Bacterial assay for the rapid assessment of antifouling and fouling release properties of coatings and materials

An assay has been developed to accurately quantify the growth and release behaviour of bacterial biofilms on several test reference materials and coatings, using the marine bacterium Cobetia marina as a model organism. The assay can be used to investigate the inhibition of bacterial growth and release properties of many surfaces when compared to a reference. The method is based upon the staining of attached bacterial cells with the nucleic acid-binding, green fluorescent SYTO 13 stain. A strong linear correlation exists between the fluorescence of the bacterial suspension measured (RFU) using a plate reader and the total bacterial count measured with epifluorescence microscopy. This relationship allows the fluorescent technique to be used for the quantification of bacterial cells attached to surfaces. As the bacteria proliferate on the surface over a period of time, the relative fluorescence unit (RFU) measured using the plate reader also shows an increase with time. This was observed on all three test surfaces (glass, Epikote and Silastic T2) over a period of 4 h of bacterial growth, followed by a release assay, which was carried out by the application of hydrodynamic shear forces using a custom-made rotary device. Different fixed rotor speeds were tested, and based on the release analysis, 12 knots was used to provide standard shear force. The assay developed was then applied for assessing three different antifouling coatings of different surface roughness. The novel assay allows the rapid and sensitive enumeration of attached bacteria directly on the coated surface. This is the first plate reader assay technique that allows estimation of irreversibly attached bacterial cells directly on the coated surface without their removal from the surface or extraction of a stain into solution.     

Application of an Escherichia coli green fluorescent protein-based lysine biosensor under nonsterile conditions and autofluorescence background

AIMS: To examine the utility of an Escherichia coli green fluorescent protein (GFP) containing biosensor for quantification of bioavailable lysine in selected feed samples under nonsterile conditions and to estimate the background fluorescence of analyzed feed samples and evaluate the risk of confounding GFP emission from the lysine assay organism. METHODS AND RESULTS: Escherichia coli lysine auxotroph GFP based biosensor was used to determine the percentage of bioavailable lysine in two samples of soybean-, cottonseed-, and meat and bone meal under nonsterile conditions. The fluorescence emitted by GFP was successfully measured using a spectrofluorimeter to monitor bacterial growth response to protein-derived lysine and lysine containing small peptides. The autofluorescence of analyzed feed samples at different concentrations could also be estimated. CONCLUSIONS: When feed protein concentrations are decreased, autofluorescence interference can be avoided. SIGNIFICANCE: The E. coli lysine auxotroph GFP-based biosensor can successfully be used for the determination of bioavailable lysine in these selected animal feed proteins under nonsterile conditions. IMPACT OF THE STUDY: E. coli GFP biosensor for lysine has potential for routine application in animal feeds.     

Green fluorescent protein-propidium iodide (GFP-PI) based assay for flow cytometric measurement of bacterial viability.

BACKGROUND: Several staining protocols have been developed for flow cytometric analysis of bacterial viability. One promising method is dual staining with the LIVE/DEAD BacLight bacterial viability kit. In this procedure, cells are treated with two different DNA-binding dyes (SYTO9 and PI), and viability is estimated according to the proportion of bound stain. SYTO9 diffuses through the intact cell membrane and binds cellular DNA, while PI binds DNA of damaged cells only. This dual-staining method allows effective separation between viable and dead cells, which is far more difficult to achieve with single staining. Although SYTO9-PI dual staining is practical for various bacterial viability analyses, the method has a number of disadvantages. Specifically, the passage of SYTO9 through the cell membrane is a slow process, which is significantly accelerated when the integrity of the cell membrane is disrupted. As a result, SYTO9 binding to DNA is considerably enhanced. PI competes for binding sites with SYTO9 and may displace the bound dye. These properties diminish the reliability of the LIVE/DEAD viability kit. In this study, we investigate an alternative method for measuring bacterial viability using a combination of green fluorescent protein (GFP) and PI, with a view to improving data reliability. METHODS: Recombinant Escherichia coli cells with a plasmid containing the gene for jellyfish GFP were stained with PI, and green and red fluorescence were measured by FCM. For comparison, cells containing the plasmid from which gfp was removed were stained with SYTO9 and PI, and analyzed by FCM. Viability was estimated according to the proportion of green and red fluorescence. In addition, bioluminescence and plate counting (other methods to assess viability) were used as reference procedures. RESULTS: SYTO9-PI dual staining of bacterial cells revealed three different cell populations: living, compromised, and dead cells. These cell populations were more distinct when the GFP-PI combination was used instead of dual staining. No differences in sensitivity were observed between the two methods. However, substitution of SYTO9 with GFP accelerated the procedure. Bioluminescence and plate counting results were in agreement with flow cytometric viability data. CONCLUSIONS: In bacterial viability analyses, the GFP-PI combination provided better distinction between current viability stages of E. coli cells than SYTO9-PI dual staining. Additionally, the overall procedure was more rapid. No marked differences in sensitivity were observed.     

Development of a whole cell green fluorescent sensor for lysine quantification

As an essential amino acid, lysine is an important component of animal and human diets and its bioavailability can depend on a variety of factors. Therefore, an accurate pre-determination of bioavailable lysine in foods and feeds is important. In this study a whole cell fluorescent biosensor for the quantification of lysine in protein sources was constructed. A gene encoding for green fluorescent protein (GFPmut3) was introduced into an E. coli lysine auxotroph genome as a part of a mini-Tn5-Km transposon. The location of the transposon was determined and the growth kinetics of the newly constructed biosensor were examined. The transposon disrupted the ybhM gene, which encodes for the synthesis of a protein with an unknown function. No effect of the transposon’s location in the genome or the expression of gfp on bacterial growth rates was observed. Based on the fluorescence emitted by GFPmut3, a standard curve after 6-h growth of the strain was generated. A correlation coefficient of 0.95 was observed when the fluorescence method was compared to the conventional optical density (OD) growth-based lysine assay. Using the newly developed lysine fluorescent whole cell sensor we determined the total lysine in casein acid hydrolyzate (7.13 ± 0.34%). When lysine added to 12 μg/ml and 30 μg/ml of casein acid hydrolyzate was quantified, recoveries of 97 ± 1.65% and 103 ± 4.66% respectively were detected. The results suggest that the microbial assay using GFP fluorescence represents a promising alternative method for the potential estimation of lysine in protein sources.     

Involvement of RNA and DNA in the staining of Escherichia coli by SYTO 13

AIMS: To assess the extent to which DNA and RNA bacterial content contributes to fluorescent response of SYTO 13. METHODS AND RESULTS: RNA and DNA of Escherichia coli 536 cells were extracted and fluorimetrically quantified to compare the different contents, throughout a 24 h culture, with their SYTO 13 fluorescence emission when analysed by the cytometer. SYTO 13 fluorescence varied depending on the stage of bacterial growth and in accordance with both DNA and RNA content. RNA content accounted for at least two-thirds of the total fluorescence of a cell. Escherichia coli cells were treated with chloramphenicol to improve their RNA content. With this treatment, both nucleic acids remained constant but there was a clear improvement in fluorescent emission. SYTO 13 fluorescence was also studied in E. coli X-1488 minicells. CONCLUSIONS: Although both nucleic acids are implicated, RNA accounts for a major part of SYTO 13 fluorescence. The fluorescence cannot be considered as a direct reflection of nucleic acid content. Other factors, such as topology or supercoiling, need to be considered. SIGNIFICANCE AND IMPACT OF THE STUDY: The results confirm the efficacy of SYTO 13 for labelling bacteria and for assessing the distinct physiological status. A better knowledge of the parameters implicated in its fluorescence emission has been achieved.     

VBNC细菌荧光观察技术问题分析及对策

以SYTO-9和PI两种荧光染料对细菌进行核染,以市售的黑色钢笔墨水替代伊拉克黑,利用手动压片方式对标本进行固定,自建一种"活的非可培养状态"(VBNC)细菌荧光显微镜观察技术。通过总结分析该项技术方法在应用过程中所出现的问题,如存在背景荧光、菌体聚集、荧光图像模糊、滤膜吸附能力差、菌体形态不鲜明、菌体荧光减淡等,并提出相应的解决策略,旨在给予相关研究者以帮助和启迪。     

Fluorescent method for monitoring cheese starter permeabilization and lysis

A fluorescence method to monitor lysis of cheese starter bacteria using dual staining with the LIVE/DEAD BacLight bacterial viability kit is described. This kit combines membrane-permeant green fluorescent nucleic acid dye SYTO 9 and membrane-impermeant red fluorescent nucleic acid dye propidium iodide (PI), staining damaged membrane cells fluorescent red and intact cells fluorescent green. For evaluation of the fluorescence method, cells of Lactococcus lactis MG1363 were incubated under different conditions and subsequently labeled with SYTO 9 and PI and analyzed by flow cytometry and epifluorescence microscopy. Lysis was induced by treatment with cell wall-hydrolyzing enzyme mutanolysin. Cheese conditions were mimicked by incubating cells in a buffer with high protein, potassium, and magnesium, which stabilizes the cells. Under nonstabilizing conditions a high concentration of mutanolysin caused complete disruption of the cells. This resulted in a decrease in the total number of cells and release of cytoplasmic enzyme lactate dehydrogenase. In the stabilizing buffer, mutanolysin caused membrane damage as well but the cells disintegrated at a much lower rate. Stabilizing buffer supported permeabilized cells, as indicated by a high number of PI-labeled cells. In addition, permeable cells did not release intracellular aminopeptidase N, but increased enzyme activity was observed with the externally added and nonpermeable peptide substrate lysyl-p-nitroanilide. Finally, with these stains and confocal scanning laser microscopy the permeabilization of starter cells in cheese could be analyzed.     

Fluorescent Method for Monitoring Cheese Starter Permeabilization and Lysis

A fluorescence method to monitor lysis of cheese starter bacteria using dual staining with the LIVE/DEAD BacLight bacterial viability kit is described. This kit combines membrane-permeant green fluorescent nucleic acid dye SYTO 9 and membrane-impermeant red fluorescent nucleic acid dye propidium iodide (PI), staining damaged membrane cells fluorescent red and intact cells fluorescent green. For evaluation of the fluorescence method, cells of Lactococcus lactis MG1363 were incubated under different conditions and subsequently labeled with SYTO 9 and PI and analyzed by flow cytometry and epifluorescence microscopy. Lysis was induced by treatment with cell wall-hydrolyzing enzyme mutanolysin. Cheese conditions were mimicked by incubating cells in a buffer with high protein, potassium, and magnesium, which stabilizes the cells. Under nonstabilizing conditions a high concentration of mutanolysin caused complete disruption of the cells. This resulted in a decrease in the total number of cells and release of cytoplasmic enzyme lactate dehydrogenase. In the stabilizing buffer, mutanolysin caused membrane damage as well but the cells disintegrated at a much lower rate. Stabilizing buffer supported permeabilized cells, as indicated by a high number of PI-labeled cells. In addition, permeable cells did not release intracellular aminopeptidase N, but increased enzyme activity was observed with the externally added and nonpermeable peptide substrate lysyl-p-nitroanilide. Finally, with these stains and confocal scanning laser microscopy the permeabilization of starter cells in cheese could be analyzed.     

A Fluorescent Gram Stain for Flow Cytometry and Epifluorescence Microscopy

The fluorescent nucleic acid binding dyes hexidium iodide (HI) and SYTO 13 were used in combination as a Gram stain for unfixed organisms in suspension. HI penetrated gram-positive but not gram-negative organisms, whereas SYTO 13 penetrated both. When the dyes were used together, gram-negative organisms were rendered green fluorescent by SYTO 13; conversely, gram-positive organisms were rendered red-orange fluorescent by HI, which simultaneously quenched SYTO 13 green fluorescence. The technique correctly predicted the Gram status of 45 strains of clinically relevant organisms, including several known to be gram variable. In addition, representative strains of gram-positive anaerobic organisms, normally decolorized during the traditional Gram stain procedure, were classified correctly by this method.Gram’s staining method is considered fundamental in bacterial taxonomy. The outcome of the Gram reaction reflects major differences in the chemical composition and ultrastructure of bacterial cell walls. The Gram stain involves staining a heat-fixed smear of cells with a rosaniline dye such as crystal or methyl violet in the presence of iodine, with subsequent exposure to alcohol or acetone. Organisms that are decolorized by the alcohol or acetone are designated gram negative.Alternative Gram staining techniques have recently been proposed. Sizemore et al. (19) reported on the use of fluorescently labeled wheat germ agglutinin. This lectin binds specifically to N-acetylglucosamine in the peptidoglycan layer of gram-positive bacteria, whereas gram-negative organisms contain an outer membrane that prevents lectin binding. Although simpler and faster than the traditional Gram stain, this method requires heat fixation of organisms.Other Gram stain techniques suitable for live bacteria in suspension have been described. Allman et al. (1) demonstrated that rhodamine 123 (a lipophilic cationic dye) rendered gram-positive bacteria fluorescent, but its uptake by gram-negative organisms was poor. This reduced uptake by gram-negative bacteria was attributed to their outer membranes. The outer membrane can be made more permeable to lipophilic cations by exposure to the chelator EDTA (4). Shapiro (18) took advantage of this fact to form the basis of another Gram stain, one which involved comparing the uptake of a carbocyanine dye before and after permeabilizing organisms with EDTA. All of these methods, however, rely on one-color fluorescence, making analysis of mixed bacterial populations difficult.An alternative to the use of stains is the potassium hydroxide (KOH) test. The method categorizes organisms on the basis of differences in KOH solubility. After exposure to KOH, gram-negative bacteria are more easily disrupted than gram-positive organisms. This technique has been used to classify both aerobic and facultatively anaerobic bacteria, including gram-variable organisms (8). In a study by Halebian et al. (9), however, this technique incorrectly classified several anaerobic strains, giving rise to the recommendation that the method should only be used in conjunction with the traditional Gram stain.In this study we demonstrate a Gram staining technique for unfixed organisms in suspension, by using clinically relevant bacterial strains and organisms notorious for their gram variability. The method uses two fluorescent nucleic acid binding dyes, hexidium iodide (HI) and SYTO 13. Sales literature (11) published by the manufacturers of HI (Molecular Probes, Inc., Eugene, Oreg.), which displays a red fluorescence, suggests that the dye selectively stains gram-positive bacteria. SYTO 13 is one of a group of cell-permeating nucleic acid stains and fluoresces green (11). These dyes have been found to stain DNA and RNA in live or dead eukaryotic cells (16). Both dyes are excited at 490 nm, permitting their use in fluorescence instruments equipped with the most commonly available light sources. We reasoned that a combination of these two dyes applied to mixed bacterial populations would result in all bacteria being labeled, with differential labeling of gram-positive bacteria (HI and SYTO 13) and gram-negative bacteria (SYTO 13 only). The different fluorescence emission wavelengths of the two dyes would ensure differentiation of gram-positive from gram-negative bacteria by either epifluorescence microscopy or flow cytometry when equipped with the appropriate excitation and emission filters. While a commercial Gram stain kit produced by Molecular Probes includes HI and an alternative SYTO dye, SYTO 9, we are unaware of any peer-reviewed publications regarding either its use or its effectiveness with traditionally gram-variable organisms.     

Rapid test for distinguishing membrane-active antibacterial agents

In the search for antibacterial agents with a novel mode-of-action (MOA) many targeted cellular and cell-free assays are developed and used to screen chemical and natural product libraries. Frequently, hits identified by the primary screens include compounds with nonspecific activities that can affect the integrity and function of bacterial membrane. For a rapid dereplication of membrane-active compounds, a simple method was established using a commercially available Live/Dead(R) Bacterial Viability Kit. This method utilized two fluorescent nucleic acid stains, SYTO9 (stains all cells green) and propidium iodide (stains cells with damaged membrane red) for the drug-treated bacterial cells. The cells were then either examined visually by fluorescence microscopy or their fluorescence emissions were recorded using a multi-label plate reader set to measure emissions at two different wavelengths. The ratio of green versus red was compared to a standard curve indicating the percentage of live versus dead bacteria. Nine known antibiotics and 14 lead compounds from various antibacterial screens were tested with results consistent with their MOA.     

AGITATION DURING INCUBATION REDUCES THE TIME REQUIRED FOR A LYSINE MICROBIOLOGICAL GROWTH ASSAY USING AN ESCHERICHIA COLI AUXOTROPHIC MUTANT

Microbiological assays involving Escherichia coli lysine auxotrophs must be optimized to facilitate routine use. Our objectives in this study were to characterize growth of an auxotrophic E. coli lysine mutant (American Type Culture Collection strain #23812) and examine the effect of agitation on E. coli mutant growth. A defined minimal salts basal medium was used and supplemented with various lysine concentrations. The E. coli lysine auxotroph responded to increasing lysine concentration with increasing optical density. When maximum optical density (MOD) was determined for the auxotroph, a linear increase was obtained as lysine concentrations were increased (R²± 0.96) for both agitation and static cultures. Growth rates were not significantly (p > 0.05) affected by lysine concentrations, cultural conditions or their combined effect. However, growth with agitation significantly (p < 0.05) reduced the assay time by shortening the lag phase and causing stationary phase to occur earlier. The values of R² (± 0.96) relatively remained constant over the range while the bacterial population were in the stationary phase. In conclusion, the lysine growth assay using the E. coli lysine auxotroph can be made more rapid by agitating the culture during incubation.     

SYTO16 labelling and flow cytometry of Mycobacterium avium

P. Ibrahim, A.S. Whiteley and M.R. Barer. 1997. Mycobacterium avium cells were harvested from agar at different stages of their growth cycle, exposed to the minimum inhibitory concentration of isoniazid (INH) for 24 h and labelled with the fluorescent nucleic acid stain SYTO16. INH exposure led to a > 10-fold increase in the intensity of labelling in the majority of cells, and revealed discrete fluorescence peaks that were consistent with development of filamentous multinucleate cells during the growth cycle. Similar enhancement of labelling was observed in unfixed INH-treated cells viewed by fluorescence microscopy. INH appears to increase the permeability of Myco. avium cells to SYTO16. A combination of growth cycle-defined inocula, labelling with the new generation of fluorescent dyes and flow cytometry provides new opportunities to study the interrelationships between growth cycle events and antimicrobial susceptibility of mycobacteria.     

In Situ Activity of Suspended and Immobilized Microbial Communities as Measured by Fluorescence Lifetime Imaging

In this study, the feasibility of fluorescence lifetime imaging (FLIM) for measurement of RNA:DNA ratios in microorganisms was assessed. The fluorescence lifetime of a nucleic acid-specific probe (SYTO 13) was used to directly measure the RNA:DNA ratio inside living bacterial cells. In vitro, SYTO 13 showed shorter fluorescence lifetimes in DNA solutions than in RNA solutions. Growth experiments with bacterial monocultures were performed in liquid media. The results demonstrated the suitability of SYTO 13 for measuring the growth-phase-dependent RNA:DNA ratio in Escherichia coli cells. The fluorescence lifetime of SYTO 13 reflected the known changes of the RNA:DNA ratio in microbial cells during different growth phases. As a result, the growth rate of E. coli cells strongly correlated with the fluorescence lifetime. Finally, the fluorescence lifetimes of SYTO 13 in slow- and fast-growing biofilms were compared. For this purpose, biofilms developed from activated sludge were grown as autotrophic and heterotrophic communities. The FLIM data clearly showed a longer fluorescence lifetime for the fast-growing heterotrophic biofilms and a shorter fluorescence lifetime for the slow-growing autotrophic biofilms. Furthermore, starved biofilms showed shorter lifetimes than biofilms supplied with glucose, indicating a lower RNA:DNA ratio in starved biofilms. It is suggested that FLIM in combination with SYTO 13 represents a useful tool for the in situ differentiation of active and inactive bacteria. The technique does not require radioactive chemicals and may be applied to a broad range of sample types, including suspended and immobilized microorganisms.     

Quantification of total and bioavailable lysine in feed protein sources by a whole-cell green fluorescent protein growth-based <Emphasis Type="Italic">Escherichia coli</Emphasis> biosensor

Using a fluorescent whole-cell Escherichia coli biosensor previously developed in our laboratory, we determined total and bioavailable lysine in four feed ingredients (soybean, cottonseed, meat and bone meal, and sorghum) and three complete feeds (chick starter and finisher, and swine starter). The same feed sources were analyzed for total lysine by high performance liquid chromatography (HPLC) and bioavailable lysine by chick bioassay. No significant differences were found between bioavailable lysine estimates for soybean, cottonseed, meat and bone meal, chick starter and finisher, and swine starter obtained by the fluorescent E. coli biosensor and chick bioassay. Except for sorghum, the E. coli biosensor estimates for total lysine were highly comparable to those obtained by HPLC. Comparisons were also conducted between conventionally performed optical density-based and the newly developed fluorescence-based lysine assay. The lack of significant differences in data obtained for total and bioavailable lysine by both detection modes indicated reliance and accuracy of the fluorescent E. coli biosensor. Overall results suggest that the microbial assay based on green fluorescent protein fluorescence represents a promising alternative method for lysine quantification.     

LIVE/DEAD BacLight : application of a new rapid staining method for direct enumeration of viable and total bacteria in drinking water.

A rapid epifluorescence staining method using the LIVE/DEAD Bacterial Viability Kit (BacLight) was applied to estimate both viable and total counts of bacteria in drinking water. BacLight is composed of two nucleic acid-binding stains: SYTO 9 and propidium iodide. SYTO 9 penetrates all bacterial membranes and stains the cells green, while propidium iodide only penetrates cells with damaged membranes, and the combination of the two stains produces red fluorescing cells. Optimal incubation conditions were found to be 15 to 20 min, at room temperature in the dark. Total (red + green) and viable (green) cells can hence be counted simultaneously. Factors affecting the staining procedure were tested (addition of glutaraldehyde, staining time, chlorine impact). In the absence of stress, BacLight viable counts were comparable and to 5-cyano-2,3-ditolyl tetrazolium (CTC) counts. BacLight total counts were comparable to acridine orange counts (differing by <0.1 log/ml). However, the increase in environmental stresses (chlorine, growth rate or temperature) induced a decrease in viability that was more pronounced for CTC and plate counts than for BacLight viable counts.     

Mechanism and use of the commercially available viability stain, BacLight.

BACKGROUND: BacLight (Molecular Probes, Eugene, OR, USA) is a popular fluorescence-based two-component stain for determining bacterial cell viability. The main purpose of this work was to fully elucidate the mechanism and to determine why it is sometimes reported that cells stain simultaneously live and dead. METHODS: Solutions of DNA were stained with the two components, propidium iodide (PI) and SYTO9, in different combinations, and fluorescence spectra were collected. RESULTS: K(PI) and K(SYTO9) were approximately 3.7 x 10(5)/M and 1.8 x 10(5)/M. SYTO9 emissions were stronger and overlapped those of PI. Fluorescence resonance energy transfer from SYTO9 to PI was observed. It was, even under normal conditions, possible for DNA bound SYTO9 to have a component in the red region equal to that of DNA bound PI. Potentially confusing emissions were also found to occur when PI was not in sufficient excess to saturate nucleic acid (>0.4 M PI to 1 M DNA base pairs). CONCLUSIONS: The mechanism is a combination of displacement of SYTO9 by PI and quenching of SYTO9 emissions by fluorescence resonance energy transfer. Confusing results can occur if the relative intensities of the stains or the concentration of PI relative to nucleic acid are not properly accounted for.     

Rapid screening method for Mycobactericidal activity of chemical germicides that uses Mycobacterium terrae expressing a green fluorescent protein gene

The slow growth of mycobacteria in conventional culture methods impedes the testing of chemicals for mycobactericidal activity. An assay based on expression of the green fluorescent protein (GFP) by mycobacteria was developed as a rapid alternative. Plasmid pBEN, containing the gene encoding a red-shifted, high-intensity GFP mutant, was incorporated into Mycobacterium terrae (ATCC 15755), and GFP expression was observed by epifluorescence microscopy. Mycobactericidal activity was assessed by separately exposing a suspension of M. terrae(pBEN) to several dilutions of test germicides based on 7.5% hydrogen peroxide, 2.4% alkaline glutaraldehyde, 10% acid glutaraldehyde, and 15.5% of a phenolic agent for contact times ranging from 10 to 20 min (22 degrees C), followed by culture of the exposed cells in broth (Middlebrook 7H9) and measurement of fluorescence every 24 h. When the fluorescence was to be compared with CFU, the samples were plated on Middlebrook 7H11 agar and incubated for 4 weeks. No increase in fluorescence or CFU occurred in cultures in which the cells had been inactivated by the germicide concentrations tested. Where the test bacterium was exposed to ineffective levels of the germicides, fluorescence increased after a lag period of 1 to 7 days, corresponding to the level of bacterial inactivation. In untreated controls, fluorescence increased rapidly to reach a peak in 2 to 4 days. A good Pearson correlation coefficient (r > or =0.85) was observed between the intensity of fluorescence and the number of CFU. The GFP-based fluorescence assay reduced the turnaround time in the screening of chemical germicides for mycobactericidal activity to < or =7 days.     

GFP-aided confocal laser scanning microscopy can monitor Agrobacterium tumefaciens cell morphology and gene expression associated with infection

We tagged Agrobacterium tumefaciens cells with a mini-Tn5 transposon containing a promoterless gene encoding a green fluorescent protein (GFP). Some of the GFP-tagged individual bacterial cells exhibited strong green fluorescence, which reflected the expression levels of the GFP-tagged genes. Those cells could be readily detected with a confocal laser scanning microscope (CLSM). We observed that the fluorescence and morphology of A. tumefaciens cells grown in plant tissues resembled those grown in a minimal medium of low pH, which is required for expression of the virulence genes responsible for tumorigenesis. This suggests that GFP-aided CLSM can be used to determine which growth medium is more representative of the nutritional conditions that a pathogen encounters in plant tissues. We also observed that the fluorescence and morphology of A. tumefaciens cells changed dramatically during the course of infection. Our data suggested that A. tumefaciens cells were probably better fed upon successful colonization. We believe that GFP-aided CLSM can help study the fate of A. tumefaciens cells inside plant tissues by monitoring cell morphology and gene expression associated with the infection process in situ.     

Microplate-based assays for the evaluation of antibacterial effects of photocatalytic coatings

Three microplate-based viability assays for assessing the antibacterial effects of photocatalytic coatings were compared to the conventional colony count method. In the experimental design, cultured Escherichia coli were exposed to photocatalysis on various TiO₂ films in the presence of either UVA or visible light. The photocatalytic effects on the bacterial physiology were determined by real-time measurements of metabolic activity (XTT assay), biomass formation in the liquid medium (growth assay), and by assessing membrane integrity (with propidium iodide and SYTO 9 fluorescent nucleic acid binding dyes—BacLight assay). All three methods proved to be more sensitive and reproducible than colony count for the evaluation of the bactericidal effect of photocatalysis, XTT, and growth assay succeeded in detecting differences in both UVA and visible light-activated photocatalytic coatings. BacLight could efficiently detect the visible light-dependent photocatalytic effect on bacteria and identify membrane damage, but resulted inadequate for evaluating the UVA-dependent antibacterial effects. The described microplate-based evaluation methods proved being more effective and rapid than the colony count assay for assessing the antibacterial effect of various photocatalytic coatings.