Microbiology and Molecular Biology Reviews, March 1999, p. 230-262, Vol. 63, No. 1
Department of Microbiology and
Guelph-Waterloo Centre for Graduate Work in Chemistry, University
of Guelph, Guelph, Ontario, Canada N1G 2W1
1092-2172/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Osmosensing by Bacteria: Signals and
Membrane-Based Sensors
SUMMARY
OSMOSENSING
SOLVENTS AND THEIR INTERACTIONS WITH BIOMOLECULES
Solvent Properties Pertinent to Osmosensing
Solvent-Macromolecule Interactions
Preferential interaction.
Hydration forces.
Pressure effects on the specificity of ligand-receptor
interactions.
Crowding and confinement.
Solvent-Membrane Interactions
Solvent effects on membrane structure.
Biomembrane permeability.
Mechanical properties of membranes.
TIMELINE OF OSMOSENSING
Phases of the Osmotic Stress Response
Cell Structure
Passive structural responses of bacteria to osmolality
changes.
Cell surface modifications and osmosensing.
Osmotic shifts, cytoplasmic composition, and nucleoid
organization.
Energy Metabolism
Osmotic upshifts.
Osmotic downshifts.
Adjustment of Cytoplasmic Solvent Composition
Cosolvent accumulation: absence of osmoprotectants.
Cosolvent accumulation: presence of osmoprotectants.
Cosolvent efflux.
Osmoregulation, Turgor Pressure, Cell Growth, and Cell
Division
Structural responses of metabolically active bacteria to
osmotic shifts.
CYTOPLASMIC MEMBRANE-BASED OSMOSENSORS
K+ Transporters of E. coli
Transporter Trk.
Transporter Kdp.
Sensor kinase KdpD.
Osmoprotectant Transporters
Glycine betaine uptake by L. monocytogenes.
Transporter BetP of C. glutamicum.
Transporter ProP of E. coli.
Mechanosensitive Channel MscL of E. coli
CONCLUSION
APPENDIX
ACKNOWLEDGMENTS
REFERENCES
SUMMARY
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Bacteria can survive dramatic osmotic shifts. Osmoregulatory responses mitigate the passive adjustments in cell structure and the growth inhibition that may ensue. The levels of certain cytoplasmic solutes rise and fall in response to increases and decreases, respectively, in extracellular osmolality. Certain organic compounds are favored over ions as osmoregulatory solutes, although K+ fluxes are intrinsic to the osmoregulatory response for at least some organisms. Osmosensors must undergo transitions between "off" and "on" conformations in response to changes in extracellular water activity (direct osmosensing) or resulting changes in cell structure (indirect osmosensing). Those located in the cytoplasmic membranes and nucleoids of bacteria are positioned for indirect osmosensing. Cytoplasmic membrane-based osmosensors may detect changes in the periplasmic and/or cytoplasmic solvent by experiencing changes in preferential interactions with particular solvent constituents, cosolvent-induced hydration changes, and/or macromolecular crowding. Alternatively, the membrane may act as an antenna and osmosensors may detect changes in membrane structure. Cosolvents may modulate intrinsic biomembrane strain and/or topologically closed membrane systems may experience changes in mechanical strain in response to imposed osmotic shifts. The osmosensory mechanisms controlling membrane-based K+ transporters, transcriptional regulators, osmoprotectant transporters, and mechanosensitive channels intrinsic to the cytoplasmic membrane of Escherichia coli are under intensive investigation. The osmoprotectant transporter ProP and channel MscL act as osmosensors after purification and reconstitution in proteoliposomes. Evidence that sensor kinase KdpD receives multiple sensory inputs is consistent with the effects of K+ fluxes on nucleoid structure, cellular energetics, cytoplasmic ionic strength, and ion composition as well as on cytoplasmic osmolality. Thus, osmoregulatory responses accommodate and exploit the effects of individual cosolvents on cell structure and function as well as the collective contribution of cosolvents to intracellular osmolality.
OSMOSENSING
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As inhabitants of natural and
artificial aqueous environments, bacteria survive dramatic changes in
extracellular osmolality (for definitions of terms used in this review,
see the glossary in Table 1). For
example, soil bacteria survive periods of low and high rainfall,
uropathogens survive urine concentration and dilution, and industrial
organisms tolerate concentrated nutrient solutions as well as the
extracellular accumulation of metabolic products. Bacteria respond both
passively and actively to changes in the osmolality of their
environment (Fig. 1 provides a summary of this phenomenon as it occurs
in Escherichia coli).
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Since the water permeability of the cytoplasmic membrane is high, imposed imbalances between turgor pressure and the osmolality gradient across the bacterial cell wall are short in duration. Changes in cell structure, organization, and composition that result from transmembrane water flux (Fig. 1, left column) trigger and are modulated by physiological responses (Fig. 1, right column). Bacteria respond to osmotic upshifts in three overlapping phases: dehydration (loss of some cell water) (phase I), adjustment of cytoplasmic solvent composition and rehydration (phase II), and cellular remodeling (phase III). Responses to osmotic downshifts are not yet well characterized, but they are also likely to proceed in three phases: water uptake (phase I), extrusion of water and cosolvents (phase II), and cytoplasmic cosolvent reaccumulation and cellular remodeling (phase III). Many of the cellular responses triggered by osmotic stimuli occur in parallel. Limited experimental evidence for sequential osmoregulatory processes (discussed below) offers insights into osmosensory mechanisms.
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Modulation of cytoplasmic solvent composition is the only response that is known to reverse the growth-inhibitory effects of osmotic shifts on bacteria. The following principles govern this process as it occurs in diverse bacteria, both gram positive and gram negative. (i) The concentrations of certain cytoplasmic solutes increase and decrease in response to osmotic up- and downshifts, respectively. (ii) There is a hierarchy of preference among the solutes that accumulate in response to an osmotic upshift. Particular zwitterionic organic cosolvents such as ectoine and glycine betaine are selected for this role over inorganic solutes such as K+. (iii) Osmoregulation of uptake, efflux, biosynthesis, and/or catabolism is required to modulate the cytoplasmic levels of these osmoregulatory solutes.
The genes and enzymes responsible for modulation of osmoregulatory solute levels have been identified in diverse bacteria. The mechanisms by which bacteria sense osmotic shifts (osmosensory mechanisms) and the basis for osmoregulatory solute selection are still poorly defined, however.
Chemosensors exploit the specificity of ligand-receptor interactions to detect the biochemistry of cellular environments, including both changes in nutrient supplies and signals with biological origins (Fig. 2). An osmosensor is a device that detects changes in extracellular water activity (direct osmosensing) or the resulting changes in cell structure (indirect osmosensing). In contrast to chemosensory mechanisms, osmosensory mechanisms cannot be based on stereospecific ligand-receptor interactions that occur at specific sites in receptor molecules. Instead, osmosensors are likely to be macromolecules that undergo changes in conformation and/or oligomerization in response to solvent changes or ensuing mechanical stimuli (Fig. 2). Our current knowledge suggests that osmosensors are located in the cytoplasmic membranes and nucleoids of bacteria. This review focuses on membrane-based osmosensors since they elicit primary osmoregulatory responses. In addition, current knowledge of the osmoregulation of transcription in bacteria has been reviewed whereas the basis for membrane-based osmosensing has not. The goals of this review are to define the properties of solvent-macromolecule and solvent-membrane interactions pertinent to osmosensing, to identify putative osmosensors and the changes that they detect, and to review our current understanding of cytoplasmic membrane-based osmosensory mechanisms in bacteria. This review is intended to stimulate broadly based research on osmosensing. Efforts have therefore been made to express microbiological, biochemical, and biophysical concepts in terms accessible to both specialists and nonspecialists. Readers who prefer textual explanations are therefore asked to accommodate the mathematical expressions preferred by those interested in the biophysical basis for osmosensing. This review also builds upon concepts developed by many other authors (2, 7, 25, 47, 74, 220, 228, 229, 256). Physiological responses to osmolality changes, to cooling-freezing, and to desiccation are related (44, 55, 124, 156). Only responses to osmolality changes are considered here, however. In addition, this review does not encompass the modifications to macromolecular structure that render organisms of the salt-in-cytoplasm type obligately halophilic (74).
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SOLVENTS AND THEIR INTERACTIONS WITH BIOMOLECULES
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Solvent Properties Pertinent to Osmosensing
For the purpose of this review, the chemical potential of water can be expressed as
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(1) |
w is the
partial molar volume of water, and P is the pressure.
Water and its solutes can be characterized by their activities (ai), where
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(2) |
i and xi are the
activity coefficient and the mole fraction of the ith
constituent, respectively, and O < ai < 1. In a physicochemically ideal system, the solvent activity approaches 1 and the solute concentrations (expressed as mole fractions or as the
more familiar molar units), which approach zero, can be used as
effective proxies for their activities. Experimental systems are often
designed to approximate ideality, so that simplifying assumptions can
be made about the resulting chemical and physical phenomena. Although
some laboratory media and some natural microbial environments can be
described as ideal solutions, many are nonideal. The aqueous
compartments within microorganisms are certainly nonideal. Nonideality
has important consequences for osmosensing (see
"Solvent-macromolecule interactions").
The ionic strength and osmotic pressure of the solvent arise through
collective contributions of ionic solutes and of all solutes,
respectively. Solutes that are present at high concentrations have the
greatest effects on osmolality and (if they are ionic) ionic strength.
The osmotic pressure of an aqueous solution (
) is defined as
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(3) |
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(4) |
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(5) |
Solvent-Macromolecule Interactions
Biologically relevant solvents, whether extracellular or intracellular, include both water and cosolvents. Cosolvents are solutes that significantly influence the behavior of water as a solvent (see "Solvent properties pertinent to osmosensing"). The term "cosolvent" is introduced here to emphasize that every solute molecule makes some contribution to solvent behavior while, more or less independently, playing a specific physiological role. Important solvent characteristics include cosolvent composition, ionic strength, osmotic pressure (or osmolality), and pH (see "Solvent properties pertinent to osmosensing"). Each cosolvent contributes, as part of the collective of cosolvents, to the osmolality of the solution. In addition, each cosolvent has particular effects on water, on its interactions with biomolecules, and hence on cellular functions. Both the osmolality and individual cosolvent effects are relevant to osmoregulatory mechanisms and their experimental study.
Diverse cosolvents are present outside and inside bacteria. NaCl and sugars (or other polyols) are the most prevalent and abundant extracellular cosolvents in natural environments. K+, organic anions, compatible solutes, proteins, and nucleic acids are the predominant intracellular cosolvents (28, 29, 74, 257, 304). The intracellular solvent of most bacteria is thus differentiated from the extracellular solvent by including a restricted array of low-molecular-weight cosolvents and a high concentration of macromolecular cosolvents. To understand osmosensing, it is necessary to deduce both the solvent changes to which osmosensors are actually exposed in vivo and the mechanisms by which those solvent changes can trigger changes in osmosensor conformation. Osmosensors located in the cytoplasmic membrane or nucleoid are exposed to the constrained solvent environments of the cytoplasm, membrane, and/or periplasm, not to the extracellular aqueous medium.
Effects of cosolvents on the behavior of water at interfaces (and their
effects on proteins) have been described, systematically but
empirically, in terms of the Hofmeister effect (41). Both ionic and nonionic cosolvents can be characterized in this way as
kosmotropic (water structure making, e.g. phosphate, ammonium, sucrose,
and glycerol), close to neutral (e.g., Na+ and
Cl
), or chaotropic (water structure breaking, e.g.,
SCN
and urea). For example, kosmotropes and chaotropes
tend to stabilize and destabilize native protein conformations,
respectively (for a further discussion, see below). Effects of
kosmotropes and chaotropes on protein function may be additive or
mutually compensatory (see, e.g., references 292 and
308).
Efforts by biochemists and biophysicists to systematically describe and explain water-cosolvent-macromolecule-solid-matrix interactions now offer additional perspectives and tools to students of osmoregulation (Fig. 3). Included are studies of (i) very low affinity ligand-receptor interactions and the impacts of diverse cosolvents on macromolecular stability, solubility, and assembly explained in terms of preferential exclusion of certain solvent constituents from macromolecular surfaces (275); (ii) the significance of hydration changes, probed by varying the osmotic pressure, for macromolecular interactions and functions (209); and (iii) the impacts of crowding and confinement on the structures and interactions of macromolecules (182, 297). Each rests on the concept that "the local environment will influence reactions taking place in a biological medium when reactants, transition state complexes, and products interact unequally with background species" (311). In the context of osmosensing, the reactants and products would be the forms ("off" and "on") of an osmosensor and the background species constituting the local environment would include water, cosolvents, and structural elements.
Preferential interaction. Some interactions among water, cosolvents, and osmosensors may be more appropriately considered in terms of preferential binding or exchange than in terms of classical binding theory. The latter describes the fractional occupancy of a specific receptor site(s) on a macromolecule by one or more ligands. This occupancy may vary from 0 (no binding) to the number of sites, which is usually small. In terms of preferential binding or exchange, the reference state is a macromolecule in pure water with all exposed surface sites hydrated (no sites occupied by cosolvent). A gradual exchange of water for cosolvent occurs as the cosolvent is added to the system. At any cosolvent concentration, the fractional occupancy of surface sites by cosolvent exceeds, matches, or falls short of the proportion (mole fraction) of cosolvent molecules in bulk solution if the surface sites prefer cosolvent over water, show no preference, or prefer water over cosolvent, respectively. This situation differs substantially from that treated by classical binding theory. Each solvent-macromolecule interaction is weak, but the number of sites involved is large and hence the potential impact of solvent-cosolvent exchange on macromolecular structure and function is also large. For example, it has been estimated that lysozyme offers a total of 266 surface sites for solvent and/or cosolvent occupancy (276). In addition, the global behavior of the macromolecule-solvent system is an average over weak interactions among macromolecule, cosolvent, and water at a large number of nonidentical surface sites.
The effects of various organic cosolvents on macromolecular solubility, conformation, and assembly can be described in terms of such preferential interactions (275, 276). A correlation is observed between cosolvent exclusion from protein surfaces and stabilization of native protein conformations and assemblies (which often have smaller surface areas exposed to solvent than denatured forms do) (Fig. 3). Steric restrictions on approach to the macromolecular surface by cosolvent molecules (as opposed to the smaller water molecules) and a thermodynamic preference of some cosolvent molecules for the bulk solvent over the solvent-protein interface are both believed to contribute to this cosolvent behavior. The protein-stabilizing effects of some compatible solutes have been described in these terms (248, 292, 308). It has been emphasized, however, that stabilization does not require preferential exclusion of the cosolvent from macromolecular surfaces. Stabilization can also be achieved if there is less preferential binding of the cosolvent to the denatured macromolecule than to the native macromolecule (as was shown for trehalose by Xie and Timasheff [301]). These observations indicate that the conformations and associations of macromolecules may be sensitive to the concentrations of cosolvents, but they also underscore the complex dependence of such phenomena on the chemical nature of cosolvent-macromolecule interactions, not on solution osmolality per se.
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Hydration forces. Osmotic pressure has been used as a tool to probe the biological significance of macromolecular hydration, i.e., of "hydration forces." For example, the addition of osmotically active, polymeric cosolvent (e.g., polyethylene glycol [PEG]) to the surrounding medium can suppress ion channel opening and alter both KD and KM for the interaction of glucose with hexokinase (209). These data are interpreted as showing that the entry of cosolvent molecules to macromolecule-associated solvent pools is restricted and that solvent compartments which differ in osmolality are thus created. Thus, macromolecules may adjust to their solvent environments by assuming states that balance the (unfavorable) exclusion of cosolvent from inaccessible solvent pools against the (unfavorable) juxtaposition of macromolecular constituents. For example, withdrawal of solvent from intramolecular pools of low osmolality may force macromolecules to adopt different (and less energetically favorable) conformations than are assumed to be present in the absence of cosolvent (Fig. 3).
For hexokinase, this adjustment appears to be related to closure of a solvent-filled protein cleft containing the active site, although more generalized dehydration of the enzyme may also be involved. As would be anticipated for a purely osmotic effect, the number of water molecules implicated in this transition depends on the size of the cosolvent molecules used to impose osmotic stress. The difference in the number of PEG-accessible water molecules between the glucose-associated and glucose-free hexokinase conformations varies from 50 to 326 as the molecular weight of PEG increases from 300 to 1,000. It then remains constant as the molecular weight of PEG increases further to 10,000 (232). Thus, a larger quantity of hexokinase-associated water is inaccessible to high-molecular-weight than to low-molecular-weight PEG. These observations suggest that protein form and function can respond on an osmotic basis to certain cosolvents, including some that are similar in molecular weight and concentration to those which impose osmotic stress or serve as compatible solutes in nature.Pressure effects on the specificity of ligand-receptor interactions. Both osmotic and hydrostatic pressures have been used to perturb DNA-protein interactions (235). Stresses imposed with osmotic and hydrostatic pressures are fundamentally different; osmotic stress causes water molecules to be transferred from cosolvent-inaccessible to cosolvent-accessible regions around or within macromolecules, whereas hydrostatic pressure changes the weight density of the entire system. For some restriction endonucleases, DNA recognition sequence specificity changed as osmotic pressure was increased by the addition of diverse, low-molecular-weight cosolvents including sucrose, glycerol, 2-propanol, and N-methylformamide (0 to 3 osmolal, but with significant effect at 1 osmolal) (see, e.g., reference 234). These effects were reversed by elevated hydrostatic pressure (up to 500 atm). They were interpreted as indicating differential involvement of water at the enzyme-DNA interface for different DNA sequences. Hydration has also been recognized as playing a critical role in carbohydrate-protein interactions (152). Such behavior differs fundamentally from that described by preferential binding or exchange theory, however. It involves small numbers of water molecules that mediate high-affinity enzyme-substrate recognition at specific enzyme and substrate sites.
Crowding and confinement. Considerations of macromolecular crowding and confinement may also be used to explain "background" effects on macromolecular structures and interactions (182, 311). They define the collective impact of macromolecules, whether soluble or present as structural elements, on cellular processes. Garner and Burg (77) have reviewed these concepts from a physiological perspective. The term "crowding" refers to effects of high collective volume occupancy by macromolecules on the structures and functions of individual macromolecular species with which they interact only weakly and nonspecifically. Such effects may be biologically significant since macromolecules occupy as much as 50% of the cytoplasmic volume. The term "confinement" refers to effects of entrapment within a subcellular matrix on macromolecular function.
Consideration of crowding and confinement effects on biological processes in terms of solution nonideality yields predictions that are supported by experimental evidence. (i) Under conditions of crowding comparable to those found in vivo, the values assumed by macromolecular association constants may be dominated by crowding effects and may exceed the corresponding values for dilute solution by up to several orders of magnitude. (ii) Compact or globular macromolecular conformations or assemblies are favored more in crowded than in dilute solutions (Fig. 3). (iii) For a given degree of crowding, effects on associations between macromolecules are expected to be much greater than effects on association of small molecules with macromolecules. (iv) Both crowding and confinement tend to enhance macromolecular associations
unlike crowding, confinement may favor the formation of
extended (linear or discoid) rather than globular aggregates. Such
considerations have led to the proposal that (some) animal cells may
regulate macromolecular crowding rather than cell volume
(183). This proposal rests on the concept that
physiologically relevant changes in extracellular water activity,
effected by modulating the extracellular concentrations of diverse
cosolvents, may be translated into changes in cytoplasmic crowding or
confinement, thereby avoiding the complication of specific
cosolvent-osmosensor interactions.
These background effects on macromolecular structures and interactions
are not mutually exclusive, and this list is not necessarily complete.
For example, phenomena other than hydration have been proposed as
origins for the mutual repulsion of macromolecular surfaces in aqueous
solution (106). It has been argued that small cosolvent
molecules may exert their effects on protein solubility, stability, and
function through excluded volume rather than through osmotic or
preferential binding effects (297). It has been proposed that water within enzyme active sites, ligand binding sites of receptors, and ion channels, like that within polymeric matrices, differs from bulk water in density and hence in its behavior as a
solvent (295, 296). Resolution of these overlapping and/or conflicting interpretations is not the objective of this review. Rather, these concepts are useful to students of osmoregulation because
they contribute to our understanding of solvent effects on cell
structure and function and because they suggest potential osmosensory
mechanisms (Table 2).
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Solvent-Membrane Interactions
The principles outlined above apply to biomembrane constituents, proteins, phospholipids, and polysaccharides, as well as to other macromolecules. Solvent effects on the assembled membrane warrant particular attention in the context of osmosensing because biomembranes constitute the semipermeable barriers which define biologically relevant osmotic compartments and some osmosensors are located within biomembranes. When an osmotic shift is imposed on a vesicular membrane system (a cell, a biomembrane vesicle, or a liposome), membrane perturbations with multiple origins occur. The following discussion is designed to enumerate these effects, indicate which may be detected by osmosensors, and provide references to pertinent experimental approaches.
Solvent effects on membrane structure.
Biophysicists
and biochemists have long been fascinated and puzzled by the diversity
and complex phase behavior of membrane phospholipids. The net charge,
hydrogen-bonding capacity, and hydration of each head group, the length
and unsaturation of each acyl chain, and the resulting overall shape of
each phospholipid molecule all contribute to the phospholipid-solvent
interactions which determine phase behavior in aqueous phospholipid
dispersions (57, 60, 78, 85, 155). Each phospholipid (and
each phospholipid mixture) has a particular propensity to form the
lamellar (L) phase characteristic of biological membranes versus
alternative arrangements, including the nonlamellar hexagonal
(HI or HII) or cubic phases. Phospholipid phase
transitions, for example, the transitions from gel (L
)
to liquid crystalline (L
) lamellar phases and from the
lamellar phase to the hexagonal phase, are characterized by phase
transition temperatures (for these examples, TM
and TH, respectively). Phospholipid phase
behavior (often indicated by phase transition temperatures) is also
influenced by amphiphiles which insert among phospholipid molecules
(e.g., alcohols, fatty acids, detergents, anaesthetics, and peptides) and by such solvent properties as pH, ionic strength, and cosolvent composition.
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Biomembrane permeability.
For small deviations from
equilibrium, the net rate of volume flux across a membrane in response
to a hydrostatic or osmotic driving force can be described in terms of
an osmotic permeability coefficient (Pf
[centimeters per second]) characteristic of the membrane and a
reflection coefficient (
[dimensionless]) characteristic of the
membrane and the cosolvent used to impose the osmotic gradient (71, 200). The flow of volume across a membrane separating compartment 1 from compartment 2 can be described as
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(6) |
w is the partial molar volume of water, R is the gas constant,
T is the temperature (Kelvin), P1 and
P2 are the hydrostatic pressures (atmospheres)
in compartments 1 and 2, respectively,
i
(0 <
i < 1) describes the relative
rates at which the ith cosolvent and water cross the
membrane (via all available pathways) and Osmi1 and Osmi2 are the osmolalities due to the
ith cosolvent in compartments 1 and 2, respectively. The
reflection coefficient (
i) varies from close
to 0 for a highly permeant cosolvent to 1 for a cosolvent which does
not cross the cell surface (e.g., glycerol has a low
and sucrose
has a high
with respect to the surface of E. coli [see
below]). The significance of
can be grasped by recognizing that a
cosolvent flux sufficiently rapid (with respect to solvent flux) to
collapse the osmolality gradient will attenuate the solvent flux
(Jv). In terms which are important for
osmosensing, Pf and
i
contribute to the rate and extent of solvent flux and hence to the
duration of imposed osmotic gradients (
Osm), the rate of change of
hydrostatic pressure (
P), and the rate at and degree to
which a plastic, membrane-bounded osmotic compartment will be deformed
in response to an osmotic shift.
Water crosses biomembranes via three pathways: the phospholipid
bilayer, aquaporins (water-selective channels), and integral membrane
proteins with other functions (e.g., transporters and channels designed
to translocate substrates other than water) (52, 284, 307).
The relative contributions of these pathways depend directly on the
numbers of the relevant proteins per unit membrane area (and, in the
case of the last pathway, the availability of substrates). The osmotic
water permeabilities of phospholipid bilayers
(Pf in the range 10
3 to
10
2 cm/s) are usually sufficient to permit equilibration
of water across the membrane on a millisecond timescale. Aquaporins
raise Pf to values in excess of
10
2 cm/s and hasten equilibration accordingly
(284). The physiological roles of aquaporins have not been
defined. However, their presence suggests that under at least some
physiological conditions, transmembrane water flux via the phospholipid
bilayer, alone, is unacceptably slow. Although water does cross
biomembranes via transporters and channels other than aquaporins, the
contributions of transporters and channels to total water flux may be
relatively small (284, 307). Our understanding of the
influence of cosolvents on the water permeabilities of phospholipid
bilayers is currently limited (see, e.g., reference
17).
Like that of water, transmembrane cosolvent flux may in principle be
either passive (occurring via the phospholipid bilayer or nonspecific
pathways involving membrane proteins) or mediated (occurring via
cosolvent-specific integral membrane proteins). Since the passive
permeabilities of biomembranes for many biologically relevant solutes
are orders of magnitude lower than that of water, solute-specific
channels or transporters are required if high solute flux rates are to
be attained (78) and the passive permeabilities for solutes
are often ignored. In fact, passive permeabilities are important for
certain biologically relevant cosolvents (e.g., glycerol, urea, and
NH3), and they may become important when cosolvents, even
those of low passive permeability, are used to impose large osmotic
gradients (133, 180). The passive permeabilities of phospholipid bilayers and biomembranes for solutes are known to increase dramatically near both TM and
TH (52, 60, 165). Although the
temperature dependence of passive membrane permeability has been
extensively explored, the degree to which absolute cosolvent levels (as
opposed to cosolvent gradients) influence cosolvent permeabilities is
not well characterized.
Mechanical properties of membranes. A membrane can be defined, mechanically, as "a material with a very small thickness in comparison with its radii of curvature which separates two adjacent, liquid-like domains and supports the stresses created by the embedding medium" (19). To understand membrane-associated osmosensors, it may become necessary to link our understanding of membrane molecular structure and dynamics (with a length scale up to a few nanometers) with our understanding of the membrane as a continuum of condensed matter (with a length scale greater than 500 nm). Barriers to the attainment of that goal include both difficulties of communication among investigators and technical limits to the study of phenomena which occur on the relevant length scale and timescale (19).
To define the mechanical properties of a biological membrane, it would be necessary to fully describe the rates and extents of shape changes that occur upon application of defined forces (stresses) in vivo. For an elastic membrane, each applied stress will result in some strain, i.e., some reversible shape change. Stress and strain are related by proportionality constants (elastic moduli) determined by material properties of the membrane:
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(7) |
TIMELINE OF OSMOSENSING
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Osmoregulators are devices that implement the response of an organism to changing environmental osmolality. An individual device may both detect and respond to solvent changes (i.e., osmosensors may also be osmoregulators). Alternatively, osmosensors and osmoregulators may be separate and communication between or among them may require additional signal transduction machinery. The high water permeability and solute selectivity of biomembranes ensure that changes in extracellular osmolality imposed with membrane-impermeant cosolvents trigger extensive changes in cell structure and chemistry. Thus, in principle osmosensors may detect changes in extracellular water activity (direct osmosensing) or they may detect and elicit responses tailored to address secondary consequences of osmotic shifts (indirect osmosensing). It is therefore expected that an array of osmosensors may detect and control a temporal cascade of cellular changes and osmoregulatory responses. As a result, the "osmotic history" of each cell will determine its response to new osmotic stimuli (76, 261).
The following discussion is designed to place putative bacterial osmosensors within the cascade of osmotically induced changes to cell structure and physiology, thereby identifying the stimuli to which each osmosensor may (or may not) respond. This process of correlation and elimination is important, since the list of stimuli which could, in principle, be detected by each osmosensor is long (Table 2).
Phases of the Osmotic Stress Response
When subjected to an increase or decrease in the osmolality of the suspending medium (an osmotic upshift or downshift, respectively), E. coli cells change as illustrated in Fig. 1. This review focuses primarily on membrane-based sensory processes that occur during phase I of this response. Phases II and III are considered only to the extent that they illustrate the physiological objectives met by osmoregulatory processes and the relevant changes in cell structure associated with steady-state exposure to media of various osmolalities. The timescales of these events have not yet been fully defined, and the interpretation of existing evidence concerning the timescales of phases I and II is subject to ambiguities (see Appendix). Nevertheless, the events listed as phase I of the response to an osmotic upshift can be considered to occur within milliseconds to minutes of an osmotic shift. Phase II (which also begins on imposition of an osmotic shift) extends until approximately 40 min or 10 to 20 min after the shift for bacteria cultivated in the absence and presence of osmoprotectants, respectively. Phase III is highly condition dependent and can extend for 1 h or more after the shift. The timescales for water uptake (phase I) and release of cytoplasmic cosolvents (phase II) after an osmotic downshift are at or below the limits of the experimental techniques that have been applied to date. These phases are complete within 1 to 2 min, however, and the ensuing cosolvent reaccumulation (phase III) is complete within 10 to 20 min after the shift. The evidence on which these estimates are based is discussed further below.
Most studies have focused on effects of osmotic upshifts or of osmotic downshifts or of steady-state adaptation to either of these conditions. Osmotic upshifts and downshifts have usually been applied to bacteria cultivated in low- and high-osmolality media, respectively. Additional insight regarding osmosensing will be gained by considering, as a continuum, cellular responses to both increases and decreases in medium osmolality (see, for example, Parker's assessment of changes in cell volume versus macromolecular crowding as triggers of regulatory volume changes in canine erythrocytes [207]). Such studies will be facilitated by examining organisms with sufficient osmotolerance to permit the application of significant osmotic shifts, both up and down, after cosolvent loading (e.g., Lactobacillus plantarum [80, 81] and Halomonas elongata).
Bacterial responses have been examined at different growth phases (usually either mid-exponential or late exponential phase) and with different growth status during the experiment (growing or partially or fully nutrient deprived). Significant cellular remodeling accompanies both long-term (minutes to hours) osmoadaptation and the transition to stationary-phase growth. Thus, the patterns outlined in Fig. 1 will be refined as the short- and long-term effects of osmolality, growth phase, and nutrient status are disentangled.
Cell Structure
Changes in extracellular osmolality may elicit changes in cell structure and/or water fluxes across the cell surface. The following discussion is designed to identify (i) osmotically induced structural changes which could be detected by membrane-based osmosensors, (ii) osmotically induced structural changes which may influence subsequent osmoregulatory responses, and (iii) experimental tools which may assist researchers in identifying osmosensors and the signals to which they respond.
Turgor pressure (
P) is defined as the hydrostatic
pressure difference which balances the osmotic pressure (or osmolality) difference between cell interior (i) and exterior
(o), rendering the chemical potentials
(µw) of intracellular and extracellular water
(see equation 1) equal at equilibrium. For cells which can be treated
as two-compartment systems:
|
(8) |
P, Osmi or
Osmo cause solvent flux across the cell surface
to reestablish the equilibrium described by equation 8. On the basis of
measured osmotic water permeability coefficients for phospholipid
bilayers and membrane vesicles, equilibration is expected to occur
within seconds of an osmotic shift (305). Since large cells
(e.g., those of plants and some eukaryotic microorganisms) can be
impaled,
P and Osmi can be
manipulated directly (89, 102, 210). Such techniques may
also be applicable to giant bacterial cells created by mutation and/or
antibiotic treatment. The rate of solvent flux and the participation of
particular cosolvents in passive cellular adjustment to osmotic shifts
are illustrated (for fluxes of small magnitude) by equation 6.
Turgor pressure imposes stress on the cell surface. For a spherical cell with a very thin surface layer against which turgor pressure is exerted, the relationship between turgor pressure and the imposed stress (S) would be
|
(9) |
Like those of phospholipid membranes, the mechanical properties of cell surfaces may in principle be described in terms of both their stiffness (or viscosity) and their elastic moduli (their ability to undergo various reversible deformations). For example, bacterial murein layers are both stiffer and more elastic than cytoplasmic membranes (as discussed by Thwaites and Mendelson [274]). For a spherical cell with a single, thin, elastic surface layer, the relationship between surface stress and strain would be
|
(10) |
A/A is the strain, i.e., the
proportional change in the cell surface area in response to the imposed
stress. In the simplest case, k would be constant regardless
of the position on the cell surface, orientation on the cell surface,
or magnitude of the applied stress.
Equations 9 and 10 can be combined to relate cell surface strain
(
A/A) and the stress imposed by turgor pressure
(
P) for a spherical cell:
|
(11) |
P]),
elastic modulus (k), and surface layer thickness
(h). (ii) Thicker cell surfaces expand and contract less
(have lower strain) under a given stress (turgor pressure
[
P]) than do thinner surfaces for a given elastic
modulus (k) and cell size (r). (iii) In cells with elastic surfaces (small k), changes in extracellular
osmolality may be accommodated by both water fluxes and adjustments in
cell surface area. The time courses of solvent flux and of structural change after an osmotic perturbation are then determined by
Pf,
i, k, and viscosity
coefficients which determine the rates at which structural changes can occur.
By using a microscopic pressure probe to measure and manipulate turgor
pressure, osmotic responses and stress-strain relationships have been
examined for large cells of plants and eukaryotic microbes. Such
analyses suggest that for those systems at least,
i, Pf, and k
can vary with turgor pressure (89, 200, 210). Cellular properties such as surface stress and elastic modulus cannot readily be
deduced from measurements of turgor pressure, osmotic gradients, and
cell dimensions for nonspherical cells or for cells in which the
elastic modulus varies with position on the cell surface, orientation
on the cell surface, or magnitude of the applied stress. Since
bacterial turgor pressure cannot readily be manipulated or measured
(see Appendix), current prospects for quantifying surface stress and
strain in whole bacteria are poor. However, pertinent information can
be obtained by examining the mechanical properties of cellular
constituents (see, e.g., reference 274).
The relationships outlined above suggest that the kinetics of
osmotically induced solvent flux, of associated structural and biochemical changes, and hence of the time constants required of
osmosensory and osmoregulatory processes can be adjusted by remodeling
the cell surface to alter its reflection coefficient (
i) and osmotic permeability coefficient
(Pf) or its mechanical properties (its stiffness
and elastic moduli). For example, reducing the reflection coefficient
for a cosolvent would facilitate its rapid equilibration across the
cytoplasmic membrane, reducing or eliminating osmotically induced water
flux. For a benign cosolvent (e.g., glycerol), such equilibration may
be more favorable to cellular survival and growth than cosolvent
exclusion. Reduction of the osmotic permeability coefficient
(Pf) would slow cellular dehydration in response
to osmotic upshifts. Adjustment of surface stiffness and elasticity,
either in general or within local cell surface regions, would modulate
the degree to which swelling, shrinkage, and shape changes occur in
response to extracellular osmolality changes.
Passive structural responses of bacteria to osmolality changes. How are the structures of bacterial cells influenced by extracellular osmolality changes? Measurements of many cellular properties are pertinent to this topic. They include turgor pressure; cellular, cytoplasmic, and periplasmic volume (cell partitioning); cell density (not population density); cell size, shape and ultrastructure; and the mechanical properties of cell surface layers. Relevant techniques are discussed in the Appendix. Some bacteria have been shown to maintain turgor pressure (references 45 and 294 and references cited therein). For example, the turgor pressure for E. coli cells cultivated in standard, defined media is believed to be 3 to 5 atm. Although the mechanical properties of bacteria have not been fully analyzed (for reasons discussed above), studies of osmotic effects on cell size and hydration offer some insight into their osmotic relations.
Building upon earlier experiments performed with a variety of bacteria, Alemohammad and Knowles (1) used turbidimetry and solute distribution measurements in parallel to assess the immediate effects of salts, sucrose, and glycerol (osmotic shifts of up to 0.55 osmolal) on resting E. coli cells. The turbidity increased over 2 min to reach a new value that was stable for 1 h after treatment with NaCl, MgCl2, or sucrose. Decreases in both cell and cytoplasmic volume were observed by measuring solute exclusion 30 min after the shift. Stopped-flow spectrophotometry was used to establish that glycerol caused only a transient (maximal at approximately 0.5 s and reversed within 8 s) increase in the turbidity of such cell suspensions. Subsequent research has confirmed that the cytoplasmic membrane has a low reflection coefficient for glycerol (
Gly), which can be further reduced by the expression of
facilitator GlpF (164).
These experiments established the time frame for the response of
E. coli cells to osmotic upshifts and validated the use of glycerol as a membrane-permeant cosolvent during studies of
osmoadaptation by E. coli. The fact that elevation of
extracellular osmolality with glycerol fails to activate an
osmoregulatory response is often taken as evidence that a change in
transmembrane osmolality gradient (or its consequence) is sensed.
However, dehydration of an osmosensor as a result of cosolvent
exclusion from intramolecular water pools could also act as a stimulus
(see "Hydration forces" above). Just as its lack of charge and its
small size allow glycerol to permeate cell membranes, they may also
allow it to penetrate intramolecular water pools and hence fail to
alter macromolecular (osmosensor) conformation.
The impact of osmotic upshifts on the cellular, cytoplasmic, and
periplasmic water content (microliters per milligram of protein or dry
weight) of nutrient-deprived bacteria (E. coli and
Salmonella only) has been assessed by applying the solute
distribution technique (1, 38, 39, 260) (see Appendix).
Decreases in both cell and cytoplasmic water contents were observed
when osmotic upshifts were imposed with solutes expected to be excluded
by the cell wall (polyglutamate) or the cytoplasmic membrane (sucrose,
MgCl2, or NaCl) but not with cytoplasmic membrane-permeant
solutes (glycerol or ethanol). Osmotic shifts imposed on
Salmonella with sucrose or on E. coli with
sucrose or NaCl were found to increase the fraction of cell water that
was periplasmic. However, the degrees and morphological consequences of
plasmolysis differed when osmotic upshifts of the same magnitude (0.365 osmolal) were imposed on E. coli cells with NaCl or sucrose
(1).
Koch (125) used turbidimetry to analyze cellular changes
5 s and 1 min after small osmotic upshifts (no larger than 0.25 osmolal) were imposed by adding arabinose or xylose to growing E. coli cells. Measurements were performed by stopped-flow
spectrophotometry and interpreted by fitting the data to a
light-scattering model in which the total intensity of the scattered
light (presumed to cover 0° through 180°) was related to cell size.
Cells were treated as solid rods (shapes obtained by averaging that of
a cylinder and that of an ellipse) of uniform and invariant refractive index (corrections for changes to the refractive index of the medium
were applied). Turbidimetric changes were reported as cell surface area
and volume changes. Arguing that the osmotic upshifts were too small to
cause plasmolysis and that the measurement times were too short to
accommodate metabolic changes (sugar uptake and/or osmoregulatory
activity), Koch reported that increasing the osmotic upshifts elicited
continuous cell surface area and volume decreases (without abrupt
changes) to approximately 40 and 50%, respectively.
Baldwin et al. (12) measured changes in the buoyant density
(density gradient centrifugation) and size (Coulter counter and light
microscopy) of nutrient-deprived E. coli cells that were
subjected to small (0.15- to 0.45-osmolar) osmotic up- and downshifts
with NaCl or sucrose. Measurements performed within 10 min of the
osmotic shift indicated that buoyant density increased as cell size
decreased in response to osmotic upshifts and that buoyant density
decreased as cell size increased in response to osmotic downshifts.
However, the volume changes were smaller than those reported by Koch
(125) (the surface area decreased 33% [microscopy] after
a 0.15-osmolar upshift, and the volume decreased 21.5% [Coulter
counter] after a 0.35-osmolar upshift). Since increased cellular
buoyant density would certainly be accompanied by an increased cellular
refractive index, Koch's assumption that all turbidity changes would
reflect changes in cell size (125) was not justified and may
have contributed to overestimation of the reported area and volume changes.
By applying phase-contrast microscopy to filamentous (ftsA
and ftsI) E. coli mutants, Koch et al.
(127) observed decreases in cell length (on average, 17%)
in response to detergent (sodium dodecyl sulfate) disruption of the
cytoplasmic membrane. The observed shrinkage was attributed to
elasticity of the murein layer. This elasticity was tested further by
measuring the angular dependence (4° through 12°) of the intensity
of unpolarized light scattered by sacculus suspensions as their pH was
titrated over a range designed to adjust the net sacculus charge from
positive through negative values (129). The data were fit to
a model of the sacculi as hollow prolate ellipsoids, assuming a
constant thickness and refractive index of the murein shell and a
constant unique axial ratio of the ellipsoids, in order to extract
changes in sacculus width, reported as sacculus area. Although
systematic changes in light scattering by the sacculi were elicited by
pH changes, their interpretation in terms of sacculus area may not be
justified. Others have also reported murein elasticity, however
(summarized in reference 58). Taken together, these
results suggest that bacterial cell surfaces are elastic, with their
elastic moduli varying with cell volume (perhaps turgor pressure) and
ionic strength.
The studies summarized above suggest that the cytoplasmic membrane and
cell wall of E. coli act as a unit structure with a sufficiently small elastic modulus (k) to permit overall
cell shrinkage in response to cellular dehydration. The available
measurements do not indicate whether turgor pressure is maintained at a
constant level as cells shrink in this way or whether it declines
steadily (see the discussion by Csonka and Hanson
[47]). It is also difficult to assess whether these
adjustments are accompanied by changes in periplasmic thickness that
might be detected by an osmosensor. The cytoplasmic membrane partitions
the cell interior into periplasmic and cytoplasmic compartments. The
osmolalities of the cytoplasm and periplasm are sometimes assumed to be
equal (228, 260), but the experimental evidence supporting
that assumption is extremely limited. If periplasmic and cytoplasmic
osmolalities were always equal, the cytoplasmic membrane would be
subject to area dilation or condensation only as a result of mechanical
stress transmitted by its linkage to the cell wall and/or the nucleoid
(see below). Assessment of the relative osmolalities of the cytoplasm
and periplasm for bacteria growing in diverse solvent environments
should be a high experimental priority.
The outer membrane is covalently linked to the murein sacculus via
lipoproteins (reference 147 and references therein)
and some porins. Although structural links between the outer membrane and the murein layer may be stronger and/or more densely spaced than
those between the cell wall and the cytoplasmic membrane, the latter
links certainly exist (147). The potential involvement of
Bayer's bridges in protein secretion and of periseptal annuli in cell
division has motivated extensive electron microscopic analyses of
plasmolyzed E. coli cells. These observations clearly indicate that membranous links between the cytoplasmic membrane and the
cell wall (Hechtian strands) are retained despite extensive plasmolysis
(118, 126, 251, 298). Limitations on fluorescence recovery
after photobleaching of periplasmic probes indicate that the periplasm
is partitioned, presumably by links between the outer and cytoplasmic
membranes (72). Woldringh et al. (299) and
Nanninga (198) emphasize the extensive linkage of cell wall and cytoplasmic membrane effected by murein biosynthetic enzyme complexes as well as the linkage of cytoplasmic membrane and nucleoid effected by cotranslational protein secretion. Thus, the cytoplasmic membrane acts as a structural link between the cytoplasm (including the
nucleoid) and the cell wall, suggesting that it may experience local
variations in mechanical stress in response to both metabolic activity
and osmotic shifts.
Plant cell plasmolysis has been examined to assess the molecular bases
for freezing and salinity tolerance (203, 219). These studies, too, reveal the presence of Hechtian strands and plasma membrane vesiculation in plasmolyzed cells. It has been suggested that
such structures may maintain the cytoplasmic membrane surface area,
cell integrity, and polarity and focus mechanical stress from the wall
to the membrane during the protoplast shrinkage that accompanies
plasmolysis. In addition, vitronectin- and fibronectin-like proteins,
present in enhanced numbers in NaCl-adapted tobacco cells, are proposed
to mediate enhanced membrane-wall adhesion (309). It would
be interesting to learn whether the incidence of wall-membrane
adhesions also varies as a function of bacterial growth medium
osmolality and/or salinity.
Cell surface modifications and osmosensing.
Does
structural remodeling of the cell surface influence the osmotic stress
response? Genetic screening has revealed genes encoding E. coli cell surface molecules whose expression is modulated by
extracellular osmolality (Table 3). In
most cases, steady-state levels of these elements have been reported
but their influence on the kinetics of the osmotic stress response has
not.
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