Streptomyces Development in Colonies and Soils

Streptomyces development was analyzed under conditions resembling those in soil. The mycelial growth rate was much lower than that in standard laboratory cultures, and the life span of the previously named first compartmentalized mycelium was remarkably increased.Streptomycetes are gram-positive, mycelium-forming, soil bacteria that play an important role in mineralization processes in nature and are abundant producers of secondary metabolites. Since the discovery of the ability of these microorganisms to produce clinically useful antibiotics (2, 15), they have received tremendous scientific attention (12). Furthermore, its remarkably complex developmental features make Streptomyces an interesting subject to study. Our research group has extended our knowledge about the developmental cycle of streptomycetes, describing new aspects, such as the existence of young, fully compartmentalized mycelia (5-7). Laboratory culture conditions (dense inocula, rich culture media, and relatively elevated temperatures [28 to 30°C]) result in high growth rates and an orderly-death process affecting these mycelia (first death round), which is observed at early time points (5, 7).In this work, we analyzed Streptomyces development under conditions resembling those found in nature. Single colonies and soil cultures of Streptomyces antibioticus ATCC 11891 and Streptomyces coelicolor M145 were used for this analysis. For single-colony studies, suitable dilutions of spores of these species were prepared before inoculation of plates containing GYM medium (glucose, yeast extract, malt extract) (11) or GAE medium (glucose, asparagine, yeast extract) (10). Approximately 20 colonies per plate were obtained. Soil cultures were grown in petri dishes with autoclaved oak forest soil (11.5 g per plate). Plates were inoculated directly with 5 ml of a spore suspension (1.5 × 10⁷ viable spores ml⁻¹; two independent cultures for each species). Coverslips were inserted into the soil at an angle, and the plates were incubated at 30°C. To maintain a humid environment and facilitate spore germination, the cultures were irrigated with 3 ml of sterile liquid GAE medium each week.The development of S. coelicolor M145 single colonies growing on GYM medium is shown in Fig. ​Fig.1.1. Samples were collected and examined by confocal microscopy after different incubation times, as previously described (5, 6). After spore germination, a viable mycelium develops, forming clumps which progressively extend along the horizontal (Fig. 1a and b) and vertical (Fig. 1c and d) axes of a plate. This mycelium is fully compartmentalized and corresponds to the first compartmentalized hyphae previously described for confluent surface cultures (Fig. 1e, f, and j) (see below) (5); 36 h later, death occurs, affecting the compartmentalized hyphae (Fig. 1e and f) in the center of the colony (Fig. ​(Fig.1g)1g) and in the mycelial layers below the mycelial surface (Fig. 1d and k). This death causes the characteristic appearance of the variegated first mycelium, in which alternating live and dead segments are observed (Fig. 1f and j) (5). The live segments show a decrease in fluorescence, like the decrease in fluorescence that occurs in solid confluent cultures (Fig. ​(Fig.11 h and i) (5, 9). As the cycle proceeds, the intensity of the fluorescence in these segments returns, and the segments begin to enlarge asynchronously to form a new, multinucleated mycelium, consisting of islands or sectors on the colony surfaces (Fig. 1m to o). Finally, death of the deeper layers of the colony (Fig. ​(Fig.1q)1q) and sporulation (Fig. ​(Fig.1r)1r) take place. Interestingly, some of the spores formed germinate (Fig. ​(Fig.1s),1s), giving rise to a new round of mycelial growth, cell death, and sporulation. This process is repeated several times, and typical, morphologically heterogeneous Streptomyces colonies grow (not shown). The same process was observed for S. antibioticus ATCC 11891, with minor differences mainly in the developmental time (not shown).Open in a separate windowFIG. 1.Confocal laser scanning fluorescence microscopy analysis of the development-related cell death of S. coelicolor M145 in surface cultures containing single colonies. Developmental culture times (in hours) are indicated. The images in panels l and n were obtained in differential interference contrast mode and correspond to the same fields as in panels k and m, respectively. The others are culture sections stained with SYTO 9 and propidium iodide. Panels c, d, k, l, p, and q are cross sections; the other images are longitudinal sections (see the methods). Panels h and i are images of the same field taken with different laser intensities, showing low-fluorescence viable hyphae in the center of the colonies that develop into a multinucleated mycelium. The arrows in panels e and s indicate septa (e) and germinated spores (s). See the text for details.Figure ​Figure22 shows the different types of mycelia present in S. coelicolor cultures under the conditions described above, depending on the compartmentalization status. Hyphae were treated with different fluorescent stains (SYTO 9 plus propidium iodide for nucleic acids, CellMask plus FM4-64 for cell membranes, and wheat germ agglutinin [WGA] for cell walls). Samples were processed as previously described (5). The young initial mycelia are fully compartmentalized and have membranous septa (Fig. 2b to c) with little associated cell wall material that is barely visible with WGA (Fig. ​(Fig.2d).2d). In contrast, the second mycelium is a multinucleated structure with fewer membrane-cell wall septa (Fig. 2e to h). At the end of the developmental cycle, multinucleated hyphae begin to undergo the segmentation which precedes the formation of spore chains (Fig. 2i to m). Similar results were obtained for S. antibioticus (not shown), but there were some differences in the numbers of spores formed. Samples of young and late mycelia were freeze-substituted using the methodology described by Porta and Lopez-Iglesias (13) and were examined with a transmission electron microscope (Fig. 2n and o). The septal structure of the first mycelium (Fig. ​(Fig.2n)2n) lacks the complexity of the septal structure in the second mycelium, in which a membrane with a thick cell wall is clearly visible (Fig. ​(Fig.2o).2o). These data coincide with those previously described for solid confluent cultures (4).Open in a separate windowFIG. 2.Analysis of S. coelicolor hyphal compartmentalization with several fluorescent indicators (single colonies). Developmental culture times (in hours) are indicated. (a, e, and i) Mycelium stained with SYTO 9 and propidium iodide (viability). (b, f, and j) Hyphae stained with Cell Mask (a membrane stain). (c, g, and l) Hyphae stained with FM 4-64 (a membrane stain). (d, h, and m) Hyphae stained with WGA (cell wall stain). Septa in all the images in panels a to j, l, and m are indicated by arrows. (k) Image of the same field as panel j obtained in differential interference contrast mode. (n and o) Transmission electron micrographs of S. coelicolor hyphae at different developmental phases. The first-mycelium septa (n) are comprised of two membranes separated by a thin cell wall; in contrast, second-mycelium septa have thick cell walls (o). See the text for details. IP, propidium iodide.The main features of S. coelicolor growing in soils are shown in Fig. ​Fig.3.3. Under these conditions, spore germination is a very slow, nonsynchronous process that commences at about 7 days (Fig. 3c and d) and lasts for at least 21 days (Fig. 3i to l), peaking at around 14 days (Fig. 3e to h). Mycelium does not clump to form dense pellets, as it does in colonies; instead, it remains in the first-compartmentalized-mycelium phase during the time analyzed. Like the membrane septa in single colonies, the membrane septa of the hyphae are stained with FM4-64 (Fig. 3j and k), although only some of them are associated with thick cell walls (WGA staining) (Fig. ​(Fig.3l).3l). Similar results were obtained for S. antibioticus cultures (not shown).Open in a separate windowFIG. 3.Confocal laser scanning fluorescence microscopy analysis of the development-related cell death and hyphal compartmentalization of S. coelicolor M145 growing in soil. Developmental culture times (in days) are indicated. The images in panels b, f, and h were obtained in differential interference contrast mode and correspond to the same fields as the images in panels a, e, and g, respectively. The dark zone in panel h corresponds to a particle of soil containing hyphae. (a, c, d, e, g, i, j, and k) Hyphae stained with SYTO 9, propidium iodide (viability stain), and FM4-64 (membrane stain) simultaneously. (i) SYTO 9 and propidium iodide staining. (j) FM4-64 staining. The image in panel k is an overlay of the images in panels i and j and illustrates that first-mycelium membranous septa are not always apparent when they are stained with nucleic acid stains (SYTO 9 and propidium iodide). (l) Hyphae stained with WGA (cell wall stain), showing the few septa with thick cell walls present in the cells. Septa are indicated by arrows. IP, propidium iodide.In previous work (8), we have shown that the mycelium currently called the substrate mycelium corresponds to the early second multinucleated mycelium, according to our nomenclature, which still lacks the hydrophobic layers characteristic of the aerial mycelium. The aerial mycelium therefore corresponds to the late second mycelium which has acquired hydrophobic covers. This multinucleated mycelium as a whole should be considered the reproductive structure, since it is destined to sporulate (Fig. ​(Fig.4)4) (8). The time course of lysine 6-aminotransferase activity during cephamycin C biosynthesis has been analyzed by other workers using isolated colonies of Streptomyces clavuligerus and confocal microscopy with green fluorescent protein as a reporter (4). A complex medium and a temperature of 29°C were used, conditions which can be considered similar to the conditions used in our work. Interestingly, expression did not occur during the development of the early mycelium and was observed in the mycelium only after 80 h of growth. This suggests that the second mycelium is the antibiotic-producing mycelium, a hypothesis previously confirmed using submerged-growth cultures of S. coelicolor (9).Open in a separate windowFIG. 4.Cell cycle features of Streptomyces growing under natural conditions. Mycelial structures (MI, first mycelium; MII, second mycelium) and cell death are indicated. The postulated vegetative and reproductive phases are also indicated (see text).The significance of the first compartmentalized mycelium has been obscured by its short life span under typical laboratory culture conditions (5, 6, 8). In previous work (3, 7), we postulated that this structure is the vegetative phase of the bacterium, an hypothesis that has been recently corroborated by proteomic analysis (data not shown). Death in confluent cultures begins shortly after germination (4 h) and continues asynchronously for 15 h. The second multinucleated mycelium emerges after this early programmed cell death and is the predominant structure under these conditions. In contrast, as our results here show, the first mycelium lives for a long time in isolated colonies and soil cultures. As suggested in our previous work (5, 6, 8), if we assume that the compartmentalized mycelium is the Streptomyces vegetative growth phase, then this phase is the predominant phase in individual colonies (where it remains for at least 36 h), soils (21 days), and submerged cultures (around 20 h) (9). The differences in the life span of the vegetative phase could be attributable to the extremely high cell densities attained under ordinary laboratory culture conditions, which provoke massive differentiation and sporulation (5-7, 8).But just exactly what are “natural conditions”? Some authors have developed soil cultures of Streptomyces to study survival (16, 17), genetic transfer (14, 17-19), phage-bacterium interactions (3), and antibiotic production (1). Most of these studies were carried out using amended soils (supplemented with chitin and starch), conditions under which growth and sporulation were observed during the first few days (1, 17). These conditions, in fact, might resemble environments that are particularly rich in organic matter where Streptomyces could conceivably develop. However, natural growth conditions imply discontinuous growth and limited colony development (20, 21). To mimic such conditions, we chose relatively poor but more balanced carbon-nitrogen soil cultures (GAE medium-amended soil) and less dense spore inocula, conditions that allow longer mycelium growth times. Other conditions assayed, such as those obtained by irrigating the soil with water alone, did not result in spore germination and mycelial growth (not shown). We were unable to detect death, the second multinucleated mycelium described above, or sporulation, even after 1 month of incubation at 30°C. It is clear that in nature, cell death and sporulation must take place at the end of the long vegetative phase (1, 17) when the imbalance of nutrients results in bacterial differentiation.In summary, the developmental kinetics of Streptomyces under conditions resembling conditions in nature differs substantially from the developmental kinetics observed in ordinary laboratory cultures, a fact that should be born in mind when the significance of development-associated phenomena is analyzed.     

Overexpression of the groESL Operon Enhances the Heat and Salinity Stress Tolerance of the Nitrogen-Fixing Cyanobacterium Anabaena sp. Strain PCC7120

The bicistronic groESL operon, encoding the Hsp60 and Hsp10 chaperonins, was cloned into an integrative expression vector, pFPN, and incorporated at an innocuous site in the Anabaena sp. strain PCC7120 genome. In the recombinant Anabaena strain, the additional groESL operon was expressed from a strong cyanobacterial P_psbA1 promoter without hampering the stress-responsive expression of the native groESL operon. The net expression of the two groESL operons promoted better growth, supported the vital activities of nitrogen fixation and photosynthesis at ambient conditions, and enhanced the tolerance of the recombinant Anabaena strain to heat and salinity stresses.Nitrogen-fixing cyanobacteria, especially strains of Nostoc and Anabaena, are native to tropical agroclimatic conditions, such as those of Indian paddy fields, and contribute to the carbon (C) and nitrogen (N) economy of these soils (22, 30). However, their biofertilizer potential decreases during exposure to high temperature, salinity, and other such stressful environments (1). A common target for these stresses is cellular proteins, which are denatured and inactivated during stress, resulting in metabolic arrest, cessation of growth, and eventually loss of viability. Molecular chaperones play a major role in the conformational homeostasis of cellular proteins (13, 16, 24, 26) by (i) proper folding of nascent polypeptide chains; (ii) facilitating protein translocation and maturation to functional conformation, including multiprotein complex assembly; (iii) refolding of misfolded proteins; (iv) sequestering damaged proteins to aggregates; and (v) solubilizing protein aggregates for refolding or degradation. Present at basal levels under optimum growth conditions in bacteria, the expression of chaperonins is significantly enhanced during heat shock and other stresses (2, 25, 32).The most common and abundant cyanobacterial chaperones are Hsp60 proteins, and nitrogen-fixing cyanobacteria possess two or more copies of the hsp60 or groEL gene (http://genome.kazusa.or.jp/cyanobase). One occurs as a solitary gene, cpn60 (17, 21), while the other is juxtaposed to its cochaperonin encoding genes groES and constitutes a bicistronic operon groESL (7, 19, 31). The two hsp60 genes encode a 59-kDa GroEL and a 61-kDa Cpn60 protein in Anabaena (2, 20). Both the Hsp60 chaperonins are strongly expressed during heat stress, resulting in the superior thermotolerance of Anabaena, compared to the transient expression of the Hsp60 chaperonins in Escherichia coli (20). GroEL and Cpn60 stably associate with thylakoid membranes in Anabaena strain PCC7120 (14) and in Synechocystis sp. strain PCC6803 (15). In Synechocystis sp. strain PCC6803, photosynthetic inhibitors downregulate, while light and redox perturbation induce cpn60 expression (10, 25, 31), and a cpn60 mutant exhibits a light-sensitive phenotype (http://genome.kazusa.or.jp/cyanobase), indicating a possible role for Cpn60 in photosynthesis. GroEL, a lipochaperonin (12, 28), requires a cochaperonin, GroES, for its folding activity and has wider substrate selectivity. In heterotrophic nitrogen-fixing bacteria, such as Klebsiella pneumoniae and Bradyrhizobium japonicum, the GroEL protein has been implicated in nif gene expression and the assembly, stability, and activity of the nitrogenase proteins (8, 9, 11).Earlier work from our laboratory demonstrated that the Hsp60 family chaperonins are commonly induced general-stress proteins in response to heat, salinity, and osmotic stresses in Anabaena strains (2, 4). Our recent work elucidated a major role of the cpn60 gene in the protection from photosynthesis and the nitrate reductase activity of N-supplemented Anabaena cultures (21). In this study, we integrated and constitutively overexpressed an extra copy of the groESL operon in Anabaena to evaluate the importance and contribution of GroEL chaperonin to the physiology of Anabaena during optimal and stressful conditions.Anabaena sp. strain PCC7120 was photoautotrophically grown in combined nitrogen-free (BG11⁻) or 17 mM NaNO₃-supplemented (BG11⁺) BG11 medium (5) at pH 7.2 under continuous illumination (30 μE m⁻² s⁻¹) and aeration (2 liters min⁻¹) at 25°C ± 2°C. Escherichia coli DH5α cultures were grown in Luria-Bertani medium at 37°C at 150 rpm. For E. coli DH5α, kanamycin and carbenicillin were used at final concentrations of 50 μg ml⁻¹ and 100 μg ml⁻¹, respectively. Recombinant Anabaena clones were selected on BG11⁺ agar plates supplemented with 25 μg ml⁻¹ neomycin or in BG11⁻ liquid medium containing 12.5 μg ml⁻¹ neomycin. The growth of cyanobacterial cultures was estimated either by measuring the chlorophyll a content as described previously (18) or the turbidity (optical density at 750 nm). Photosynthesis was measured as light-dependent oxygen evolution at 25 ± 2°C by a Clark electrode (Oxy-lab 2/2; Hansatech Instruments, England) as described previously (21). Nitrogenase activity was estimated by acetylene reduction assays, as described previously (3). Protein denaturation and aggregation were measured in clarified cell extracts containing ∼500 μg cytosolic proteins treated with 100 μM 8-anilino-1-naphthalene sulfonate (ANS). The pellet (protein aggregate) was solubilized in 20 mM Tris-6 M urea-2% sodium dodecyl sulfate (SDS)-40 mM dithiothreitol for 10 min at 50°C. The noncovalently trapped ANS was estimated using a fluorescence spectrometer (model FP-6500; Jasco, Japan) at a λ_excitation of 380 nm and a λ_emission of 485 nm, as described previously (29).The complete bicistronic groESL operon (2.040 kb) (GenBank accession no. {"type":"entrez-nucleotide","attrs":{"text":"FJ608815","term_id":"222354886","term_text":"FJ608815"}}FJ608815) was PCR amplified from PCC7120 genomic DNA using specific primers (Table ​(Table1)1) and the amplicon cloned into the NdeI-BamHI restriction sites of plasmid vector pFPN, which allows integration at a defined innocuous site in the PCC7120 genome and expression from a strong cyanobacterial P_psbA1 promoter (6). The resulting construct, designated pFPNgro (Table ​(Table1),1), was electroporated into PCC7120 using an exponential-decay wave form electroporator (200 J capacitive energy at a full charging voltage of 2 kV; Pune Polytronics, Pune, India), as described previously (6). The electroporation was carried out at 6 kV cm⁻¹ for 5 ms, employing an external autoclavable electrode with a 2-mm gap. The electroporation buffer contained high concentrations of salt (10 mM HEPES, 100 mM LiCl, 50 mM CaCl₂), as have been recommended for plant cells (23) and other cell types (27). The electrotransformants, selected on BG11⁺ agar plates supplemented with 25 μg ml⁻¹ neomycin by repeated subculturing for at least 25 weeks to achieve complete segregation, were designated AnFPNgro.

TABLE 1.

Plasmids, strains, and primers used in this study

Ganglioside GT1b Is a Putative Host Cell Receptor for the Merkel Cell Polyomavirus

The Merkel cell polyomavirus (MCPyV) was identified recently in human Merkel cell carcinomas, an aggressive neuroendocrine skin cancer. Here, we identify a putative host cell receptor for MCPyV. We found that recombinant MCPyV VP1 pentameric capsomeres both hemagglutinated sheep red blood cells and interacted with ganglioside GT1b in a sucrose gradient flotation assay. Structural differences between the analyzed gangliosides suggest that MCPyV VP1 likely interacts with sialic acids on both branches of the GT1b carbohydrate chain. Identification of a potential host cell receptor for MCPyV will aid in the elucidation of its entry mechanism and pathophysiology.Members of the polyomavirus (PyV) family, including simian virus 40 (SV40), murine PyV (mPyV), and BK virus (BKV), bind cell surface gangliosides to initiate infection (2, 8, 11, 15). PyV capsids are assembled from 72 pentamers (capsomeres) of the major coat protein VP1, with the internal proteins VP2 and VP3 buried within the capsids (7, 12). The VP1 pentamer makes direct contact with the carbohydrate portion of the ganglioside (10, 12, 13) and dictates the specificity of virus interaction with the cell. Gangliosides are glycolipids that contain a ceramide domain inserted into the plasma membrane and a carbohydrate domain that directly binds the virus. Specifically, SV40 binds to ganglioside GM1 (2, 10, 15), mPyV binds to gangliosides GD1a and GT1b (11, 15), and BKV binds to gangliosides GD1b and GT1b (8).Recently, a new human PyV designated Merkel cell PyV (MCPyV) was identified in Merkel cell carcinomas, a rare but aggressive skin cancer of neuroendocrine origin (3). It is as yet unclear whether MCPyV is the causative agent of Merkel cell carcinomas (17). A key to understanding the infectious and transforming properties of MCPyV is the elucidation of its cellular entry pathway. In this study, we identify a putative host cell receptor for MCPyV.Because an intact infectious MCPyV has not yet been isolated, we generated recombinant MCPyV VP1 pentamers in order to characterize cellular factors that bind to MCPyV. VP1 capsomeres have been previously shown to be equivalent to virus with respect to hemagglutination properties (4, 16), and the atomic structure of VP1 bound to sialyllactose has demonstrated that the capsomere is sufficient for this interaction (12, 13). The MCPyV VP1 protein (strain w162) was expressed and purified as described previously (1, 6). Briefly, a glutathione S-transferase-MCPyV VP1 fusion protein was expressed in Escherichia coli and purified using glutathione-Sepharose affinity chromatography. The fusion protein was eluted using glutathione and cleaved in solution with thrombin. The thrombin-cleaved sample was then rechromatographed on a second glutathione-Sepharose column to remove glutathione transferase and any uncleaved protein. The unbound VP1 was then chromatographed on a P-11 phosphocellulose column, and peak fractions eluting between 400 and 450 mM NaCl were collected. The purified protein was analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), followed by Coomassie blue staining (Fig. ​(Fig.1A,1A, left) and immunoblotting using an antibody (I58) that generally recognizes PyV VP1 proteins (Fig. ​(Fig.1A,1A, right) (9). Transmission electron microscopy (Philips CM10) analysis confirmed that the purified recombinant MCPyV VP1 formed pentamers (capsomeres), which did not assemble further into virus-like particles (Fig. ​(Fig.1B).1B). In an initial screening of its cell binding properties, we tested whether the MCPyV VP1 pentamers hemagglutinated red blood cells (RBCs). The MCPyV VP1 pentamers were incubated with sheep RBCs and assayed as previously described (5). SV40 and mPyV recombinant VP1 pentamers served as negative and positive controls, respectively. We found that MCPyV VP1 hemagglutinated the RBCs with the same efficiency as mPyV VP1 (protein concentration/hemagglutination unit) (Fig. ​(Fig.1C,1C, compare rows B and C from wells 1 to 11), suggesting that MCPyV VP1 engages a plasma membrane receptor on the RBCs. The recombinant murine VP1 protein used for comparison was from the RA strain, a small plaque virus (4). Thus, MCPyV VP1 has the hemagglutination characteristics of a small plaque mPyV (12, 13).Open in a separate windowFIG. 1.Characterization of MCPyV VP1. Recombinant MCPyV VP1 forms pentamers and hemagglutinates sheep RBCs. (A) Coomassie blue-stained SDS-PAGE and an immunoblot of the purified recombinant MCPyV VP1 protein are shown. Molecular mass markers are indicated. (B) Electron micrograph of the purified MCPyV VP1. MCPyV VP1 (shown in panel A) was diluted to 100 μg/ml and absorbed onto Formvar/carbon-coated copper grids. Samples were washed with phosphate-buffered saline, stained with 1% uranyl acetate, and visualized by transmission electron microscopy at 80 kV. Bar = 20 nm. (C) Sheep RBCs (0.5%) were incubated with decreasing concentrations of purified recombinant SV40 VP1 (row A), mPyV VP1 (row B), and MCPyV VP1 (row C). Wells 1 to 11 contain twofold serial dilutions of protein, starting at 2 μg/ml (well 1). Well 12 contains buffer only and serves as a negative control. Well 7 (rows B and C) corresponds to 128 hemagglutination units per 2 μg/ml VP1 protein.To characterize the chemical nature of the putative receptor for MCPyV, total membranes from RBCs were purified as described previously (15). The plasma membranes (30 μg) were incubated with MCPyV VP1 (0.5 μg) and floated on a discontinuous sucrose gradient (15). After fractionation, the samples were analyzed by SDS-PAGE, followed by immunoblotting with I58. VP1 was found in the bottom of the gradient in the absence of the plasma membranes (Fig. ​(Fig.2A,2A, first panel). In the presence of plasma membranes, a fraction of the VP1 floated to the middle of the gradient (Fig. ​(Fig.2A,2A, second panel), supporting the hemagglutination results that suggested that MCPyV VP1 binds to a receptor on the plasma membrane.Open in a separate windowFIG. 2.MCPyV VP1 binds to a protease-resistant, sialic acid-containing receptor on the plasma membrane. (A) Purified recombinant MCPyV VP1 was incubated with or without the indicated plasma membranes. The samples were floated in a discontinuous sucrose gradient, and the fractions were collected from the top of the gradient, subjected to SDS-PAGE, and immunoblotted with the anti-VP1 antibody I58. (B) Control and proteinase K-treated plasma membranes were subjected to SDS-PAGE, followed by Coomassie blue staining. (C) HeLa cells treated with proteinase K (4 μg/ml) were incubated with MCPyV at 4°C, and the resulting cell lysate was probed for the presence of MCPyV VP1. (D) As described in the legend to panel C, except 293T cells were used. (E) Purified MCPyV VP1 was incubated with plasma membranes pretreated with or without α2-3,6,8 neuraminidase and analyzed as described in the legend to panel A.To determine whether the receptor is a protein or a lipid, plasma membrane preparations (30 μg) were incubated with proteinase K (Sigma), followed by analysis with SDS-PAGE and Coomassie blue staining. Under these conditions, the majority of the proteins in the plasma membranes were degraded by the protease (Fig. ​(Fig.2B,2B, compare lanes 1 and 2). Despite the lack of proteins, the proteinase K-treated plasma membranes bound MCPyV VP1 as efficiently as control plasma membranes (Fig. ​(Fig.2A,2A, compare the second and third panels), demonstrating that MCPyV VP1 interacts with a protease-resistant receptor. The absence of VP1 in the bottom fraction in Fig. ​Fig.2A2A (third panel) is consistent with the fact that the buoyant density of the membranes is lowered by proteolysis. Of note, a similar result was seen with binding of the mPyV to the plasma membrane (15). Binding of MCPyV to the cell surface of two human tissue culture cells (i.e., HeLa and 293T) was also largely unaffected by pretreatment of the cells with proteinase K (Fig. 2C and D, compare lanes 1 and 2), further indicating that a nonproteinaceous molecule on the plasma membrane engages the virus.We next determined whether the protease-resistant receptor contains a sialic acid modification. Plasma membranes (10 μg) were incubated with a neuraminidase (α2-3,6,8 neuraminidase; Calbiochem) to remove the sialic acid groups. In contrast to the control plasma membranes, the neuraminidase-treated membranes did not bind MCPyV VP1 (Fig. ​(Fig.2E,2E, compare first and second panels), indicating that the MCPyV receptor includes a sialic acid modification.Gangliosides are lipids that contain sialic acid modifications. We asked if MCPyV VP1 binds to gangliosides similar to other PyV family members. The structures of the gangliosides used in this analysis (gangliosides GM1, GD1a, GD1b, and GT1b) are depicted in Fig. ​Fig.3A.3A. To assess a possible ganglioside-VP1 interaction, we employed a liposome flotation assay established previously (15). When liposomes (consisting of phosphatidyl-choline [19 μl of 10 mg/ml], -ethanolamine [5 μl of 10 mg/ml], -serine [1 μl of 10 mg/ml], and -inositol [3 μl of 10 mg/ml]) were incubated with MCPyV VP1 and subjected to the sucrose flotation assay, the VP1 remained in the bottom fraction (Fig. ​(Fig.3B,3B, first panel), indicating that VP1 does not interact with these phospholipids. However, when liposomes containing GT1b (1 μl of 1 mM), but not GM1 (1 μl of 1 mM) or GD1a (1 μl of 1 mM), were incubated with MCPyV VP1, the vesicles bound this VP1 (Fig. ​(Fig.3B).3B). A low level of virus floated partially when incubated with liposomes containing GD1b (Fig. ​(Fig.3B),3B), perhaps reflecting a weak affinity between MCPyV and GD1b. Importantly, MCPyV binds less efficiently to neuraminidase-treated GT1b-containing liposomes than to GT1b-containing liposomes (Fig. ​(Fig.3B,3B, sixth panel), suggesting that the GT1b sialic acids are involved in virus binding. This finding is consistent with the ability of neuraminidase to block MCPyV binding to the plasma membrane (Fig. ​(Fig.2E).2E). The level of virus flotation observed in the neuraminidase-treated GT1b-containing liposomes is likely due to the inefficiency of the neuraminidase reaction with a high concentration of GT1b used to prepare the vesicles.Open in a separate windowFIG. 3.MCPyV VP1 binds to ganglioside GT1b. (A) Structures of gangliosides GM1, GD1a, GD1b, and GT1b. The nature of the glycosidic linkages is indicated. (B) Purified MCPyV VP1 protein was incubated with liposomes only or with liposomes containing the indicated gangliosides. The samples were analyzed as described in the legend to Fig. ​Fig.2A.2A. Where indicated, GT1b-containing liposomes were pretreated with α2-3,6,8 neuraminidase and analyzed subsequently for virus binding. (C to E) The indicated viruses were incubated with liposomes and analyzed as described in the legend to panel B.As controls, GM1-containing liposomes bound SV40 (Fig. ​(Fig.3C),3C), GD1a-containing liposomes bound mPyV (Fig. ​(Fig.3D),3D), and GD1b-containing liposomes bound BKV (Fig. ​(Fig.3E),3E), demonstrating that the liposomes were functionally intact. We note that, while all of the MCPyV VP1 floated when incubated with liposomes containing GT1b (Fig. ​(Fig.3B,3B, sixth panel), a significant fraction of SV40, mPyV, and BKV VP1 remained in the bottom fraction despite being incubated with liposomes containing their respective ganglioside receptors (Fig. 3C to E, second panels). This result is likely due to the fact that in contrast to MCPyV, which are assembled as pentamers (Fig. ​(Fig.1B),1B), the SV40, mPyV, and BKV used in these experiments are fully assembled particles: their larger and denser nature prevents efficient flotation. Nonetheless, we conclude that MCPyV VP1 binds to ganglioside GT1b efficiently.The observation that GD1a does not bind to MCPyV VP1 suggests that the monosialic acid modification on the right branch of GT1b (Fig. ​(Fig.3A)3A) is insufficient for binding. Similarly, the failure of GD1b to bind MCPyV VP1 suggests that the sialic acid on the left arm of GT1b is necessary for binding. Together, these observations suggest that MCPyV VP1 interacts with sialic acids on both branches of GT1b (Fig. ​(Fig.4).4). A recent structure of SV40 VP1 in complex with the sugar portion of GM1 (10) demonstrated that although SV40 VP1 binds both branches of GM1 (Fig. ​(Fig.4),4), only a single sialic acid in GM1 is involved in this interaction. In the case of mPyV, structures of mPyV VP1 in complex with different carbohydrates (12, 13) revealed that the sialic acid-galactose moiety on the left branch of GD1a (and GT1b) is sufficient for mPyV VP1 binding (Fig. ​(Fig.4).4). Although no structure of BKV in complex with the sugar portion of GD1b (or GT1b) is available, in vitro binding studies (8) have suggested that the disialic acid modification on the right branch of GD1b (and GT1b) is responsible for binding BKV VP1 (Fig. ​(Fig.4).4). Thus, it appears that the unique feature of the MCPyV VP1-GT1b interaction is that the sialic acids on both branches of this ganglioside are likely involved in capsid binding.Open in a separate windowFIG. 4.A potential model of the different VP1-ganglioside interactions (see the text for discussion).The identification of a potential cellular receptor for MCPyV will facilitate the study of its entry mechanism. An important issue for further study is to determine whether MCPyV targets Merkel cells preferentially, and if so, whether GT1b is found in higher levels in these cells to increase susceptibility.     

Efficient Production of 2-Oxobutyrate from 2-Hydroxybutyrate by Using Whole Cells of Pseudomonas stutzeri Strain SDM

2-Oxobutyrate is an important intermediate in the chemical, drug, and food industries. Whole cells of Pseudomonas stutzeri SDM, containing NAD-independent lactate dehydrogenases, effectively converted 2-hydroxybutyrate into 2-oxobutyrate. Under optimal conditions, the biocatalytic process produced 2-oxobutyrate at a high concentration (44.4 g liter⁻¹) and a high yield (91.5%).2-Oxobutyrate (2-OBA) is used as a raw material in the synthesis of chiral 2-aminobutyric acid, isoleucine, and some kinds of medicines (1, 8). There is no suitable starting material for 2-OBA production by chemical synthesis; therefore, the development of innovative biotechnology-based techniques for 2-OBA production is desirable (12).2-Hydroxybutyrate (2-HBA) is cheaper than 2-OBA and can be substituted for 2-OBA in the production of isoleucine, as reported previously (9, 10). The results of those studies also indicated that it might be possible to produce 2-OBA from 2-HBA by a suitable biocatalytic process. In the presence of NAD, NAD-dependent 2-hydroxybutyrate dehydrogenase can catalyze the oxidation of 2-HBA to 2-OBA (4). However, due to the high cost of pyridine cofactors (11), it is preferable to use a biocatalyst that directly catalyzes the formation of 2-OBA from 2-HBA without any requirement for NAD as a cofactor.In our previous report, we confirmed that NAD-independent lactate dehydrogenases (iLDHs) in the pyruvate-producing strain Pseudomonas stutzeri SDM (China Center for Type Culture Collection no. M206010) could oxidize lactate and 2-HBA (6). Therefore, in addition to pyruvate production from lactate, P. stutzeri SDM might also have a potential application in 2-OBA production.To determine the 2-OBA production capability of P. stutzeri SDM, the strain was first cultured at 30°C in a minimal salt medium (MSM) supplemented with 5.0 g liter⁻¹ dl-lactate as the sole carbon source (5). The whole-cell catalyst was prepared by centrifuging the medium and resuspending the cell pellet, and biotransformation was then carried out under the following conditions using 2-HBA as the substrate and whole cells of P. stutzeri SDM as the biocatalyst: 2-HBA, 10 g liter⁻¹; dry cell concentration, 6 g liter⁻¹; buffer, 100 mM potassium phosphate (pH 7.0); temperature, 30°C; shaking speed, 300 rpm. After 4 h of reaction, the mixture was analyzed by high-performance liquid chromatography (HPLC; Agilent 1100 series; Hewlett-Packard) using a refractive index detector (3). The HPLC system was fitted with a Bio-Rad Aminex HPX-87 H column. The mobile phase consisted of 10 mM H₂SO₄ pumped at 0.4 ml min⁻¹ (55°C). Biotransformation resulted in the production of a compound that had a retention time of 19.57 min, which corresponded to the peak of authentic 2-OBA (see Fig. S1 in the supplemental material).After acidification and vacuum distillation, the new compound was analyzed by negative-ion mass spectroscopy. The molecular ion ([M − H]⁻, m/z 101.1) signal of the compound was consistent with the molecular weight of 2-OBA, i.e., 102.1 (see Fig. S2 in the supplemental material). These results confirmed that 2-HBA was oxidized to 2-OBA by whole cells of P. stutzeri SDM.To investigate whether iLDHs are responsible for 2-OBA production in the above-described biocatalytic process, 2-HBA oxidation activity in P. stutzeri SDM was probed by native polyacrylamide gel electrophoresis. After electrophoresis, the gels were soaked in a substrate solution [50 mM Tris-HCl buffer (pH 8.0) containing 0.1 mM phenazine methosulfate, 0.1 mM 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide, and 1 mM l-lactate, dl-lactate, or dl-2-HBA] and gently shaken. As shown in Fig. ​Fig.1,1, d- and l-iLDH migrated as two bands with distinct mobilities. The activities responsible for d- and l-2-HBA oxidation were located at the same positions as the d- and l-iLDH activities, respectively. No other bands responsible for d- and l-2-HBA oxidation were detected. Moreover, the dialysis of the crude cell extract did not lead to loss of 2-HBA oxidation activity and the addition of 10 mM NAD⁺ could not stimulate the reaction (see Table S1 in the supplemental material). These results implied that in the biocatalytic system, 2-HBA was oxidized to 2-OBA by iLDHs present in P. stutzeri SDM.Open in a separate windowFIG. 1.Activity staining of iLDHs after native polyacrylamide gel electrophoresis with lactate or 2-HBA as the substrate.Although the SDM strain could not use 2-HBA or 2-OBA for growth (see Fig. S3 in the supplemental material), 2-HBA might induce some of the enzymes responsible for 2-OBA production in the biocatalytic process. To exclude this possibility, the SDM strain was cultured in MSM containing dl-lactate or pyruvate as the sole carbon source. As shown in Fig. ​Fig.2,2, the enzyme activities that catalyzed lactate and 2-HBA oxidation were simultaneously present in the cells cultured on lactate and were absent in those cultured on pyruvate. After the lactate or pyruvate was exhausted, 5.05 g liter⁻¹ dl-2-HBA was added to the medium. It was observed that dl-2-HBA was efficiently converted to 2-OBA in the medium containing dl-lactate (Fig. ​(Fig.2a).2a). No 2-OBA production was detected in the medium containing pyruvate. Because 2-HBA addition did not induce the enzymes involved in 2-HBA oxidation (Fig. 2a and b), we concluded that the iLDHs induced by dl-lactate catalyzed 2-HBA oxidation in this biocatalytic process.Open in a separate windowFIG. 2.Time course of P. stutzeri SDM growth on media containing dl-lactate (a) and pyruvate (b). 2-HBA was added to the medium after the exhaustion of lactate or pyruvate. Symbols: ▴, lactate; ▵, pyruvate; •, 2-HBA; ○, 2-OBA; ▪, cell density; ▧, iLDHs activity with dl-lactate as the substrate; ▒, iLDHs activity with dl-2-HBA as the substrate.iLDHs could catalyze the oxidation of the substrate in a flavin-dependent manner and might use membrane quinone as the electron acceptor. Unlike the oxidases, which directly use the oxygen as the electron acceptor, this substrate oxidation mechanism could prevent the formation of H₂O₂ (see Fig. S4 in the supplemental material). The P. stutzeri SDM strain efficiently converted dl-2-HBA to 2-OBA with high yields (4.97 g liter⁻¹ 2-OBA was produced from 5.05 g liter⁻¹ dl-2-HBA); therefore, 2-OBA production by this strain can be a valuable and technically feasible process. To increase the efficiency of P. stutzeri SDM in the biotechnological production of 2-OBA, the conditions for biotransformation using whole cells of P. stutzeri SDM were first optimized. The influence of the reaction pH and 2-HBA concentration on 2-OBA production was determined in 100 mM phosphate buffer containing whole cells harvested from the medium containing dl-lactate as the sole carbon source. The reaction was initiated by adding the whole cells and 2-HBA at 37°C, followed by incubation for 10 min. After stopping the reaction by adding 1 M HCl, the 2-OBA concentration was determined by HPLC.As shown in Fig. ​Fig.3a,3a, ​,2-OBA2-OBA production was highest at pH 7.0. Under acidic or alkaline conditions, the transformation of 2-HBA to 2-OBA decreased. The optimal 2-HBA concentration was found to be 0.4 M, as shown in Fig. ​Fig.3b.3b. 2-OBA production increased as the 2-HBA concentration increased up to about 0.4 M and decreased thereafter. The concentration of the whole-cell catalyst was then optimized using 0.4 M 2-HBA as the substrate at pH 7.0. As shown in Fig. ​Fig.3c,3c, the highest 2-OBA concentration was obtained with 20 g (dry cell weight [DCW]) liter⁻¹ of P. stutzeri SDM. The 2-OBA concentration decreased with any increase beyond this cell concentration.Open in a separate windowFIG. 3.Optimization of the biocatalysis conditions. (a) Effect of pH on 2-OBA production activity. (b) Effect of 2-HBA concentrations on 2-OBA production activity. (c) Effect of the concentration of P. stutzeri SDM on biotransformation. OD, optical density.After optimizing the biocatalytic conditions, we studied the biotechnological production of 2-OBA from 2-HBA by using the whole-cell catalyst P. stutzeri SDM. As shown in Fig. ​Fig.4,4, when 20 g (DCW) liter⁻¹ P. stutzeri SDM was used as the biocatalyst, 48.5 g liter⁻¹ 2-HBA was biotransformed into 44.4 g liter⁻¹ 2-OBA in 24 h.Open in a separate windowFIG. 4.Time course of production of 2-OBA from 2-HBA under the optimum conditions. Symbols: ▪, 2-OBA; •, 2-HBA.Biocatalytic production of 2-OBA was carried out using crotonic acid, propionaldehyde, 1,2-butanediol, or threonine as the substrate (2, 7, 8, 12). Resting cells of the strain Rhodococcus erpi IF0 3730 produced 15.7 g liter⁻¹ 2-OBA from 20 g liter⁻¹ 1,2-butanediol, which is the highest reported yield of 2-OBA to date (8). By using the whole-cell catalyst P. stutzeri SDM, it was possible to produce 2-OBA at a high concentration (44.4 g liter⁻¹) and a high yield (91.5%). Due to the simple composition of the biocatalytic system (see Fig. S5 in the supplemental material), 2-HBA and 2-OBA could be easily separated on a column using a suitable resin. Separation of 2-OBA from the biocatalytic system was relatively inexpensive. The biocatalytic process presented in this report could be a promising alternative for the biotechnological production of 2-OBA.      

A Photostable Green Fluorescent Protein Variant for Analysis of Protein Localization in Candida albicans

Fusions to the green fluorescent protein (GFP) are an effective way to monitor protein localization. However, altered codon usage in Candida species has delayed implementation of new variants. Examination of three new GFP variants in Candida albicans showed that one has higher signal intensity and increased resistance to photobleaching.The human fungal pathogen Candida albicans can cause severe infections, particularly in immunocompromised patients. Important insights into its pathogenesis have been obtained by analyzing fusions to green fluorescent protein (GFP) (8). Although GFP tagging has been very successful, many fusion proteins are not easily detected. New GFP variants with improved fluorescence and protein folding properties have been identified by genetic approaches in other organisms (2, 7, 8). However, these GFP variants have not been assessed in C. albicans and related species, presumably because of the added difficulties of attempting heterologous expression in C. albicans.To adapt GFP for effective use in C. albicans, Cormack et al. introduced three types of codon changes: the S65G S72A mutations to enhance fluorescence; the CTG codon 201 change to TTG, since CUG is translated as Ser instead of Leu in C. albicans; and the optimization of the other codons for translation in C. albicans (1). This variant, known as YeGFP3, was introduced into convenient vectors for creating gene fusions in C. albicans (4). Another version of eGFP known as mut2 (S65A V68L S72A Q80R) was adapted for C. albicans by changing the CTG codon but without further codon optimization (5). These obstacles to heterologous expression in C. albicans have presumably delayed implementation of newer versions of GFP. Therefore, in this study three different GFP variants were introduced into YeGFP3 and examined for function in C. albicans.The GFP variants were constructed using standard methods to introduce changes in the coding sequence of YeGFP3. In brief, mutagenic oligonucleotides were used to prime PCR synthesis of a plasmid carrying YeGFP3, the template DNA was then destroyed by digestion with DpnI, and then the resulting DNA was transformed into Escherichia coli. DNA sequencing (carried out by the Stony Brook University DNA Sequencing Facility) confirmed that the correct substitutions were present. The mutant GFP genes were then released as PstI-AscI fragments and then were subcloned to replace the corresponding GFP fragment of plasmid pFa-GFP-URA3 (6), which carries a PCR cassette module for creating GFP fusions in C. albicans. Because of the large number of changes, the mutants were given the more convenient names of CaGFPα (F64L S65T F99S M153T V163A), CaGFPβ (F64L S65T N149K M153T I167T; also known as emerald), and CaGFPγ (F64L S65C V163A I167T). The CaGFPγ was also introduced into vectors that contain selectable markers HIS1 and ARG4 (6). DNA sequences used to design primers for creating GFP fusions in C. albicans were as follows: forward primer, 5′ (region of homology)-GGTGCTGGCGCAGGTGCTTC-3′, and reverse primer, 5′ (region of homology)-TCTGATATCATCGATGAATTCGAG-3′.CDC11-GFP fusion genes were created in C. albicans by homologous recombination, as described previously (4, 6). In brief, long oligonucleotide primers with homology to the 3′ end of the CDC11 open reading frame were used to prime PCR synthesis of each of the corresponding GFP variant genes plus an adjacent selectable marker gene (URA3). These DNA elements were then introduced into C. albicans cells and allowed to recombine with the homologous region of the CDC11 gene in C. albicans to create the CDC11-GFP fusion genes. Sequences used for the design of PCR primers to amplify the pFa-GFP plasmids are shown above. Cells carrying the indicated CDC11-GFP fusion gene were grown overnight in log phase in synthetic medium (yeast nitrogen base plus amino acids and dextrose). Cdc11-GFP fluorescence intensity was analyzed with an Olympus BH2 microscope equipped with a Zeiss AxioCam camera run by Openlab software. The relative GFP signal was determined by measuring the intensity of GFP fluorescence of the septin ring and then subtracting the fluorescence of an area immediately adjacent to each ring. All samples were visualized under the same conditions.Samples were prepared for Western blot analysis by resuspending cells in TNE lysis buffer (10 mM Tris base, 1 mM EDTA, 100 mM NaCl) with 100× protease mix (40 mg/ml pepstatin A, 40 mg/ml aprotinin, 20 mg/ml leupeptin) and then agitating in the presence of glass beads. The supernatant was collected after low-speed centrifugation at 3,000 rpm for 1 min, protein concentrations were determined by the bicinchoninic acid (BCA) protein assay (Pierce), and then equal amounts of protein extract were separated by gel electrophoresis and transferred to a Protran nitrocellulose membrane (Whatman GmbH). The blots were incubated with mouse anti-GFP (Millipore), rabbit anti-glucose-6-phosphate dehydrogenase (anti-G6PD; Sigma), or rabbit anti-Cdc11 (Santa Cruz Biotechnology) primary antibodies; washed; and then incubated with either goat anti-mouse IRDye 800cw or goat anti-rabbit IRDye 680 (Li-Cor Biosciences, Lincoln, NE). The immunoreactive proteins were visualized with a Li-Cor fluorescence scanner run by Odyssey software.Three new GFP variants based on YeGFP3 were constructed by introducing mutations predicted to improve either the fluorescence properties or protein folding (2, 7, 8). Because multiple changes were introduced into each variant, they were given the more convenient names of CaGFPα, CaGFPβ, and CaGFPγ (see above). The key mutations in CaGFPα and CaGFPβ have been described previously (2, 7, 8), but CaGFPγ represents a novel combination of mutations. The 3 new GFP variants plus the YeGFP3 and mut2 versions were compared by fusing them to the C terminus of the Cdc11 septin protein (3). The Cdc11 protein was selected because its restricted localization to the bud neck facilitated microscopic analysis and comparison of fluorescence properties. CDC11-GFP fusion genes were constructed in strain BWP17 (9) using PCR-generated modules with a URA3 selectable marker, as described previously (4, 6).Cells were grown in synthetic medium overnight to log phase at both 30°C and 37°C, temperatures that are commonly used to propagate C. albicans and that may affect the folding properties of GFP. GFP fluorescence was then analyzed by quantifying the intensity of the septin rings in digital images (Fig. ​(Fig.1A).1A). Septin rings were analyzed only if they were obviously in focus and at the same stage of the cell cycle (large budded). CaGFPγ gave a slightly stronger signal than the other variants, which was most obvious at 30°C (Fig. ​(Fig.1A).1A). At least two independent clones were analyzed for each CDC11-GFP variant, and the two gave similar results (data not shown).Open in a separate windowFIG. 1.Properties of Cdc11-GFP fusion proteins. Cells were grown to log phase overnight at the indicated temperature, and then Cdc11-GFP fluorescence was analyzed. (A) Signal intensity for the different versions of Cdc11-GFP was compared in three independent assays in which 50 septin rings per assay were quantified for each different Cdc11-GFP. The average fluorescence intensity was normalized to 100 for Cdc11-YeGPF3. The Cdc11-CaGFPγ variant gave a significantly stronger signal than the other variants (P < 0.001). (B) Western blot analysis comparing the levels of Cdc11-GFP produced in the indicated strains. The lane labeled “neg” refers to the negative-control strain (BWP17) that lacks GFP. Blots were probed with anti-GFP to detect Cdc11-GFP, anti-glucose-6-phosphate dehydrogenase (αG6PD) as a control, and anti-Cdc11 to detect the untagged version of Cdc11.The levels of the Cdc11-GFP proteins at both 30°C and 37°C were compared on two independent Western blots using anti-GFP antibody (Fig. ​(Fig.1B).1B). The relative levels of Cdc11-mut2GFP and Cdc11-CaGFPα were the lowest, consistent with their lower fluorescence intensity. The lower levels of Cdc11-mut2GFP are consistent with the fact that the codons in the mut2 version of GFP were not optimized for expression in C. albicans (5). The Cdc11-YeGFP3 and Cdc11-CaGFPγ were present at higher levels, and the Cdc11-CaGFPβ was produced at even slightly higher levels, consistent with reports that this latter version of GFP (also known as emerald) has improved folding properties (7). The Cdc11-GFP variants did not affect the production of the untagged Cdc11 protein (Fig. ​(Fig.1B1B).Photobleaching is also an important factor for GFP (7), especially in time-lapse studies or Z-stack analysis of different optical sections of cells. Photostability of the GFP variants was examined by taking pictures at 4-s intervals during 1 min of continuous exposure to the fluorescence excitation lamp (Fig. 2A and B). The fluorescence of YeGFP3, mut2GFP, and CaGFPα fused to Cdc11 decayed to 50% of original intensity within 15 to 30 s, and the rate of photobleaching was even higher for CaGFPβ. In contrast, Cdc11-CaGFPγ showed extended photostability at both 30°C and 37°C (half-life [t_1/2] of ∼2 min). Similar results were also obtained for CaGFPγ fused to the Golgi protein Vrg4 (data not shown), although the standard deviations were larger because the mobile Golgi compartments frequently moved out of the focal plane during the time course (data not shown). On a practical level, the Cdc11-GFPγ fluorescence was readily detectable after several minutes of continuous exposure (Fig. ​(Fig.2C),2C), demonstrating its clear advantage for allowing more time to observe protein localization before photobleaching becomes significant.Open in a separate windowFIG. 2.Photostability of GFP variants. (A and B) Relative fluorescence intensity of the GFP variants at 4-s intervals over a time course of 1 min of continuous exposure to the fluorescence excitation lamp after growth at 30°C (A) and at 37°C (B). CaGFPγ showed the best photostability (t_1/2 of ∼2 min). The relative fluorescence was normalized to 100 for each Cdc11-GFP variant at the start of the time course. The results represent the average of three independent assays in which three septin rings were analyzed for each mutant. Error bars indicate standard deviations. (C) Cells carrying Cdc11 fused to YeGFP3 or CaGFPγ were continuously exposed to the fluorescence excitation lamp, and then images of septin rings were captured at the indicated times.Altogether, Cdc11-CaGFPγ had the best overall properties based on protein levels, signal intensity, and photostability in C. albicans. The higher level of Cdc11-CaGFPβ production was apparently offset by increased photobleaching, resulting in no overall advantage for this variant. The Cdc11-CaGFPα was produced at relatively low levels, and it was less photostable compared to the other versions. Thus, CaGFPγ is a novel GFP variant that offers improved features for the study of protein localization in C. albicans and will likely also be useful for expression in other species.     

Production of Recombinant Proteins in the lon-Deficient BL21(DE3) Strain of Escherichia coli in the Absence of the DnaK Chaperone

To eliminate unavoidable contamination of purified recombinant proteins by DnaK, we present a unique approach employing a BL21(DE3) ΔdnaK strain of Escherichia coli. Selected representative purified proteins remained soluble, correctly assembled, and active. This finding establishes DnaK dispensability for protein production in BL21(DE3), which is void of Lon protease, key to eliminating unfolded proteins.Obtaining substantial amounts of pure protein is essential in innumerable biological studies and indispensable to the biochemical characterization of proteins. The ease of growth, well-characterized genetics, and the large number of tools for gene expression have long made Escherichia coli the organism of choice for protein overproduction. The BL21(DE3) strain is widely used for recombinant protein production because of its engineered capacity to produce T7 polymerase and its deficiency in Lon and OmpT proteases.DnaK is an abundant protein (about 1% of the total protein of E. coli) (17) that interacts with a wide range of newly synthesized polypeptides (28) and assists their proper folding and assembly into oligomers by preventing protein aggregation. DnaK, together with ClpB ATPase, is also required to disaggregate preformed protein aggregates (12, 20), and it participates in the degradation of damaged proteins by Lon and ClpP (26, 27).The inactivation of dnaK has been shown previously to increase the insoluble fractions of certain aggregation-prone recombinant proteins (7), and DnaK alleviates the aggregation of certain heterologous proteins when coproduced with the protein of interest (8). However, fruitful coproduction of recombinant proteins with chaperones has been challenged by recent findings demonstrating that chaperones increase the solubility but not necessarily the quality of proteins (13, 15).The DnaK binding site, a five-residue hydrophobic core flanked by two basic residue-enriched regions, occurs on average every 36 residues in protein sequences (25). One consequence is unwanted DnaK contamination of recombinant proteins during purification in E. coli, even after several chromatographic steps (1, 3, 11, 14, 16, 21, 23).One challenge in protein purification is to obtain the highest level of purity in the fewest steps. Biologically active impurities can jeopardize research or therapeutic applications even if present in trace amounts. One approach developed to circumvent DnaK contamination is extensive washing of columns with ATP since DnaK in its ATP-bound state has low affinity for protein (3). However, this strategy lengthens the purification procedure, is expensive, and is of inconsistent effectiveness (1, 14).In an attempt to eliminate DnaK contamination, we have investigated whether recombinant proteins could be produced in the absence of DnaK. Toward that end, we constructed a ΔdnaK derivative of the extensively employed E. coli B host strain BL21(DE3). The consequences of the absence of DnaK for the production, solubilities, correct assembly, and activities of several recombinant proteins in BL21(DE3) have been studied. Obtaining a BL21(DE3) ΔdnaK strain has allowed us to elucidate to what extent such a major E. coli chaperone is indispensable to protein overproduction in the particular genetic context of an E. coli strain that lacks Lon, an ATP-dependent protease responsible for degrading unfolded proteins (10).dnaK in BL21(DE3) was inactivated by the introduction of a null allele, ΔdnaK::Kan, from the E. coli PopC4617 strain by P1 transduction (see Table S1 in the supplemental material). Transductants were selected at 30°C in Luria-Bertani medium complemented with kanamycin. The absence of dnaK was verified by colony PCR using the specific primers dnaK-Nter (5′-GGTAAAATAATTGGTATCGACCTGG-3′) and dnaK-Cter (5′-GTCTTTGACTTCTTCAAATTCAGCG-3′) (see Fig. S1 in the supplemental material). Immunoblotting using an anti-DnaK antibody showed that the obtained transductant (EN2) did not produce DnaK (see Fig. S1 in the supplemental material). The EN2 strain has been deposited at the Collection Nationale de Culture de Microorganismes at the Institut Pasteur (with identification number CNCM I-3863).E. coli K-12 dnaK mutants usually have a narrow range of permissive temperatures for growth (around 30°C) and exhibit multiple cellular defects, such as impaired cell division and the inhibition of DNA and RNA synthesis (4, 6, 18, 22). Inactivating dnaK in the genetic background of BL21(DE3), an E. coli B strain which is already deficient in OmpT and Lon proteases, did not lead to a dramatic difference in the exponential growth rate at 30°C, but at stationary phase, EN2 cells exhibited slightly reduced ability to form colonies on plates (data not shown). As expected for dnaK mutants, EN2 cells demonstrated impaired growth at 42°C (data not shown). Inactivating dnaK in BL21(DE3) did not induce major morphological defects, and EN2 cells were never found to form long filaments, as dnaK mutants with other genetic backgrounds have previously been reported to do (5) (data not shown). Therefore, the EN2 strain can easily be cultivated at 30°C.We next investigated whether inactivating dnaK in BL21(DE3) would impair the production and solubilities of different recombinant proteins (whose features are summarized in Table S2 in the supplemental material). These proteins belong to organisms of different kingdoms, and their molecular masses range from 19 to 51 kDa; therefore, they potentially correspond to DnaK substrates since the masses of polypeptides interacting with DnaK range from 14 to 90 kDa (28). Many of them exist as oligomers and may require the assistance of DnaK for proper assembly. These proteins were also chosen for their different levels of production and solubility in E. coli. Four of them (CpxP, ClpP1, PA28α, and proteasome-activating nucleotidase [PAN]) are totally soluble, and two of them (ClpP2 and green fluorescent protein [GFP]) are aggregation prone and may require the presence of DnaK to prevent their aggregation. Importantly, all these proteins were contaminated by DnaK when purified from E. coli (see Fig. ​Fig.33).Open in a separate windowFIG. 3.Purification of recombinant proteins in the absence of DnaK. Aliquots of 10 μg of CpxP, ClpP1, ClpP2, GFP, and PAN purified from BL21(DE3) cells (lanes 1, 3, 5, 7, and 9) or EN2 cells (lanes 2, 4, 6, 8, and 10) were loaded onto an SDS-12% PAGE gel. (A) Proteins were revealed by Coomassie blue staining. (B and C) DnaK (B) and GroEL (C) were detected by Western blotting. Sizes of molecular mass markers (lanes MW) are given in kilodaltons and indicated to the left of the gel. The asterisk indicates the position of DnaK on the gel. The two major bands in the purified PAN sample correspond to full-length 50-kDa His-PAN and the 40-kDa PAN fragment resulting from the internal initiation of translation, which copurify as oligomeric complexes (30). The gels shown are representative of results from at least three independent experiments.The production of recombinant proteins in exponentially growing BL21(DE3) and EN2 cells in Luria-Bertani medium at 30°C was induced with 1 mM isopropyl-β-d-thiogalactopyranoside (IPTG) for 2 h. The same biomasses of BL21(DE3) and EN2 cells were sonicated in 1 ml of lysis buffer (50 mM Tris, pH 7.5, 100 mM KCl, 1 mM dithiothreitol). Soluble proteins were separated from aggregated proteins and cellular debris by 30 min of centrifugation at 14,000 × g and 4°C. Pellets containing protein aggregates were resuspended in 1 ml of Tris, pH 7.5, containing 1% sodium dodecyl sulfate (SDS). Total extracts and soluble and insoluble fractions were analyzed by SDS-polyacrylamide gel electrophoresis (PAGE) (Fig. ​(Fig.11).Open in a separate windowFIG. 1.Levels of production and solubility of recombinant proteins in the absence of DnaK. Aliquots of 10 μg of total extracts (T-un and T) and soluble (S) and insoluble (P) fractions from uninduced (T-un) and IPTG-induced (T, S, and P) BL21(DE3) and EN2 cells overexpressing CpxP (A), ClpP1 (B), PA28α (C), ClpP2 (D), GFP (E), or PAN (F) were analyzed by SDS-12% PAGE on gels stained by Coomassie blue. Sizes of molecular mass markers (lanes MW) are given in kilodaltons and indicated to the left of each gel. Arrowheads indicate the positions of recombinant proteins. The gels shown are representative of results from at least three independent experiments.The levels of production of all tested proteins in EN2 and BL21(DE3) cells were similar, as demonstrated by the protein amounts in total extracts (Fig. ​(Fig.1,1, lanes T). Moreover, dnaK inactivation did not affect the solubilities of recombinant proteins, even those such as CpxP (Fig. ​(Fig.1A),1A), ClpP1 (Fig. ​(Fig.1B),1B), and PA28α (Fig. ​(Fig.1C)1C) produced in high amounts or those such as ClpP2 (Fig. ​(Fig.1D)1D) and GFP (Fig. ​(Fig.1E)1E) prone to aggregation. These findings were surprising since the function of the DnaK chaperone is to prevent protein aggregation during synthesis and to cooperate with DnaJ, GrpE, and ClpB in the disaggregation of aggregates. It seems that, even for aggregation-prone recombinant proteins, solubility may not necessarily be dependent on endogenous DnaK. This finding may reflect the different folding requirements of specific proteins. Another explanation may be the presence of another chaperone with an overlapping conjoint function. In fact, a consequence of the absence of DnaK in cells is higher levels of production of heat shock proteins such as GroEL/GroES (29). Consistent with these data, EN2 cells produced higher amounts of GroEL than BL21(DE3) cells (data not shown), and these higher amounts may compensate for the absence of DnaK in preventing protein aggregation, as was shown previously for endogenous E. coli proteins and other recombinant proteins (8, 28). An abundance of different chaperones playing nonspecialized roles in recombinant protein folding in E. coli cells may permit toleration of the loss of DnaK, without impairing cell capacity as a protein production factory.Since the examined proteins could fold and assemble independently of DnaK, we next tested whether a protein known to interact with DnaK could be produced in the absence of this chaperone. Nemo, the IκB kinase complex regulatory component of the NF-κB signaling pathway in eukaryotes, was shown previously to tightly bind and be contaminated by DnaK when produced in E. coli (1). When recombinant His-tagged Nemo was produced in BL21(DE3) under our conditions, it was barely detectable on electrophoresis gel (Fig. 2A and B). However, immunodetection using an anti-His₆ antibody (Roche) at a 1:2,000 dilution showed that the absence of DnaK resulted in an increase in Nemo production (Fig. ​(Fig.2D).2D). When Nemo was produced in higher amounts, most of the protein was found in the soluble fraction, indicating that it could be produced as a soluble species in the absence of DnaK (Fig. ​(Fig.2D,2D, lane 5). Increased production of Nemo in the absence of DnaK could be explained by a role of this chaperone in Nemo degradation. Producing Nemo in a BL21(DE3) strain that is deficient in the protease ClpP did not increase its cellular amount (data not shown), indicating that if Nemo was degraded in a DnaK-dependent manner in BL21(DE3) (which already lacks Lon protease), ClpP was not responsible for this proteolysis or the absence of ClpP was compensated for by another protease.Open in a separate windowFIG. 2.Levels of production and solubility of recombinant Nemo in the absence of DnaK. Aliquots of 10 μg (A and C) or 20 μg (B and D) of total extracts (T) and soluble (S) and insoluble (P) fractions from uninduced and IPTG-induced BL21(DE3) and EN2 cells overexpressing Nemo were loaded onto an SDS-10% PAGE gel. Proteins were detected by Coomassie blue staining (A and C), and His-tagged Nemo was detected by Western blotting (B and D). Sizes of molecular mass markers (lanes MW) are given in kilodaltons and indicated to the left of each gel. The gels shown are representative of results from at least three independent experiments.We next tested whether recombinant proteins produced in the absence of DnaK would remain soluble and active during their purification. Samples of 200 ml of cells overproducing CpxP, ClpP1, ClpP2, or GFP or 500 ml of PAN-overproducing cells were sonicated in 2 ml of lysis buffer (50 mM NaH₂PO₄, pH 8.0, 300 mM NaCl, 10 mM imidazole). The soluble fraction obtained after 30 min of centrifugation at 38,000 × g and 4°C was loaded onto 400 μl of nickel-nitrilotriacetic acid resin, and His-tagged proteins were purified according to the recommendations of the resin manufacturer (Qiagen). After elution, His-tagged proteins were dialyzed against 50 mM Tris, pH 7.5, concentrated, and analyzed by electrophoresis.By this procedure, recombinant proteins were purified to the levels of homogeneity indicated in Fig. ​Fig.3A.3A. Samples of 10 μg of purified proteins were used for the immunodetection of contamination by DnaK (using an anti-DnaK antibody from Stressgen at a 1:2,000 dilution). We found that DnaK in BL21(DE3) cells contaminated all preparations of purified recombinant proteins, albeit to different extents (Fig. ​(Fig.3B,3B, lanes 1, 3, 5, 7, and 9). As expected, dnaK inactivation prevented such contamination (Fig. ​(Fig.3B,3B, lanes 2, 4, 6, 8, and 10). It is noteworthy that most of the proteins purified from BL21(DE3) were also contaminated by GroEL, although this contamination was minor. In EN2 cells, where GroEL expression is increased, we did not systematically observe greater contamination by GroEL (Fig. ​(Fig.3C).3C). Moreover, CpxP and GFP, the proteins that exhibited the greatest DnaK contamination, were not the most contaminated by GroEL, and GroEL did not copurify with PAN in the absence of DnaK. Thus, the absence of DnaK did not necessarily lead to a higher level of contamination by GroEL.Despite the absence of DnaK, all purified recombinant proteins remained soluble even after being concentrated. Since some aggregates are soluble and solubility does not always guarrantee a native active conformation (15, 19), the activities (when readily measurable) or native conformations of some of the purified proteins were examined. One microgram of purified PAN was used to measure ATP hydrolysis at 55°C as described earlier (2). PAN proteins purified from BL21(DE3) and EN2 cells had comparable ATPase activities, with means ± standard errors of 762.33 ± 145.51 and 968.32 ± 198.85 nmol mg⁻¹ h⁻¹ (n = 3), respectively. The fluorescence emission spectrum (at an excitation wavelength of 400 nm) of GFP purified from EN2 cells was indistinguishable from that of GFP purified from BL21(DE3) cells (Fig. ​(Fig.4A),4A), indicating that GFP remained correctly folded when produced in the absence of DnaK. CpxP, a component of the Cpx signal transduction pathway, was the protein that exhibited the greatest DnaK contamination (11). It self-associates into dimers (M. Miot and J.-M. Betton, unpublished data), and to test its correct assembly, 100 μl of purified CpxP at 1 mg/ml in a buffer of 25 mM Tris, pH 7.5, and 150 mM NaCl was loaded onto a size exclusion chromatography column (Superdex 200 HR10/30; GE Healthcare) and eluted with the same buffer at a flow rate of 0.5 ml/min. Recombinant CpxP purified from BL21(DE3) eluted at a volume of 14.98 ml (Fig. ​(Fig.4B),4B), corresponding to a species with an apparent molecular mass of 39.85 kDa (a dimer of Cpx). Recombinant CpxP purified from EN2 eluted at a nearly identical volume of 14.97 ml (Fig. ​(Fig.4B).4B). Thus, the absence of DnaK did not alter CpxP dimeric assembly and did not produce any soluble higher-molecular-mass aggregate species.Open in a separate windowFIG. 4.Folding and assembly of proteins in the absence of DnaK. (A) Fluorescence emission spectra of 8-μg/ml GFP preparations purified from BL21(DE3) and EN2 cells, recorded with an FP-6200 spectrofluorimeter (Jasco) at a scan rate of 250 nm min⁻¹ using a bandwidth of 5 nm for both excitation and emission beams. The spectra shown are representative of results from at least two independent experiments. (B) Size exclusion chromatograms for CpxP proteins purified from BL21(DE3) and EN2 cells. Arrowheads indicate the elution volumes of the standards, and their masses are given in kilodaltons. The chromatograms shown are representative of results from at least three independent experiments.Altogether, these findings indicate that high levels of correctly folded, assembled, and active soluble recombinant proteins can be produced in the absence of endogenous DnaK chaperone in BL21(DE3). Surprisingly, our study showed that the inactivation of dnaK in BL21(DE3), which does not contain Lon, did not result in an increase in the aggregation of recombinant proteins, as was seen previously in E. coli K-12 (24). It seems that in BL21(DE3) cells, and in E. coli B cells in general, factors other than DnaK and Lon may be fundamental in managing the accumulation of aggregated proteins. Through the detailed characterization of a BL21(DE3) ΔdnaK strain and testing of the production of proteins of different natures, origins, and sizes, including aggregation-prone proteins, our study demonstrates that this EN2 strain offers a strategy that can be generally and extensively used to avoid unwanted contamination by DnaK. In addition, since DnaK has ATPase activity, the EN2 strain is particularly well suited for the production and purification of recombinant ATPases, eliminating the undifferentiable ATPase contamination. Given that GroEL, another major chaperone in E. coli, has also been found to contaminate purified recombinant proteins (9), it would be of additional interest to find conditions under which both dnaK and groEL could be eliminated in the BL21(DE3) strain without impairing its survival and its remarkable protein factory capacities.     

Simple,Fast, and Sensitive Method for Quantification of Tellurite in Culture Media

A fast, simple, and reliable chemical method for tellurite quantification is described. The procedure is based on the NaBH₄-mediated reduction of TeO₃²⁻ followed by the spectrophotometric determination of elemental tellurium in solution. The method is highly reproducible, is stable at different pH values, and exhibits linearity over a broad range of tellurite concentrations.The tellurium oxyanion tellurite is toxic for most organisms, making important its accurate assessment. Several methods for quantifying tellurite have been described to date. However, most of them are rather complicated and require sophisticated equipment and in some cases the detection is not quite sensitive enough to allow the assessment of TeO₃²⁻ concentrations below 50 μg/ml (200 μM). For example, the analytical determination of tellurium (Te) oxyanions by atomic absorption spectrometry (AAS) is hampered by poor sensitivity. Where flame or electrothermal AAS routinely yields detection limits of less than 10 ppb for iron (16), normal flame AAS tellurium detection limits are 100 to 1,000 times higher and require pretreatment to achieve the +IV oxidation state before analysis (11).On the other hand, hydride generation AAS (HGAAS) is used to achieve ppb-level detection limits for Se and Te as well as arsenic and antimony among others. For Te the volatile hydride gas H₂Te is generated by first converting the metalloid to the +IV oxidation state and then by chemical reduction to the gaseous hydride using—almost universally—sodium borohydride (NaBH₄). In automated HGAAS systems, an inert purge gas sweeps the volatile hydride formed in a glass reaction vessel into a quartz cell heated by the AAS flame where gaseous hydride decomposition and atomization occur. Though tellurite reduction, precipitation, and detection methods have been reported (3, 17), they are temporally relatively unstable and pH dependent.Since tellurium is toxic and environmentally important (7, 8), determining low concentrations in bacterial cultures is very desirable and a simple analysis without pretreatment steps that could quickly establish total metalloid oxyanion content in a liquid sample would be a plus. Here we report a new method for the determination of tellurite in bacterial culture media. This procedure is based on the NaBH₄ reduction of tellurite to the elemental form, which is analyzed spectrophotometrically at 500 nm or 320 nm (see below), by which the light scattered by the particles of elemental metalloid in solution is measured. While the detection limits do not compare to those of HGAAS (14) or capillary electrophoresis (13), they do approach those of old flame AAS but involve a much simpler and quicker procedure requiring only one reagent and a spectrophotometer to determine total content of solutions of +IV oxyanions in solution. Linear calibration range, method development time and probe stability, effect of sample pH, common interferences, and detection limits were investigated.Calibration curves to determine K₂TeO₃ concentrations in routinely used microbiological culture media such as Luria-Bertani (LB) or M9 minimal medium amended with 0.2% glucose (15) were constructed. A set of solutions containing increasing concentrations of K₂TeO₃ (Sigma) were prepared in LB or M9 culture medium, and the tellurium oxyanion was quantitatively reduced using freshly prepared 3.5 mM NaBH₄ (final concentration).The reaction was carried out at 60°C for 10 min (bubbling was overcome by vortexing), and after 5 min at room temperature, the optical density at 500 nm (OD₅₀₀) was determined spectrophotometrically as described previously (4, 5, 9, 12). Blanks contained no borohydride. Figure ​Figure11 shows that in both media good curve linearity was obtained, with r² values of 0.9740 and 0.9963 for LB and M9, respectively. Tellurite concentrations lower than 1 μg/ml or higher than 200 μg/ml were also tested, but OD₅₀₀ values were close to the spectrophotometer error limit at low concentrations or nonlinear above 200 μg/ml (not shown). Thus, the NaBH₄ method allows determination of a wide range of tellurite concentrations in a fast and simple way. Tellurite concentrations lower than 50 μg/ml in both rich and minimal media can be easily determined; the experimental error was about 10%, similar to that reported for the diethyl dithiocarbamate (DDTC) tellurite method (17).Open in a separate windowFIG. 1.Calibration curves to determine K₂TeO₃ concentrations in LB (A) (R² = 0.9963) or M9 minimal (B) (R² = 0.9740) medium. Optical density at 500 nm was determined after reducing the oxyanion with sodium borohydride. Error bars denote 1 standard deviation of three replicates.To analyze the resulting solutions after tellurite reduction by NaBH₄, absorption/scattering spectra were determined. Figure ​Figure22 shows that spectra from LB and those from M9 after tellurite reduction are quite different, which may be a consequence of the different chemical compositions of these culture media. In both cases, absorption spectra showed linearity between optical density at 500 nm and tellurite concentration in the sample. However, high tellurite concentrations (∼100 μg/ml) caused a loss of linearity in LB medium.Open in a separate windowFIG. 2.Absorption spectra after reducing samples of LB (A) or M9 (B) culture medium containing increasing tellurite concentrations with 3.5 mM NaBH₄. Tellurite concentrations used were 20, 40, 60, 80, and 100 (LB) and 2, 4, 6, 8, and 10 (M9) μg/ml. (Inset) Calibration curve in M9 medium using the absorbance maxima at 320 nm.Figure ​Figure2B2B shows that in M9 medium there is a zone around 320 nm exhibiting higher optical density than that at 500 nm, which represents an advantage in the determination of tellurite in chemically defined culture media. This is reflected in a wider range of measurable concentrations at 320 nm (Fig. ​(Fig.2B,2B, inset), as well as in a higher sensitivity of the method as determined by the slope of the calibration curve. The product of tellurite reduction by NaBH₄ showed good stability at both wavelengths in rich and minimal culture media (not shown).Since in M9 medium the method allows the determination of minor tellurite concentrations (1 to 20 μg/ml), it would be of great help in assessing tellurite uptake in tellurite-sensitive microorganisms whose MICs range from 1 to 10 μg/ml. Sulfur-containing salts, commonly present in culture media as sulfites and sulfates, did not interfere with our NaBH₄ method for tellurite assessment at concentrations up to 0.5 M (not shown).As shown in Fig. ​Fig.3,3, tellurite assessment was not affected by the pH of the culture medium. In fact, linearity was observed in a wide pH range with minor slope changes in LB. Similar results were obtained with M9 medium, although tellurite assessment was not possible at pH values higher than 7.0 because of the formation of a precipitate. This may be due to an interaction of the phosphate salts present in the medium and some charged (2⁺) chemical species forming at alkaline pH values, as has been reported earlier (17).Open in a separate windowFIG. 3.Effect of pH in determining tellurite concentrations in LB (A) and M9 minimal (B) media.To date, the most commonly used procedure for determining tellurite in culture media is that involving the spectrophotometric determination (340 nm) of the complex that forms between tellurite and diethyl dithiocarbamate (17). This procedure has been used to assess tellurite uptake by the phototrophic bacterium Rhodobacter capsulatus, which is naturally resistant to K₂TeO₃ (MIC, ∼1.4 mM) (2, 3). However, K₂TeO₃ uptake studies in highly sensitive cells such as Escherichia coli (MIC, ∼4 μM) are difficult to carry out because of the low concentrations of toxicant present in the culture medium, far below the detection limit of the DDTC procedure (17).In this context and for testing the applicability of our method in vivo, we used the tellurite-sensitive bacterium E. coli BW25113 (10) and the tellurite-resistant Aeromonas caviae ST (5, 6). An overnight culture of E. coli BW25113 in M9 minimal medium was diluted 100-fold with fresh M9 supplemented with 0.2% glucose and grown at 37°C with shaking. When the OD₆₀₀ was 0.1, the culture was amended with 20 μg/ml K₂TeO₃ (arrow, Fig. ​Fig.4A).4A). Then aliquots were taken at the indicated times and cells were centrifuged at 8,500 × g for 3 min; supernatants were used to assess extracellular tellurite by our NaBH₄ method. While added tellurite did not affect bacterial growth (Fig. ​(Fig.4A),4A), the remaining tellurite in the supernatant dropped approximately to one-third after 3 h (Fig. ​(Fig.4B).4B). Tellurite determinations were validated using, in parallel, the DDTC method (not shown).Open in a separate windowFIG. 4.Tellurite uptake by Escherichia coli. Time zero in panel B represents the moment of tellurite addition.Regarding the tellurite-resistant bacterium A. caviae ST, a 1:100 dilution of an overnight culture was inoculated into fresh LB medium and the OD₆₀₀ was recorded at the indicated times. When the OD₆₀₀ was ∼0.4, the culture was amended with tellurite (400 μg/ml final concentration) (Fig. ​(Fig.5A,5A, arrow) and the remaining tellurite in the supernatants was assessed as described above. Figure ​Figure5B5B shows that in 4 h ∼27% of the toxic oxyanion was removed from the culture medium.Open in a separate windowFIG. 5.Tellurite uptake by Aeromonas caviae ST. See the text for details.In summary and in comparison to the DDTC procedure, the NaBH₄ method described here allows more sensitive determination of the initial tellurite concentrations as well as the continuous uptake of the toxicant by tellurite-sensitive and tellurite-resistant microorganisms. This method should be of great help in future studies aimed at unveiling the tellurite reductase activity exhibited by some metabolic enzymes such as nitrate reductase (1), catalase (4), and the pyruvate dehydrogenase complex (5, 6). These studies are currently being carried out in our laboratory.     

Acetolactate Synthase from Bacillus subtilis Serves as a 2-Ketoisovalerate Decarboxylase for Isobutanol Biosynthesis in Escherichia coli

A pathway toward isobutanol production previously constructed in Escherichia coli involves 2-ketoacid decarboxylase (Kdc) from Lactococcus lactis that decarboxylates 2-ketoisovalerate (KIV) to isobutyraldehyde. Here, we showed that a strain lacking Kdc is still capable of producing isobutanol. We found that acetolactate synthase from Bacillus subtilis (AlsS), which originally catalyzes the condensation of two molecules of pyruvate to form 2-acetolactate, is able to catalyze the decarboxylation of KIV like Kdc both in vivo and in vitro. Mutational studies revealed that the replacement of Q487 with amino acids with small side chains (Ala, Ser, and Gly) diminished only the decarboxylase activity but maintained the synthase activity.We have previously shown that 2-keto acids generated from amino acid biosynthesis can serve as precursors for the Ehrlich degradation pathway (15) to higher alcohols (3). In order to produce isobutanol, the valine biosynthesis pathway was used to generate 2-ketoisovalerate (KIV), the precursor to valine, which was then converted to isobutanol via a decarboxylation and reduction step (Fig. ​(Fig.1A).1A). The entire pathway to isobutanol from glucose is shown in Fig. ​Fig.1A.1A. To produce isobutanol, we overexpressed five genes, alsS (Bacillus subtilis), ilvC (Escherichia coli), ilvD (E. coli), kdc (Lactococcus lactis), and ADH2 (Saccharomyces cerevisiae) (Fig. ​(Fig.1A).1A). This E. coli strain produced 6.8 g/liter isobutanol in 24 h (Fig. ​(Fig.1B)1B) and more than 20 g/liter in 112 h (3). More recently, we have found that an alcohol dehydrogenase (Adh) encoded by yqhD on the E. coli genome can convert isobutyraldehyde to isobutanol efficiently (5) (Fig. ​(Fig.1B1B).Open in a separate windowFIG. 1.Schematic representation of the pathway for isobutanol production. (A) The Kdc-dependent synthetic pathway for isobutanol production. (B) Isobutanol production with the Kdc-dependent and -independent synthetic pathways. IlvC, acetohydroxy acid isomeroreductase; IlvD, dihydroxy acid dehydratase. (C) Enzymatic reaction of Als, Ahbs, and Kdc activities.One key reaction in the production of isobutanol is the conversion of KIV to isobutyraldehyde catalyzed by 2-ketoacid decarboxylase (Kdc) (Fig. ​(Fig.1C).1C). Since E. coli does not have Kdc, kdc from L. lactis was overexpressed. Kdc is a nonoxidative thiamine PP_i (TPP)-dependent enzyme and is relatively rare in bacteria, being more frequently found in plants, yeasts, and fungi (8, 19). Several enzymes with Kdc activity have been found, including pyruvate decarboxylase, phenylpyruvate decarboxylase (18), branched-chain Kdc (8, 19), 2-ketoglutarate decarboxylase (10, 17, 20), and indole-3-pyruvate decarboxylase (13).In this work, unexpectedly, we find that Kdc is nonessential for E. coli to produce isobutanol (Fig. ​(Fig.1).1). An E. coli strain overexpressing only alsS (from B. subtilis), ilvC, and ilvD (both from E. coli) is still able to produce isobutanol. Since E. coli is not a natural producer of isobutanol, it cannot be detected from the culture media in any unmodified strain. We identify that AlsS from B. subtilis, which was introduced in E. coli for acetolactate synthesis (Als), catalyzes the decarboxylation of 2-ketoisovalerate like Kdc both in vivo and in vitro. AlsS is part of the acetoin synthesis pathway and catalyzes the aldo condensation of two molecules of pyruvate to 2-acetolactate (Als activity) (Fig. ​(Fig.1C)1C) (11). The overall reaction catalyzed by AlsS is irreversible because of CO₂ evolution. The first step in catalysis is the ionized thiazolium ring of TPP reacting with the first pyruvate, followed by decarboxylation. This intermediate then reacts with the second pyruvate. Deprotonation followed by C-C bond breakage produces 2-acetolactate. In this work, mutational approaches were used to assess the importance of Q487 in the Kdc activity of AlsS.