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Is it true that in an enzyme-catalyzed reaction, the rate of reaction does not vary with varying osmotic pressure?
Instead of just a true or false answer, why does it, or why does it not vary with osmotic pressure?
This paper is a compelling example of an effect of osmotic pressure on enzyme function. Restriction enzymes are normally very selective in the base sequences of sites at which they cut DNA, but in some conditions they show cutting at off-target sites, called "star activity." "Star activity" increased for some restriction enzymes almost linearly with osmotic pressure generated by a range of non-ionic solutes. Supporting the idea that the higher osmotic pressure was directly responsible, increased hydrostatic pressure reversed the effect of increased osmotic pressure.
Increasing the osmotic pressure is essentially decreasing the chemical activity of the water in the system. So it's not surprising that any enzyme reaction might be affected, particularly if the structures of active sites of enzymes are altered as suggested by the authors of the paper cited above.
In order to achieve initiation of DNA replication in vitro at the E. coli chromosomal origin of replication, Arthur Kornberg's lab had to include polyethylene glycol (PEG) in their reactions. The proposed mechanism for this enhancement was termed "molecular crowding" which was thought to increase the local concentration of the substrates.
A similar effect has been seen in reactions with T4 DNA ligase. Blunt-ended ligations are more efficient in buffers containing PEG.
Concentrated dye diffuses along the concentration gradient until reaching equilibrium (no net movement). Diffusion is the net movement of a substance from high concentration to low concentration. This difference in concentration is referred to as a concentration gradient . This movement does not require any external energy, but uses the free energy intrinsic to the system.
Osmotic Pressure Triggered Rapid Release of Encapsulated Enzymes with Enhanced Activity
In this study, a single-step microfluidic approach is reported for encapsulation of enzymes within microcapsules with ultrathin polymeric shell for controlled release triggered by an osmotic shock. Using a glass capillary microfluidic device, monodisperse water-in-oil-in-water double emulsion droplets are fabricated with enzymes in the core and an ultrathin middle oil layer that solidifies to produce a consolidated inert polymeric shell with a thickness of a few tens to hundreds of nanometers. Through careful design of microcapsule membranes, a high percentage of cargo release, over 90%, is achieved, which is triggered by osmotic shock when using poly(methyl methacrylate) as the shell material. Moreover, it is demonstrated that compared to free enzymes, the encapsulated enzyme activity is maintained well for as long as 47 days at room temperature. This study not only extends industrial applications of enzymes, but also offers new opportunities for encapsulation of a wide range of sensitive molecules and biomolecules that can be controllably released upon applying osmotic shock.
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2. Materials and Methods
2.1. Study Species
The experimental fish were 1-year-old, pool-bred fish (40 g, 16.57.61𠂜m fork length). The fish were held in a glass-steel circular cylinder (1 m diameter) containing 300 L aerated water (pH 7.51, salinity 0.28‰, total alkalinity 1.39 mM, total hardness 1.10 mM, total phosphorus 0.11 mg/L, total nitrogen 2.95 mg/L, total iron 0.002 mg/L, SO4 2− 31.19 mg/L, Ca 2+ 19.81 mg/L, and Mg 2+ 2.03 mg/L) at room temperature. The fish were acclimated to experimental conditions for 3𠂝. The analytical reagents (AR) were NaCl and NaHCO3.
2.2. Experimental Design
First, we evaluated the individual effect of salinity or alkalinity. The fish were divided into four treatment groups for each factor: 5, 8, 11, and 14 g/L NaCl and 15, 30, 45, and 60 mM NaHCO3. Second, we evaluated the combined effects of salinity and alkalinity. The fish were divided into 9 treatment groups to test all combinations of salinity (5, 8, and 11 g/L) and alkalinity (15, 30, and 45 mM), based on orthogonal table L9 (3 4 ) ( Table 1 ). Each treatment group consisted of three replicates with eight specimens per replicate, and the number of the experimental fish was 480. The experiment was performed for 60 days without aeration at 22 ± 0.5ଌ. A third of the water was changed daily and the corresponding concentration was adjusted with a pre-prepared master mix.
2.3. Sample Collection and Testing
After the experiment, the fish were anaesthetized by immersion in Tricaine Methanesulphonate (MS-222) at 200 mg/L. Approximately 1.5𠄲.0 mL blood was drawn from the caudal vessels, just behind the anal fin, with a 2.5 mL syringe. The samples were centrifuged at 4000 ×g for 10 min and the plasma was collected and frozen at ଌ until required. After collecting the blood, the fish were put into an ice box. The gills were then removed and washed with cold saline water (0.7 g/L), the surface moisture was dried with filter paper, and the gill filament was then accurately weighed to the nearest 0.2 g. The gill tissue was then homogenized in 1.8 mL saline water according to weight and volume and centrifuged at 1200 ×g for 10 min, and the clear supernatant was eluted for testing. Plasma and gill filament test samples were taken from three fish.
Plasma Na + , Cl − , and K + concentrations were measured by using the detection kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China). The urea nitrogen was spectrophotometrically measured with Nessler's reagent as the chromogenic reagent according to Lu export method . The activity of Na + /K + -ATPase was determined by Spectrophotometry using ATP as substrate thus, 1 μmol inorganic phosphorus generated by ATPase enzyme decomposition is equal to one ATPase enzyme activity unit (μmol Pi/mg prot/h). Coomassie blue staining and calf plasma protein as a standard were used for protein determination . Plasma osmotic pressure was determined using an autofreezing point osmometer (Labsun Technology Development Co. Ltd., Beijing, China).
2.4. Statistical Analysis
Count experimental data and significance tests were performed in SPSS 11.5 software (SPSS Inc., Chicago, IL, USA). Each parameter was analyzed separately by one-way ANOVA. Comparisons between different groups and between the treatment groups and control were performed by ANOVA followed by Duncan's post hoc test.
Bacterial responses to osmotic challenges
Exposure to diverse environments is a hallmark of microbial life. Microbes are everywhere collectively, microbes experience everything. They live inside and outside eukaryotic hosts, in soil, water, and air at diverse planetary sites. They may exist as individuals (planktonic growth) or aggregates, and form biofilms on biotic and abiotic surfaces. Some survive gradual or abrupt, temporal or spatial transitions between different environments.
Our understanding of microbial responses to osmotic challenges is based on intensive studies of representative bacteria, archaea, and eukaryotic microbes. This Perspective focuses on bacterial responses to osmotic challenges. Among the representative bacteria for which the osmotic stress response is well characterized, Escherichia coli lives in terrestrial and aquatic environments as well as in the meninges and the intestinal and urinary tracts of mammals. Bacillus subtilis and Corynebacterium glutamicum are soil bacteria (C. glutamicum is also used to manufacture fine chemicals), and Halomonas elongata was isolated from a solar saltern (Wood, 2011a). Some bacteria can survive in pure water and grow at a water activity (aW) near 1, many thrive within human tissues (e.g., human blood, aW of 0.995) or in seawater (aW of 0.98), whereas others can only inhabit hypersaline environments with water activities as low as 0.75. Further examples, discussed below, illustrate the range of environments and environmental variations to which bacteria respond.
Bacteria are bounded by semipermeable cytoplasmic membranes, often including aquaporins. Most are also surrounded by a rigid, elastic, and porous cell wall (the murein or peptidoglycan layer) that determines cell shape. The cell wall of Gram-negative bacteria (such as E. coli) is bounded by an outer lipid membrane that includes porins like those of mitochondria. The area between the outer and cytoplasmic membranes is denoted the “periplasm.” The integrity and hydration of the cell and its compartments are dictated by their solute contents and the osmotic pressures of their environments (discussed in Altendorf et al., 2009). A decrease in external osmotic pressure causes water influx and swelling or even lysis, whereas an increase in external osmotic pressure causes water efflux and dehydration. Water fluxes simultaneously, and almost instantaneously, perturb many cellular properties. These include cell volume (or the relative volumes of the cytoplasm and periplasm) turgor pressure cell wall strain and cytoplasmic membrane tension as well as individual uncharged solute, salt ion, and biopolymer concentrations. Cells exposed consistently to a very high osmotic pressure must maintain correspondingly high cytoplasmic solute concentrations. Evidence suggests that the regulation of cytoplasmic composition and hydration is a key objective of cellular homeostasis (Wood, 2011b).
Common themes emerged as researchers characterized the osmoadaptive mechanisms of bacteria representing diverse phylogenetic groups (Wood, 2011a, and references cited therein). Cells respond to variations in external osmotic pressure by accumulating or releasing solutes, thereby attenuating water fluxes. Those solutes include inorganic ions (often K + ), and organic molecules denoted “osmolytes” (Fig. 1). The latter are selected to minimally perturb cellular functions, even after accumulating to high (up to molar) concentrations. In turn, organisms have adapted to tolerate osmoregulatory solute accumulation. In the extreme, some halophiles accumulate KCl to molar concentrations, and their proteins function only in high salt environments. Osmoregulatory solutes accumulate via active transport or synthesis if the osmotic pressure rises and are released via mechanosensitive channels if the osmotic pressure falls. Multiple enzymes, transporters, and channels with redundant functions and specificities mediate solute accumulation and release from each organism (e.g., Fig. 2). The abundance of most osmoregulatory systems is controlled transcriptionally (Altendorf et al., 2009 Krämer, 2010). Translational regulation, mediated by small regulatory RNAs, is emerging as an important determinant of bacterial cell wall structure that may also influence the levels of osmoregulatory systems.
Care must be taken to differentiate osmotic stress from parallel, solute-specific effects that dominate particular environments. For example, bacteria inhabiting seawater face a higher osmotic pressure than those inhabiting most freshwater environments. Salts predominate in seawater, and marine organisms simultaneously face both a high osmotic pressure and a high Na + concentration. Na + fluxes are also implicated in pH homoeostasis. Distinctions are also drawn between bacteria adapted to environments with extreme and stable osmotic pressures (e.g., sea water, salt lakes) and those experiencing osmotic pressure variations (e.g., those inhabiting estuarine waters or colonizing mammalian intestinal tracts).
What cellular systems limit bacterial cell and population growth rates under osmotic stress? How are osmotic stress responses orchestrated over time and space?
Solute accumulation powerfully stimulates bacterial growth at high osmotic pressure, and solute release allows cells to survive osmotic downshocks. Thus, studies of bacterial osmoregulation have focused on the enzymes, transporters, and channels mediating solute accumulation and release (Krämer, 2010 Kung et al., 2010 Wood, 2011b) (Fig. 2). However, we do not fully understand how increasing osmotic pressure would limit bacterial cell or population growth in the absence of solute accumulation.
The evolution of bacterial cell and population size, protonmotive force, DNA replication, protein synthesis, and solute content were documented both after osmotic shifts and during steady-state culture of E. coli at various osmotic pressures, in the absence or presence of osmoprotective solutes (Wood, 1999 Cayley and Record, 2004 Altendorf et al., 2009). Such studies revealed that the population growth rate is directly proportional to cytoplasmic hydration, and that accumulating solutes differentially affect cellular rehydration and population growth. K + glutamate accumulation partially rehydrates cells and perturbs protein–nucleic acid interactions. It thereby offsets the impact of increased macromolecular crowding on cellular processes but does not restore growth to its pre-stress rate. In contrast, organic osmolytes rehydrate the cytoplasm and restore growth to an extent that correlates with their preferential exclusion from biopolymer surfaces (Cayley and Record, 2004) (discussed further below).
In contrast to our understanding of other stresses (e.g., oxidative stress Imlay, 2013), we don’t know what cellular properties or processes limit population growth rate when cells dehydrate. It was widely assumed that osmoregulation is necessary because turgor pressure is essential for cell wall expansion and cell growth. However, evidence contradicts that assumption (e.g., E. coli Cayley and Record, 2003 Rojas et al., 2014), and other cellular properties may be critical. Single-cell imaging techniques are now elucidating how osmotic stress affects cell growth and development (e.g., Pilizota and Shaevitz, 2013 Rojas et al., 2014), the composition and biophysical properties of the cytoplasm and cell membranes (Mika and Poolman, 2011 Sochacki et al., 2011 Wood, 2011b Sévin and Sauer, 2014), and the subcellular locations of osmoregulatory systems (Romantsov et al., 2010).
Respiration, the synthesis of precursor metabolites, replication, transcription, and translation are obvious candidates for growth rate limitation (Wood, 1999). Individual strains within a species vary widely in osmotic stress tolerance (e.g., Kunin et al., 1992 Murdock et al., 2014). Analysis of new strains obtained via directed evolution and of naturally occurring variants may reveal what modifications, to what systems, extend the range of cytoplasmic hydration tolerated by an organism.
The application of high throughput “omic” technologies and cell sorting are also opening new avenues of investigation. Such tools can elucidate the orchestration of osmoadaptive mechanisms after an osmotic shift or during steady-state growth at various osmotic pressures (Withman et al., 2013, and other studies cited therein). They can also show how osmotic stress affects phenotypic variation within a microbial population. Analyses of bacterial community composition suggest that the bacterial lineages inhabiting marine and freshwater ecosystems are phylogenetically distinct, and that the capacity for osmoadaptation may be a primary determinant of that divergence (Walsh et al., 2013). Organisms adapted to a stable, high salinity marine environment may face particular barriers when transitioning to a more variable estuarine or fresh water environment. Such studies have relied heavily on genomic comparisons and annotations. Key tests of these ideas may be devised by combining physiological experiments with phylogenetic approaches.
How do proteins detect and respond to osmotic pressure variations?
Membrane proteins implicated in bacterial osmoregulation became the paradigms for the study of osmosensing because they retain osmotic pressure–dependent activities after purification and reconstitution in proteoliposomes (Poolman et al., 2004). Proteoliposome-based studies provided critical evidence that mechanosensitive channels and osmosensing transporters detect and respond to osmotic pressure changes in their phospholipid environments, without input from other cellular components. Studies of bacterial systems provided seminal evidence that mechanosensitive channels open in response to forces exerted by the lipid bilayer (Teng et al., 2015). Analyses of MscL and MscS continue to elucidate mechanosensory mechanisms (Iscla and Blount, 2012 Naismith and Booth, 2012). The signal(s) to which osmosensing transporters respond remains less clear, however.
ProP of E. coli, BetP of C. glutamicum, and OpuA of Lactococcus lactis serve as paradigms for the study of osmosensing (Wood, 2011b) (Fig. 3). They represent different phylogenetic groups and energy-coupling mechanisms. ProP is a proton symporter and a member of the major facilitator superfamily, BetP is a Na + symporter and a member of the betaine-choline-carnitine transporter family, and OpuA is an ATP-hydrolyzing ATP-binding cassette (ABC) transporter. Available data suggest that each is similar in structure and transport mechanism to its paralogues that are not osmosensors.
The rate of osmolyte uptake via each transporter (A) is a sigmoid function of the osmotic pressure (Π) or osmolality (Π/RT, where R is the gas constant and T is the temperature). Such data have been fit to an arbitrary relationship that implies no particular activation mechanism:
where Amax is the asymptotic uptake rate, B is a constant inversely proportional to the slope of the response curve, and Π1/2/RT is the osmolality at which activity is half-maximal. In this relationship, Π1/2/RT can be replaced with any property that varies in parallel with the osmolality (e.g., the calculated concentration of a luminal solute in proteoliposomes). Proteoliposome data have also been fit to the Hill equation:
where Kion is the ion concentration required to attain half-maximal activity, and n is a constant related to the slope of the curve (Mahmood et al., 2006).
To understand osmosensing, we must learn what cellular property is detected by an osmosensor and understand how variations to that property modulate osmosensor structure and function (Wood, 1999). In principle, an osmosensor would trigger a homeostatic response upon detecting deviations from a “set point” of such a critical property. Experiments performed with cells and proteoliposomes ruled out turgor pressure and membrane strain as determinants of osmosensing transporter activity (Poolman et al., 2004). Proteoliposome systems were then exploited to further assess the impacts of the external and luminal solvents on the activity of each osmosensing transporter.
Merits and liabilities of proteoliposome systems.
The interpretation of proteoliposome data are supported by evidence that secondary transporters ProP and BetP reconstitute predominantly with their cytoplasmic surfaces facing the lumen, and the direction of transport is determined by an imposed ion motive force. Studies of ABC transporter OpuA exploit the fact that the direction of transport can be controlled by supplying ATP in either the external or the luminal medium (Wood, 2011b). To date, functional tests have been the primary indicators of osmosensing (i.e., solute uptake assays as opposed to spectroscopic indicators of transporter conformation). The requirements to maintain the membrane permeability barrier and to meet energy requirements for transport restrict the range of luminal and external solvent compositions accessible for these studies. The Amax values obtained with proteoliposomes are variable because transporter purification, reconstitution, and solute loading are intrinsically variable procedures. A recent comparison of the molecular activities of BetP in cells and proteoliposomes indicated that only 2.4% of BetP molecules in proteoliposomes were active (Maximov et al., 2014), reinforcing the need for careful interpretation of proteoliposome data. In contrast, Π1/2/RT, B, and Kion values are independent of transporter quantity, more reproducible, and hence presumed to be more reliable indicators.
Solvent effects on biopolymer structures.
Current knowledge of solvent effects on biopolymer structures provides a useful context for the analysis of osmosensory mechanisms. Soluble proteins and DNA have been the primary foci of such studies, which explore the thermodynamic nonideality inherent to physiological systems and their models (Record et al., 1998b, 2013). Solvent additives can affect biopolymer processes by binding as ligands at specific sites, via preferential interactions with buried or exposed biopolymer surfaces (Hofmeister effects, involving both uncharged and charged solutes) and via conformation-specific, Coulombic interactions with fixed biopolymer charges (charged solutes only). Thus, solutes may act individually (high affinity ligand binding at one or a few specific sites) and/or collectively (weak interactions at many sites).
Collective solute effects modulate the equilibrium constant (K) for any process that changes the amount of biopolymer surface interacting with a solute. The magnitudes and functional forms of these collective effects are determined by the nature of the solute excluded from or concentrated at the biopolymer surface (particularly whether it is charged or uncharged) and of the exposed or buried biopolymer surface (e.g., that of a high charge density polyelectrolyte like DNA or a low to no charge density biopolymer like a typical protein). When salt concentrations are low, the collective effects are primarily Coulombic (salts weaken charge–charge interactions). When salt or uncharged solute concentrations are high, their contributions to Hofmeister effects become dominant (e.g., Fig. 4 Record et al., 2013). The salt concentration ranges over which Coulombic and Hofmeister effects dominate for DNA and protein processes differ because DNA is a polyelectrolyte.
If a process changes the amount of uncharged or weakly charged biopolymer surface exposed to a preferentially interacting solute, then the free energy (or the logarithm of the equilibrium constant, K) for that process is a linear function of the solute concentration with a proportionality constant (the thermodynamic m-value) that reflects the properties of the solute and the magnitude of the exposed or buried biopolymer surface. Such effects are very weak at low solute concentration. In contrast, if a process changes the amount of a (polyanionic) DNA surface exposed to ionic solutes, then the logarithm of the equilibrium constant (K) varies with a power of the logarithm of the ion concentration. Such Coulombic effects are large even at low salt concentrations. The latter analysis supersedes the Debye–Hückel approximation, based on ionic strengths calculated as a function of ion concentrations and valencies, which has much more limited application. Principles governing protein–membrane interactions have not been analyzed in this way, but interactions of proteins with polyanionic membrane surfaces can be expected to share characteristics with protein–DNA interactions.
The principles outlined above were established primarily with in vitro systems. There is also evidence that cytoplasmic solutes collectively influence cellular processes, particularly as osmotic pressure changes alter cytoplasmic hydration (Record et al., 1998a,b). Small cytoplasmic solutes (e.g., K + , glutamate, and other metabolites) are preferentially excluded from nonpolar biopolymer surfaces that become exposed in unfolding. Increasing concentrations of these solutes will favor conformational changes that bury nonpolar surfaces (Record et al., 2013). At the same time, condensation of K + as a DNA counterion impedes processes involving protein–DNA interactions. In addition, increased concentrations of cytoplasmic biopolymers favor folding, especially if folding is coupled to oligomerization, by an excluded volume effect (sometimes denoted as “macromolecular crowding” Cayley and Record, 2004).
Conceptual framework for the analysis of osmosensing.
In proteoliposomes, osmosensory transporters become active as luminal solute concentrations approach 0.5 M. This suggests that both Coulombic and Hofmeister effects may participate in transporter activation and that resolution of Coulombic and Hofmeister effects will be challenging (c.f. Fig. 4). The sigmoid relationship between A and Π1/2/RT implies that transporter molecules are systematically converted from an inactive to an active conformation as the osmolality increases:
If so, the fraction of transporter active at a particular osmolality may be represented by:
where A is the initial rate of substrate uptake at a given osmolality, and Amax is the asymptotic initial rate approached at high osmolality. Then the equilibrium constant K for this transition at a particular osmolality is:
If the activating conformational change were triggered only by solute exclusion from nonpolar transporter surfaces that were exposed in the inactive and buried in the active transporter (a Hofmeister effect), the logarithm of the equilibrium constant K would be expected to vary linearly with the solute concentration X (Record et al., 2013):
In this equation, K0 would be the equilibrium constant at X = 0, where the transporter activity is undetectably small, and m/RT would be a thermodynamic parameter (the thermodynamic m-value) characteristic of the solute and the conformational change. To obtain m and K0 for transporter activation, values of A at each X would be fit to the following combined relationship:
The m-values for an array of solutes would follow the Hofmeister series (a ranking of solutes according to their effects on diverse biopolymer processes Record et al., 2013). Eq. 7 has the same form as Eq. 1, but it provides a thermodynamic interpretation of the resulting parameters. If the activating conformational change were triggered only by interactions of ions with charged surfaces (a Coulombic effect), the logarithm of the equilibrium constant K would be expected to vary with a power (n) of the logarithm of the solute concentration X (Record et al., 2013):
A relationship analogous to Eq. 7 would then reflect the dependence of ln K on [ln X] n , and ln K0 would be the value of ln K at an ion concentration (X) of 1 M. It is critical to note that reliable estimates of m/RT, the most informative parameter, can only be obtained from data that define the full range of f values.
Proteoliposome-based analysis of osmosensing by ProP, BetP, and OpuA (Wood, 2011b).
All tested membrane-impermeant solutes had similar effects on transporter activity when applied to attain the same osmolality at the external transporter surface. This response was phospholipid sensitive: the osmolality at which each transporter activates was a direct function of the anionic lipid content of the host membrane (both in vitro and in vivo). All three transporters became active as inorganic ions were concentrated at their cytoplasmic surfaces from ∼0.1 to ∼0.5 M. Differences emerged when diverse ions were used, however.
ProP activity correlated with luminal cation concentration but not luminal K + concentration. For proteoliposomes loaded with K phosphate plus the K salts of various anions, the osmolality yielding half-maximal ProP activity (Π1/2/RT) followed the Hofmeister series. ProP activity was enhanced when proteoliposomes were loaded with high molecular weight polymers (polyethyleneglycols or bovine serum albumin) at concentrations that simulated the volume exclusion occurring in the bacterial cytoplasm (Culham et al., 2012). Culham et al. (2012) concluded that ProP activity is determined by the concentrations of Hofmeister anions and macromolecular crowding.
Internal K + phosphate, glutamate or chloride, Rb + , or Cs + chloride activated BetP to varying degrees, whereas Na + (the coupling ion), NH4 + , or choline chloride did not. K + salts yielded the strongest stimulations (Krämer, 2010). However, K + dependence did not fully account for the osmotic activation of BetP in vivo, leading Maximov et al. (2014) to conclude that BetP senses K + concentration and a signal from the membrane. The effects of crowding agents on BetP activity have not been reported.
The rate of glycine betaine uptake via OpuA was enhanced similarly by K + , Na + , Li + , or NH4 + chloride. OpuA was further activated by MgCl2 and BaCl2 and inhibited by RbCl and CsCl. Ions and a large polyethyelene glycol (PEG600) acted synergistically to stimulate substrate-dependent ATP hydrolysis by OpuA in nanodiscs (Karasawa et al., 2013). Karasawa et al. (2013) concluded that OpuA responds synergistically to the ionic strength and macromolecular crowding.
These reports evoke critical roles for electrolytes, for a membrane with a polyanionic surface, and possibly for cytoplasmic volume exclusion in transporter activation. All are likely to result from some combination of collective Coulombic and Hofmeister effects of luminal solutes on changes to cytoplasm-exposed membrane and transporter surfaces. Unfortunately, the reported data are insufficient to clearly delineate the relative contributions of Coulombic and Hofmeister effects.
It is challenging to deduce the structural mechanism of osmosensing because membrane proteins are refractory to structural analysis. An impressive series of crystal structures has made enormous contributions to our understanding of the transport mechanism for BetP and related systems (Perez et al., 2014). However, conformational differences between inactive and osmotically activated BetP conformers remain to be defined. Data outlined above suggest that BetP is a chemosensor, possessing one or more cytoplasm-exposed, K + -specific regulatory sites, but those sites have not been identified. By comparison, our structural knowledge of ProP and OpuA is limited (Fig. 3).
We do know that each transporter is an oligomer (ProP and OpuA are dimers BetP is a trimer Wood, 2011b). The role of oligomerization in osmosensing by BetP has been explored experimentally but remains uncertain (Becker et al., 2014). The roles of oligomerization for ProP and OpuA remain unknown. Each transporter possesses an extended cytoplasmic C terminus (Fig. 3). The C termini of some ProP orthologues form antiparallel, intermolecular coiled-coils, whereas the extended C termini of other orthologues do not include coiled-coil motifs. The C terminus of BetP forms a long α helix that mediates inter-monomer interactions within BetP trimers, and the C termini of the two ATP-binding subunits of OpuAA include dual cystathionine-β-synthase domains with anionic tails. Structural changes to the C-terminal domains modulate the osmoregulatory response (they shift the osmolalities at which the transporters become active). It has been proposed that the cytoplasmic C termini mediate osmosensing via salt-sensitive interactions with other transporter elements (protein–protein interactions) and/or the polyanionic membrane surface. Osmotically induced variations in membrane surface charge density would also modulate protein–membrane interactions. Each of these interactions would have a characteristic thermodynamic signature, and clear dominance of Coulombic or Hofmeister effects would support distinct structural models. Thus, osmosensing may provide a paradigm for the regulation of membrane protein structure and function through protein–solvent interactions, involving solute exclusion from or accumulation at extensive protein and/or membrane surfaces.
This Perspective series includes articles by Andersen , Sachs and Sivaselvan , and Haswell and Verslues .
Clinical biochemistry of the gastrointestinal tract
Clinical aspects of carbohydrate absorption
Although inherited disorders of sucrase-isomaltase and of transport proteins causing glucose-galactose malabsorption and fructose malabsorption are recognized, they are very uncommon. Adult lactase deficiency, however, is common in humans and may be regarded as a normal finding in many ethnic groups. Malabsorption of carbohydrates predisposes to osmotic diarrhoea with excessive flatus, abdominal distension and discomfort (‘griping’ pains).
Lactase activity is highest in the earliest months of life, but levels of the enzyme usually decline in all races after weaning. The prevalence of adult lactase deficiency is highly variable between races so, for example, 5–15% of northern Europeans have demonstrable hypolactasia or alactasia, but > 70% of Africans, Asians and especially Inuits have deficiency of the enzyme. However, the vast majority of affected individuals are asymptomatic, because overall completeness of disaccharide hydrolysis depends on the amount ingested, intestinal dilution and transit times, as well as the state of the enzyme (reduced or absent enzyme activity).
In practice, the diagnosis is often made by a therapeutic trial of a diet low in lactose, but metabolic tests are available (see below) and the diagnosis can be confirmed by assaying lactase activity in small bowel biopsy samples (though this is rarely required). Congenital lactase deficiency (i.e. lactase deficiency that is present at birth) has been described, but is extremely rare. The form of enzyme deficiency discussed above is usually known as primary or ‘genetic’ lactase deficiency. However, lactase deficiency may also complicate mucosal diseases of the small intestine and is then referred to as ‘secondary’. This secondary lactase deficiency is a regular feature of coeliac disease, and is also seen in tropical sprue, IBD, radiation enteritis, chronic alcoholism with malnutrition and the enteropathy associated with acquired immune deficiency syndrome (AIDS). It may also accompany infections such as acute gastroenteritis and giardiasis, but usually resolves following resolution or successful treatment of the illness.
Osmoregulators and Osmoconformers
Persons lost at sea without any fresh water to drink are at risk of severe dehydration because the human body cannot adapt to drinking seawater, which is hypertonic in comparison to body fluids. Organisms such as goldfish that can tolerate only a relatively narrow range of salinity are referred to as stenohaline. About 90 percent of all bony fish are restricted to either freshwater or seawater. They are incapable of osmotic regulation in the opposite environment. It is possible, however, for a few fishes like salmon to spend part of their life in fresh water and part in sea water. Organisms like the salmon and molly that can tolerate a relatively wide range of salinity are referred to as euryhaline organisms. The opposite of euryhaline organisms are stenohaline ones, which can only survive within a narrow range of salinities. Most freshwater organisms are stenohaline, and will die in seawater, and similarly most marine organisms are stenohaline, and cannot live in fresh water.
Osmoconformers match their body osmolarity to their environment actively or passively. Most marine invertebrates are osmoconformers, although their ionic composition may be different from that of seawater. Osmoregulators tightly regulate their body osmolarity, which always stays constant, and are more common in the animal kingdom. Osmoregulators actively control salt concentrations despite the salt concentrations in the environment. An example is freshwater fish.
Some fish have evolved osmoregulatory mechanisms to survive in all kinds of aquatic environments. When they live in fresh water, their bodies tend to take up water because the environment is relatively hypotonic, as illustrated in Figure 2. In such hypotonic environments, these fish do not drink much water. Instead, they pass a lot of very dilute urine, and they achieve electrolyte balance by active transport of salts through the gills.
Figure 2. Osmoregulation in a freshwater environment. (credit: modification of work by Duane Raver, NOAA)
When they move to a hypertonic marine environment, these fish start drinking sea water they excrete the excess salts through their gills and their urine, as illustrated in Figure 3. Most marine invertebrates, on the other hand, may be isotonic with sea water (osmoconformers). Their body fluid concentrations conform to changes in seawater concentration. Cartilaginous fishes’ salt composition of the blood is similar to bony fishes however, the blood of sharks contains the organic compounds urea and trimethylamine oxide (TMAO). This does not mean that their electrolyte composition is similar to that of sea water. They achieve isotonicity with the sea by storing large concentrations of urea. These animals that secrete urea are called ureotelic animals. TMAO stabilizes proteins in the presence of high urea levels, preventing the disruption of peptide bonds that would occur in other animals exposed to similar levels of urea. Sharks are cartilaginous fish with a rectal gland to secrete salt and assist in osmoregulation.
Figure 3. Osmoregulation in a saltwater environment. (credit: modification of work by Duane Raver, NOAA)
Dialysis is a medical process of removing wastes and excess water from the blood by diffusion and ultrafiltration. When kidney function fails, dialysis must be done to artificially rid the body of wastes. This is a vital process to keep patients alive. In some cases, the patients undergo artificial dialysis until they are eligible for a kidney transplant. In others who are not candidates for kidney transplants, dialysis is a life-long necessity.
Dialysis technicians typically work in hospitals and clinics. While some roles in this field include equipment development and maintenance, most dialysis technicians work in direct patient care. Their on-the-job duties, which typically occur under the direct supervision of a registered nurse, focus on providing dialysis treatments. This can include reviewing patient history and current condition, assessing and responding to patient needs before and during treatment, and monitoring the dialysis process. Treatment may include taking and reporting a patient’s vital signs and preparing solutions and equipment to ensure accurate and sterile procedures.
AS Biology OCR
Cofactor - a Substance that is Required by Enzymes for them to work.
Enzyme Inhibitor - a Substance that Stops Enzymes doing their job. (some are harmful, others can be used for medicines).
Cofactors and Coenzymes
Some Enzymes will Only Work if there is Another Substance Attached to it. These are Non-Proteins and are called Cofactors.
Some of these are Inorganic Molecules (or Ions). They work by Helping the Enzyme and Substrate to Bind Together. They Don't directly Participate in the Reaction, so are Not Used up or Change in any way. An example would be Chloride Ions (Cl-) which are Cofactors for Amylase (in Saliva)
Other Cofactors are Organic, these ones are called Coenzymes. Unlike the Inorganic Cofactors, these Do Participate in Reactions and Are Changed by it (They're like a second substrate). They often act as Carriers, moving Chemical Groups between Different Enzymes. They're Continually Recycled during this process. Vitamins are often Sources of Coenzymes.
If a Cofactor is Tightly Bound to the Enzyme, it's known as a Prosthetic Group. For Example, Zinc Ions: (Zn2+) are a Prosthetic Group for Carbonic Anhydrase (An Enzyme in Red Blood Cells) which Catalyses the Production of Carbonic Acid from Water and Carbon Dioxide.The Zinc Ions are a Permanent part of the Enzyme's Active Site.
Enzyme Activity can be Prevented by Enzyme Inhibitors, which are Molecules that Bind to the Enzyme they Inhibit. There are two types of Inhibitors, Competitive and Non-Competitive:
Step 1, Find the Concentration of Sucrose
To do this, look up the atomic weights of the elements in the compound:
From the periodic table:
C = 12 g/mol
H = 1 g/mol
O = 16 g/mol
Use the atomic weights to find the molar mass of the compound. Multiply the subscripts in the formula times the atomic weight of the element. If there is no subscript, it means one atom is present.
molar mass of sucrose = 12(12) + 22(1) + 11(16)
molar mass of sucrose = 144 + 22 + 176
molar mass of sucrose = 342
nsucrose = 13.65 g x 1 mol/342 g
nsucrose = 0.04 mol
Msucrose = nsucrose/Volumesolution
Msucrose = 0.04 mol/(250 mL x 1 L/1000 mL)
Msucrose = 0.04 mol/0.25 L
Msucrose = 0.16 mol/L
The response and osmotic pressure regulation mechanism of Haliotis discus hannai (Mollusca, Gastropoda) to sudden salinity changes
Salinity is one of the critical ecological factors which will impact the growth and development of marine shellfish. With the rapid expansion of aquaculture area for Haliotis discus hannai, the frequent summer rainstorms in South China, the influx of freshwater, or the strong volatility of seawater in coastal areas, inner bays have placed abalone into the dynamic environment where the salinity is changing drastically. This work examined the effects of sudden salinity changes on abalone’s survival, osmotic pressure regulation, energy metabolism, and related gene expression by simulating the salinity changes of water for breeding H. discus hannai caused by heavy storm. The salinity was gradually reduced from 30 to 20, then kept at 20 for 48 h, followed by gradual increase to 30, and was kept at 30 for 48 h. Samples were taken at 6, 12, 36, 60, 66, 72, 96, and 120 h after the start of the experiment, respectively. Results showed that the survival rate of abalone at 120 h was significantly lower than that at any other time except at 96 h (P < 0.05). With the decrease and increase of salinity, the hemolymph osmotic pressure and the concentration of Na + , K + , Ca 2+ , Cl − in the hemolymph also followed the same trend, while the concentration of hemocyanin, total soluble protein, taurine, and free amino acids showed an inverse trend. The activity of Na + /K + -ATPase also increased then declined with salinity changes. Except at 0, 6, and 12 h, the activity of Na + /K + -ATPase in the salinity-changing group was significantly higher than that in the control group (P < 0.05). In the salinity-changing group, the activity of pyruvate kinase, succinate dehydrogenase, and malate dehydrogenase reached a maximum at 72 h, but no significant difference was found at the end of the experiment compared to the control group (P > 0.05). The expression levels of catalase, thioredoxin peroxidase, sigma-glutathione-s-transferase, and Mu-glutathione-s-transferase significantly rose with the salinity changes, and were significantly higher than that in the control group up to the end of the experiment (P < 0.05). As sudden salinity changes may cause some abalone deaths, the enhanced activity of related enzymes and the increase of gene expression levels might be one of the effective methods for an organism to respond to salinity stress and regulate osmotic pressure.
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