2002) Biochim. Biophys. Acta 1601, 208–214[Medline] [Order article via Infotrieve]
  30. Herald, V. L., Heazlewood, J. L., Day, D. A., and Millar, A. H. (2003) FEBS Lett. 537, 96–100[CrossRef][Medline] [Order article via Infotrieve]
  31. Frederick, A. F., Kay, L. E., and Prestegard, J. H. (1988) FEBS Lett. 238, 43–48[CrossRef][Medline] [Order article via Infotrieve]
  32. Flaman, A. S., Chen, J. M., Van Iderstine, S. C., and Byers, D. M. (2001) J. Biol. Chem. 276, 35934–35939[Abstract/Free Full Text]
  33. Lambalot, R. H., and Walsh, C. T. (1997) Methods Enzymol. 279, 254–262[CrossRef][Medline] [Order article via Infotrieve]
  34. Rock, C. O., Cronan, J. E., Jr., and Armitage, I. M. (1981) J. Biol. Chem. 256, 2669–2674[Abstract/Free Full Text]
  35. Fice, D., Shen, Z., and Byers, D. M. (1993) J. Bacteriol. 175, 1865–1870[Abstract/Free Full Text]
  36. Gong, H., and Byers, D. M. (2003) Biochem. Biophys. Res. Commun. 302, 35–40[CrossRef][Medline] [Order article via Infotrieve]
  37. Sweet, C. R., Lin, S., Cotter, R. J., and Raetz, C. R. H. (2001) J. Biol. Chem. 276, 19565–19574[Abstract/Free Full Text]
  38. Andrade, M. A., Chacon, P., Merelo, J. J., and Moran, F. (1993) Protein Eng. 6, 383–390[Abstract/Free Full Text]
  39. Flugel, R. S., Hwangbo, Y., Lambalot, R. H., Cronan, J. E., Jr., and Walsh, C. T. (2000) J. Biol. Chem. 275, 959–968[Abstract/Free Full Text]
  40. McAllister, K. A., Peery, R. B., and Zhao, G. (2006) J. Bacteriol. 188, 4737–4748[Abstract/Free Full Text]
  41. Anderson, M. S., Bull, H. G., Galloway, S. M., Kelly, T. M., Mohan, S., Radika, K., and Raetz, C. R. H. (1993) J. Biol. Chem. 268, 19858–19865[Abstract/Free Full Text]
  42. Wright, P. E., and Dyson, H. J. (1999) J. Mol. Biol. 293, 321–331[CrossRef][Medline] [Order article via Infotrieve]
  43. Jain, N. U., Wyckoff, T. J. O., Raetz, C. R. H., and Prestegard, J. H. (2004) J. Mol. Biol. 343, 1379–1389[CrossRef][Medline] [Order article via Infotrieve]
  44. Abita, J. P., Lazdunski, M., and Ailhaud, G. (1971) Eur. J. Biochem. 23, 412–420[Medline] [Order article via Infotrieve]
  45. Kim, Y., Kovrigin, E. L., and Eletr, Z. (2006) Biochem. Biophys. Res. Commun. 341, 776–783[CrossRef][Medline] [Order article via Infotrieve]
  46. Jiang, Y., Chan, C. H., and Cronan, J. E., Jr. (2006) Biochemistry 45, 10008–10019[CrossRef][Medline] [Order article via Infotrieve]
  47. Schulz, H., Weeks, G., Toomey, R. E., Shapiro, M., and Wakil, S. J. (1969) J. Biol. Chem. 244, 6577–6583[Abstract/Free Full Text]
  48. Pettersen, E. F., Goddard, T. D., Huang, C. C., Couch, G. S., Greenblatt, D. M., Meng, E. C., and Ferrin, T. E. (2004) J. Comput. Chem. 25, 1605–1612[CrossRef][Medline] [Order article via Infotrieve]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
D. I. Chan, T. Stockner, D. P. Tieleman, and H. J. Vogel
Molecular Dynamics Simulations of the Apo-, Holo-, and Acyl-forms of Escherichia coli Acyl Carrier Protein
J. Biol. Chem., November 28, 2008; 283(48): 33620 - 33629.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
N. R. De Lay and J. E. Cronan
In Vivo Functional Analyses of the Type II Acyl Carrier Proteins of Fatty Acid Biosynthesis
J. Biol. Chem., July 13, 2007; 282(28): 20319 - 20328.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
282/7/4494    most recent
M608234200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Gong, H.
Right arrow Articles by Byers, D. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Gong, H.
Right arrow Articles by Byers, D. M.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2007 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement