Overexpression of biotin synthase and biotin ligase is required for efficient generation of sulfur-35 labeled biotin in E. coli

<p>Abstract</p> <p>Background</p> <p>Biotin is an essential enzyme cofactor that acts as a CO₂carrier in carboxylation and decarboxylation reactions. The <it>E. coli </it>genome encodes a biosynthetic pathway that produces biotin from pimeloyl-CoA in four enzymatic steps. The final step, insertion of sulfur into desthiobiotin to form biotin, is catalyzed by the biotin synthase, BioB. A dedicated biotin ligase (BirA) catalyzes the covalent attachment of biotin to biotin-dependent enzymes. Isotopic labeling has been a valuable tool for probing the details of the biosynthetic process and assaying the activity of biotin-dependent enzymes, however there is currently no established method for ³⁵S labeling of biotin.</p> <p>Results</p> <p>In this study, we produced [³⁵S]-biotin from Na³⁵SO₄and desthiobiotin with a specific activity of 30.7 Ci/mmol, two orders of magnitude higher than previously published methods. The biotinylation domain (<it>Pf</it>BCCP-79) from the <it>Plasmodium falciparum </it>acetyl-CoA carboxylase (ACC) was expressed in <it>E. coli </it>as a biotinylation substrate. We found that overexpression of the <it>E. coli </it>biotin synthase, BioB, and biotin ligase, BirA, increased <it>Pf</it>BCCP-79 biotinylation 160-fold over basal levels. Biotinylated <it>Pf</it>BCCP-79 was purified by affinity chromatography, and free biotin was liberated using acid hydrolysis. We verified that we had produced radiolabeled biologically active [<it>D</it>]-biotin that specifically labels biotinylated proteins through reuptake in <it>E. coli</it>.</p> <p>Conclusions</p> <p>The strategy described in our report provides a simple and effective method for the production of [³⁵S]-biotin in <it>E. coli </it>based on affinity chromatography.</p

Increased prevalence of the pfdhfr/phdhps quintuple mutant and rapid emergence of pfdhps resistance mutations at codons 581 and 613 in Kisumu, Kenya

<p>Abstract</p> <p>Background</p> <p>Anti-malarial drug resistance in Kenya prompted two drug policy changes within a decade: sulphadoxine-pyrimethamine (SP) replaced chloroquine (CQ) as the first-line anti-malarial in 1998 and artemether-lumefantrine (AL) replaced SP in 2004. Two cross-sectional studies were conducted to monitor changes in the prevalence of molecular markers of drug resistance over the period in which SP was used as the first-line anti-malarial. The baseline study was carried out from 1999-2000, shortly after implementation of SP, and the follow-up study occurred from 2003-2005, during the transition to AL.</p> <p>Materials and methods</p> <p>Blood was collected from malaria smear-positive, symptomatic patients presenting to outpatient centers in Kisumu, Kenya, during the baseline and follow-up studies. Isolates were genotyped at codons associated with SP and CQ resistance. <it>In vitro </it>IC₅₀values for antifolates and quinolones were determined for isolates from the follow-up study.</p> <p>Results</p> <p>The prevalence of isolates containing the <it>pfdhfr </it>N51I/C59R/S108N/<it>pfdhps </it>A437G/K540E quintuple mutant associated with SP-resistance rose from 21% in the baseline study to 53% in the follow-up study (p < 0.001). Isolates containing the <it>pfdhfr </it>I164L mutation were absent from both studies. The <it>pfdhps </it>mutations A581G and A613S/T were absent from the baseline study but were present in 85% and 61%, respectively, of isolates from the follow-up study. At follow-up, parasites with mutations at five <it>pfdhps </it>codons, 436, 437, 540, 581, and 613, accounted for 39% of isolates. The CQ resistance-associated mutations <it>pfcrt </it>K76T and <it>pfmdr1 </it>N86Y rose from 82% to 97% (p = 0.001) and 44% to 76% (p < 0.001), respectively, from baseline to follow-up.</p> <p>Conclusions</p> <p>During the period in which SP was the first-line anti-malarial in Kenya, highly SP-resistant parasites emerged, including isolates harboring <it>pfdhps </it>mutations not previously observed there. SP continues to be widely used in Kenya; however, given the highly resistant genotypes observed in this study, its use as a first-line anti-malarial should be discouraged, particularly for populations without acquired immunity to malaria. The increase in the <it>pfcrt </it>K76T prevalence, despite efforts to reduce CQ use, suggests that either these efforts are not adequate to alleviate CQ pressure in Kisumu, or that drug pressure is derived from another source, such as the second-line anti-malarial amodiaquine.</p

The Amidase Domain of Lipoamidase Specifically Inactivates Lipoylated Proteins In Vivo

BACKGROUND:In the 1950s, Reed and coworkers discovered an enzyme activity in Streptococcus faecalis (Enterococcus faecalis) extracts that inactivated the Escherichia. coli and E. faecalis pyruvate dehydrogenase complexes through cleavage of the lipoamide bond. The enzyme that caused this lipoamidase activity remained unidentified until Jiang and Cronan discovered the gene encoding lipoamidase (Lpa) through the screening of an expression library. Subsequent cloning and characterization of the recombinant enzyme revealed that lipoamidase is an 80 kDa protein composed of an amidase domain containing a classic Ser-Ser-Lys catalytic triad and a carboxy-terminal domain of unknown function. Here, we show that the amidase domain can be used as an in vivo probe which specifically inactivates lipoylated enzymes. METHODOLOGY/PRINCIPAL FINDINGS:We evaluated whether Lpa could function as an inducible probe of alpha-ketoacid dehydrogenase inactivation using E. coli as a model system. Lpa expression resulted in cleavage of lipoic acid from the three lipoylated proteins expressed in E. coli, but did not result in cleavage of biotin from the sole biotinylated protein, the biotin carboxyl carrier protein. When expressed in lipoylation deficient E. coli, Lpa is not toxic, indicating that Lpa does not interfere with any other critical metabolic pathways. When truncated to the amidase domain, Lpa retained lipoamidase activity without acquiring biotinidase activity, indicating that the carboxy-terminal domain is not essential for substrate recognition or function. Substitution of any of the three catalytic triad amino acids with alanine produced inactive Lpa proteins. CONCLUSIONS/SIGNIFICANCE:The enzyme lipoamidase is active against a broad range of lipoylated proteins in vivo, but does not affect the growth of lipoylation deficient E. coli. Lpa can be truncated to 60% of its original size with only a partial loss of activity, resulting in a smaller probe that can be used to study the effects of alpha-ketoacid dehydrogenase inactivation in vivo

Lipoic Acid Metabolism in Microbial Pathogens

Summary: Lipoic acid [(R)-5-(1,2-dithiolan-3-yl)pentanoic acid] is an enzyme cofactor required for intermediate metabolism in free-living cells. Lipoic acid was discovered nearly 60 years ago and was shown to be covalently attached to proteins in several multicomponent dehydrogenases. Cells can acquire lipoate (the deprotonated charge form of lipoic acid that dominates at physiological pH) through either scavenging or de novo synthesis. Microbial pathogens implement these basic lipoylation strategies with a surprising variety of adaptations which can affect pathogenesis and virulence. Similarly, lipoylated proteins are responsible for effects beyond their classical roles in catalysis. These include roles in oxidative defense, bacterial sporulation, and gene expression. This review surveys the role of lipoate metabolism in bacterial, fungal, and protozoan pathogens and how these organisms have employed this metabolism to adapt to niche environments

Primer sequences.

<p>Primer sequences.</p

Growth and lipoylation of Lpa constructs containing only the amidase domain.

<p>(A) Growth assay of <i>E. coli</i> cells expressing Lpa_t471 (closed triangles) and Lpa_t521 (open squares) compared to expression of Lpa (open triangles) and vector alone (open circles). The OD₆₀₀ of cultures was measured at the time of induction with IPTG and at 2, 4, 6, and 10 hours post-induction. Error bars represent the standard deviation of three replicates from a representative growth assay. (B) Anti-lipoic acid and streptavidin affinity blots to determine levels of lipoylation and biotinylation in <i>E. coli</i> expressing empty vector, Lpa, and Lpa truncation mutants. (C) Densitometry analysis of PDH and KDH lipoylation in cells expressing Lpa and Lpa truncation mutants. The fraction of lipoylation in cells expressing Lpa or the Lpa active site mutants relative to cells expressing vector alone was determined as described in <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0007392#pone-0007392-g004" target="_blank">Figure 4C</a>. Error bars represent the SEM for the fraction of lipoylation for three independent western blots. (D) Solubility of Lpa and Lpa truncation mutants expressed at 20°C and 37°C. After induction of protein expression, cells were grown at 20°C for 10 hours or 37°C for four hours. Lpa in the insoluble (I) and soluble (S) fractions was analyzed by anti-His western blot.</p

Growth of lipoylation deficient <i>E. coli</i> expressing Lpa and Lpa active site mutants.

<p>Growth assay of TM136 strain cells <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0007392#pone.0007392-Morris1" target="_blank">[16]</a> expressing Lpa (squares), Lpa S235A (upward triangles), and Lpa S259A (circles). Growth curves at 37°C (solid symbols) and 20°C (open symbols are shown with cultures induced with 0.4 mM IPTG highlighted in red. In each condition, the growth curve corresponding to cells expressing wild type Lpa is shown with a thickened line.</p

Uptake and incorporation of ³⁵S-lipoic acid in the presence of Lpa.

<p>(A) KER176 cells that are auxotrophic for lipoic acid were transformed with plasmids encoding wild type Lpa (Lpa) or the active site nucleophile mutant Lpa S259A and were grown in minimal medium supplemented with ³⁵S-lipoic acid. After a 10 hour induction at 20°C, cell samples were normalized by OD₆₀₀ and protein extracts were separated by SDS-PAGE and analyzed by autoradiography. The assignment of the labeled species to the three lipoylated proteins in <i>E. coli</i>, the PDH, KDH, and H-protein, is indicated. (B) Scintillation counting was used to quantify ³⁵S-lipoic acid taken up by the KER176 cultures shown in <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0007392#pone-0007392-g002" target="_blank">Figure 2A</a>. Counts per minute (CPM) correspond to the uptake of cells from 5 µl of culture with an OD₆₀₀ of 1.1.</p

Lipoylation and biotinylation sites.

<p>(A) Amino acid sequence alignments of lipoylation and biotinylation sites in <i>E. coli</i>. The lysine that is involved in lipoic acid or biotin attachment is marked in bold. Residues corresponding to conserved glycine and glutamine residues are shaded <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0007392#pone.0007392-Fujiwara2" target="_blank">[33]</a>. Residues forming the biotinylation consensus site are underlined. (B) ClustalW comparison of amino acid sequences surrounding the site of lipoate attachment for lipoylated proteins found in <i>E. faecalis</i>. The substitution of the Glu three residues amino-terminal to the lipoyl lysine with Gln (underlined residues) is a common motif in the BCDH E2.</p

Lpa expression at 20 and 37°C.

<p>(A) Growth assay of <i>E. coli</i> cells expressing lipoamidase at 20°C (open triangles) and 37°C (closed triangles) relative to cells containing vector alone at 20°C (open circles) and 37°C (closed circles). The OD₆₀₀ of cultures was measured at the time of induction with IPTG and then 2, 4, 6, and 10 hours post-induction. Error bars represent the standard deviation of three replicates. (B) Anti-His western blot of cells expressing Lpa at 20°C and 37°C. Samples were taken from cultures at the time of induction and at 2, 4, 6, and 10 hours post-induction.</p