Effectiveness of SYTOX Green Stain for Bacterial Viability Assessment

The effectiveness of SYTOX Green nucleic acid stain for measuring bacterial viability was tested on starved populations of Escherichia coli and Salmonella typhimurium. This stain underestimates the fraction of dead cells within starved populations containing cells with damaged nucleic acids or membranes. Its application to natural samples should be considered with caution.     

Development of a short-term assay based on the evaluation of the plasma membrane integrity of the alga Pseudokirchneriella subcapitata

Membrane integrity has been used as a criterion for the definition of cell viability. In the present work, staining conditions (time and dye concentration) for the evaluation of membrane integrity in a fluorescence microplate reader, using the membrane-impermeant nucleic-acid dye SYTOX Green, were optimized. Incubating Pseudokirchneriella subcapitata algal cells with 0.5?μmol/l SYTOX Green for 40?min allowed a clear discrimination between live (intact plasma membrane) and dead cells (with compromised plasma membrane). Algal cell suspensions, labelled with SYTOX Green, exhibited a green fluorescence proportional to the fraction of the cells with a permeabilized plasma membrane. The optimized staining conditions were used to assess the toxicity of 1-pentanol on P. subcapitata in a short-term exposure (6?h) assay. The loss of membrane integrity in the cell population increased with the concentration of 1-pentanol. The 6-h EC(10) and EC(50) values were 7,617?mg/l 1-pentanol (95?% confidence limits 4,670-9,327) and 12,818?mg/l 1-pentanol (95?% confidence limits 10,929-15,183), respectively. The developed microplate-based short-term assay can be useful in the high-throughput screening of toxics or environmental samples using the alga P. subcapitata.     

Use of SYTOX green dye in the flow cytometric analysis of bacterial phagocytosis

BACKGROUND: Fluorescein isothiocyanate (FITC) is used widely to label the targets used in flow cytometric phagocytosis assays. Unfortunately, the fluorescence intensity of phagocytosed FITC-labeled targets is influenced by changes in intracellular pH level, making quantitative measurements with this fluorophore problematic. We describe the use of SYTOX green nucleic acid stain to measure phagocytosis by flow cytometry. METHODS: Suspensions of isopropyl alcohol-permeabilized Escherichia coli DH5alpha were stained with the SYTOX green dye and then incubated with resident peritoneal macrophages. The samples were analyzed by flow cytometry and phagocytosis was determined by gating the cells. RESULTS: Results are expressed as percentage of phagocyte-associated green fluorescent cells. The validity of the method was shown by the effects of a phagocytosis inhibitor (incubation at 4 degrees C) or enhancer (gamma interferon [IFN- gamma] treatment) being accurately assessed with this assay. CONCLUSIONS: The method described was reproducible and provides an advantageous alternative to the use of FITC to label bacteria for the flow cytometric measurement of target uptake by phagocytic cells.     

Use of a real-time polymerase chain reaction thermocycler to study bacterial cell permeabilization by antimicrobial peptides

Antimicrobial peptides are good leads to develop new antibiotics, but knowledge of their mode of action is a prerequisite. Destruction of the microbial membranes through a detergent-like mechanism is one of these modes of action. This is usually studied by using a fluorescent nucleic acid stain such as SYTOX Green, which is impermeable to living cells. Using a simple protocol based on the use of a standard real-time thermocycler, we confirmed that the actions of the antimicrobial peptides LL-37 and magainin 2 on bacterial cells are different.     

CELLULAR DNA CONTENT OF MARINE PHYTOPLANKTON USING TWO NEW FLUOROCHROMES: TAXONOMIC AND ECOLOGICAL IMPLICATIONS1

Two new fluorochromes, PicoGreen® and SYTOX Green? stain (Molecular Probes, Inc.), are useful with flow cytometry for quantitative detection of cellular DNA in a variety of marina phytoplankton. The basic instrument configuration of modern low-power flow cytometers (15 mW, 488 nm excitation) is sensitive enough to detect the DNA signal in nearly all of the 121 strains (from 12 taxonomic classes)examined. The major advantages of these dyes over others are 1)suitability for direct use in seawater, 2)green fluorescence emission of the DNA-dye complex (wavelength 525 ± 15 nm) showing no overlap with the autofluorescence of the plankton pigments in the red band, 3) high fluorescence yield of the DNA-dye complex with an increase in fluorescence > 100-fold compared to the unstained cell, and 4)dyes can be used to quantify double-stranded DNA. The high sensitivity allowed the quantification of the DNA of the smallest known phyto-plankter (Prochlorococcus) as well as bacteria found in some of the algal cultures. Of the 12 taxonomic classes tested, only the 3 Nannochloropsis spp. (Eustagmatophyceae) stained poorly, and a few members of the Chlorophyceae and Pelagophyceae showed poor staining occasionally. In general, maximal fluorescence was achieved within 15 min after addition of the dye. Although the PicoGreen dye stained some living phytoplankton species, preservation is recommended for quantitation. SYTOX Green did not stain live cells. The combination of the dyes, therefore, allows the discrimination between live and dead cells in some algal groups (Prochlorococcus, diatoms, prasinophytes, and pelagophytes). Paraformaldehyde was preferred over glutaraldehyde for fixation to avoid (induced) green autofluorescence. Total DNA values measured in 90 algal species (ca. 121 strains) varied by a factor of 20,000. The lowest values were found in Prochlorococcus and the highest in a large dinoflagellate (Prorocentrum micans). DNA content appears to be a scaleable cell component covarying with the carbon and nitrogen contents of the phytoplankton cells. This covariation allows the total DNA content to be used as an accurate, independent estimate of total cell carbon biomass in unicellular pelagic phytoplankton.     

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.     

Fluorescence microplate-based assay for tumor necrosis factor activity using SYTOX Green stain

We have developed a simple, sensitive, fluorescence microplate-based assay for tumor necrosis factor (TNF) biological activity. The assay employs SYTOX Green nucleic acid stain to detect TNF-induced cell necrosis in actinomycin D sensitized cultured cell lines. SYTOX Green stain is a cationic unsymmetrical cyanine dye that is excluded from live cells but can readily penetrate cells with compromised cell membranes. Upon binding to cellular nucleic acids, the dye exhibits a large enhancement in fluorescence, which is monitored at fluorescein wavelengths. We detected 2.5 pg/mL and quantitated 25-500 pg/mL recombinant murine (rm) and recombinant human (rh) TNF-alpha, using mouse fibroblast-derived WEHI 164, WEHI 13var, and L929 cell lines. The procedure can also be used to detect agents that modulate TNF activity. We demonstrated complete inhibition of rhTNF-alpha using monoclonal anti-human TNF-alpha antibody and determined that approximately 20 ng/mL antibody was sufficient to neutralize 50% of the biological activity of 250 pg/mL rhTNF-alpha in these cell lines. Reagents are added in a single step, followed by a 6- to 8-h incubation period, during which the cytokine exhibits its effects. There are no wash steps, and the assay is readily amenable to automation and high-throughput screening procedures.     

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.     

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.     

Nonperturbative Imaging of Nucleoid Morphology in Live Bacterial Cells during an Antimicrobial Peptide Attack

Studies of time-dependent drug and environmental effects on single, live bacterial cells would benefit significantly from a permeable, nonperturbative, long-lived fluorescent stain specific to the nucleoids (chromosomal DNA). The ideal stain would not affect cell growth rate or nucleoid morphology and dynamics, even during laser illumination for hundreds of camera frames. In this study, time-dependent, single-cell fluorescence imaging with laser excitation and a sensitive electron-multiplying charge-coupled-device (EMCCD) camera critically tested the utility of “dead-cell stains” (SYTOX orange and SYTOX green) and “live-cell stains” (DRAQ5 and SYTO 61) and also 4′,6-diamidino-2-phenylindole (DAPI). Surprisingly, the dead-cell stains were nearly ideal for imaging live Escherichia coli, while the live-cell stains and DAPI caused nucleoid expansion and, in some cases, cell permeabilization and the halting of growth. SYTOX orange performed well for both the Gram-negative E. coli and the Gram-positive Bacillus subtilis. In an initial application, we used two-color fluorescence imaging to show that the antimicrobial peptide cecropin A destroyed nucleoid-ribosome segregation over 20 min after permeabilization of the E. coli cytoplasmic membrane, reminiscent of the long-term effects of the drug rifampin. In contrast, the human cathelicidin LL-37, while similar to cecropin A in structure, length, charge, and the ability to permeabilize bacterial membranes, had no observable effect on nucleoid-ribosome segregation. Possible underlying causes are suggested.     

A dual fluorescence flow cytometric analysis of bacterial adherence to mammalian host cells

Flow cytometry has provided a powerful tool for analyzing bacteria-host cell associations. Established approaches have used bacteria, labeled either directly with fluorochromes or indirectly with fluorescently conjugated antibodies, to detect these associations. Although useful, these techniques are consistently unable to include all host cells in the analysis while excluding free, aggregated bacteria. This study describes a new flow cytometry method of assessing bacterial adherence to host cells based on direct fluorescent labeling of both bacteria and host cells. Eukaryotic host cells were labeled with PKH-26, a red fluorescent dye, and bacteria were labeled with fluorescein isothiocyanate, a green fluorescent dye. The red host cells were gated and the mean green fluorescence intensity (MFI) of these red cells was determined. We used MFI values obtained from control samples (unlabeled and labeled host cells with unlabeled bacteria) to eliminate contributions due to autofluorescence. The final MFI values represent fluorescence of host cells resulting from the adherent bacteria. Because all red fluorescent cells are analyzed, this method includes all the eukaryotic cells for analysis but excludes all free or aggregated bacteria that are not bound to target cells.     

Pyronin Y as a fluorescent stain for paraffin sections

Pyronin Y has long been used, in combination with other dyes such as Methyl Green, as a differential stain for nucleic acids in paraffin tissue sections. It also forms fluorescent complexes with double-stranded nucleic acids, especially RNA, enabling semi-quantitative analysis of cellular RNA in flow cytometry. However, the possibility of using pyronin Y as a fluorescent stain for paraffin tissue sections has rarely been investigated. We herein report that in sections stained with Methyl Green–pyronin Y, red blood cells, elastic fibre of blood vessels, zymogen granules of pancreatic acinar cells, surface membrane of heptocytes and kidney tubular cells showed strikingly strong green and/or red fluorescence, while the nuclei of cells appeared non-fluorescent. The use of confocal laser-scanning microscope greatly improved the resolution and selectivity of the fluorescent images. Staining with pyronin Y alone gave similar results in terms of fluorescence properties of the specimens. Pretreatment of paraffin sections with RNase significantly reduced cytoplasmic pyronin Y staining as judged by transmission light microscopy, but it had little effect on the fluorescence intensity of red blood cells, elastic fibres and zymogenbreak granules.     

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.     

Detection of bacterial contamination in starch and resin-based papermaking chemicals using fluorescence techniques

Rapid fluorescence techniques were evaluated for the detection of bacterial contaminants in papermaking chemicals including starch and the resin-based sizes and starch slurries used in the paper industry. Viable and non-viable bacterial cells were visualised by fluorescent probes and detected by epifluorescence microscopy and flow cytometry. The best discrimination ability was obtained with the fluorescent probes LIVE/DEAD and SYBR Green, based on the staining of cellular nucleic acid, and ChemChrome V3, which demonstrated cellular enzymatic activity. The process samples had to be diluted and filtered before fluorescence staining and analysis because they were viscous and contained solid particles. Fluorescence microscopic counts of bacteria in highly contaminated process samples were similar to plate counts, but flow cytometric enumeration of bacterial cells in process samples yielded 2- to 10-fold lower counts compared with plate counts, depending on the consistency of the sample. The detection limits in flow cytometric analysis and in epifluorescence microscopy were 10³–10⁶ cells ml⁻¹ and 10⁵–10⁶ cells ml⁻¹, respectively. Intrinsic bacterial contamination was detectable with fluorescence techniques and highly contaminated process samples could be analysed with fluorescence methods. Electronic Publication     

A Novel Staining Protocol for Multiparameter Assessment of Cell Heterogeneity in Phormidium Populations (Cyanobacteria) Employing Fluorescent Dyes

Bacterial populations display high heterogeneity in viability and physiological activity at the single-cell level, especially under stressful conditions. We demonstrate a novel staining protocol for multiparameter assessment of individual cells in physiologically heterogeneous populations of cyanobacteria. The protocol employs fluorescent probes, i.e., redox dye 5-cyano-2,3-ditolyl tetrazolium chloride, ‘dead cell’ nucleic acid stain SYTOX Green, and DNA-specific fluorochrome 4′,6-diamidino-2-phenylindole, combined with microscopy image analysis. Our method allows simultaneous estimates of cellular respiration activity, membrane and nucleoid integrity, and allows the detection of photosynthetic pigments fluorescence along with morphological observations. The staining protocol has been adjusted for, both, laboratory and natural populations of the genus Phormidium (Oscillatoriales), and tested on 4 field-collected samples and 12 laboratory strains of cyanobacteria. Based on the mentioned cellular functions we suggest classification of cells in cyanobacterial populations into four categories: (i) active and intact; (ii) injured but active; (iii) metabolically inactive but intact; (iv) inactive and injured, or dead.     

TwinGFP, a marker for cell cycle analysis in transiently transfected cells

Green fluorescent protein (GFP) is widely used as a marker to identify transfected cells either by fluorescence microscopy or flow cytometry. However, cell cycle analysis with propidium iodide typically employs ethanol for cell permeabilization. During this treatment, soluble GFPs generally leak out of cells, probably due to their small size. We have now significantly improved cellular retention by creating an in-frame fusion of two GFP DNA sequences, thereby generating a double-sized GFP (TwinGFP, 57 kDa). Permeabilized HeLa cells transfected with pTwinGFP showed a strong green fluorescent signal localized throughout the cells that could easily be detected by fluorescence microscopy and flow cytometry, in contrast to cells transfected with a standard single GFP construct. The experiment indicates that protein size constitutes the major determinant of the loss of fluorescence in permeabilized cells. As a proof of principle, pTwinGFP was cotransfected with the p53 tumor suppressor gene into HeLa cells, and cells transiently expressing p53 could be identified and phenotypically characterized by flow cytometry.     

Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4'',6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.     

Fluorophores for live cell imaging of AGT fusion proteins across the visible spectrum

O6-alkylguanine-DNA alkyltransferase (AGT) fusion proteins can be specifically and covalently labeled with fluorescent O6-benzylguanine (O6-BG) derivatives for multicolor live cell imaging approaches. Here, we characterize several new BG fluorophores suitable for in vivo AGT labeling that display fluorescence emission maxima covering the visible spectrum from 472 to 673 nm, thereby extending the spectral limits set by fluorescent proteins. We show that the photostability of the cell-permeable dyes BG Rhodamine Green (BG505) and CP tetramethylrhodamine (CP-TMR) is in the range of enhanced green fluorescent protein (EGFP) and monomeric red fluorescent protein (mRFP), and that BG diethylaminomethyl coumarin (BGDEAC), a derivative of coumarin, is even more stable than enhanced cyan fluorescent protein (ECFP). Due to the increasing number of new BG derivatives with interesting fluorescence properties, such as far-red emission, fluorescence labeling of AGT fusion proteins is becoming a versatile alternative to existing live cell imaging approaches.     

Effect of moderate electric field frequency and growth stage on the cell membrane permeability of Lactobacillus acidophilus

Changes in growth kinetics and metabolic activity of microorganisms under the presence of a moderate electric field (MEF) have been hypothesized as being due to temporary permeabilization of cell membranes. We investigated herein the effects of frequency and growth stage on cell membrane permeabilization of Lactobacillus acidophilus OSU 133 during MEF fermentation. Cells were stained with two fluorescent nucleic acid stains: the green, nonselective, cell membrane permeable SYTO 9, and the red, cell membrane impermeable propidium iodide (PI). Fluorescence exhibition post‐treatment was assessed using fluorescence microscopy. Total plate counting was done to determine whether or not the permeabilized population represented live cells. Fermentation treatments investigated were conventional (control) and MEF (2 V/cm, 45, 60, 1,000, 10,000 Hz) at 30°C. Studies were conducted at 45 Hz for lag, exponential, and stationary phases of growth. Low frequency MEF treated cells exhibited significantly greater numbers of red cell counts than conventional treatments; further, no significant differences existed in viable counts between MEF and conventional treatments, suggesting that the red counts represent permeabilized live cells. MEF treatments at the early stage of bacterial growth at 45 Hz exhibited the maximum permeabilization followed by treatments at 60 Hz. MEF treated samples at frequencies higher than 60 Hz did not exhibit red fluorescence. Cells at lag phase showed the greatest susceptibility to permeabilization followed by those at exponential phase. No evidence of electroporation was observed during the stationary phase. To our knowledge, these observations provide the first evidence that cell membrane permeabilization occurs under the presence of electric fields as low as those under MEF. © 2009 American Institute of Chemical Engineers Biotechnol. Prog., 2009     

Rapid,in situ detection of Agrobacterium tumefaciens attachment to leaf tissue

Attachment of the plant pathogen Agrobacterium tumefaciens to host plant cells is an early and necessary step in plant transformation and agroinfiltration processes. However, bacterial attachment behavior is not well understood in complex plant tissues. Here we developed an imaging‐based method to observe and quantify A. tumefaciens attached to leaf tissue in situ. Fluorescent labeling of bacteria with nucleic acid, protein, and vital dyes was investigated as a rapid alternative to generating recombinant strains expressing fluorescent proteins. Syto 16 green fluorescent nucleic acid stain was found to yield the greatest signal intensity in stained bacteria without affecting viability or infectivity. Stained bacteria retained the stain and were detectable over 72 h. To demonstrate in situ detection of attached bacteria, confocal fluorescent microscopy was used to image A. tumefaciens in sections of lettuce leaf tissue following vacuum‐infiltration with labeled bacteria. Bacterial signals were associated with plant cell surfaces, suggesting detection of bacteria attached to plant cells. Bacterial attachment to specific leaf tissues was in agreement with known leaf tissue competencies for transformation with Agrobacterium. Levels of bacteria attached to leaf cells were quantified over time post‐infiltration. Signals from stained bacteria were stable over the first 24 h following infiltration but decreased in intensity as bacteria multiplied in planta. Nucleic acid staining of A. tumefaciens followed by confocal microscopy of infected leaf tissue offers a rapid, in situ method for evaluating attachment of A. tumefaciens' to plant expression hosts and a tool to facilitate management of transient expression processes via agroinfiltration. © 2012 American Institute of Chemical Engineers Biotechnol. Prog., 2012