Microbial analysis at the single-cell level: tasks and techniques

Journal of Microbiological Methods 42 (2000) 3–16

Journal of Microbiological Methods www.elsevier.com / locate / jmicmeth

Microbial analysis at the single-cell level: tasks and techniques Howard M. Shapiro* Howard M. Shapiro, M.D., P.C., 283 Highland Ave., West Newton, MA 02465 -2513, USA Accepted 28 April 2000

Abstract The heterogeneity of microorganisms themselves is orders of magnitude greater than the heterogeneity of perspectives from which they are contemplated by human observers. Even closely related species may exhibit marked differences in biochemistry and behavior, and, under many conditions, similar, striking heterogeneity may exist within a clonal population of organisms which, in the aggregate, occupy too small a region of space to be visible to the unaided human eye. Using methods of microscopy, microspectrophotometry, and cytometry developed and refined since the 1960s, it is now possible to characterize the physiology and pharmacology of individual microorganisms, and, in many cases, to isolate organisms with selected characteristics for culture and / or further analysis. These methods include fluorescent and confocal microscopy, scanning and image cytometry, and flow cytometry. Fluorescence measurements are particularly important in single-cell analysis; they allow demonstration and quantification of cells’ nucleic acid content and sequence, of the presence of specific antigens, and of physiologic characteristics such as enzyme activity and membrane potential. Multiparameter cytometry, combined with cell sorting, provides insight into population heterogeneity and allows selected cells to be separated for further analysis and culture. The technology is applicable to a wide range of problems in contemporary microbiology, including strain selection and the development of antimicrobial agents.  2000 Elsevier Science B.V. All rights reserved. Keywords: Bacteria; Flow cytometry; Fluorescent dyes; Membrane potential

1. Introduction Different people see microorganisms from different perspectives. To evolutionary and molecular biologists, microbes are relatives, with whom we set up correspondence. To biotechnologists, they are workers, to be employed and, perhaps, exploited. To environmental microbiologists, they may be merely scenery, or analogous to canaries in coal mines, but they are generally viewed as good neighbors if we

*Tel.: 1 1-617-965-6044 or 1 1-617-783-8392; fax: 1 1-617244-7110. E-mail address: [email protected] (H.M. Shapiro).

have good fences. To clinical, food, and sanitary microbiologists, and to the defense establishment, microorganisms are enemies to be tracked, contained, and killed, and to leaders of rogue states and terrorist organizations, they are useful tools which are much easier to get through airports than are firearms and explosives. That said, it is clear that the heterogeneity of microorganisms themselves is orders of magnitude greater than the heterogeneity of perspectives from which they are contemplated by human observers. Even closely related species may exhibit marked differences in biochemistry and behavior, and, under many conditions, similar, striking heterogeneity may exist within a clonal population of organisms which, in the aggregate, occupy

0167-7012 / 00 / $ – see front matter  2000 Elsevier Science B.V. All rights reserved. PII: S0167-7012( 00 )00167-6

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too small a region of space to be visible to the unaided human eye. Without microscopy, humans would likely be unaware of the existence of the microbial world. However, from van Leeuwenhoek’s time until the late 20th Century, it was relatively difficult to study microorganisms at the single-cell level. Even in modern laboratories, most analyses of growth and metabolism are carried out using bulk, macroscopic measurements of populations, which limit the degree to which microbial behavior can be understood. Using methods of microscopy, microspectrophotometry, and cytometry developed and refined since the 1960s, it is now possible to characterize the physiology and pharmacology of individual microorganisms, and, in many cases, to isolate organisms with selected characteristics for culture and / or further analysis. This paper will present a relatively brief, updated discussion of some tasks in microbiology amenable to single-cell analysis, and the applicable methods, measurements, and reagents. There is neither room nor need for an extensive treatment of the subject material here; for further details, the reader is referred to the author’s book (Shapiro, 1995) and a comprehensive, indispensable review by Davey and Kell (1996), as well as to the papers which follow in this issue of The Journal of Microbiological Methods.

2. Tasks in microbiology — heterogeneity makes them harder Microbiologists analyze specimens for a variety of reasons. In the simplest cases, it is necessary only that microorganisms be detected in a sample; at the next level of complexity, the organisms must be counted, explicitly or by the use of some surrogate indicator of the number present. Beyond this, an organism in pure culture may need to be identified and / or characterized as to its growth, metabolism, viability, and, in clinical situations, interaction with antimicrobial agents. Finally, it may be necessary to detect, identify, count and characterize each of several organisms in a mixed population. Even the simplest task, that of detection of microorganisms, may demand examination of several aspects of the specimen. An observer using a micro-

scope will consider the size, shape, and motility, or lack thereof, of particles within the field of view, in order to decide whether they are or are not microorganisms; the job becomes more difficult as the sample contains increasing amounts of inorganic or organic particulates in the microbial size range. In this instance, it may be necessary to use stains to demonstrate the presence of nucleic acid and / or enzyme activity or other chemical evidence of metabolism in order to discriminate organisms from other particles. The same considerations apply when instrumental methods of single-cell analysis are used for examination of a specimen as when the microscope is the only apparatus available. An electronic cell counter, set to detect and count particles in the size range from below 1 to several mm, can successfully identify all patients with urinary tract infections (where infection is defined as the presence of 10 5 or more CFU / ml urine), but cannot discriminate between urine samples from infected patients and samples from uninfected individuals which contain large numbers of particles. While the electronic counter is ineffective for screening, an instrument that measures both particle size and enzyme activity or nucleic acid content, doing what is referred to in cytometry as multiparameter analysis, can be substantially more effective. Additional challenges are presented by the heterogeneity of microbial populations. Even in pure cultures derived from a single organism, different cells may exhibit chemical differences, by virtue of being in different phases of the reproductive cycle or in different physiologic states due to differences in nutrient availability or other environmental conditions. Also, mutation rates are such that a colony containing more than a few hundred thousand cells almost certainly contains genetically different individuals. Single-cell analysis provides the best, and, in some cases, the only means of characterizing heterogeneity within populations.

3. Characterization of cells — parameters and probes In the jargon of single-cell analysis, cellular characteristics, such as size, nucleic acid content, and membrane potential, are usually referred to as pa-

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rameters, a term also used for the physical characteristics, such as absorption, light scattering, and fluorescence intensity measured by instruments such as flow cytometers. The intended meaning can usually be inferred from context. With respect to cellular characteristics, an intrinsic parameter is one that can be measured without the use of reagents; an extrinsic parameter requires the use of a reagent, or probe, for its measurement. Cellular parameters are also characterized as structural or functional; size, DNA content, and the presence and copy number of an antigen or nucleic acid sequence are structural parameters, while internal pH, membrane potential, and enzyme activity are functional parameters. The distinction between structural and functional parameters may blur at its edges, but the concept has been generally useful. Electronic (Coulter) counters, sensitive to the change in electrical impedance resulting from the passage of a particle through a saline-filled orifice, have been used to count bacteria (Kubitschek, 1958) and even viruses (De Blois et al., 1974). However, most instruments used for single-cell analysis of microorganisms make optical measurements. While light absorption is the easiest physical parameter to measure in eukaryotic cells, and absorption spectra, particularly in the infrared region, may provide specific information about chemical composition, absorption measurements of individual bacteria, which are near the resolution limit of optical microscopes, are difficult, while measurements of individual virions, which are below the resolution limit, are impossible. Orsini et al. (2000) (this issue) describe the use of Fourier transform infrared (FTIR) microspectrophotometry to characterize differences in glucose metabolism in different areas of Candida albicans microcolonies, but their measurements include contributions from substantial numbers of cells. Light scattered by, and fluorescence emitted from, particles below the resolution limit of optical microscopes can be seen by a human observer, or measured by an electronic detector, such as a photomultiplier tube (PMT), photodiode, or charge-coupled device (CCD). In the 1920s, a device called the ultramicroscope, with dark-field illumination, made it possible to observe individual virus particles based on their light scattering. In the 1940s, what is generally recognized as the first flow cytometer, built

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by Gucker et al. (1947), detected light scattering from bacteria as small as 0.6 mm in aerosols. More recently, Hercher et al. (1979) used a specially designed flow cytometer to measure light scattering from individual virions of bacteriophage T2 and other viruses. Hara et al. (1991) and Hennes and Suttle (1995), using epifluorescence microscopy, observed and counted single viruses stained with nucleic acid-binding dyes, and Marie et al. (1999) detected fluorescence signals from similarly stained marine viruses using a standard commercial flow cytometer. Raman scattering, which reflects an interaction between electronic and vibrational energy states, can be detected from small regions of space, and can therefore provide information about the infrared absorption spectra of single microorganisms which could not be obtained directly using FT-IR microscopy. Raman microscopy was used by Schuster et al. (2000) (this issue) to characterize metabolic heterogeneity among organisms in cultures of Clostridium acetobutylicum. For particles above the resolution limit for optical microscopy, i.e., those with dimensions exceeding about one-half the wavelength of the incident light, the intensity of light scattered at small angles (0.5– 108) to the incident illuminating beam (a physical parameter referred to as small angle or forward scatter) is strongly dependent on the size of the object, but also strongly dependent on the refractive index. The size dependence of forward scatter is not completely monotonic; thus, an instrument may produce larger scatter signals from 5 mm plastic particles than from 6 mm particles with the same refractive index. It is, in general, inadvisable to compare sizes of particles with different refractive indices, although forward scatter measurements can be, and have been, calibrated for determination of biomass of bacteria, even in mixed populations (Robertson et al., 1998). In many flow cytometers, forward scatter signals from bacteria are near or below the noise level, and any mismatch of refractive index between the sheath fluid and the fluid in which cells are suspended adds noise to the measurement; it is therefore desirable to correct such index mismatches in order to maximize the chance of obtaining usable signals. The intensity of light scattered at large angles (158 and above) to the illuminating beam (a physical

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parameter referred to as large angle, wide-angle, 908, or orthogonal scatter), while size dependent, also depends strongly on the surface roughness and / or internal granularity of the particles in question. In some cases, large angle light scattering signals provide a good indication of total protein content (Steen and Boye, 1980). Allman et al. (1990) demonstrated changes in orthogonal scatter signals with protein and storage granule content in Azotobacter vinelandii, and correlated these qualitatively with ultrastructural changes observed by electron microscopy. However, some structural changes, such as vacuole formation, can produce anomalous effects on both orthogonal and forward scatter signals (Dubelaar et al., 1987). Both scatter and fluorescence signals are sensitive to the polarization of the illumination source and to the geometry of the optical measurement system used. The importance of polarization in flow cytometry is just beginning to be appreciated (Asbury et al., 2000), and may be more significant for cytometry of microorganisms than for cytometry of eukaryotes, since the cell walls of many microorganisms are birefringent. Although some problems remain to be solved, including those relating to polarization effects, quantitative cytometric fluorescence measurements are closer to being standardized than are measurements of light scattering. An entire issue of the journal Cytometry (Vol. 33, No. 2, October 1998) was devoted to this topic (Lenkei et al., 1998). Fluorescence measurements and fluorescent probes are widely used in static and flow cytometry and in confocal microscopy. Fluorescent probes allow measurement of the widest variety of extrinsic cellular parameters, and the fluorescence of phycobiliproteins, which, when extracted from algae, are used as fluorescent labels for extrinsic probe measurements, is also an intrinsic cellular characteristic of cyanobacteria, and helps to classify phytoplankton populations (Trask et al., 1982).

4. Fluorescence — excited states and ground rules In order for an atom or molecule to emit fluorescence, it must first absorb light at a wavelength

shorter than or equal to the wavelength of the emitted light, raising an electron to a higher energy level, known as an excited state. The process of absorption requires only about a femtosecond. Fluorescence occurs when the electron loses all or some of the absorbed energy by light emission. The period between absorption and emission is known as the fluorescence lifetime; this is typically on the order of a few nanoseconds for fluorescent organic compounds, but is notably longer (hundreds of microseconds) for some materials, e.g. lanthanide chelates. In almost all cases, some of the excitation energy is lost nonradiatively, by transitions between different vibrational energy levels of the electronic excited state; this requires that the emitted energy be less than the energy absorbed, meaning that the fluorescence emission will be at a longer wavelength than the excitation. The difference between the absorption and emission wavelengths (usually defined as the difference between the principal absorption and emission maxima in the fluorescence spectrum) is known as the Stokes’ shift, in honor of George Stokes, who first described fluorescence in the mid-1800s. Typical Stokes’ shifts are no more than a few tens of nanometers. Despite the use of the term wavelength in the preceding paragraph, fluorescence is an intrinsically quantum mechanical process; the absorbed and emitted energy are in the form of photons. The likelihood that a molecule will absorb is quantified in its absorption cross-section and extinction coefficient. The quantum yield and quantum efficiency of fluorescence are, respectively, the number and percentage of photons emitted per photon absorbed; they typically increase with the cross-section and extinction coefficient, but are also dependent on the relative likelihoods of the excited molecule losing energy via fluorescence emission and nonradiative mechanisms. The quantum yields of some dyes used in cytometry are quite high, above 0.5, but it should be noted that quantum yield, particularly for an organic material, is affected by the chemical environment in which the molecule finds itself. If an excited molecule that might otherwise fluoresce instead loses energy nonradiatively, for example by collision with solvent molecules, it is said to be quenched; once returned to the electronic ground state, it can be reexcited. However, there is usually a finite prob-

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ability that light absorption will be followed by a change in molecular structure, making further cycles of fluorescence excitation and emission impossible; this is called (photo)bleaching. In principle, increasing the illumination intensity can increase the intensity of light scattering signals without limit. However, this is not even theoretically possible for fluorescence signals, because, at some level of illumination, all the available molecules will be in excited states, leaving no more to be excited if illumination intensity is further increased. This condition of photon saturation is often reached in cytometers which use laser powers of 100 mW or more; bleaching, which may also make the dependence of emission intensity on excitation intensity less than linear, is noticeable at power levels of tens of milliwatts. Saturation and bleaching are discussed by van den Engh and Farmer (1992). In recent years, the technique of multiphoton excitation (Denk and Svoboda, 1997; Masters et al., 1999) has come into vogue for studies of intrinsic fluorescence and fluorescent probes in living cells and tissues. A fluorescent molecule which would normally require a single photon of a given energy level for fluorescence excitation can also be excited by two photons with half the requisite energy level (e.g., two red photons instead of one ultraviolet (UV) photon), three with one third that energy level, and so on, provided the multiple photons arrive almost simultaneously. This condition can be achieved using brief, very high power pulses of laser light. Even so, the requisite intensity is typically achieved in only a very small volume of the subject under study. This is beneficial for two reasons; first, illumination and collection of light from a thin layer of the specimen provides better spatial localization of cellular structures, because there is less interfering fluorescence from above and below the focal plane of the optical system. Second, there is very little bleaching of fluorescent material, and little damage to cellular structures, in the areas above and below the region being observed, whereas these are problems with conventional fluorescence excitation. At present, a principal limitation of multiphoton excitation techniques is their requirement for very expensive lasers. When an excited molecule is in close proximity (typically no more than a few nanometers) to another molecule, particularly when the absorption spectrum

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of the second molecule overlaps the emission spectrum of the first, nonradiative energy transfer (fluorescence resonance energy transfer, or FRET) from the excited (donor) molecule to the nearby acceptor molecule may occur, followed by fluorescence emission from the acceptor in its emission region. In the intact photosynthetic apparatus of algae and cyanobacteria, absorbed light in the blue-green and green spectral regions is made usable for photosynthesis by a series of intra- and intermolecular energy transfers via phycobiliproteins to chlorophyll, without subsequent emission. Once extracted from phycobilisomes, individual phycobiliproteins, such as phycoerythrin, behave like fluorescent molecules with large Stokes’ shifts; the acceptor moieties in the phycobiliprotein continue to quench the donor molecules, which initially absorb the light, but emit fluorescence instead of transferring energy nonradiatively to a different phycobiliprotein with appropriate spectral characteristics. Phycobiliproteins themselves are useful for fluorescent labeling of otherwise nonfluorescent reagents such as antibodies and lectins; it is now also common practice to attempt to improve on nature by conjugating dyes to phycobiliproteins to add an additional phase of energy transfer and further shift the emission spectrum. For example, phycoerythrin (PE) absorbs blue-green and green light and emits yellow light (emission maximum about 575 nm); a so-called tandem conjugate, PE–Cy5, which is phycoerythrin to which the covalent fluorescent dye Cy5 (which absorbs in the yellow and orange spectral regions) has been conjugated, emits red light (emission maximum about 675 nm). It is also possible to use the FRET phenomenon to estimate distances between different molecules in or on cells, using a donor dye (which might be attached to an antibody) to label one structure and an acceptor to label the other. An accessible review of FRET is provided by ¨ 0 si et al. (1998). Szollo

5. Cytometric technology A variety of cytometric techniques can be employed to study microorganisms using fluorescent probes. Confocal microscopy, with or without multiphoton excitation, provides the highest resolution

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images, and is capable of the most precise spatial localization of fluorescence, therefore yielding the most information about cell structure. Lower resolution techniques, such as conventional fluorescence microscopy, scanning laser cytometry (Kamentsky et al., 1997; Darzynkiewicz et al., 1999), and volumetric capillary cytometry (Dietz et al., 1996) are also applicable; they provide relatively minimal information about structure but, like confocal microscopy, they allow repeated observation of a particular cell or cells over a period of time. Conventional flow cytometry (Shapiro, 1995) is capable of analysis of tens of thousands of cells per second, but permits only one observation of each cell, although flow cytometers with sorting capabilities can separate cells of interest for further analysis. Newly developed microfluidic flow cytometers (Fu et al., 1999) may combine the best features of flow and static cytometers; while their analysis rate is typically no more than a few hundred cells per second, they can incorporate not only sorting capability, but the ability to add and mix reagents, and to observe individual cells over time. The slow (down to zero, and reversible) flow rates used with the microfluidic devices also make it possible to collect much weaker fluorescence signals than can be measured in conventional flow cytometers, permitting analysis of DNA fragments stained with appropriate dyes (Chou et al., 1999) and, potentially, measurement of fluorescent antibodies and nucleic acid stains bound to viruses, which could be sorted. Microfluidic cytometers use closed fluidic systems for sorting, an advantage in working with pathogens. Steen (2000) (this issue) has discussed the past, present, and future of flow cytometric technology for microbial analysis. Cytometric fluorescence measurements require a relatively intense light source, which may be an arc lamp or a laser. Mercury arc lamps have strong spectral lines in the UV (366 nm), violet (405 nm), blue (436 nm), green (546 nm) and yellow (577 nm) regions. The lasers most commonly used in commercial apparatus are air-cooled argon ion lasers emitting 10–25 mW of blue-green light at 488 nm; some low-power argon lasers can also emit green light at 515 nm and lower power levels at several other wavelengths between 457 nm (blue-violet) and 515 nm. Higher-power air- and water-cooled argon lasers can provide tens to hundreds of milliwatts of UV

(351 and 363 nm) and blue-violet (457 nm) light as well as the blue and green lines. Krypton ion lasers emit over a very broad spectral range, with higherpower, water-cooled systems producing UV (350 and 356 nm) and violet (406–422 nm) light as well as blue (three lines near 470 nm), green (520 and 530 nm), yellow (568 nm), red (647 and 676 nm), and infrared (IR) (752 and 799 nm) light. Air-cooled argon ion lasers containing a mixture of argon and krypton can emit at any of the argon or krypton wavelengths from the blue to the far red; they are often used in confocal microscopes. Air-cooled helium–cadmium (He–Cd) lasers can produce tens of milliwatts of UV (325 and 355 nm) and blue-violet (442 nm) light, while air-cooled helium–neon (He–Ne) lasers emit as much as 50 mW in the red region at 633 nm, and can also be made to emit green (543 nm), yellow (594 nm), and orange (611 nm) light at lower power levels. When power levels of 10 mW and below are sufficient, red (635 nm) diode lasers may replace red He–Ne lasers; diode lasers, which are small, inexpensive, and extremely energy-efficient, are also available with far red (675–720 nm) and IR (750–900 nm) emission. Diode lasers emitting approximately 5 mW near 400 nm have recently become available; these can be used to excite some dyes that are typically used with UV illumination (Shapiro, 2000). Other solid-state lasers are also becoming useful in cytometry. Frequency-doubled, diode-pumped yttrium aluminum garnet (YAG) lasers, emitting at 532 nm, are competitive in cost with 488 nm argon ion lasers, but are more energy-efficient, and have lower optical noise, which is a desirable feature for improving light scattering measurement sensitivity; they are also more efficient than 488 nm lasers in exciting phycoerythrin and its tandem conjugates, providing increased fluorescence measurement sensitivity. Doubled Alexandrite lasers have produced UV output, and a solid-state 488 nm laser, which may provide a more compact and energy-efficient replacement for the argon ion laser, has recently been described (Gitin et al., 2000). The power of cytometric apparatus has been increased, as the cost has been reduced, by improvements in electronics and computers. Instruments commonly incorporate personal computers for data acquisition and analysis, and a shift from analog to

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digital signal processing electronics (Shapiro et al., 1998) should further improve performance. Data analysis techniques used in cytometry are beyond the scope of the present review, but are discussed by Shapiro (1995) and Davey and Kell (1996).

6. Microbial cytometry — probes and problems The most valuable single reference on fluorescent probes for microscopy, cytometry, and related analyses is the Handbook of Fluorescent Probes and

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Research Chemicals (Haugland, 1996), the catalog of Molecular Probes, Inc. (Eugene, OR, USA); the current (7th) edition of this is, at present, only available on CD-ROM and on the World Wide Web (http: / / www.probes.com). Table 1 presents a brief summary of microbial parameters measurable by cytometry and the probes that can, where necessary, be used for their measurement. Most cytometric instrumentation was developed for analysis of eukaryotic, predominantly mammalian, cells, and what is known about the behavior of many of the reagents in now common use was

Table 1 Microbial parameters measurable by flow cytometry Parameter Intrinsic structural parameters (no probe) Cell size Cytoplasmic granularity, vacuoles, etc. Birefringence Pigment content (e.g., photosynthetic pigments) Intrinsic functional parameter (no probe) Redox state Extrinsic structural parameters DNA content DNA base ratio

Measurement method and probe if used Small angle light scattering, electronic impedance (DC) Large angle light scattering, electronic impedance (AC) Polarized light scattering Fluorescence Fluorescence (endogenous pyridine and flavin nucleotides)

Antigens Lipids Surface sugars (lectin binding sites)

Fluorescence (e.g., DAPI, Hoechst dyes, mithramycin) Fluorescence (A–T and G–C preference dyes; e.g., Hoechst 33342 and chromomycin A 3 ) Fluorescence (labeled oligonucleotides) Fluorescence (e.g., pyronin Y (with DNA blocked)) Fluorescence (ethidium, propidium, asymmetric cyanines) Fluorescence (covalent or ionic bonded acid dyes), large angle light scattering Fluorescence (labeled antibodies) Fluorescence (e.g., Nile red) Fluorescence (labeled lectins)

Extrinsic functional parameters Surface receptors Surface charge Membrane fusion / turnover Membrane integrity (‘viability’) Membrane permeability (dye / drug / substrate uptake / efflux) Intracellular receptors Enzyme activity Oxidative metabolism Sulfhydryl groups / glutathione DNA synthesis Membrane potential Cytoplasmic [Ca 21 ] Intracellular pH

Fluorescence (labeled ligands) Fluorescence (labeled polyionic molecules) Fluorescence (tracking dyes) Fluorescence (e.g., propidium, fluorescein esters) Fluorescence (e.g., Hoechst 33342, rhodamine 123, cyanines, anthracyclines, labeled substrate analogs) Fluorescence (labeled ligands) Fluorescence (fluorogenic substrates) Fluorescence (e.g., CTC) Fluorescence (e.g., monochlorobimane) Fluorescence (anti-BrUdR antibodies; dye mixtures; SBIP) Fluorescence (e.g., cyanines, oxonols, rhodamine 123) Fluorescence ratio (e.g., indo-1) Fluorescence ratio (e.g., BCECF, SNAFL, SNARF)

Nucleic acid sequence RNA content (double-stranded) Total double-stranded nucleic acid Total protein

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learned largely from observation of the interaction of those reagents with those cells. The most important lesson learned to date in the history of microbial cytometry is that bacteria are not just little eukaryotes. The uptake of, and efflux of, dyes, drugs, and other reagents by and from bacterial cells are affected by the structure of the cell wall, and by the presence of pores and pumps which may or may not be analogous to those found in mammalian cells. Thus, while we can say with some confidence whether or not a particular dye behaves as a vital stain in mammalian cells (i.e., enters without fixation of the cell or permeabilization of the cytoplasmic membrane), and, if not, what chemical treatments must be applied in order for the cells to be stained, it is dangerous to conclude that even well-characterized dyes behave in bacteria as they do in mammalian cells. The outer membrane of Gram-negative bacteria excludes most lipophilic or hydrophobic molecules, among which are included drugs such as the tetracyclines and reagents such as cyanine dyes, used for estimation of membrane potential (see Shapiro, 1995); while chemical agents such as ethylene diamine tetraacetic acid (EDTA) may be used to permeabilize the outer membrane to drugs and dyes with at least transient retention of some metabolic function, the characteristics of the permeabilized bacteria are distinct from those of organisms in the native state. While Gram-positive organisms may take up a somewhat wider range of reagents without additional chemical treatment, the results of Walberg et al. (1998), cited by Steen (2000) (this issue), show substantial variability in patterns of uptake of nucleic acid binding dyes by three species of Gram-positive bacteria. Some cytometric techniques require that cells be made permeable to macromolecules, as when intracellular structures are stained with fluorescent antibodies, or when RNAse is added to remove interference by double-stranded RNA with stoichiometric staining of DNA by dyes such as ethidium and propidium. Methodology for permeabilizing mammalian cells, however well-tested, is unlikely to be reliable across the range of microbial species. Although it is relatively simple to distinguish single microbial cells from cell aggregates under the microscope, or in static cytometric apparatus with

relatively high resolution, it is more difficult to do so in flow cytometers. The techniques used in flow cytometry to identify eukaryotic cell aggregates rely either on measurement of pulse shape or on demonstration of hyperdiploid DNA content; the small size of microbes and the relatively large variance of light scattering and DNA content distributions make it difficult to apply either approach. This can interfere with accurate estimation of parameters such as membrane potential and membrane permeability, where one would ideally like to determine the concentration, rather than the total amount, of a dye present in cells. My colleagues and I (Novo et al., 1999, 2000) have recently described techniques for accurate measurement of membrane potential and permeability which use the ratio of two size-dependent fluorescence measurements to eliminate most of the effects of size variation from the parameter measurement; there are similar ratiometric methods for measurement of pH and cytoplasmic calcium ion concentration (see Haugland, 1996). The size of microorganisms, which are typically two to three orders of magnitude smaller than eukaryotic cells, is itself an impediment to some cytometric fluorescence measurements. Although typical instruments may reliably detect fewer than 1000 molecules of fluorescent material in or on a cell, measurements at such low levels typically exhibit large variances due to photoelectron statistics; thus, some microbial constituents may be undetectable or measurable with only limited precision.

6.1. Nucleic acid staining A large number of fluorescent dyes can be used to stain DNA and / or RNA. However, relatively few of them are specific for DNA, and most of these are sensitive to base composition (A–T / G–C ratio). DAPI (49,6-diamidino-2-phenylindole) and the Hoechst bis-benzimidazole dyes 33258 and 33342 increase fluorescence approximately 100 times when bound to A–T triplets in DNA. These dyes are UV-excited, with emission maxima in the blue spectral region between 450 and 500 nm. Chromomycin A 3 and the closely related mithramycin exhibit increased fluorescence on binding to G–C pairs in DNA; they are excited by violet or

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blue-violet light and emit in the green between 525 and 550 nm. The combination of Hoechst 33258 and chromomycin A 3 has been used with dual excitationbeam flow cytometers to discriminate the majority of human chromosomes based on differences in DNA base composition, and to demonstrate differences in base composition among bacterial species (see Shapiro, 1995, pp. 262–264). Dyes such as ethidium bromide (EB) and propidium iodide (PI) increase fluorescence on binding to double-stranded nucleic acid, whether DNA or RNA, and the same property is shared by a large number of asymmetric cyanine nucleic acid stains (e.g., the TO-PRO- and TOTO-series, SYTO-series, Pico Green, etc.) introduced by Molecular Probes. These dyes can be used to stain total nucleic acid in microorganisms; specific staining of DNA would require RNAse treatment. Since a typical metabolically active bacterium contains about five times as much double-stranded ribosomal RNA as DNA, this may be problematic. Relatively specific staining of double-stranded (predominantly ribosomal) RNA in mammalian cells can be achieved using a combination of pyronin Y (excitation by blue-green or green light; emission in the yellow around 575 nm), which stains RNA, with one of the Hoechst dyes or with methyl green, which bind to DNA and prevent DNA staining by pyronin Y (see Shapiro, 1995, pp. 268–270). In a dual excitation-beam instrument, DNA and RNA content can be estimated simultaneously from pyronin Y and Hoechst dye fluorescence. Pyronin Y has been used for RNA content estimation in microorganisms.

6.2. Fluorescent labels and their uses Until both flow cytometers and monoclonal antibodies became widely available in the early 1980s, the most widely used fluorescent label was fluorescein, usually conjugated to proteins as the isothiocyanate (FITC); second labels were not often needed. Fluorescein is nearly optimally excited at 488 nm, the argon ion laser wavelength commonly used in flow cytometers and confocal microscopes, and emits in the green near 525 nm. While rhodamine dyes had been used for two-color immunofluorescence analysis by microscopy, they

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were not suitable for 488 nm excitation. A small number of studies were done with yellow-excited dyes, which needed a second excitation beam, making flow cytometers substantially more expensive. Then, Oi et al. (1982) reported that algal phycobiliproteins could be used as highly efficient fluorescent labels with large Stokes’ shifts. Phycoerythrin (PE), which absorbs well at 488 nm and is maximally excited by green light, emits in the yellow near 575 nm; tandem conjugates of PE with other dyes emit at longer wavelengths (PE–Texas red, near 610 nm; PE–Cy5, near 675 nm; PE–Cy5.5, near 700 nm; PE–Cy7, near 770 nm). The phycobiliprotein allophycocyanin (APC) absorbs maximally in the red near 650 nm, and emits near 660 nm; it is well excited by red diode and He–Ne lasers. Tandem conjugates of APC with Cy5.5 and Cy7 emit in the far red and near infrared, as do the PE conjugates with the same dyes. A principal disadvantage of phycobiliproteins as fluorescent labels is their large size; with a molecular weight near 240,000, PE binding increases the molecular weight of an immunoglobulin G antibody by about 150%. This may not be an issue when labeled antibodies or lectins are used to stain cell surface structures, but becomes one when it is necessary to use labeled reagents to demonstrate intracellular constituents. A number of lower molecular weight labels have been developed for this purpose, including symmetric cyanines (Mujumdar et al., 1993) and the Alexa series (see the Molecular Probes handbook on CD-ROM). Natarajan and Srienc (2000) (this issue) describe the use of a glucose analog with a low molecular weight fluorescent label for flow cytometric analysis of glucose uptake in single cells of E. coli. Low molecular weight labels are also typically used with oligonucleotide probes for demonstration and quantification of specific nucleic acid sequences in cells. In microorganisms, 16S ribosomal RNA probes are particularly useful, as the several thousand ribosomes normally present per cell provide built-in amplification; these probes are also useful for classification of mixed populations, and particularly so when many of the species in a sample are not culturable. The use of labeled nucleic acid probes has been reviewed by Porter and Pickup (2000) (this issue).

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6.3. Microdroplets, growth, and multiplex analysis Weaver et al. (1984) described a technique for encapsulating individual microorganisms or eukaryotic cells in small droplets of a material such as agarose gel, which can facilitate several types of cytometric analyses. When cells in suspension culture divide, it is impossible to keep track of their lineage; when cells in gel microdroplets divide, the daughter cells form a microcolony within the droplet. Since the gel matrix passes nutrients and low molecular weight dyes, cells in droplets can be cultured for a period of time, then stained for nucleic acid and analyzed, at which point an estimate of cell numbers or colony sizes and, therefore, of growth can be obtained. Products secreted by cells may be retained in an appropriately formulated gel, and then quantified by flow cytometry. Katsuragi et al. (2000) (this issue) demonstrate an advantage of this approach. They subjected microcolonies in gel microdroplets to a relatively destructive analytical technique to determine thiamin production, but were able to recover and culture high-producing strains from the small fraction of surviving cells in the sorted microdroplets. Cell growth can also be determined with the use of tracking dyes, which bind tightly to membranes, by incorporation in the lipid bilayer, or covalently to intracellular proteins. If cells are treated with a tracking dye and then washed, the amount of dye per cell will be diluted by half with each successive cell division; this allows relatively precise estimation of the distribution of generational age in a cell population. Nebe-von-Caron et al. (2000) (this issue) show an example. Flow cytometers have been used for chemical analysis. In a so-called sandwich assay, an antibody against the analyte of interest is bound to plastic beads, to which the sample is then added. Thereafter, with a wash step optionally interspersed, a fluorescent antibody to the same analyte is added and the fluorescence of the beads is measured in a flow cytometer. This can provide a highly sensitive assay, since the cytometer can detect a few thousand molecules of analyte bound to a bead. However, the technique becomes most useful when several assays are run in a single tube, using beads that can be discriminated from one another on the basis of their

size, color, or both; this is referred to as multiplex analysis (Fulton et al., 1997). An instrument recently developed by Luminex Corporation (Austin, TX, USA) can perform as many as 100 assays simultaneously, using different proportions of two redexcited fluorescent dyes to color-code beads, and PE-labeled antibodies, nucleotides, or other ligands, excited by a 532 nm YAG laser, to quantify analytes. Multiplex analysis may be usable in combination with tracking dyes or the gel microdroplet technique to facilitate flow cytometric assays that would otherwise be infeasible because of the requirement to analyze a large number of aliquots of sample. Antibiotic susceptibility testing provides an example. It is typically necessary to monitor growth (or death) of a pathogen in the presence of one or two dosage levels of 10 or more antibiotics; since it is difficult to run a single sample in a flow cytometer in less than 30 s, it would take at least 10 min to analyze the 20 aliquots included in a single test, giving a relatively low throughput level. However, one could incorporate a different dye mixture into the microdroplets used with each antibiotic and dose level, and monitor colony growth in gel microdroplets, using a single nucleic acid stain; the microdroplets could then be mixed, allowing the assay to be read by running only a single sample through the cytometer. If sufficient precision were obtainable, it might be possible to apply multiplexing at the single-cell level, e.g. by using a tracking dye to follow cell growth, and covalent labels or tightly bound nucleic acid dyes in various color combinations to identify cells cultured with different antibiotics.

6.4. Fluorescent proteins: mild-mannered reporters During the 1980s several approaches were taken to demonstrating the introduction of genes into prokaryotic and eukaryotic cells. If the gene for a protein or enzyme foreign to the transfected cells were included in the vector, transfection could be inferred when the protein or enzyme could be detected on or in the target cells using a fluorescent antibody or a fluorescent enzyme substrate. A substantial advance was made by Chalfie et al. (1994), who found that the gene for the green fluorescent protein from the jellyfish Aequorea could be used as a reporter without the need for additional reagents.

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Although the native green fluorescent protein is not well suited for excitation by the 488 nm lasers commonly used in flow cytometers, mutants have been developed (Cubitt et al., 1995) which are better and brighter, and which now permit detection of several reporter proteins by virtue of differences in emission spectra. Papers by Gordon et al. (2000), De Wulf et al. (2000), and Porro et al. (2000) (all in this issue) deal with various aspects and applications of reporter gene technology.

6.5. Approaches to determination of cell viability: dye exclusion and membrane potential Antibiotic susceptibility testing as performed at the bulk level typically relies on determination of the viability of microorganisms as reflected by growth in numbers. It is possible to quantify the number of microorganisms in a sample using a cytometer if the volume of the analyzed sample is known. This condition is satisfied in the simplest way when a counting chamber is used with a microscope or scanning cytometer, or when a constant volume pump is used to deliver sample to a flow cytometer. It is also possible to calculate the volume of sample analyzed in a flow cytometer relatively simply by adding readily identifiable plastic beads at a known concentration to the sample. Counts of microorganisms, however, may or may not correlate with viable counts obtained by plating. The colony forming unit (CFU) may be a single organism, or an aggregate containing one or more viable cells with or without associated nonviable cells. For a variety of reasons, both microbiologists and cytometrists would like to be able to characterize microorganisms as viable at the single-cell level. A number of criteria for viability have been suggested; impermeability of the membrane to dyes such as propidium is one, and the presence of metabolic activity, as indicated by the production and retention of fluorescent product from a nonfluorescent enzyme substrate or by maintenance of a membrane potential, is another. Nebe-von-Caron et al. (2000) (this issue) have dealt at length with viability, applying cell sorting to determine which cells (or aggregates) in a multiparameter measurement space will grow when sorted onto a solid medium.

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Propidium (usually available as the iodide (PI)) and ethidium (usually available as the bromide (EB)) are structurally similar dyes containing a phenanthridinium ring; both bind, with fluorescence enhancement, to double-stranded nucleic acids. However, ethidium has only a single positive charge; its Nalkyl group is an ethyl group. Ethidium and other dyes with a single delocalized positive charge are membrane-permeant; i.e., they cross the intact cytoplasmic membranes of both prokaryotic and eukaryotic cells, although the dyes may be pumped out by efflux pumps. Propidium has a propyl group with a quaternary ammonium as its N-alkyl group, and thus bears a double positive charge. It, and a number of dyes which also bear quaternary ammonium groups and more than one positive charge (e.g., TO-PRO-l, TO-PRO-3, and Sytox Green, all from Molecular Probes), are generally believed to be membraneimpermeant; i.e., they are excluded by prokaryotic and eukaryotic cells with intact cytoplasmic membranes. Cells which take up propidium and other multiply charged dyes are usually considered to be nonviable, although transient permeability to these dyes can be induced by certain chemical and physical treatments, e.g. electroporation, with subsequent recovery of membrane integrity and viability. Thus, staining (or the lack thereof) with propidium is the basis of a so-called dye exclusion test of viability. Acid dyes, such as trypan blue and eosin, are also membrane-impermeant and are used in dye exclusion tests. A variation on the dye exclusion test employs a nonfluorescent, membrane-permeant substrate for an intracellular enzyme, which crosses intact or damaged cell membranes and which is then enzymatically cleaved to form a fluorescent, impermeant (or slowly permeant) product. The product is retained in cells with intact membranes, and quickly lost from putatively nonviable cells with damaged membranes. One commonly used substrate is diacetylfluorescein, also called fluorescein diacetate (FDA), which yields the slowly permeant fluorescein; nonfluorescent esters of some fluorescein derivatives are better for dye exclusion tests because their products are less permeant (see Haugland, 1996). Another substrate is 5-cyano-2,3-ditolyl tetrazolium chloride (CTC) (see Davey and Kell, 1996), which, being reduced by various dehydrogenases to a fluorescent formazan

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product, provides an indication of the activity of the respiratory chain as well as of membrane integrity. Bacteria normally maintain an electrical potential gradient of over 100 mV across the cytoplasmic membrane, with the interior negative with respect to the exterior. Charged dyes that are sufficiently lipophilic to pass readily through the lipid bilayer portion of the membrane will partition across the membrane in response to the potential gradient. Positively charged lipophilic dyes, such as rhodamine 123 and the cyanines, will be concentrated inside cells that maintain membrane potential, while negatively charged lipophilic dyes, such as the oxonols, will be excluded. Thus, if two cells of the same volume, one with a transmembrane potential gradient and one without, were equilibrated with a cyanine dye, the cell with the gradient would contain more dye than the one without; if the cells were equilibrated with an oxonol dye, the cell without the gradient would contain more dye. However, cells of different sizes might contain different amounts of dye, irrespective of their membrane potentials. It is the concentration of dye, rather than the amount, in the cell that reflects membrane potential; what the flow cytometer measures is the amount, not the concentration. My colleagues and I (Novo et al., 1999) found that when the oxacarbocyanine dye DiOC 2 (3) was added to cells at much higher concentrations than are normally used for flow cytometric estimation of membrane potential, we were able to detect red (near 610 nm) fluorescence in addition to the green (near 515 nm) fluorescence normally emitted by this dye; we believe the red fluorescence to be due to the formation of dye aggregates. The green fluorescence is dependent on cell size, but independent of membrane potential; the red fluorescence, however, is both size- and potential-dependent. The ratio of red and green fluorescence, which largely eliminates the dependence on size, provides a substantially more accurate and precise measurement of bacterial membrane potential than was previously obtainable by flow cytometry, and calls into question some results previously obtained using cyanine, rhodamine, and oxonol dyes. In theory, oxonol dyes, in particular, should produce little or no staining of cells with normal membrane potentials and brighter staining of cells in which the potential gradient no longer exists. However, we believe that the increased oxonol

fluorescence seen in the heat-killed and alcohol-fixed bacteria often used as zero-potential controls may reflect changes in size and in lipid and protein chemistry resulting from these treatments, as well as changes in membrane potential. In our hands, the effects of less drastic measures, such as nutrient deprivation, which lower membrane potential, were detectable by our ratiometric method but produced no change in oxonol fluorescence.

6.6. Cytometry-aided approaches to antimicrobial drug design As Nebe-von-Caron et al. (2000) (this issue) have elegantly demonstrated, multiparameter cytometry can provide substantial information on the dynamics of microbial populations. My colleagues and I (Novo et al., 2000) took a different approach from theirs to simultaneous measurement of membrane potential and membrane integrity in cultures of Staphylococcus aureus exposed to antibiotics. We used a duallaser flow cytometer, exciting the green (525 nm) and red (613 nm) fluorescence of DiOC 2 (3) at 488 nm and the far red (675 nm) fluorescence of the impermeant nucleic acid dye TO-PRO-3 at 633 nm. The ratio of red to green DiOC 2 (3) fluorescence provided a size-independent measurement of membrane potential, while the ratio of far red TO-PRO-3 fluorescence to green DiOC 2 (3) fluorescence provided a similarly size-independent indicator of loss of membrane integrity. As can be seen from Fig. 1, a culture exposed to a sublethal dose of the betalactam antibiotic amoxicillin for 2 h shows a substantial population of what appear to be cells that have maintained normal or near-normal membrane potential, but have also become permeable to TOPRO-3. This population disappears over the course of another 2 h. While some of the events in the population may represent aggregates of viable and nonviable cells, we believe that the antibiotic treatment transiently permeabilizes the membranes of some viable cells, allowing TO-PRO-3 and similar dyes, such as propidium, to enter. Although Nebevon-Caron et al. found that cells that took up propidium did not grow when sorted onto agar plates, it is likely that propidium, at the concentrations used, would itself be toxic. Thus, it appears that further work will be needed to better define the cytometric correlates of viability in culture.

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Fig. 1. Effects of a sublethal dose of amoxicillin on cultures of Staphylococcus aureus (ATCC 29213). The vertical axis represents normalized permeability, as defined by the logarithm of the ratio of far red (675 nm) TO-PRO-3 fluorescence to green (525 nm) DiOC 2 (3) fluorescence. The horizontal axis represents membrane potential, as indicated by the logarithm of the ratio of red (613 nm) DiOC 2 (3) fluorescence to green (525 nm) DiOC 2 (3) fluorescence (see Novo et al., 2000).

6.7. A cytometry-aided approach to antimicrobial drug development Transient permeabilization may provide an approach to the development of agents active against microorganisms that are resistant to multiple antibiotics. If an antibiotic or other drug(s), while not lethal to such pathogens, can transiently permeabilize them, they could be killed by a variety of toxic agents rendered otherwise membrane impermeant, e.g. by addition of a quaternary ammonium group. These agents would not enter intact mammalian cells, and should therefore exhibit minimal toxicity to the host. Multiparameter cytometry has provided the initial evidence for this hypothesis; it should be invaluable in testing it.

Acknowledgements I thank Douglas Kell and David Novo for their thoughts on this subject, which are always stimulating, even when they do not agree with mine.

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